FRONTIERS COPYRIGHT STATEMENT

© Copyright 2007-2019 Frontiers Media SA. All rights reserved.

All content included on this site, such as text, graphics, logos, button icons, images, video/audio clips, downloads, data compilations and software, is the property of or is licensed to Frontiers Media SA (“Frontiers”) or its licensees and/or subcontractors. The copyright in the text of individual articles is the property of their respective authors, subject to a license granted to Frontiers.

The compilation of articles constituting this e-book, wherever published, as well as the compilation of all other content on this site, is the exclusive property of Frontiers. For the conditions for downloading and copying of e-books from Frontiers’ website, please see the Terms for Website Use. If purchasing Frontiers e-books from other websites or sources, the conditions of the website concerned apply.

Images and graphics not forming part of user-contributed materials may not be downloaded or copied without permission.

Individual articles may be downloaded and reproduced in accordance with the principles of the CC-BY licence subject to any copyright or other notices. They may not be re-sold as an e-book.

As author or other contributor you grant a CC-BY licence to others to reproduce your articles, including any graphics and third-party materials supplied by you, in accordance with the Conditions for Website Use and subject to any copyright notices which you include in connection with your articles and materials.

All copyright, and all rights therein, are protected by national and international copyright laws.

The above represents a summary only. For the full conditions see the Conditions for Authors and the Conditions for Website Use.

ISSN 1664-8714
ISBN 978-2-88963-142-1
DOI 10.3389/978-2-88963-142-1

About Frontiers

Frontiers is more than just an open-access publisher of scholarly articles: it is a pioneering approach to the world of academia, radically improving the way scholarly research is managed. The grand vision of Frontiers is a world where all people have an equal opportunity to seek, share and generate knowledge. Frontiers provides immediate and permanent online open access to all its publications, but this alone is not enough to realize our grand goals.

Frontiers Journal Series

The Frontiers Journal Series is a multi-tier and interdisciplinary set of open-access, online journals, promising a paradigm shift from the current review, selection and dissemination processes in academic publishing. All Frontiers journals are driven by researchers for researchers; therefore, they constitute a service to the scholarly community. At the same time, the Frontiers Journal Series operates on a revolutionary invention, the tiered publishing system, initially addressing specific communities of scholars, and gradually climbing up to broader public understanding, thus serving the interests of the lay society, too.

Dedication to Quality

Each Frontiers article is a landmark of the highest quality, thanks to genuinely collaborative interactions between authors and review editors, who include some of the world’s best academicians. Research must be certified by peers before entering a stream of knowledge that may eventually reach the public - and shape society; therefore, Frontiers only applies the most rigorous and unbiased reviews.

Frontiers revolutionizes research publishing by freely delivering the most outstanding research, evaluated with no bias from both the academic and social point of view.

By applying the most advanced information technologies, Frontiers is catapulting scholarly publishing into a new generation.

What are Frontiers Research Topics?

Frontiers Research Topics are very popular trademarks of the Frontiers Journals Series: they are collections of at least ten articles, all centered on a particular subject. With their unique mix of varied contributions from Original Research to Review Articles, Frontiers Research Topics unify the most influential researchers, the latest key findings and historical advances in a hot research area! Find out more on how to host your own Frontiers Research Topic or contribute to one as an author by contacting the Frontiers Editorial Office: researchtopics@frontiersin.org

PRODUCING, SENSING AND RESPONDING TO CELLULAR STRESS IN IMMUNITY

Topic Editors:

Heitor A. Paula-Neto, Universidade Federal do Rio de Janeiro, Brazil

Renata Meirelles Pereira, Universidade Federal do Rio de Janeiro, Brazil

Leticia A. Carneiro, Universidade Federal do Rio de Janeiro, Brazil

Cellular stress, being considered as any disturbance in cellular physiology, is a fundamental aspect of tissue and body capacity to adapt to the ever changing environment. It also surges as a consequence of tissue injury or invasion of the body by pathogens. Since the immune system was developed to sense and respond to these deleterious processes, it is reasonable to consider that immune cells are capable of sensing and responding to signs of cellular stress. Moreover, cells of the immune system undergo cellular stress during an immune response. This Research Topic presents a series of articles focusing on how cellular stress influences the outcome of immune responses, covering not only how cellular stress can be a fundamental process during immune cell activation and function, but also how cells of the immune system are capable of sensing and being influenced by factors produced by stressed cells.

Citation: Paula-Neto, H. A., Pereira, R. M., Carneiro, L. A., eds. (2019). Producing, sensing and responding to cellular stress in immunity. Lausanne: Frontiers Media. doi: 10.3389/978-2-88963-142-1

Table of Contents

Editorial: Producing, Sensing and Responding to Cellular Stress in Immunity

Heitor A. Paula-Neto, Renata M. Pereira and Leticia A. M. Carneiro

REVIEWS

Natural Killer Cell Response to Chemotherapy-Stressed Cancer Cells: Role in Tumor Immunosurveillance

Alessandra Zingoni, Cinzia Fionda, Cristiana Borrelli, Marco Cippitelli, Angela Santoni and Alessandra Soriani

S100 Proteins as an Important Regulator of Macrophage Inflammation

Chang Xia, Zachary Braunstein, Amelia C. Toomey, Jixin Zhong and Xiaoquan Rao

Sensing Bacterial-Induced DNA Damaging Effects via Natural Killer Group 2 Member D Immune Receptor: From Dysbiosis to Autoimmunity and Carcinogenesis

J. Luis Espinoza and Mika Minami

Regulation of Cytokine Production by the Unfolded Protein Response; Implications for Infection and Autoimmunity

Judith A. Smith

Metabolic Reprogramming in Modulating T Cell Reactive Oxygen Species Generation and Antioxidant Capacity

Josephin N. Rashida Gnanaprakasam, Ruohan Wu and Ruoning Wang

Autophagy and its Interaction With Intracellular Bacterial Pathogens

Mariana da Silva Siqueira, Renato de Moraes Ribeiro and Leonardo H. Travassos

Integrated Stress Responses to Bacterial Pathogenesis Patterns

Larissa O. C. P. Rodrigues, Rodrigo S. F. Graça and Leticia A. M. Carneiro

Pattern Recognition Receptors and the Host Cell Death Molecular Machinery

Gustavo P. Amarante-Mendes, Sandy Adjemian, Laura Migliari Branco, Larissa C. Zanetti, Ricardo Weinlich and Karina R. Bortoluci

ORIGINAL ARTICLE

Systems Approach Reveals Nuclear Factor Erythroid 2-Related Factor 2/Protein Kinase R Crosstalk in Human Cutaneous Leishmaniasis

Áislan de Carvalho Vivarini, Teresa Cristina Calegari-Silva, Alessandra Mattos Saliba, Viviane Sampaio Boaventura, Jaqueline França-Costa, Ricardo Khouri, Tim Dierckx, Karina Luiza Dias-Teixeira, Nicolas Fasel, Aldina Maria Prado Barral, Valéria Matos Borges, Johan Van Weyenbergh and Ulisses Gazos Lopes

EDITORIAL
published: 06 September 2019
doi: 10.3389/fimmu.2019.02053

Editorial: Producing, Sensing and Responding to Cellular Stress in Immunity

Heitor A. Paula-Neto[1][*], Renata M. Pereira[2][*] and Leticia A. M. Carneiro[2][*]

1Faculdade de Farmácia, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil

2Instituto de Microbiologia Paulo de Góes, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil

Approved by:
Francesca Granucci, University of Milano-Bicocca, Italy

*Correspondence: Heitor A. Paula-Neto, heitorapneto@gmail.com
Renata M. Pereira, renata.pereira@micro.ufrj.br
Leticia A. M. Carneiro, leticiac@micro.ufrj.br

Specialty section: This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 31 July 2019
Accepted: 14 August 2019
Published: 06 September 2019

Citation: Paula-Neto HA, Pereira RM and Carneiro LAM (2019) Editorial: Producing, Sensing and Responding to Cellular Stress in Immunity. Front. Immunol. 10:2053. doi: 10.3389/fimmu.2019.02053

Keywords: stress response, homeostasis, immune response, host-pathogen, innate immunity, UPR, autphagy, integrated stress response

Editorial on the Research Topic
Producing, Sensing and Responding to Cellular Stress in Immunity

Stress, meaning any disturbance of the internal environment of a cell, can result not only from external stimuli but also from physiological processes such as the intrinsic free radical production by the metabolic functioning of mitochondria. Stressors can threaten the cell and therefore mechanisms were selected throughout evolution to cope with and adapt to cell stress. Since the immune system is, ultimately, a system to sense and respond to stress posed by tissue damage, cell injury, and/or pathogens, it is reasonable to assume that all those cell-autonomous pathways involved in stress response also play a key role in immunity. This Frontiers in Immunology Research Topic focuses on different stress responses and their role in host–pathogen interaction and immunity.

A clear example of intrinsic stress is given by the fundamental role of reactive oxygen species (ROS) in T-cell activation. Gnanaprakasam et al. discuss how T-cells use their antioxidant machinery to fine-tune ROS activity so that it is sufficient to activate and polarize T-cells, but well-controlled to not result in cell damage. Other non-infectious process that impacts immunity is cell death. Controlled forms of cell death are an ancestral mechanism involved in key aspects of the physiology of multicellular organisms, including the elimination of unwanted, damaged, or infected cells. Amarante-Mendes et al. provide an overview on the three major types of molecularly controlled forms of cells death—apoptosis, necroptosis, and pyroptosis—that participate in host defense through the elimination of infected cells. Furthermore, the authors discuss how these events are both regulated by signals derived from PRRs as well as a source of danger-associated molecular patterns (DAMPS) that trigger immune responses through PRRs.

The concept of DAMPs is also key to understand how self-molecules can alert the immune system that homeostasis has been compromised. Among well-known DAMPs are the chromatin-associated protein HMGB1, extracellular purine metabolites, and S100 proteins. S100 proteins are a family of cytosolic proteins with a plethora of functions in cellular homeostasis that, when released from the cell as a result of tissue damage or cellular stress, can serve as DAMPs. Xia et al. explore this aspect of S100 proteins and how it interferes with different steps of inflammatory responses including their functions as DAMPs, on macrophage migration and on tissue repair.

Apart from its role in physiological processes, such as the removal of dead cells, the immune system is well-recognized for its function in host defense, interaction with microbes, and immune surveillance. During an infection, it is critical for the host to properly assess the potential threat posed by a given pathogen. In this sense, stress response pathways can be instrumental in providing the cell with the ability to sense alterations on homeostasis and tissue damage caused during infections. Rodrigues et al. discuss how the highly conserved integrated stress response (ISR) can shape the host response to bacterial pathogens. By sensing alterations to cellular homeostasis, rather than the bacteria itself, the ISR initiates a cellular program that includes transcription of key genes, profound alterations in translation of new proteins, and cell-autonomous antimicrobial mechanisms, such as autophagy. Smith also discussed how cellular stress induced by invading pathogens (virus and bacteria) is sensed, focusing on the impacts in protein folding induced by infection. Unfolded protein response (UPR) contributes to host defense through cytokine induction. The downside of the enhancement of host response is that UPR response has been increasingly recognized in a variety of autoimmune and inflammatory diseases.

Although the host cell is partly prepared to induce pathways that intervene in infections by sensing changes in homeostasis, pathogens like Leishmania parasites can adapt to these pathways and even benefit from them. Vivarini et al. demonstrated that Leishmania amazonensis induces the activation of the transcriptional factor Nrf2 (Nuclear factor erythroid 2-related factor 2), a master regulator of phase II defense gene expression that protect cells from oxidative stress. The authors show that Nrf2 knockdown promotes oxidative stress and impairs parasite survival in macrophages. Using the combination of in vitro, ex vivo, and in silico approach, the group shows Nrf2/PKR cross-talk and reveals a central role of Nrf2 in human cutaneous leishmaniasis. Nrf2 activation by L. amazonensis also required PI3K/Akt signaling and autophagy mechanisms. Autophagy is also the focus of a review by Siqueira et al. This process, known as a cellular mechanism to recycle organelles or digest intracellular contents in times of energy shortage, also plays a key role in immunity against intracellular pathogens.

Besides its interaction with pathogenic microorganisms, the immune system is also involved in the interaction and control of commensal microbiota, which, in turn, play a major role in instructing the immune system and maintaining homeostasis. Dysbiosis, which is a disruption of the normal microbiota–host relationship, has been associated with a myriad of human diseases including metabolic disorders, autoimmunity, and cancer. Espinoza and Minamo compile evidence suggesting that dysbiosis triggers DNA damage response, either by producing genotoxins or by promoting chronic inflammation, leading to overexpression of NKG2D-L in stressed cells. Consequently, these cells are tagged to be eliminated by Natural Killer (NK) cells and various subsets of T cells, which could be linked to autoimmunity and carcinogenesis.

On the other hand, the ability of immune cells, in particular NK cells, to detect cells displaying signs of stress is crucial for tumor immunosurveillance. This has been used to design new therapies that not only have antiproliferative and cytotoxic effects but also boost antitumor immunity by rendering tumor cells better targets for NK cells. Zingoni et al. discuss how cellular stress pathways induced in various tumors by different chemotherapeutic regimens can stimulate NK cells' effector function and provide new therapeutic approaches.

This collection of review and original papers is an invitation for the reader to appreciate the view of the immune system as a platform designed for the sensing, detection, and response to stress (in a broad definition), that uses all the stress-coping machinery selected by evolution to ensure an appropriate interaction with environmental challenges and host survival.

AUTHOR CONTRIBUTIONS

All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Paula-Neto, Pereira and Carneiro. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

MINI REVIEW
published: 25 September 2017
doi: 10.3389/fimmu.2017.01194

Natural Killer Cell Response to Chemotherapy-Stressed Cancer Cells: Role in Tumor Immunosurveillance

Alessandra Zingoni[1]*, Cinzia Fionda[1], Cristiana Borrelli1,2, Marco Cippitelli[1], Angela Santoni1,3 and Alessandra Soriani[1]*

1 Department of Molecular Medicine, Sapienza University of Rome, Laboratory Affiliated to Istituto Pasteur Italia – Fondazione Cenci Bolognetti, Rome, Italy

2 Center for Life Nano Science@Sapienza, Istituto Italiano di Tecnologia, Rome, Italy

3 Neuromed I.R.C.C.S. – Istituto Neurologico Mediterraneo, Pozzilli, Italy

OPEN ACCESS

Edited by:

Leticia A. Carneiro, Federal University of Rio de Janeiro, Brazil

Reviewed by:

Angelo A. Manfredi, Vita-Salute San Raffaele University, Italy
Ofer Mandelboim, Hebrew University of Jerusalem, Israel
Martin Hernán Bonamino, Instituto Nacional do Câncer, Brazil

*Correspondence:

Alessandra Zingoni
alessandra.zingoni@uniroma1.it;
Alessandra Soriani
alessandra.soriani@uniroma1.it

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 19 July 2017
Accepted: 08 September 2017
Published: 25 September 2017

Citation:

Zingoni A, Fionda C, Borrelli C, Cippitelli M, Santoni A and Soriani A (2017) Natural Killer Cell Response to Chemotherapy-Stressed Cancer Cells: Role in Tumor Immunosurveillance. Front. Immunol. 8:1194. doi: 10.3389/fimmu.2017.01194

Natural killer (NK) cells are innate cytotoxic lymphoid cells that actively prevent neoplastic development, growth, and metastatic dissemination in a process called cancer immunosurveillance. An equilibrium between immune control and tumor growth is maintained as long as cancer cells evade immunosurveillance. Therapies designed to kill cancer cells and to simultaneously sustain host antitumor immunity are an appealing strategy to control tumor growth. Several chemotherapeutic agents, depending on which drugs and doses are used, give rise to DNA damage and cancer cell death by means of apoptosis, immunogenic cell death, or other forms of non-apoptotic death (i.e., mitotic catastrophe, senescence, and autophagy). However, it is becoming increasingly clear that they can trigger additional stress responses. Indeed, relevant immunostimulating effects of different therapeutic programs include also the activation of pathways able to promote their recognition by immune effector cells. Among stress-inducible immunostimulating proteins, changes in the expression levels of NK cell-activating and inhibitory ligands, as well as of death receptors on tumor cells, play a critical role in their detection and elimination by innate immune effectors, including NK cells. Here, we will review recent advances in chemotherapy-mediated cellular stress pathways able to stimulate NK cell effector functions. In particular, we will address how these cytotoxic lymphocytes sense and respond to different types of drug-induced stresses contributing to anticancer activity.

Keywords: natural killer cells, immunochemotherapy, cancer, stress, natural killer cell activating ligands, damage-associated molecular patterns, death receptors, PDL-1

INTRODUCTION

Natural killer (NK) cells represent a crucial component of antitumor innate immune response displaying cytotoxic functions and secreting several cytokines/chemokines (1, 2).

Natural killer cell cytotoxic activity regulation depends on an integrated interplay between inhibitory receptors and numerous activating receptors acting in concert to efficiently eliminate tumor cells.

Relevant activating receptors for tumor cell recognition are NKG2D that recognizes MICA/B and ULBPs proteins, orthologs of the mouse RAE1 molecules, DNAM-1 that binds two ligands named poliovirus receptor (PVR/CD155) and Nectin-2 (CD112), and the receptors NKp30, NKp44, and NKp46 belonging to the natural cytotoxicity receptors and shown to interact with a broad spectrum of ligands (3).

Natural killer cells also express inhibitory receptors for molecules of the major histocompatibility complex (MHC) class I, which are Ly49 receptors in mice, killer cell immunoglobulin-like receptors (KIRs) that bind to HLA-A, -B, and -C molecules in humans, and the CD94-NKG2A heterodimer in both species (4). In addition, NK cells express two inhibitory receptors for PVR, called TACTILE (CD96) and TIGIT, that counterbalance the DNAM-1-mediated activation of NK cells (5).

The activation of NK cells leads to the release of cytotoxic granules containing perforin and various granzymes and to cytokine production, most prominently interferon-γ (IFN-γ) (6–8). In addition, the expression at the cell surface of death-inducing ligands belonging to the tumor necrosis factor (TNF) family, such as Fas ligand (FasL) and TNF-related apoptosis-inducing ligand (TRAIL), also drives the activation of the caspase enzymatic cascade through the binding to the death receptors (DRs), namely, Fas, DR4 (TRAIL-RI), and DR5 (TRAIL-RII), on target cells (9, 10).

More recently, immunological checkpoint molecules commonly associated with T cells, such as CTLA-4 and PD-1, have been described on NK cells as negative regulators of their immune function (11–13).

Conventional chemotherapies were initially designed to produce antiproliferative or cytotoxic effects on dividing tumor cells. However, as result of numerous demonstrations indicating that an endogenous antitumor immunity is essential for complete remission during tumor therapy (14–16) several antineoplastic drugs, even at low doses, have been reconsidered also as potential immunomodulatory agents (17).

In this context, it has becoming always more evident that dying or stressed cells release or expose stress molecules, called damage-associated molecular patterns (DAMPs) that can alert the immune system (18). Moreover, many chemotherapy-mediated stress pathways modulate the expression of NK cell activating and inhibitory ligands, rendering tumor cells more immunogenic.

In this review, we will summarize the effects of different chemotherapeutic agents on the activity of NK cells, emphasizing the immunomodulatory effects of both conventional and low concentrations of drugs at the interface between stressed or dying cancer cells and the immune system, in the attempt of exploiting them for therapeutic purposes.

REGULATION OF NK CELL-ACTIVATING AND -INHIBITORY LIGAND EXPRESSION BY CHEMOTHERAPEUTIC DRUGS

A number of evidence indicate that chemotherapy-induced sensitization of tumor cells to immune effectors plays an important role in anticancer therapy. Indeed, different types of drug-induced stresses can modulate the expression of NK cell-activating/or -inhibitory ligands on cancer cells thus affecting their recognition and elimination by NK cells (Table 1). Besides genotoxic drugs or radiotherapy, many other pharmacological compounds already approved for the treatment of different malignancies or entered in clinical trials have been described to increase NK cell-activating ligand expression (19–27). Moreover, most of these drugs are also able to downregulate NK cell-inhibitory ligand expression, so that different and multiple mechanisms concur to make tumor cells more susceptible to NK cell-mediated lysis (28–32).

TABLE 1 | Chemotherapy-induced pathways and molecular targets able to modulate natural killer (NK) cell activating ligands and PDL-1 on cancer cells.

In the case of genotoxic drugs or DNA replication inhibitors, the mechanisms regulating the NKp30 ligand B7-H6 expression on human cancer cells remain largely unknown (23), while much evidence indicate a major role for the DNA damage response (DDR) pathway in the upregulation of the stimulatory ligands for the NKG2D and DNAM-1 immunoreceptors. In addition, ionizing radiations represent classical stimuli to induce NKG2D ligand upregulation, through the induction of the DDR (33). The activation of the kinases ATM/ATR and the production of reactive oxygen species converge on the E2F1 factor able to activate MICA, MICB, and PVR transcription on multiple myeloma (MM) cells by doxorubicin and melphalan (34). On the other hand, a different pathway governing NKG2DLs expression by chemicals known to induce genotoxic stress has been characterized in murine lymphoma cells: DDR drives to the presence of cytosolic DNA and to STING/TBK1-dependent activation of the transcription factor IRF3, responsible for the upregulation of RAE1 expression (35). Interestingly, in murine leukemia cells, concomitantly to NKG2D ligand upregulation, DDR-activating therapeutic agents cause a loss of the inhibitory NK cell ligand Clr-b, thus enhancing the cytotoxicity mediated by NKRP1B+ NK cells (36).

Non-lethal heat shock mimicking hyperthermia therapy can promote NKG2DL expression both in human and murine cancer cells but with different mechanisms. MICA and MICB upregulation occurs at the transcriptional level via HSF1 activation (37) and, with a similar mechanism, MICA and MICB expression on MM cells is enhanced by HSP90 chaperone inhibitors that activate this transcription factor (21). In a different way, increased surface expression of the mouse NKG2D ligand Mult1 depends on the inhibition of protein ubiquitination and lysosomal degradation (38).

Treatment of different tumor cell types with epigenetic drugs, like histone deacetylase inhibitors (HDACi) and DNA-methyltransferase inhibitors (DNMTi) (25–27, 39–43), leads to the upregulation of NKG2DLs and PVR surface levels, although it downregulates B7-H6 expression (44). For DNMTi the molecular mechanisms underlying NKG2DLs upregulation are still unclear, while different pathways cooperate in the regulation of these molecules in response to HDACi, and this might depend on the type of tumor and the dose of the drug used. In particular, valproic acid (VPA) has been reported to upregulate MICA/B with a mechanism dependent on PI3K/Akt pathway in pancreatic cancer cells (40), while the involvement of ERK in MICA/B and ULBP2 upregulation in response to VPA has been shown in MM cells (45). Moreover, Yang and colleagues proposed that the capability of the HDACi suberoylanilide-hydroxamic acid (SAHA) to increase MICA expression in hepatoma cancer cells is dependent on miR-17-92 cluster (46).

In MM cells, the bromodomain and extra terminal domain inhibitors (BETi) and immunomodulatory drugs (IMiDs) can block the repressive activity of the transcription factors IRF4 and IKZF1/3 on MICA and PVR promoters (19, 47). In addition, both these therapeutic agents can downregulate the expression of PD-L1 on cancer cells (28, 29, 31, 32). Indeed, BETi interrupt the activity of the epigenetic reader protein BRD4 on PD-L1 promoter region, by significantly reducing both the constitutive and IFN-γ inducible expression of this ligand. In this regard, the downstream mediators of IFN-γ signaling, JAK kinases, can be pharmacologically blocked to negatively regulate PD-L1 expression in cancer cells (48). Furthermore, drugs disrupting RAF/MEK/ERK signaling pathway, such as Sorafenib and the TLR3 agonists poly-IC, can synergistically reduce the percentage of tumor cells expressing PD-L1 and enhance NK and T cell activation in a mouse model of hepatocarcinoma (49).

Regarding drugs that disrupt the microtubule assembly, sub-lethal doses of Vincristine can activate p38 MAPK and regulate NKG2DL expression both at transcriptional and posttranscriptional level in MM cells (50). Moreover, Cytochalasin D, nocodazole, and docetaxel can enhance NKG2D, DNAM-1, and NKp30 ligands on tumor cell surface, with MICA upregulation being dependent on both DNA damage and endoplasmic reticulum (ER) stress response (51).

Different studies have been done by using proteasome inhibitors in MM cells. In this regard, low doses of bortezomib can induce the upregulation of both NKG2D and DNAM-1 ligands (22, 52, 53), and in accordance with these data, Jinushi and colleagues reported a DDR-ATM-dependent upregulation of MICA surface levels (24). On the other hand, no significant change in NKG2DL expression was observed upon bortezomib treatment by Shi and colleagues (30). Interestingly, the latter study described the capability of bortezomib to downregulate HLA class I surface expression by sensitizing MM cells to NK cell–mediated lysis (30).

Chemotherapeutic agents can also contribute to the posttranslational regulation of NK activating ligand expression by promoting the release of soluble NKG2DLs through the modulation of the expression and activity of metalloproteinases (MMP) and ADAM enzymes on cancer cells (54). Although an increased stimulation of the shedding process in response to genotoxic agents has been reported (55), some studies using different drugs describe an inhibitory effect. Indeed, gemcitabine treatment impaired ULBP2 shedding through downregulation of ADAM10 in pancreatic cancer (56). Likewise, the hypomethylating agents, azacitidine and decitabine, reduced MICA, MICB, and ULBP2 release in AML by increasing TIMP3 expression, a potent inhibitor of MMP family (57).

Thus, antitumor therapeutics can work also as activators of different “stress pathways” that enhance tumor sensitivity to NK cell cytolysis by modulating the expression of the activating and inhibitory ligands on tumor cells.

MODULATION OF DRs BY CANCER THERAPEUTIC AGENTS

Many cancer therapeutic drugs can induce DR expression and redistribution (58) (Table 2). Several studies described a role for different types of HDACi in the upregulation of TRAIL receptors on various malignant tumor cells (59–63). In this context, SAHA and trichostatin A (TSA) were shown to increase cell-surface expression of DR4 and DR5 in human MM cell lines (64). A study from Insinga et al. showed that different DR and their ligands (i.e., TRAIL, DR5, FasL, and Fas) are upregulated by HDACi on leukemic cells, but not in the normal counterpart of hematopoietic progenitors, promoting tumor apoptosis through the activation of the DR pathway (65).

TABLE 2 | Chemotherapy-induced pathways and molecular targets able to modulate death receptors (DRs) on cancer cells.

A number of studies showed that bortezomib upregulated surface expression of TRAIL receptors on a variety of human tumor cell lines, enhancing their susceptibility to NK cell lysis with a mechanism mainly dependent on TRAIL (66). In another model, a bortezomib-treated murine renal carcinoma cell line is more susceptible to both NK-cell perforin/granzyme and recombinant TRAIL-mediated apoptosis, resulting in enhanced caspase-8 activity (67). Indeed, in human non-small cell lung cancer cells this drug has been shown to trigger TRAIL-induced apoptosis via DR5 upregulation (68). Several pieces of evidence reported that another proteasome inhibitor, namely, MG132, increases DR5 expression cooperating in establishing apoptosis in several cancer cells (69–71).

DR4 and DR5 were demonstrated to be DNA damaging-inducible and p53-regulated genes (72–76). Accordingly, many DNA damaging chemotherapeutic agents can regulate DR expression, rendering cancer cells more sensitive to DR-elicited apoptosis (74, 75, 77–81).

Altogether, these results suggest that the extrinsic apoptotic pathway has an important role in chemotherapy-induced apoptosis through the promotion of DRs-mediated recognition by cytotoxic lymphocytes. In addition, chemotherapies can promote the cell death by regulating the balance between pro- and antiapoptotic proteins toward apoptosis. Many evidence show that drugs may control the cell intrinsic apoptosis by altering Bax and Bcl-2 expression in different tumor cells (82–86).

CHEMOTHERAPY-INDUCED DAMPs ALERTING NK CELLS

Many anticancer chemotherapies increase the immunogenic potential of cancer cells mainly through the establishment of immunogenic cell death, or other forms of non-apoptotic death, including autophagy, and the release of the so-called DAMPs, such as high-mobility group box 1 proteins (HMGB1), ATP, heat shock proteins (HSPs), and the ER chaperone calreticulin (87).

Damage-associated molecular patterns are intracellularly sequestered in normal physiological conditions, but they can be actively secreted or aberrantly exposed on the cell surface under conditions of cellular stress.

Engagement of various target receptors present on immune cells by DAMPs leads to the elicitation of a potent antitumor immunity. Mostly, DAMPs have been proposed to activate local APCs, thus promoting the adaptive immune system. For example, both HSP70 and HMGB1 boost dendritic cell (DC) maturation through toll-like receptor 4, favoring the induction of antigen-specific T cell-mediated antitumor immune responses (88, 89). Less is known about DAMP contribution to NK cell stimulation; thus, we will focus the attention on HMGB1 and HSPs, due to their ability to exert different effects on NK cell-mediated functions.

High-mobility group box 1 protein is an endogenous nuclear factor released both by activated immune cells or injured non-immune cells, and in the extracellular milieu acts as a DAMP alerting the immune system to danger and triggering immune response activation through the interaction either with multiple TLRs and the receptor for advanced glycation end products (RAGE), expressed on a variety of cells (90). In this regard, the chemotherapeutic agent cyclophosphamide has been recently shown to facilitate NK cell activation through a process involving HMGB1 release in a glioma mouse model (91). Accordingly, it was demonstrated that in HMGB1-deficient tumors, different innate immune cells, including NK cells, have impaired ability to reach the tumor tissue in response to DNA alkylating agent treatments (92). In addition, HMGB1 can be released by NK cells and can stimulate NK cell chemotaxis through RAGE, thus further amplifying their response to tumors (93) and can also play an important role in the cross-talk between NK and DC, by promoting DC maturation (94, 95). Interestingly, HGMB1 can induce autophagy (96), which may control the regulation of the innate and adaptive immune responses contributing to enhance antigen processing and presentation (97).

Heat shock proteins are localized in most intracellular compartments where they act as molecular chaperone by supporting protein folding and transport across membranes. Several studies demonstrated an unusual HSP70 cell membrane localization on transformed tumor cells (98–100). As already mentioned, stressful conditions can cause HSPs mobilization to the plasma membrane, or their release from cells, thus acting as potent danger signals. In this respect, therapeutic treatments including radio and chemotherapy have been shown to produce an augmentation of HSP70 cell-surface expression on tumor cells (101, 102). Several studies have shown that membrane-bound HSP70 directly promotes NK cell mediated cytotoxicity in vitro (103, 104) and in vivo (105) thus, there is an increasing interest in the therapeutic potential of targeting HSP70. Interestingly, Elsner and colleagues have shown a synergistic potentiating effect of two stress-inducible immunological danger signals HSP70 and NKG2D ligands on cytotoxicity of human (106) and mouse NK cells (107), suggesting that the drug-mediated upregulation of activating ligands and HSP70 on the cancer cell surface might be an encouraging strategy aimed at promoting the antitumor NK cell responses. Moreover, several pieces of evidence demonstrate that extracellular-located HSPs can be associated to extracellular vesicles (108–112), and a number of chemotherapeutic agents, including etoposide (109), melphalan (110), cisplatin, and 5-fluorouracil (112), have been shown to stimulate an enhanced secretion of exosomes from different types of cancer cells. Notably, colon carcinoma-derived HSP70 associated to exosomes can stimulate NK cell migration and cytotoxic activity (108). In addition, we have recently demonstrated that HSP70 on the surface of MM-derived exosomes triggers NK cell-mediated IFN-γ production through a mechanism dependent on TLR2 (110).

DIRECT EFFECTS OF CHEMOTHERAPY ON NK CELL-MEDIATED FUNCTIONS

Alterations of NK cell activities upon administration of chemotherapeutic drugs can be different in terms of cytotoxicity and immunoregulatory activity; indeed, standard chemotherapeutic protocols used in the treatment of cancer patients mainly suppress NK cell-mediated killing against cancer cells and their cytokine production. However, several studies aimed at analyzing the NK cell behavior in patients undergoing cytotoxic chemotherapy have demonstrated different and variable effects depending on both the type and the dose of the drug used.

In this regard, by producing IFN-γ, NK cells induce CD8+ T cells to become CTLs, and also help to differentiate CD4+ T cells toward a Th1 response. Moreover, NK cell-derived cytokines might also regulate antitumor antibody production by B cells. Thus, therapeutic strategies able to preserve NK functions in cancer patients are of pivotal importance, particularly those eligible for monoclonal antibody-based treatments. In this context, metronomic low cyclophosphamide (CTX) regimen was shown to potently stimulate NK functions in terms of cytokine production and antitumor immunity (18). A number of drugs, including bortezomib, genotoxic agents, and epigenetic drugs, exert immunosuppressive effects at high concentrations, whereas at sub-lethal doses, they can render tumor cells more immunogenic without affecting the immune cell activity (113). As an example, low doses of bortezomib capable of stimulating NK cell activating ligand expression on MM (22, 52), do not alter NK cell degranulation against sensitive targets (52). In another study, low concentrations of bortezomib reduced IFN-γ production without affecting NK cell cytotoxicity (114). Moreover, a combination of bortezomib with exogenous cytokine treatment enhanced the cytotoxic effects of NK cells against cancer cells in two different models (115, 116). The treatment of NK cells with sub-lethal doses of doxorubicin, able to upregulate NKG2D and DNAM-1 ligands on MM cells, does not change the capacity of NK cell to degranulate in response to target cells, as well as the ability to produce IFN-γ (34). Although the wide range of HDACi, structurally different from each other, can have both stimulatory and inhibitory effects on immune cell function, the most of them (i.e., romidepsin, vorinostat, TSA, and VPA) have been shown to suppress NK cell activity at therapeutically relevant concentrations (117–119). However, some reports describe a beneficial effect on NK cells as for the narrow-spectrum HDACi entinostat that can increase NKG2D expression on NK cells without affecting their cytotoxic activity (120). Furthermore, a recent study demonstrates that the HDACi panobinostat has the capability to potentiate the antitumor effects of trastuzumab by stimulating the antibody-dependent cell-mediated cytotoxicity (ADCC) mediated by NK cells (121). Regarding the DNTMi decitabine and 5-azacytidine, treatment of NK cells leads to increased reactivity toward different tumor cells (122, 123), while another study describes that 5-azacytidine exposure compromises their activity in AML and MDS patients (124).

Immunomodulatory drugs (lenalidomide, pomalidomide, and thalidomide) exert strong immunomodulatory effects involving both innate and adaptive immunity. In particular, these compounds activate both NK and T cells by inducing their proliferation, cytokine production, and cytotoxic activity (125) and promising clinical trials have been reported their use for the treatment of hematological malignancies, such as myeloma, lymphoma, and leukemia, as well as of solid tumors (126–128). Interestingly, Lagrue and colleagues demonstrated that lenalidomide enhances NK cell response (IFN-γ production and cytotoxicity) by augmenting actin remodeling, thus rendering them able to respond to lower densities of activating ligands on tumor cells (126). Furthermore, lenalidomide has synergistic effects on NK cell functions when used in combination with monoclonal antibodies able to promote ADCC that are already approved in therapeutic protocols, such as rituximab or elotuzumab (129, 130); indeed, novel strategies in the treatment of MM combines the use of lenalidomide and the anti-inhibitory KIR antibody (IPH2101) (131, 132).

CONCLUSION

The modulation of the expression and/or the release of stress molecules has emerged as a new paradigm of the therapeutic possibilities associated with the use of chemotherapy (Figure 1). In this context, the characterization of novel drugs and regulatory pathways activated by cellular stress modifiers able to affect tumor growth and, at the same time, to improve the activities mediated by cytotoxic lymphocytes such as NK cells, will importantly contribute to the developing field of chemo-immunotherapy.

FIGURE 1 | Antitumor efficacy of chemotherapy. Chemotherapeutic agents activate molecular pathways eliciting upregulation and/or the release of stress molecules that promote tumor cell recognition and elimination by natural killer (NK) cells. Moreover, chemotherapy can also downregulate the expression of ligands such as PD-L1 and major histocompatibility complex (MHC)-I of inhibitory receptors.

AUTHOR CONTRIBUTIONS

AZ, CF, CB, MC, ASantoni, and ASoriani contributed equally to writing and critically revised the paper.

ACKNOWLEDGMENTS

This work was supported by Italian Association for Cancer Research (AIRC Investigator Grant cod. 16014 and AIRC 5x1000 cod. 9962) and grants by the “Sapienza” University of Rome (RM116154C8F24748).

REFERENCES

1. Caligiuri MA. Human natural killer cells. Blood (2008) 112(3):461–9. doi:10.1182/blood-2007-09-077438

2. Vivier E, Raulet DH, Moretta A, Caligiuri MA, Zitvogel L, Lanier LL, et al. Innate or adaptive immunity? The example of natural killer cells. Science (2011) 331(6013):44–9. doi:10.1126/science.1198687

3. Lanier LL. NK cell recognition. Annu Rev Immunol (2005) 23:225–74. doi:10.1146/annurev.immunol.23.021704.115526

4. Parham P. MHC class I molecules and KIRs in human history, health and survival. Nat Rev Immunol (2005) 5(3):201–14. doi:10.1038/nri1570

5. Stanietsky N, Simic H, Arapovic J, Toporik A, Levy O, Novik A, et al. The interaction of TIGIT with PVR and PVRL2 inhibits human NK cell cytotoxicity. Proc Natl Acad Sci U S A (2009) 106(42):17858–63. doi:10.1073/pnas.0903474106

6. Vivier E, Tomasello E, Baratin M, Walzer T, Ugolini S. Functions of natural killer cells. Nat Immunol (2008) 9(5):503–10. doi:10.1038/ni1582

7. Krzewski K, Coligan JE. Human NK cell lytic granules and regulation of their exocytosis. Front Immunol (2012) 3:335. doi:10.3389/fimmu.2012.00335

8. Voskoboinik I, Smyth MJ, Trapani JA. Perforin-mediated target-cell death and immune homeostasis. Nat Rev Immunol (2006) 6(12):940–52. doi:10.1038/nri1983

9. Wallin RP, Screpanti V, Michaelsson J, Grandien A, Ljunggren HG. Regulation of perforin-independent NK cell-mediated cytotoxicity. Eur J Immunol (2003) 33(10):2727–35. doi:10.1002/eji.200324070

10. Smyth MJ, Cretney E, Kelly JM, Westwood JA, Street SE, Yagita H, et al. Activation of NK cell cytotoxicity. Mol Immunol (2005) 42(4):501–10. doi:10.1016/j.molimm.2004.07.034

11. Alvarez IB, Pasquinelli V, Jurado JO, Abbate E, Musella RM, de la Barrera SS, et al. Role played by the programmed death-1-programmed death ligand pathway during innate immunity against Mycobacterium tuberculosis. J Infect Dis (2010) 202(4):524–32. doi:10.1086/654932

12. Norris S, Coleman A, Kuri-Cervantes L, Bower M, Nelson M, Goodier MR. PD-1 expression on natural killer cells and CD8(+) T cells during chronic HIV-1 infection. Viral Immunol (2012) 25(4):329–32. doi:10.1089/vim.2011.0096

13. Stojanovic A, Fiegler N, Brunner-Weinzierl M, Cerwenka A. CTLA-4 is expressed by activated mouse NK cells and inhibits NK Cell IFN-gamma production in response to mature dendritic cells. J Immunol (2014) 192(9):4184–91. doi:10.4049/jimmunol.1302091

14. Zitvogel L, Apetoh L, Ghiringhelli F, Andre F, Tesniere A, Kroemer G. The anticancer immune response: indispensable for therapeutic success? J Clin Invest (2008) 118(6):1991–2001. doi:10.1172/JCI35180

15. Zitvogel L, Kepp O, Kroemer G. Immune parameters affecting the efficacy of chemotherapeutic regimens. Nat Rev Clin Oncol (2011) 8(3):151–60. doi:10.1038/nrclinonc.2010.223

16. Zitvogel L, Apetoh L, Ghiringhelli F, Kroemer G. Immunological aspects of cancer chemotherapy. Nat Rev Immunol (2008) 8(1):59–73. doi:10.1038/nri2216

17. Shurin MR, Naiditch H, Gutkin DW, Umansky V, Shurin GV. ChemoImmunoModulation: immune regulation by the antineoplastic chemotherapeutic agents. Curr Med Chem (2012) 19(12):1792–803. doi:10.2174/092986712800099785

18. Bracci L, Schiavoni G, Sistigu A, Belardelli F. Immune-based mechanisms of cytotoxic chemotherapy: implications for the design of novel and rationale-based combined treatments against cancer. Cell Death Differ (2014) 21(1):15–25. doi:10.1038/cdd.2013.67

19. Abruzzese MP, Bilotta MT, Fionda C, Zingoni A, Soriani A, Vulpis E, et al. Inhibition of bromodomain and extra-terminal (BET) proteins increases NKG2D ligand MICA expression and sensitivity to NK cell-mediated cytotoxicity in multiple myeloma cells: role of cMYC-IRF4-miR-125b interplay. J Hematol Oncol (2016) 9(1):134. doi:10.1186/s13045-016-0362-2

20. Fionda C, Malgarini G, Soriani A, Zingoni A, Cecere F, Iannitto ML, et al. Inhibition of glycogen synthase kinase-3 increases NKG2D ligand MICA expression and sensitivity to NK cell-mediated cytotoxicity in multiple myeloma cells: role of STAT3. J Immunol (2013) 190(12):6662–72. doi:10.4049/jimmunol.1201426

21. Fionda C, Soriani A, Malgarini G, Iannitto ML, Santoni A, Cippitelli M. Heat shock protein-90 inhibitors increase MHC class I-related chain A and B ligand expression on multiple myeloma cells and their ability to trigger NK cell degranulation. J Immunol (2009) 183(7):4385–94. doi:10.4049/jimmunol.0901797

22. Soriani A, Zingoni A, Cerboni C, Iannitto ML, Ricciardi MR, Di Gialleonardo V, et al. ATM-ATR-dependent up-regulation of DNAM-1 and NKG2D ligands on multiple myeloma cells by therapeutic agents results in enhanced NK-cell susceptibility and is associated with a senescent phenotype. Blood (2009) 113(15):3503–11. doi:10.1182/blood-2008-08-173914

23. Cao G, Wang J, Zheng X, Wei H, Tian Z, Sun R. Tumor therapeutics work as stress inducers to enhance tumor sensitivity to natural killer (NK) cell cytolysis by up-regulating NKp30 ligand B7-H6. J Biol Chem (2015) 290(50):29964–73. doi:10.1074/jbc.M115.674010

24. Jinushi M, Vanneman M, Munshi NC, Tai YT, Prabhala RH, Ritz J, et al. MHC class I chain-related protein A antibodies and shedding are associated with the progression of multiple myeloma. Proc Natl Acad Sci U S A (2008) 105(4):1285–90. doi:10.1073/pnas.0711293105

25. Zhu Z, Lu X, Jiang L, Sun X, Zhou H, Jia Z, et al. STAT3 signaling pathway is involved in decitabine induced biological phenotype regulation of acute myeloid leukemia cells. Am J Transl Res (2015) 7(10):1896–907.

26. Tang KF, He CX, Zeng GL, Wu J, Song GB, Shi YS, et al. Induction of MHC class I-related chain B (MICB) by 5-aza-2’-deoxycytidine. Biochem Biophys Res Commun (2008) 370(4):578–83. doi:10.1016/j.bbrc.2008.03.131

27. Rohner A, Langenkamp U, Siegler U, Kalberer CP, Wodnar-Filipowicz A. Differentiation-promoting drugs up-regulate NKG2D ligand expression and enhance the susceptibility of acute myeloid leukemia cells to natural killer cell-mediated lysis. Leuk Res (2007) 31(10):1393–402. doi:10.1016/j.leukres.2007.02.020

28. Hogg SJ, Vervoort SJ, Deswal S, Ott CJ, Li J, Cluse LA, et al. BET-bromodomain inhibitors engage the host immune system and regulate expression of the immune checkpoint ligand PD-L1. Cell Rep (2017) 18(9):2162–74. doi:10.1016/j.celrep.2017.02.011

29. Zhu H, Bengsch F, Svoronos N, Rutkowski MR, Bitler BG, Allegrezza MJ, et al. BET bromodomain inhibition promotes anti-tumor immunity by suppressing PD-L1 expression. Cell Rep (2016) 16(11):2829–37. doi:10.1016/j.celrep.2016.08.032

30. Shi J, Tricot GJ, Garg TK, Malaviarachchi PA, Szmania SM, Kellum RE, et al. Bortezomib down-regulates the cell-surface expression of HLA class I and enhances natural killer cell-mediated lysis of myeloma. Blood (2008) 111(3):1309–17. doi:10.1182/blood-2007-03-078535

31. Gorgun G, Samur MK, Cowens KB, Paula S, Bianchi G, Anderson JE, et al. Lenalidomide enhances immune checkpoint blockade-induced immune response in multiple myeloma. Clin Cancer Res (2015) 21(20):4607–18. doi:10.1158/1078-0432.CCR-15-0200

32. Giuliani M, Janji B, Berchem G. Activation of NK cells and disruption of PD-L1/PD-1 axis: two different ways for lenalidomide to block myeloma progression. Oncotarget (2017) 8(14):24031–44. doi:10.18632/oncotarget.15234

33. Gasser S, Orsulic S, Brown EJ, Raulet DH. The DNA damage pathway regulates innate immune system ligands of the NKG2D receptor. Nature (2005) 436(7054):1186–90. doi:10.1038/nature03884

34. Soriani A, Iannitto ML, Ricci B, Fionda C, Malgarini G, Morrone S, et al. Reactive oxygen species- and DNA damage response-dependent NK cell activating ligand upregulation occurs at transcriptional levels and requires the transcriptional factor E2F1. J Immunol (2014) 193(2):950–60. doi:10.4049/jimmunol.1400271

35. Lam AR, Le Bert N, Ho SS, Shen YJ, Tang ML, Xiong GM, et al. RAE1 ligands for the NKG2D receptor are regulated by STING-dependent DNA sensor pathways in lymphoma. Cancer Res (2014) 74(8):2193–203. doi:10.1158/0008-5472.CAN-13-1703

36. Fine JH, Chen P, Mesci A, Allan DS, Gasser S, Raulet DH, et al. Chemotherapy-induced genotoxic stress promotes sensitivity to natural killer cell cytotoxicity by enabling missing-self recognition. Cancer Res (2010) 70(18):7102–13. doi:10.1158/0008-5472.CAN-10-1316

37. Venkataraman GM, Suciu D, Groh V, Boss JM, Spies T. Promoter region architecture and transcriptional regulation of the genes for the MHC class I-related chain A and B ligands of NKG2D. J Immunol (2007) 178(2):961–9. doi:10.4049/jimmunol.178.2.961

38. Nice TJ, Coscoy L, Raulet DH. Posttranslational regulation of the NKG2D ligand Mult1 in response to cell stress. J Exp Med (2009) 206(2):287–98. doi:10.1084/jem.20081335

39. Diermayr S, Himmelreich H, Durovic B, Mathys-Schneeberger A, Siegler U, Langenkamp U, et al. NKG2D ligand expression in AML increases in response to HDAC inhibitor valproic acid and contributes to allorecognition by NK-cell lines with single KIR-HLA class I specificities. Blood (2008) 111(3):1428–36. doi:10.1182/blood-2007-07-101311

40. Shi P, Yin T, Zhou F, Cui P, Gou S, Wang C. Valproic acid sensitizes pancreatic cancer cells to natural killer cell-mediated lysis by upregulating MICA and MICB via the PI3K/Akt signaling pathway. BMC Cancer (2014) 14:370. doi:10.1186/1471-2407-14-370

41. Armeanu S, Bitzer M, Lauer UM, Venturelli S, Pathil A, Krusch M, et al. Natural killer cell-mediated lysis of hepatoma cells via specific induction of NKG2D ligands by the histone deacetylase inhibitor sodium valproate. Cancer Res (2005) 65(14):6321–9. doi:10.1158/0008-5472.CAN-04-4252

42. Kato N, Tanaka J, Sugita J, Toubai T, Miura Y, Ibata M, et al. Regulation of the expression of MHC class I-related chain A, B (MICA, MICB) via chromatin remodeling and its impact on the susceptibility of leukemic cells to the cytotoxicity of NKG2D-expressing cells. Leukemia (2007) 21(10):2103–8. doi:10.1038/sj.leu.2404862

43. Pfeiffer MM, Burow H, Schleicher S, Handgretinger R, Lang P. Influence of histone deacetylase inhibitors and DNA-methyltransferase inhibitors on the NK cell-mediated lysis of pediatric B-lineage leukemia. Front Oncol (2013) 3:99. doi:10.3389/fonc.2013.00099

44. Fiegler N, Textor S, Arnold A, Rolle A, Oehme I, Breuhahn K, et al. Downregulation of the activating NKp30 ligand B7-H6 by HDAC inhibitors impairs tumor cell recognition by NK cells. Blood (2013) 122(5):684–93. doi:10.1182/blood-2013-02-482513

45. Wu X, Tao Y, Hou J, Meng X, Shi J. Valproic acid upregulates NKG2D ligand expression through an ERK-dependent mechanism and potentially enhances NK cell-mediated lysis of myeloma. Neoplasia (2012) 14(12):1178–89. doi:10.1593/neo.121236

46. Yang H, Lan P, Hou Z, Guan Y, Zhang J, Xu W, et al. Histone deacetylase inhibitor SAHA epigenetically regulates miR-17-92 cluster and MCM7 to upregulate MICA expression in hepatoma. Br J Cancer (2015) 112(1):112–21. doi:10.1038/bjc.2014.547

47. Fionda C, Abruzzese MP, Zingoni A, Cecere F, Vulpis E, Peruzzi G, et al. The IMiDs targets IKZF-1/3 and IRF4 as novel negative regulators of NK cell-activating ligands expression in multiple myeloma. Oncotarget (2015) 6(27):23609–30. doi:10.18632/oncotarget.4603

48. Bellucci R, Martin A, Bommarito D, Wang K, Hansen SH, Freeman GJ, et al. Interferon-gamma-induced activation of JAK1 and JAK2 suppresses tumor cell susceptibility to NK cells through upregulation of PD-L1 expression. Oncoimmunology (2015) 4(6):e1008824. doi:10.1080/2162402X.2015.1008824

49. Ho V, Lim TS, Lee J, Steinberg J, Szmyd R, Tham M, et al. TLR3 agonist and Sorafenib combinatorial therapy promotes immune activation and controls hepatocellular carcinoma progression. Oncotarget (2015) 6(29):27252–66. doi:10.18632/oncotarget.4583

50. Soriani A, Borrelli C, Ricci B, Molfetta R, Zingoni A, Fionda C, et al. p38 MAPK differentially controls NK activating ligands at transcriptional and post-transcriptional level on multiple myeloma cells. Oncoimmunology (2017) 6(1):e1264564. doi:10.1080/2162402X.2016.1264564

51. Acebes-Huerta A, Lorenzo-Herrero S, Folgueras AR, Huergo-Zapico L, Lopez-Larrea C, Lopez-Soto A, et al. Drug-induced hyperploidy stimulates an antitumor NK cell response mediated by NKG2D and DNAM-1 receptors. Oncoimmunology (2016) 5(2):e1074378. doi:10.1080/2162402X.2015.1074378

52. Niu C, Jin H, Li M, Zhu S, Zhou L, Jin F, et al. Low-dose bortezomib increases the expression of NKG2D and DNAM-1 ligands and enhances induced NK and gammadelta T cell-mediated lysis in multiple myeloma. Oncotarget (2017) 8(4):5954–64. doi:10.18632/oncotarget.13979

53. Soriani A, Fionda C, Ricci B, Iannitto ML, Cippitelli M, Santoni A. Chemotherapy-elicited upregulation of NKG2D and DNAM-1 ligands as a therapeutic target in multiple myeloma. Oncoimmunology (2013) 2(12):e26663. doi:10.4161/onci.26663

54. Zingoni A, Vulpis E, Nardone I, Soriani A, Fionda C, Cippitelli M, et al. Targeting NKG2D and NKp30 ligands shedding to improve NK cell-based immunotherapy. Crit Rev Immunol (2016) 36(6):445–60. doi:10.1615/CritRevImmunol.2017020166

55. Zingoni A, Cecere F, Vulpis E, Fionda C, Molfetta R, Soriani A, et al. Genotoxic stress induces senescence-associated ADAM10-dependent release of NKG2D MIC ligands in multiple myeloma cells. J Immunol (2015) 195(2):736–48. doi:10.4049/jimmunol.1402643

56. Lin X, Huang M, Xie F, Zhou H, Yang J, Huang Q. Gemcitabine inhibits immune escape of pancreatic cancer by down regulating the soluble ULBP2 protein. Oncotarget (2016) 7(43):70092–9. doi:10.18632/oncotarget.11780

57. Raneros AB, Puras AM, Rodriguez RM, Colado E, Bernal T, Anguita E, et al. Increasing TIMP3 expression by hypomethylating agents diminishes soluble MICA, MICB and ULBP2 shedding in acute myeloid leukemia, facilitating NK cell-mediated immune recognition. Oncotarget (2017) 8(19):31959–76. doi:10.18632/oncotarget.16657

58. Elrod HA, Sun SY. Modulation of death receptors by cancer therapeutic agents. Cancer Biol Ther (2008) 7(2):163–73. doi:10.4161/cbt.7.2.5335

59. Kim YH, Park JW, Lee JY, Kwon TK. Sodium butyrate sensitizes TRAIL-mediated apoptosis by induction of transcription from the DR5 gene promoter through Sp1 sites in colon cancer cells. Carcinogenesis (2004) 25(10):1813–20. doi:10.1093/carcin/bgh188

60. Nakata S, Yoshida T, Horinaka M, Shiraishi T, Wakada M, Sakai T. Histone deacetylase inhibitors upregulate death receptor 5/TRAIL-R2 and sensitize apoptosis induced by TRAIL/APO2-L in human malignant tumor cells. Oncogene (2004) 23(37):6261–71. doi:10.1038/sj.onc.1207830

61. Hirai S, Endo S, Saito R, Hirose M, Ueno T, Suzuki H, et al. Antitumor effects of a sirtuin inhibitor, tenovin-6, against gastric cancer cells via death receptor 5 up-regulation. PLoS One (2014) 9(7):e102831. doi:10.1371/journal.pone.0102831

62. Guo F, Sigua C, Tao J, Bali P, George P, Li Y, et al. Cotreatment with histone deacetylase inhibitor LAQ824 enhances Apo-2L/tumor necrosis factor-related apoptosis inducing ligand-induced death inducing signaling complex activity and apoptosis of human acute leukemia cells. Cancer Res (2004) 64(7):2580–9. doi:10.1158/0008-5472.CAN-03-2629

63. Shankar S, Singh TR, Fandy TE, Luetrakul T, Ross DD, Srivastava RK. Interactive effects of histone deacetylase inhibitors and TRAIL on apoptosis in human leukemia cells: involvement of both death receptor and mitochondrial pathways. Int J Mol Med (2005) 16(6):1125–38. doi:10.3892/ijmm.16.6.1125

64. Fandy TE, Shankar S, Ross DD, Sausville E, Srivastava RK. Interactive effects of HDAC inhibitors and TRAIL on apoptosis are associated with changes in mitochondrial functions and expressions of cell cycle regulatory genes in multiple myeloma. Neoplasia (2005) 7(7):646–57. doi:10.1593/neo.04655

65. Insinga A, Monestiroli S, Ronzoni S, Gelmetti V, Marchesi F, Viale A, et al. Inhibitors of histone deacetylases induce tumor-selective apoptosis through activation of the death receptor pathway. Nat Med (2005) 11(1):71–6. doi:10.1038/nm0205-233a

66. Lundqvist A, Abrams SI, Schrump DS, Alvarez G, Suffredini D, Berg M, et al. Bortezomib and depsipeptide sensitize tumors to tumor necrosis factor-related apoptosis-inducing ligand: a novel method to potentiate natural killer cell tumor cytotoxicity. Cancer Res (2006) 66(14):7317–25. doi:10.1158/0008-5472.CAN-06-0680

67. Lundqvist A, Yokoyama H, Smith A, Berg M, Childs R. Bortezomib treatment and regulatory T-cell depletion enhance the antitumor effects of adoptively infused NK cells. Blood (2009) 113(24):6120–7. doi:10.1182/blood-2008-11-190421

68. Liu X, Yue P, Chen S, Hu L, Lonial S, Khuri FR, et al. The proteasome inhibitor PS-341 (bortezomib) up-regulates DR5 expression leading to induction of apoptosis and enhancement of TRAIL-induced apoptosis despite up-regulation of c-FLIP and survivin expression in human NSCLC cells. Cancer Res (2007) 67(10):4981–8. doi:10.1158/0008-5472.CAN-06-4274

69. Kabore AF, Sun J, Hu X, McCrea K, Johnston JB, Gibson SB. The TRAIL apoptotic pathway mediates proteasome inhibitor induced apoptosis in primary chronic lymphocytic leukemia cells. Apoptosis (2006) 11(7):1175–93. doi:10.1007/s10495-006-8048-9

70. He Q, Huang Y, Sheikh MS. Proteasome inhibitor MG132 upregulates death receptor 5 and cooperates with Apo2L/TRAIL to induce apoptosis in Bax-proficient and -deficient cells. Oncogene (2004) 23(14):2554–8. doi:10.1038/sj.onc.1207351

71. Yoshida T, Shiraishi T, Nakata S, Horinaka M, Wakada M, Mizutani Y, et al. Proteasome inhibitor MG132 induces death receptor 5 through CCAAT/enhancer-binding protein homologous protein. Cancer Res (2005) 65(13):5662–7. doi:10.1158/0008-5472.CAN-05-0693

72. Wu GS, Burns TF, McDonald ER III, Jiang W, Meng R, Krantz ID, et al. KILLER/DR5 is a DNA damage-inducible p53-regulated death receptor gene. Nat Genet (1997) 17(2):141–3. doi:10.1038/ng1097-141

73. Takimoto R, El-Deiry WS. Wild-type p53 transactivates the KILLER/DR5 gene through an intronic sequence-specific DNA-binding site. Oncogene (2000) 19(14):1735–43. doi:10.1038/sj.onc.1203489

74. Guan B, Yue P, Clayman GL, Sun SY. Evidence that the death receptor DR4 is a DNA damage-inducible, p53-regulated gene. J Cell Physiol (2001) 188(1):98–105. doi:10.1002/jcp.1101

75. Liu X, Yue P, Khuri FR, Sun SY. p53 upregulates death receptor 4 expression through an intronic p53 binding site. Cancer Res (2004) 64(15):5078–83. doi:10.1158/0008-5472.CAN-04-1195

76. Shetty S, Graham BA, Brown JG, Hu X, Vegh-Yarema N, Harding G, et al. Transcription factor NF-kappaB differentially regulates death receptor 5 expression involving histone deacetylase 1. Mol Cell Biol (2005) 25(13):5404–16. doi:10.1128/MCB.25.13.5404-5416.2005

77. Muller M, Wilder S, Bannasch D, Israeli D, Lehlbach K, Li-Weber M, et al. p53 activates the CD95 (APO-1/Fas) gene in response to DNA damage by anticancer drugs. J Exp Med (1998) 188(11):2033–45. doi:10.1084/jem.188.11.2033

78. Sheikh MS, Burns TF, Huang Y, Wu GS, Amundson S, Brooks KS, et al. el-Deiry WS: p53-dependent and -independent regulation of the death receptor KILLER/DR5 gene expression in response to genotoxic stress and tumor necrosis factor alpha. Cancer Res (1998) 58(8):1593–8.

79. Gibson SB, Oyer R, Spalding AC, Anderson SM, Johnson GL. Increased expression of death receptors 4 and 5 synergizes the apoptosis response to combined treatment with etoposide and TRAIL. Mol Cell Biol (2000) 20(1):205–12. doi:10.1128/MCB.20.1.205-212.2000

80. Nagane M, Pan G, Weddle JJ, Dixit VM, Cavenee WK, Huang HJ. Increased death receptor 5 expression by chemotherapeutic agents in human gliomas causes synergistic cytotoxicity with tumor necrosis factor-related apoptosis-inducing ligand in vitro and in vivo. Cancer Res (2000) 60(4):847–53.

81. Wen J, Ramadevi N, Nguyen D, Perkins C, Worthington E, Bhalla K. Antileukemic drugs increase death receptor 5 levels and enhance Apo-2L-induced apoptosis of human acute leukemia cells. Blood (2000) 96(12):3900–6.

82. Srivastava RK, Mi QS, Hardwick JM, Longo DL. Deletion of the loop region of Bcl-2 completely blocks paclitaxel-induced apoptosis. Proc Natl Acad Sci U S A (1999) 96(7):3775–80. doi:10.1073/pnas.96.7.3775

83. Eliopoulos AG, Kerr DJ, Herod J, Hodgkins L, Krajewski S, Reed JC, et al. The control of apoptosis and drug resistance in ovarian cancer: influence of p53 and Bcl-2. Oncogene (1995) 11(7):1217–28.

84. Siemer S, Ornskov D, Guerra B, Boldyreff B, Issinger OG. Determination of mRNA, and protein levels of p53, MDM2 and protein kinase CK2 subunits in F9 cells after treatment with the apoptosis-inducing drugs cisplatin and carboplatin. Int J Biochem Cell Biol (1999) 31(6):661–70. doi:10.1016/S1357-2725(99)00020-5

85. Tolis C, Peters GJ, Ferreira CG, Pinedo HM, Giaccone G. Cell cycle disturbances and apoptosis induced by topotecan and gemcitabine on human lung cancer cell lines. Eur J Cancer (1999) 35(5):796–807. doi:10.1016/S0959-8049(98)00425-0

86. Wang S, Wang Z, Boise L, Dent P, Grant S. Loss of the bcl-2 phosphorylation loop domain increases resistance of human leukemia cells (U937) to paclitaxel-mediated mitochondrial dysfunction and apoptosis. Biochem Biophys Res Commun (1999) 259(1):67–72. doi:10.1006/bbrc.1999.0669

87. Garg AD, Galluzzi L, Apetoh L, Baert T, Birge RB, Bravo-San Pedro JM, et al. Molecular and translational classifications of DAMPs in immunogenic cell death. Front Immunol (2015) 6:588. doi:10.3389/fimmu.2015.00588

88. Apetoh L, Ghiringhelli F, Tesniere A, Obeid M, Ortiz C, Criollo A, et al. Toll-like receptor 4-dependent contribution of the immune system to anticancer chemotherapy and radiotherapy. Nat Med (2007) 13(9):1050–9. doi:10.1038/nm1622

89. Chen T, Guo J, Han C, Yang M, Cao X. Heat shock protein 70, released from heat-stressed tumor cells, initiates antitumor immunity by inducing tumor cell chemokine production and activating dendritic cells via TLR4 pathway. J Immunol (2009) 182(3):1449–59. doi:10.4049/jimmunol.182.3.1449

90. Lotze MT, Tracey KJ. High-mobility group box 1 protein (HMGB1): nuclear weapon in the immune arsenal. Nat Rev Immunol (2005) 5(4):331–42. doi:10.1038/nri1594

91. Wu J, Waxman DJ. Metronomic cyclophosphamide eradicates large implanted GL261 gliomas by activating antitumor Cd8+ T-cell responses and immune memory. Oncoimmunology (2015) 4(4):e1005521. doi:10.1080/2162402X.2015.1005521

92. Guerriero JL, Ditsworth D, Catanzaro JM, Sabino G, Furie MB, Kew RR, et al. DNA alkylating therapy induces tumor regression through an HMGB1-mediated activation of innate immunity. J Immunol (2011) 186(6):3517–26. doi:10.4049/jimmunol.1003267

93. Parodi M, Pedrazzi M, Cantoni C, Averna M, Patrone M, Cavaletto M, et al. Natural Killer (NK)/melanoma cell interaction induces NK-mediated release of chemotactic High Mobility Group Box-1 (HMGB1) capable of amplifying NK cell recruitment. Oncoimmunology (2015) 4(12):e1052353. doi:10.1080/2162402X.2015.1052353

94. Semino C, Angelini G, Poggi A, Rubartelli A. NK/iDC interaction results in IL-18 secretion by DCs at the synaptic cleft followed by NK cell activation and release of the DC maturation factor HMGB1. Blood (2005) 106(2):609–16. doi:10.1182/blood-2004-10-3906

95. Semino C, Ceccarelli J, Lotti LV, Torrisi MR, Angelini G, Rubartelli A. The maturation potential of NK cell clones toward autologous dendritic cells correlates with HMGB1 secretion. J Leukoc Biol (2007) 81(1):92–9. doi:10.1189/jlb.0306172

96. Tang D, Kang R, Livesey KM, Cheh CW, Farkas A, Loughran P, et al. Endogenous HMGB1 regulates autophagy. J Cell Biol (2010) 190(5):881–92. doi:10.1083/jcb.200911078

97. Valdor R, Macian F. Autophagy and the regulation of the immune response. Pharmacol Res (2012) 66(6):475–83. doi:10.1016/j.phrs.2012.10.003

98. Multhoff G, Botzler C, Wiesnet M, Muller E, Meier T, Wilmanns W, et al. A stress-inducible 72-kDa heat-shock protein (HSP72) is expressed on the surface of human tumor cells, but not on normal cells. Int J Cancer (1995) 61(2):272–9. doi:10.1002/ijc.2910610222

99. Gehrmann M, Schmetzer H, Eissner G, Haferlach T, Hiddemann W, Multhoff G. Membrane-bound heat shock protein 70 (Hsp70) in acute myeloid leukemia: a tumor specific recognition structure for the cytolytic activity of autologous NK cells. Haematologica (2003) 88(4):474–6.

100. Pfister K, Radons J, Busch R, Tidball JG, Pfeifer M, Freitag L, et al. Patient survival by Hsp70 membrane phenotype: association with different routes of metastasis. Cancer (2007) 110(4):926–35. doi:10.1002/cncr.22864

101. Kleinjung T, Arndt O, Feldmann HJ, Bockmuhl U, Gehrmann M, Zilch T, et al. Heat shock protein 70 (Hsp70) membrane expression on head-and-neck cancer biopsy-a target for natural killer (NK) cells. Int J Radiat Oncol Biol Phys (2003) 57(3):820–6. doi:10.1016/S0360-3016(03)00629-1

102. Gehrmann M, Pfister K, Hutzler P, Gastpar R, Margulis B, Multhoff G. Effects of antineoplastic agents on cytoplasmic and membrane-bound heat shock protein 70 (Hsp70) levels. Biol Chem (2002) 383(11):1715–25. doi:10.1515/BC.2002.192

103. Gehrmann M, Schonberger J, Zilch T, Rossbacher L, Thonigs G, Eilles C, et al. Retinoid- and sodium-butyrate-induced decrease in heat shock protein 70 membrane-positive tumor cells is associated with reduced sensitivity to natural killer cell lysis, growth delay, and altered growth morphology. Cell Stress Chaperones (2005) 10(2):136–46. doi:10.1379/CSC-88R1.1

104. Gross C, Holler E, Stangl S, Dickinson A, Pockley AG, Asea AA, et al. An Hsp70 peptide initiates NK cell killing of leukemic blasts after stem cell transplantation. Leuk Res (2008) 32(4):527–34. doi:10.1016/j.leukres.2007.03.027

105. Multhoff G, Pfister K, Botzler C, Jordan A, Scholz R, Schmetzer H, et al. Adoptive transfer of human natural killer cells in mice with severe combined immunodeficiency inhibits growth of Hsp70-expressing tumors. Int J Cancer (2000) 88(5):791–7. doi:10.1002/1097-0215(20001201)88:5<791::AID-IJC17>3.0.CO;2-I

106. Elsner L, Flugge PF, Lozano J, Muppala V, Eiz-Vesper B, Demiroglu SY, et al. The endogenous danger signals HSP70 and MICA cooperate in the activation of cytotoxic effector functions of NK cells. J Cell Mol Med (2010) 14(4):992–1002. doi:10.1111/j.1582-4934.2009.00677.x

107. Elsner L, Muppala V, Gehrmann M, Lozano J, Malzahn D, Bickeboller H, et al. The heat shock protein HSP70 promotes mouse NK cell activity against tumors that express inducible NKG2D ligands. J Immunol (2007) 179(8):5523–33. doi:10.4049/jimmunol.179.8.5523

108. Gastpar R, Gehrmann M, Bausero MA, Asea A, Gross C, Schroeder JA, et al. Heat shock protein 70 surface-positive tumor exosomes stimulate migratory and cytolytic activity of natural killer cells. Cancer Res (2005) 65(12):5238–47. doi:10.1158/0008-5472.CAN-04-3804

109. Lv LH, Wan YL, Lin Y, Zhang W, Yang M, Li GL, et al. Anticancer drugs cause release of exosomes with heat shock proteins from human hepatocellular carcinoma cells that elicit effective natural killer cell antitumor responses in vitro. J Biol Chem (2012) 287(19):15874–85. doi:10.1074/jbc.M112.340588

110. Vulpis E, Cecere F, Molfetta R, Soriani A, Fionda C, Peruzzi G, et al. Genotoxic stress modulates the release of exosomes from multiple myeloma cells capable of activating NK cell cytokine production: role of HSP70/TLR2/NF-kB axis. Oncoimmunology (2017) 6(3):e1279372. doi:10.1080/2162402X.2017.1279372

111. Lancaster GI, Febbraio MA. Exosome-dependent trafficking of HSP70: a novel secretory pathway for cellular stress proteins. J Biol Chem (2005) 280(24):23349–55. doi:10.1074/jbc.M502017200

112. Gobbo J, Marcion G, Cordonnier M, Dias AMM, Pernet N, Hammann A, et al. Restoring anticancer immune response by targeting tumor-derived exosomes with a HSP70 peptide aptamer. J Natl Cancer Inst (2016) 108(3):djv330. doi:10.1093/jnci/djv330

113. Pellom ST Jr, Dudimah DF, Thounaojam MC, Sayers TJ, Shanker A. Modulatory effects of bortezomib on host immune cell functions. Immunotherapy (2015) 7(9):1011–22. doi:10.2217/imt.15.66

114. Armeanu S, Krusch M, Baltz KM, Weiss TS, Smirnow I, Steinle A, et al. Direct and natural killer cell-mediated antitumor effects of low-dose bortezomib in hepatocellular carcinoma. Clin Cancer Res (2008) 14(11):3520–8. doi:10.1158/1078-0432.CCR-07-4744

115. Wang X, Feng X, Wang J, Shao N, Ji C, Ma D, et al. Bortezomib and IL-12 produce synergetic anti-multiple myeloma effects with reduced toxicity to natural killer cells. Anticancer Drugs (2014) 25(3):282–8. doi:10.1097/CAD.0000000000000058

116. Khan T, Stauffer JK, Williams R, Hixon JA, Salcedo R, Lincoln E, et al. Proteasome inhibition to maximize the apoptotic potential of cytokine therapy for murine neuroblastoma tumors. J Immunol (2006) 176(10):6302–12. doi:10.4049/jimmunol.176.10.6302

117. Ni L, Wang L, Yao C, Ni Z, Liu F, Gong C, et al. The histone deacetylase inhibitor valproic acid inhibits NKG2D expression in natural killer cells through suppression of STAT3 and HDAC3. Sci Rep (2017) 7:45266. doi:10.1038/srep45266

118. Ogbomo H, Michaelis M, Kreuter J, Doerr HW, Cinatl J Jr. Histone deacetylase inhibitors suppress natural killer cell cytolytic activity. FEBS Lett (2007) 581(7):1317–22. doi:10.1016/j.febslet.2007.02.045

119. Kelly-Sell MJ, Kim YH, Straus S, Benoit B, Harrison C, Sutherland K, et al. The histone deacetylase inhibitor, romidepsin, suppresses cellular immune functions of cutaneous T-cell lymphoma patients. Am J Hematol (2012) 87(4):354–60. doi:10.1002/ajh.23112

120. Zhu S, Denman CJ, Cobanoglu ZS, Kiany S, Lau CC, Gottschalk SM, et al. The narrow-spectrum HDAC inhibitor entinostat enhances NKG2D expression without NK cell toxicity, leading to enhanced recognition of cancer cells. Pharm Res (2015) 32(3):779–92. doi:10.1007/s11095-013-1231-0

121. Medon M, Vidacs E, Vervoort SJ, Li J, Jenkins MR, Ramsbottom KM, et al. HDAC inhibitor panobinostat engages host innate immune defenses to promote the tumoricidal effects of trastuzumab in HER2+ tumors. Cancer Res (2017) 77(10):2594–606. doi:10.1158/0008-5472.CAN-16-2247

122. Schmiedel BJ, Arelin V, Gruenebach F, Krusch M, Schmidt SM, Salih HR. Azacytidine impairs NK cell reactivity while decitabine augments NK cell responsiveness toward stimulation. Int J Cancer (2011) 128(12):2911–22. doi:10.1002/ijc.25635

123. Sohlberg E, Pfefferle A, Andersson S, Baumann BC, Hellstrom-Lindberg E, Malmberg KJ. Imprint of 5-azacytidine on the natural killer cell repertoire during systemic treatment for high-risk myelodysplastic syndrome. Oncotarget (2015) 6(33):34178–90. doi:10.18632/oncotarget.6213

124. Schonefeldt C, Sockel K, Wehner R, Sopper S, Wolf D, Wermke M, et al. Azacytidine impairs NK cell activity in AML and MDS patients undergoing MRD-based pre-emptive treatment after allogeneic stem cell transplantation. Blood Cancer J (2013) 3:e136. doi:10.1038/bcj.2013.35

125. Sehgal K, Das R, Zhang L, Verma R, Deng Y, Kocoglu M, et al. Clinical and pharmacodynamic analysis of pomalidomide dosing strategies in myeloma: impact of immune activation and cereblon targets. Blood (2015) 125(26):4042–51. doi:10.1182/blood-2014-11-611426

126. Lagrue K, Carisey A, Morgan DJ, Chopra R, Davis DM. Lenalidomide augments actin remodeling and lowers NK-cell activation thresholds. Blood (2015) 126(1):50–60. doi:10.1182/blood-2015-01-625004

127. Acebes-Huerta A, Huergo-Zapico L, Gonzalez-Rodriguez AP, Fernandez-Guizan A, Payer AR, Lopez-Soto A, et al. Lenalidomide induces immunomodulation in chronic lymphocytic leukemia and enhances antitumor immune responses mediated by NK and CD4 T cells. Biomed Res Int (2014) 2014:265840. doi:10.1155/2014/265840

128. Shortt J, Hsu AK, Johnstone RW. Thalidomide-analogue biology: immunological, molecular and epigenetic targets in cancer therapy. Oncogene (2013) 32(36):4191–202. doi:10.1038/onc.2012.599

129. Sidaway P. Haematological cancer: rituximab enhances responses to lenalidomide. Nat Rev Clin Oncol (2017) 14(2):70. doi:10.1038/nrclinonc.2016.209

130. Gormley NJ, Ko CW, Deisseroth A, Nie L, Kaminskas E, Kormanik N, et al. FDA drug approval: elotuzumab in combination with lenalidomide and dexamethasone for the treatment of relapsed or refractory multiple myeloma. Clin Cancer Res (2017). doi:10.1158/1078-0432.CCR-16-2870

131. Benson DM Jr, Cohen AD, Jagannath S, Munshi NC, Spitzer G, Hofmeister CC, et al. A phase I trial of the anti-KIR antibody IPH2101 and lenalidomide in patients with relapsed/refractory multiple myeloma. Clin Cancer Res (2015) 21(18):4055–61. doi:10.1158/1078-0432.CCR-15-0304

132. Nijhof IS, Lammerts van Bueren JJ, van Kessel B, Andre P, Morel Y, Lokhorst HM, et al. Daratumumab-mediated lysis of primary multiple myeloma cells is enhanced in combination with the human anti-KIR antibody IPH2102 and lenalidomide. Haematologica (2015) 100(2):263–8. doi:10.3324/haematol.2014.117531

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Zingoni, Fionda, Borrelli, Cippitelli, Santoni and Soriani. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

REVIEW
published: 05 January 2018
doi: 10.3389/fimmu.2017.01908

S100 Proteins As an Important Regulator of Macrophage Inflammation

Chang Xia1,2, Zachary Braunstein[3], Amelia C. Toomey[4], Jixin Zhong[2]* and Xiaoquan Rao[2]*

1 College of Health Science and Nursing, Wuhan Polytechnic University, Wuhan, China

2 Cardiovascular Research Institute, Case Western Reserve University, Cleveland, OH, United States

3 Boonshoft School of Medicine, Wright State University, Dayton, OH, United States

4 Department of Health Sciences, University of Missouri, Columbia, MO, United States

OPEN ACCESS

Edited by:

Leticia A. Carneiro, Universidade Federal do Rio de Janeiro, Brazil

Reviewed by:

Charles E. McCall, Wake Forest Baptist Medical Center, United States
Penghua Yang, University of Maryland, Baltimore, United States

*Correspondence:

Jixin Zhong
jixin.zhong@case.edu;
Xiaoquan Rao
jannet18@163.com

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 06 October 2017
Accepted: 14 December 2017
Published: 05 January 2018

Citation:

Xia C, Braunstein Z, Toomey AC, Zhong J and Rao X (2018) S100 Proteins As an Important Regulator of Macrophage Inflammation. Front. Immunol. 8:1908. doi: 10.3389/fimmu.2017.01908

The S100 proteins, a family of calcium-binding cytosolic proteins, have a broad range of intracellular and extracellular functions through regulating calcium balance, cell apoptosis, migration, proliferation, differentiation, energy metabolism, and inflammation. The intracellular functions of S100 proteins involve interaction with intracellular receptors, membrane protein recruitment/transportation, transcriptional regulation and integrating with enzymes or nucleic acids, and DNA repair. The S100 proteins could also be released from the cytoplasm, induced by tissue/cell damage and cellular stress. The extracellular S100 proteins, serving as a danger signal, are crucial in regulating immune homeostasis, post-traumatic injury, and inflammation. Extracellular S100 proteins are also considered biomarkers for some specific diseases. In this review, we will discuss the multi-functional roles of S100 proteins, especially their potential roles associated with cell migration, differentiation, tissue repair, and inflammation.

Keywords: S100 proteins, inflammation, tissue repair, biomarkers, inflammatory disease, macrophages

INTRODUCTION

The S100 proteins, belonging to a calcium-binding cytosolic protein family, are composed of 25 known members (1–4). They have a broad range of intracellular and extracellular functions encompassing regulation cell apoptosis, proliferation, differentiation, migration, energy metabolism, calcium balance, protein phosphorylation, and inflammation (5–8).

Based on their functional roles, s100 proteins are categorized into three main subgroups: S100 proteins that only exert intracellular functions, S100 proteins that have both intracellular and extracellular roles, and S100 proteins that mainly possess extracellular effects (7). The S100 proteins within the first subgroup only exert intracellular functions. For example, S100A1 is predominantly expressed in striated muscle (especially cardiac muscle) (9) and only exert intracellular regulatory effects such as regulating SR Ca2+ recycle and enhancing the gain of the calcium-induced calcium release (CICR) cascade (10–12). In addition to intracellular roles, some S100 proteins are released into the extracellular environment and may exert extracellular functions. S100B in this subgroup was known to directly interact with nuclear Dbf2-related protein kinase (NDR kinase) and block the recruitment of its substrates to NDR kinase (13). Furthermore, extracellular S100B could also activate extracellular signal-regulated protein kinase (ERK) and NFκB in chondrocytes by binding to its cell surface receptor, receptor for advanced glycation end products (RAGE) (14). The third subgroup of S100 proteins such as S100A15 mainly exerts extracellular regulatory functions. These of S100 proteins are considered as potential therapeutic targets for various human disorders, including arthritis, cancer, and Alzheimer’s disease (15, 16).

S100 proteins are involved in multiple intracellular functions which include: interacting with intracellular receptors or molecule subunits (17), membrane protein recruitment and transportation, transcriptional regulation (18, 19), regulating enzymes, nucleic acids, and DNA repair (20, 21) (Figure 1). There are two critical steps for S100 protein activation: Ca2+ binding (22) and homo- or hetero-dimer formation (23). Each S100 protein forming the dimer participates in ion (Ca2+, Zn2+, or Cu2+) binding. Ca2+ also contributes to the formation of S100 protein oligomers, especially calprotectin (S100A8/A9 tetramer) (22, 24, 25).

FIGURE 1 | Involvement of S100 proteins in stress and inflammation-mediated responses. Cell stress or inflammation induce the release of S100 proteins to acellular compartment where they bind cell surface receptors such as RAGE, TLR4, CD147, and GPCR. The interactions between S100 proteins and their receptors activate intracellular signaling pathways such as AP1 and NFκB, which further initiates multiple cellular processes such as cell differentiation, migration, apoptosis, proliferation, and inflammation. AP1, activator protein 1; ERK, extracellular signal-regulated protein kinase; GPCR, G-protein-coupled receptor; IL-1, interleukin 1; IL-7, interleukin 7; IκBα, nuclear factor of kappa light polypeptide gene enhancer in B-cells inhibitor alpha; JNK, c-Jun N-terminal kinase; P38, p38 mitogen-activated protein kinase; RAGE, receptor for advanced glycation end products; TLR4, toll-like receptor 4; Traf2, TNF receptor-associated factor 2.

When released to the extracellular space, S100 proteins have crucial activities in the regulation of immune homeostasis, post-traumatic injury, and inflammation. S100 proteins trigger inflammation through interacting with receptors RAGE and TLR4 (26). Increasing evidence has demonstrated that calprotectin (S100A8/A9) is an endogenous agonist of TLR4 (26). Binding to TLR4 initiates a signaling cascade and regulates inflammation, cell proliferation, differentiation, and tumor development in an NF-κB-dependent manner (8, 26–28). Apart from TLR4, RAGE has also been suggested to bind S100 proteins such as S100A7, S100A12, S100A8/A9, and S100B (27, 29–31). By interacting with RAGE, S100 proteins activate NF-κB, inducing the production of pro-inflammatory cytokines leading to the migration of neutrophils, monocytes, and macrophages (30, 31). In addition to the NF-κB pathway, MAP kinase-mediated signaling is also induced by S100 proteins such as S100P (32, 33). Interestingly, S100A6 activates RAGE and promotes apoptosis, while S100B inactivates RAGE by interacting with the basic fibroblast growth factor and its receptor (14, 34). Extracellular S100 proteins may regulate the apoptosis, proliferation, differentiation, and migration of a number of cell types including monocytes, macrophages, neutrophils, lymphocytes, myoblast, epithelial cells, endothelial cells, smooth muscle cells, neurons, and fibroblasts. In this review, we aim to summarize the immune regulatory role of S100 proteins and their potential involvement in inflammatory regulation, tissue repair, and tumorigenesis.

S100 GENES AND MOLECULAR STRUCTURE

Each S100 family protein is encoded by a separate gene. Most S100 genes are located within the chromosome 1q21 with a few exceptions. For example, S100A11P is located within chromosome 7q22-q3, S100B in located within chromosome 21q22, S100P is located in chromosome 4p16, S100G is located in chromosome Xp22, and S100Z is located with chromosome 5q13 (5). The sequence homology among S100 proteins varies from 22 to 57%, which is mainly due to the variance at the hinge region and C-terminus, the regions associated with their function (35).

S100 proteins are small proteins with a molecular weight of 10–12 kDa. Each S100 protein consists of two EF-hand helix–loop–helix structural motifs, which are arranged in a back-to-back manner and linked with a flexible hinge (23). The activity of the proteins is regulated by metal ions (such as calcium, zinc, and copper), which modulates the folding and oligomerization of the protein (36, 37).

EXPRESSION PATTERN AND REGULATION

Epigenetic mechanisms play a key role in the regulation of S100 protein expression. S100A3, S100A10, S10011, and S100P could be detected in various medulloblastoma cell lines treated with DNA de-methylation (38). It is reported that DNA hypomethylation could induce S100A6 overexpression in gastric cancer. Lower levels of CpG methylation in the first intron and second exon regions of the S100A6 gene, accompanied by higher levels of acetylated histone H3 binding to the promoter, have been reported in the gastric cancer tissues (39). Lower methylation in the proximal promoter region of the S100P gene was also found in prostate cancer cell lines (40). The expression of S100 proteins may also be regulated by micro RNAs, although further studies are needed to provide direct evidence. NFAT5, a transcription factor that initiates S100A4 expression (41), is regulated by miR-568 (42).

The expression of S100 proteins is strictly regulated to maintain immune homeostasis (7, 43). S100A8 and S100A9 are predominately expressed in monocytes, neutrophils, and dendritic cells (44, 45). However, they are also expressed in various other types of cells upon activation, such as fibroblasts (46), mature macrophages (47), vascular endothelial cells (48–50), and keratinocytes (51). In neutrophils, 45% of the cytosolic proteins are constituted with S100A8 and S100A9, whereas the proportion is only 1% in monocytes (52). The expression levels in different monocyte subsets also vary. The level of S100A8 mRNA is higher in classical CD14+/CD16− human monocytes when compared to non-classical CD14+CD16+ monocytes (47).

Increasing evidence indicates that the expression of most S100 proteins is different between physiological and pathological conditions. The expression of S100A8 and S100A9 could be upregulated by a number of conditions such as oxidative stress, specific cytokines, and growth factors in many types of cells (53). S100A12 is mainly expressed in neutrophils, monocytes, and early macrophages (53, 54), but it can also be detected in endothelial cells, keratinocytes, epithelial cells, and pro-inflammatory macrophages under inflammatory conditions (51, 55–58). In human epidermal keratinocytes, interleukin (IL)-1α induces a significant increase of S100A9 expression by the p38 MAPK pathway (59). The expression of S100A5 is upregulated in bladder cancers (60). Pro-inflammatory cytokines could increase S100A7 expression in human breast cancer (61). IL-17, IL-22, and bacterial products (e.g., flagellin) can enhance S100A7 expression in keratinocytes (62). IL-6 and IL-8 released from myofibroblasts could also trigger the upregulation of S100A8/A9 in tumor-infiltrated myeloid cells (63). S100A9 was significantly higher in the peripheral blood in patients with implant-associated osteomyelitis. S100A9 expressing cells were also increased in tissue biopsies from patients with implant infections, compared with the non-infected individuals (64).

S100 PROTEINS FUNCTION AS DAMAGE-ASSOCIATED MOLECULAR PATTERN (DAMP) MOLECULES

In addition to serving as calcium-binding proteins, S100 proteins were later discovered as DAMP molecules (26, 65, 66). DAMPs were considered as a series of intracellular molecules linked with cell death and tissue damage through inducing a rapid inflammatory response or production biologically active molecules (67, 68). DAMPs are biomolecules that are released from damaged or stressed cells and could act as endogenous danger signal to activate inflammatory response (69). S100 proteins could be released from the cells after cell damage/stress or activation of phagocytes such as neutrophils and macrophages. The extracellular S100 proteins then become danger signals and activate immune cells and endothelial cells by binding to the pattern recognition receptors such as TLRs and RAGE.

They play an important role in modulating inflammatory responses (70). Once released from the cell, calprotectins function as an endogenous agonist to bind TLR4 (S100A8/A9 and S100A12) (26) and RAGE (S100A8/A9 and S100A7) [(6, 31, 71) #3535]. In the site of inflammation, calprotectin acts as a chemotactic factor by inducing neutrophils adhesion (72). Furthermore, S100A8/A9 induces apoptosis and autophagy in various cell types such as lymphocytes, macrophages, endothelial cells, and tumor cells (73). It has been shown that reactive oxygen species (ROS) is the critical factor in S100A8/A9-induced cell death and involves BNIP3. The increase of ROS production in mitochondria subsequently causes mitochondrial damage and lysosomal activation (73).

S100 PROTEINS IN MACROPHAGES MIGRATION, INVASION, AND DIFFERENTIATION

It is widely accepted that macrophages contribute to immune defense, immune regulation, and tissue repair. Based on their cytokine production and activation conditions, macrophages are categorized into two populations: pro-inflammatory M1 (classically activated macrophage) and anti-inflammatory M2 (alternatively activated macrophage). Calprotectin could induce pro-inflammatory cytokine production in monocytes and macrophages through NF-κB and p38 MAPK pathways (74). An increasing number of findings demonstrate that S100 proteins contribute to the adhesion and migration of leukocytes. For example, the release of S100A8/A9 has been suggested to facilitate monocyte and neutrophil transmigration (75, 76). The S100A8/A9 heterodimer enhances the expression of β2 integrin CD11b and the ability of adhesion in phagocytes (72, 77). Moreover, the response of S100A9−/− monocytes to chemotaxis was reduced when compared with wild-type cells. For example, IL 8-induced CD11b upregulation was abolished in S100A9−/− monocytes and neutrophils (78). S100A4 has also been shown to interact with cytoskeletal proteins to promote cell migration and deletion of s100a4, which leads to the deficiency of macrophage migration and chemotactic reactions (79–81). S100A12 induced the production of pro-inflammatory cytokine IL-6 and IL-8 in both a dose-dependent and time-dependent manner. This was critical to regulate the recruitment of monocytes and TNF-α release (82).

The intimate relationship between macrophages and cancer cells plays a crucial role in tumor growth and metastasis. Tumor associated macrophages influence tumor growth by modulating local inflammation, inhibiting antitumor immunity, and stimulating angiogenesis (83–85). It is commonly accepted that macrophages contribute to tumor growth and invasion. They are recruited to the site of tumors via chemoattractants such as CCL3-8, vascular endothelial growth factor (VEGF), and macrophage inflammatory protein-1 alpha (MIP-1α) (86). The monocytes or macrophages tend to differentiate into the M2 macrophage phenotype rather than the tumoricidal M1 phenotype, producing pro-tumor cytokines, such as macrophage colony-stimulating factor, IL-10, IL-4, and IL-13 (83, 87, 88). S100A10 was shown to mediate the migration of macrophages to the tumor site. Tumor growth was reduced in S100A10-null mice, compared with wild-type mice, and was accompanied by less macrophages within the tumor. There were many macrophages throughout the tumor in wild-type mice, where macrophages were observed only around the absolute tumor tissue border in S100A10-null mice (89). Intraperitoneal injection of wild-type macrophages restored macrophage density within the tumor, but injection of S100A10-deficient macrophages did not. Interestingly, intratumoral injection of macrophages of either genotype could rescue tumor growth, suggesting that S100A10−/− macrophages still have the ability to stimulate tumor growth but lack the ability to invade into the tumor (89). Another study showed that S100A10 deficiency decreased plasmin generation and matrix metalloproteinase 9 activation in macrophages, both of which are associated with macrophage invasion and migration (90).

Downregulation of S100A8 and S100A9 is associated with the differentiation of myeloid cells toward dendritic cells and macrophages (91, 92). S100A8 and S100A9 are co-expressed in fetal myeloid progenitors, with its expression level associated with the development of the myeloid lineage (93). They are highly expressed in monocytes and neutrophils. However, the expressions of S100A8 and S100A9 are lost when monocytes terminally differentiate into tissue macrophages (93). Recent data have shown that S100A8 can be induced by oxidative stress in macrophages in an IL-10-dependent manner (51). Interestingly, S100A8/A9 has also been shown to control the cell cycle (94). S100A9 inhibited myeloid cells differentiation through generation of ROS (92). S100A9 is able to induce the differentiation of monocytes toward the osteoclast type in in vitro culture experiments and S100A9 derived from neutrophils and S100A9-induced osteoclast generation were considered as important reasons for bone degradation in infectious osteomyelitis (95). S100A8 and S100A9 have also been shown to mediate the arresting effect of TNF-α on the differentiation of immature myeloid-derived suppressor cells into dendritic cells and macrophages in a RAGE-dependent manner (96). Consistent with this finding, IL-6 and IL-8 released from myofibroblasts in tumor microenvironment upregulate S100A8/S100A9 in myeloid cells and induce the differentiation of myeloid cells into S100A8/S100A9-expressing myeloid-derived suppressor cells and M2 macrophages (63).

ROLE OF S100 PROTEINS IN TISSUE REPAIR

Damage-associated molecular pattern molecules play a critical role in tissue repair. S100A7, S100A8/A9, S100A12, and S100A15, well-documented DAMPs, have been shown to participate inflammatory tissue damage and tissue repair. The link between S100A12 and the severity of coronary and carotid atherosclerosis has been evidenced by multiple human studies (97–99). S100A7 is highly expressed in the skin, and the expression is increased in inflamed skin, which has been shown to be induced by pro-inflammatory cytokines (IL-17 and IL-22) and bacterial products such as flagellin (62), that the increase of S100A7 has been associated with multiple inflammatory skin diseases, such as psoriasis and atopic dermatitis (62, 100). Similarly, the expression of S100A15 was amplified in the epidermis of psoriatic lesions and acted as chemoattractants for immune cells (101). S100A8/A9 exerts anti-inflammatory function in healthy state, while oxidative stress-associated pathological conditions activate their pro-inflammatory functions (102). Increased plasma S100A8/A9 levels have been associated with atherogenesis, plaque vulnerability, myocardial infarction (MI), cardiovascular death, and heart failure. In a mouse model of angiotensin-induced cardiac damage, it was shown that S100A8/A9 released by granulocytes upregulated pro-inflammatory gene expression and induced the release of cytokines and chemokines in a RAGE-dependent manner. This process promoted myocardial tissue inflammation and fibrotic scar formation (103, 104). In a mouse model of collagenase-induced arthritis, the expression of S100A8 and S100A9 in synovial was upregulated in wild-type mice. In addition, S100a9−/− mice were protected from collagenase-induced synovitis, cartilage degradation, and osteophyte formation (105, 106). S100A9 antibodies could block the accumulation of fibroblasts and decrease fibrosis in local inflammatory microenvironment (104). In contrast, S100A1 or S100A4, released following MI, has a beneficial effect following heart injury by promoting muscle tissue repair and maintaining contractility (107, 108).

Binding of S100B to RAGE and the subsequent increase of angiogenic factor VEGF have been shown to be essential in the development of macular degeneration (109). In addition, S100B activates the Ras-MEK-ERK1/2-NF-κB pathway in neural cells and leads to the activation of small GTPases, Rac1/Cdc 42, and neurite growth (110). In vascular smooth muscle cells, S100B induces the upregulation of ROS and recruits JAK2 and STAT3, which results in the proliferation of vascular smooth muscle cells (111). Similarly, S100B also increased cellular proliferation though activating the Phosphatidylinositol-4,5-bisphosphate 3-kinase-AKT pathway in a RAGE-dependent manner (14). On the other hand, S100B could induce apoptosis by increasing production of ROS and the release of cytochrome-c from mitochondria (110). High levels of S100B are released from injured cardiomyocytes following MI and could promote cell apoptosis through RAGE. Also, S100B released from injured skeletal muscle tissue could stimulate myoblast proliferation but inhibit myoblast differentiation by activating bFGF/FGFR1 signaling (112, 113). However, the regeneration effects of S100B on the injured myoblasts are strongly dependent on cell density, because it triggers RAGE, but not bFGF/FGFR1 signaling, at an early stage of low-density myoblast differentiation (114).

THE ROLE OF S100 PROTEINS IN INFLAMMATORY DISEASES

S100 proteins, particularly calgranulins, play a significant role in mediating innate and acquired immune responses, which contribute to the development of chronic inflammatory diseases.

Calgranulins are associated with joint inflammation in patients with rheumatoid arthritis (RA) (115). The level of S100A8/A9 in the serum and synovial fluid was significantly increased in RA (116, 117). Recent findings showed that S100A8/A9 was upregulated in early but not late phase osteoarthritis (OA) (118). S100A8/A9 plasma levels were increased at baseline in human OA participants. Meanwhile, osteophyte size was drastically reduced in S100A9−/− mice-induced OA (106). It has also been confirmed that S100A8/A9 contributes to cartilage degradation and development of inflammatory arthritis in an antigen-induced arthritis model (119). Similar to S100A8/A9, human S100A7 and S100A15 were first confirmed as over-expressed in psoriatic plaques (120). Increasing evidence supports an association of S100A7 with several inflammatory skin diseases, including psoriasis and atopic dermatitis (62, 100). Evidence strongly indicates that S100A8/A9 levels are higher in hypercalprotectinemia, an extremely rare inflammatory disorder (121–123). Although the mechanism is still unclear, it is possible that the releasing of extracellular S100A8/A9 is dysregulated, which accounts for the abnormal increase of calprotectin and subsequent hyperactive inflammatory reaction. It is suggested that S100 proteins are involved in interacting with both the immune system and the pathogen. S100A12 plays a key role in fighting infections. For example, it has been shown that S100A12 plays a critical role in anti-parasite responses (124). In addition to directly killing Mycobacterium tuberculosis and Mycobacterium leprae, S10012 is also required for TLR2/1L- and IFN-γ-induced antimicrobial activity against Mycobacterium (125). Haley et al. also showed that S100A12 can help to repress the biogenesis and activity of H. pylori cag type IV secretion system by binding nutrient zinc, which results in suppressed bacterial growth and viability (126).

S100A8, S100A9, and S100A12 are abundantly expressed by neutrophils. Evidence indicates that these three members of S100 proteins are released by neutrophils, inducing MUC5AC production in airway epithelial cells through activating TLR4 and RAGE signaling pathway. This reveals the relationship between chronic neutrophilic inflammation and obstructive airway diseases such as severe asthma, COPD, and cystic fibrosis (127). In correlation with their role in the development of chronic inflammation, S100A8/A9 also participates in the hyperglycemia-induced increase of myelopoiesis occurring in a RAGE-dependent manner in diabetic mice (128). Interestingly, the amount of circulating monocytes and neutrophils were decreased when antidiabetic treatment normalized the glycemic index of Ldlr−/− atherosclerotic mice, which might explain the increased severity of atherosclerosis found in patients with diabetes (128). In accordance with these findings, increased serum concentrations of S100A8/A9 were detected in obese individuals (129). Furthermore, the expression of the macrophage marker CD68 was increased in the visceral adipose tissue (130). Some research of dipeptidyl peptidase-4 inhibitors for the treatment of type 2 diabetes mellitus indicates that vildagliptin could increase the mRNA expression levels of S100A9 and TNF-α in human hepatocytes. In addition, it may induce the release of S100A8/A9 complex from HL-60 cells via TNF-α-independent manner, which might be a contributing factor of vildagliptin-associated liver dysfunction (131).

S100 PROTEINS AS BIOMARKERS IN SPECIFIC DISEASES

Extracellular S100 proteins are involved in the activation of G protein-coupled receptors, heparan sulfate proteoglycans or N-Glycans, and scavenger receptors in autocrine and paracrine manners (132, 133). Since S100A proteins can be detected in body fluids, such as urine, cerebrospinal fluid, serum, sputum, and feces, extracellular S100 proteins are considered as biomarkers associated with certain diseases (134–137).

It has been suggested that S100A12, S100A8/A9, and S100B are linked to specific diseases and conditions such as auto-inflammatory diseases, stroke, and trauma (138). The level of S100A12 in the blood is increased in the patients with diabetes, which is correlated with a higher risk of cardiovascular disease development (139). Bogdanova et al. detected the serum concentration of S100A12 and other acute-phase inflammatory markers in thirty-five patients with periodic disease (PD) (140). The level of S100A12 in PD was significantly higher compared to other familial periodic fevers. S100A12 was more sensitive to assess the subclinical activity of autoinflammatory diseases, when compared to other inflammatory biomarkers such as neutrophil counts, fibrinogen, C-reactive protein (CRP), and erythrocyte sedimentation rate (140). Similarly, the serum concentrations of S100A12, as a novel biomarker, were shown to be upregulated in patients with Familial Mediterranean fever in comparison to controls (141).

The plasma concentrations of S100A9 were significantly higher in patients with implant-associated infectious osteomyelitis when compared to patients with sterile inflammation or healthy individuals. In addition, S100A9 was associated with osteoclast generation and bone degradation. Therefore, it could serve as a novel diagnostic marker to aid in the differential diagnosis (95). Similarly, serum levels of S100A8 and S100A9 were dramatically increased in IL-1Ra−/− mice and contributed to bone erosion, cartilage damage, and synovial inflammation. Thus, they can be considered as a systemic or local biomarker to evaluate the extent of inflammation and inflammatory joint destruction in seronegative arthritis (142). It was shown that the expression of S100A8/A9 was high in human atherosclerotic lesions and the blood levels were also increased in the patients with coronary artery diseases (CAD), which implied S100A8/A9 might act as a biomarker for cardiovascular events (143). Recent research has shown similar findings that serum S100A8/A9 levels were elevated in 178 CAD patients with unstable angina pectoris or acute myocardial infarction, and the level of S100A8/A9 was significantly positively linked with CRP (P < 0.01) (144). These clinical data suggest that S100A8/A9 may become a novel biomarker for CAD (139).

In addition, more studies explored the value S100A8/A9 as a predictive biomarker for autoimmune diseases. In RA, S100A8/A9 was suggested as a potential biomarker in predicting clinical response to monitor treatment (145, 146). Some clinical investigations have indicated that S100A8/A9 levels might be a more sensitive predictor for monitoring synovial inflammation in RA patients when compared with other markers such as CRP levels (147).

The study by Shakeri et al. suggested that S100 B protein could be used as a posttraumatic biomarker for predicting brain death in severely injured patients with exclusive head trauma during the first 6 h after trauma, but found no relationship between S100 B levels and death (148). Pelinka et al. confirmed that in vitro S100 B concentrations increased significantly in rats with femoral fractures but not head injury (149). Interestingly, adverse results indicated that there was no difference in S100 B concentrations between patients with and without head injury (150). S100B has also been considered as a prognostic marker of the acute phase of neurologic damage (151), predicting the outcome of traumatic brain injury and large volume cerebral infarction (152, 153). The level of serum S100B in ischemic stroke implied a worse outcome secondary to the stroke (154, 155). This research demonstrates that S100 B is correlated to trauma and a worse long-term outcome. S100B has recently been confirmed to be associated with some genetic disorders and was found to be over-expressed in patients with Down syndrome (156, 157). There was also study showing that S100B may be one of the best biomarkers of melanoma (158).

S100 PROTEINS AS THERAPEUTIC TARGETS IN DISEASE

Although direct clinical evidence is limited, increasing studies indicate that S100 proteins may also serve as a therapeutic target for certain disease conditions. As mentioned above, S100 proteins are involved in a number of diseases including inflammatory disease. It has been reported that multiple anti-allergic drugs such as amlexanox, cromolyn, and tranilast are able to bind S100A12 and S100A13, and block downstream RAGE signaling (159). Therefore, these drugs may serve as a therapeutic approach to target S100 proteins. Multiple S100 proteins such as S100A4 (160) and S100B (161) have been shown to participate in the neoplastic disorders by binding to P53 and suppressing its phosphorylation (162). Therefore, efforts are being made to restore P53 function by targeting S100 proteins (163). In an in vitro study, Most et al. demonstrated that extracellular S100A1 is endocytosed by the neonatal ventricular cardiomyocytes protects cardiomyocytes from 2-deoxyglucose and oxidative stress-induced apoptosis via activation of ERK (164). Adeno-associated virus-mediated S100A1 gene transfer in failing cardiomyocytes was also shown to be able to restore the contractile function, suggesting a potential implication of AAV-mediated S100A1 gene therapy in heart failure (165, 166). Despite the promising potentials, the feasibility and safety of these approaches and issues such as how to control and keep expression levels in the therapeutic window need to be further investigated (166).

CONCLUSION

Evidence strongly supports that S100 proteins, as a remarkable multifunctional proteins family, are involved in the regulation of several important biological processes such as the inflammatory response, protecting the intra- and extracellular environments during infection, cell proliferation and differentiation, tumor growth and metastasis, cell apoptosis, energy, and glutathione metabolism.

However, the activities of all members S100 proteins depend on the cell-specific expression patterns and binding targets even the local microenvironment. Extracellular effects of S100 proteins interact with receptors including TLR-4, RAGE, and heparan sulfate proteoglycans during infection and inflammation which associated with the pathogenesis of inflammatory such as autoimmune disease, infectious diseases, allergy, tumorigenesis and metastasis, and anti-microbial disease. Extracellular S100 proteins can also contribute to the regulation of tissue development and regeneration or repair, which is essential for elucidating their role in the pathological procession of tissue damage, cell apoptosis, or tissue repair.

Although growing evidence has begun to show the regulation of S100 proteins in detail which improves our understanding of how immune homeostasis is maintained during the development of S100 protein-associated disease, there are certain gaps in our understanding of the role of S100 proteins in pathophysiology. Among 25 known members of S100 family, only limited number of S100 proteins such as S100A8 and S100A9 have been well documented and the functional roles of other members are underappreciated. In addition, further studies are required to fully reveal the underlying mechanisms by which S100 proteins participate in a variety of disease conditions. For instance, a role of S100P has been reported in leukemia (167), while the exact function of S100P in leukemia and the signal pathways involved in this process are not completely understood. Also, the direct clinical evidence of the therapeutic potential of S100 proteins is limited at current stage. Therefore, future directions in this area could focus on the development of therapeutic approaches targeting S100 proteins, verification of the therapeutic potential of S100 proteins in both preclinical and clinical settings, and elucidation of the underlying mechanisms.

AUTHOR CONTRIBUTIONS

CX and JZ reviewed the literature and wrote the first draft. ZB, AT, JZ, and XR reviewed the literature and finalized the manuscript.

FUNDING

This work was supported by grants from National Natural and Scientific Foundation of China (81670431 and 81101247), NIH (K99ES026241 and K01DK105108), AHA (17GRNT33670485), and Natural Science Foundation of Zhejiang Province (No. Y2110580). These funding supports did not lead to any conflict of interests regarding the publication of this manuscript.

REFERENCES

1. Moore BW. A soluble protein characteristic of the nervous system. Biochem Biophys Res Commun (1965) 19:739–44. doi:10.1016/0006-291X(65)90320-7

2. Moore BW, McGregor D. Chromatographic and electrophoretic fractionation of soluble proteins of brain and liver. J Biol Chem (1965) 240:1647–53.

3. Moore BW, Perez VJ, Gehring M. Assay and regional distribution of a soluble protein characteristic of the nervous system. J Neurochem (1968) 15:265–72. doi:10.1111/j.1471-4159.1968.tb11610.x

4. Potts BC, Smith J, Akke M, Macke TJ, Okazaki K, Hidaka H, et al. The structure of calcyclin reveals a novel homodimeric fold for S100 Ca(2+)-binding proteins. Nat Struct Biol (1995) 2:790–6. doi:10.1038/nsb0995-790

5. Marenholz I, Heizmann CW, Fritz G. S100 proteins in mouse and man: from evolution to function and pathology (including an update of the nomenclature). Biochem Biophys Res Commun (2004) 322:1111–22. doi:10.1016/j.bbrc.2004.07.096

6. Leclerc E, Fritz G, Vetter SW, Heizmann CW. Binding of S100 proteins to RAGE: an update. Biochim Biophys Acta (2009) 1793:993–1007. doi:10.1016/j.bbamcr.2008.11.016

7. Donato R, Cannon BR, Sorci G, Riuzzi F, Hsu K, Weber DJ, et al. Functions of S100 proteins. Curr Mol Med (2013) 13:24–57. doi:10.2174/1566524011307010024

8. Gross SR, Sin CG, Barraclough R, Rudland PS. Joining S100 proteins and migration: for better or for worse, in sickness and in health. Cell Mol Life Sci (2014) 71:1551–79. doi:10.1007/s00018-013-1400-7

9. Zimmer DB, Landar A. Analysis of S100A1 expression during skeletal muscle and neuronal cell differentiation. J Neurochem (1995) 64:2727–36. doi:10.1046/j.1471-4159.1995.64062727.x

10. Most P, Bernotat J, Ehlermann P, Pleger ST, Reppel M, Borries M, et al. S100A1: a regulator of myocardial contractility. Proc Natl Acad Sci U S A (2001) 98:13889–94. doi:10.1073/pnas.241393598

11. Most P, Pleger ST, Volkers M, Heidt B, Boerries M, Weichenhan D, et al. Cardiac adenoviral S100A1 gene delivery rescues failing myocardium. J Clin Invest (2004) 114:1550–63. doi:10.1172/JCI21454

12. Most P, Boerries M, Eicher C, Schweda C, Volkers M, Wedel T, et al. Distinct subcellular location of the Ca2+-binding protein S100A1 differentially modulates Ca2+-cycling in ventricular rat cardiomyocytes. J Cell Sci (2005) 118:421–31. doi:10.1242/jcs.01614

13. Bhattacharya S, Large E, Heizmann CW, Hemmings B, Chazin WJ. Structure of the Ca2+/S100B/NDR kinase peptide complex: insights into S100 target specificity and activation of the kinase. Biochemistry (2003) 42:14416–26. doi:10.1021/bi035089a

14. Leclerc E, Fritz G, Weibel M, Heizmann CW, Galichet A. S100B and S100A6 differentially modulate cell survival by interacting with distinct RAGE (receptor for advanced glycation end products) immunoglobulin domains. J Biol Chem (2007) 282:31317–31. doi:10.1074/jbc.M703951200

15. Marshak DR, Pena LA. Potential role of S100 beta in Alzheimer’s disease: an hypothesis involving mitotic protein kinases. Prog Clin Biol Res (1992) 379:289–307.

16. Griffin KA, Bidani AK. Progression of renal disease: renoprotective specificity of renin-angiotensin system blockade. Clin J Am Soc Nephrol (2006) 1:1054–65. doi:10.2215/CJN.02231205

17. Donato R. S100: a multigenic family of calcium-modulated proteins of the EF-hand type with intracellular and extracellular functional roles. Int J Biochem Cell Biol (2001) 33:637–68. doi:10.1016/S1357-2725(01)00046-2

18. Komatsu K, Kobune-Fujiwara Y, Andoh A, Ishiguro S, Hunai H, Suzuki N, et al. Increased expression of S100A6 at the invading fronts of the primary lesion and liver metastasis in patients with colorectal adenocarcinoma. Br J Cancer (2000) 83:769–74. doi:10.1054/bjoc.2000.1356

19. Melle C, Ernst G, Schimmel B, Bleul A, Von Eggeling F. Colon-derived liver metastasis, colorectal carcinoma, and hepatocellular carcinoma can be discriminated by the Ca(2+)-binding proteins S100A6 and S100A11. PLoS One (2008) 3:e3767. doi:10.1371/journal.pone.0003767

20. Berge G, Maelandsmo GM. Evaluation of potential interactions between the metastasis-associated protein S100A4 and the tumor suppressor protein p53. Amino Acids (2011) 41:863–73. doi:10.1007/s00726-010-0497-3

21. Mishra SK, Siddique HR, Saleem M. S100A4 calcium-binding protein is key player in tumor progression and metastasis: preclinical and clinical evidence. Cancer Metastasis Rev (2012) 31:163–72. doi:10.1007/s10555-011-9338-4

22. Brophy MB, Hayden JA, Nolan EM. Calcium ion gradients modulate the zinc affinity and antibacterial activity of human calprotectin. J Am Chem Soc (2012) 134:18089–100. doi:10.1021/ja307974e

23. Fritz G, Botelho HM, Morozova-Roche LA, Gomes CM. Natural and amyloid self-assembly of S100 proteins: structural basis of functional diversity. FEBS J (2010) 277:4578–90. doi:10.1111/j.1742-4658.2010.07887.x

24. Korndorfer IP, Brueckner F, Skerra A. The crystal structure of the human (S100A8/S100A9)2 heterotetramer, calprotectin, illustrates how conformational changes of interacting alpha-helices can determine specific association of two EF-hand proteins. J Mol Biol (2007) 370:887–98. doi:10.1016/j.jmb.2007.04.065

25. Hayden JA, Brophy MB, Cunden LS, Nolan EM. High-affinity manganese coordination by human calprotectin is calcium-dependent and requires the histidine-rich site formed at the dimer interface. J Am Chem Soc (2013) 135:775–87. doi:10.1021/ja3096416

26. Vogl T, Tenbrock K, Ludwig S, Leukert N, Ehrhardt C, Van Zoelen MA, et al. Mrp8 and Mrp14 are endogenous activators of toll-like receptor 4, promoting lethal, endotoxin-induced shock. Nat Med (2007) 13:1042–9. doi:10.1038/nm1638

27. Turovskaya O, Foell D, Sinha P, Vogl T, Newlin R, Nayak J, et al. RAGE, carboxylated glycans and S100A8/A9 play essential roles in colitis-associated carcinogenesis. Carcinogenesis (2008) 29:2035–43. doi:10.1093/carcin/bgn188

28. Sorci G, Riuzzi F, Giambanco I, Donato R. RAGE in tissue homeostasis, repair and regeneration. Biochim Biophys Acta (2013) 1833:101–9. doi:10.1016/j.bbamcr.2012.10.021

29. Hofmann MA, Drury S, Fu C, Qu W, Taguchi A, Lu Y, et al. RAGE mediates a novel proinflammatory axis: a central cell surface receptor for S100/calgranulin polypeptides. Cell (1999) 97:889–901. doi:10.1016/S0092-8674(00)80801-6

30. Gebhardt C, Riehl A, Durchdewald M, Nemeth J, Furstenberger G, Muller-Decker K, et al. RAGE signaling sustains inflammation and promotes tumor development. J Exp Med (2008) 205:275–85. doi:10.1084/jem.20070679

31. Ghavami S, Rashedi I, Dattilo BM, Eshraghi M, Chazin WJ, Hashemi M, et al. S100A8/A9 at low concentration promotes tumor cell growth via RAGE ligation and MAP kinase-dependent pathway. J Leukoc Biol (2008) 83:1484–92. doi:10.1189/jlb.0607397

32. Maletzki C, Bodammer P, Breitruck A, Kerkhoff C. S100 proteins as diagnostic and prognostic markers in colorectal and hepatocellular carcinoma. Hepat Mon (2012) 12:e7240. doi:10.5812/hepatmon.7240

33. Arumugam T, Ramachandran V, Sun D, Peng Z, Pal A, Maxwell DS, et al. Designing and developing S100P inhibitor 5-methyl cromolyn for pancreatic cancer therapy. Mol Cancer Ther (2013) 12:654–62. doi:10.1158/1535-7163.MCT-12-0771

34. Sorci G, Riuzzi F, Arcuri C, Giambanco I, Donato R. Amphoterin stimulates myogenesis and counteracts the antimyogenic factors basic fibroblast growth factor and S100B via RAGE binding. Mol Cell Biol (2004) 24:4880–94. doi:10.1128/MCB.24.11.4880-4894.2004

35. Boom A, Pochet R, Authelet M, Pradier L, Borghgraef P, Van Leuven F, et al. Astrocytic calcium/zinc binding protein S100A6 over expression in Alzheimer’s disease and in PS1/APP transgenic mice models. Biochim Biophys Acta (2004) 1742:161–8. doi:10.1016/j.bbamcr.2004.09.011

36. Heizmann CW, Fritz G, Schafer BW. S100 proteins: structure, functions and pathology. Front Biosci (2002) 7:d1356–68. doi:10.2741/A846

37. Chazin WJ. Relating form and function of EF-hand calcium binding proteins. Acc Chem Res (2011) 44:171–9. doi:10.1021/ar100110d

38. Lindsey JC, Lusher ME, Anderton JA, Gilbertson RJ, Ellison DW, Clifford SC. Epigenetic deregulation of multiple S100 gene family members by differential hypomethylation and hypermethylation events in medulloblastoma. Br J Cancer (2007) 97:267–74. doi:10.1038/sj.bjc.6603852

39. Wang XH, Zhang LH, Zhong XY, Xing XF, Liu YQ, Niu ZJ, et al. S100A6 overexpression is associated with poor prognosis and is epigenetically up-regulated in gastric cancer. Am J Pathol (2010) 177:586–97. doi:10.2353/ajpath.2010.091217

40. Wang Q, Williamson M, Bott S, Brookman-Amissah N, Freeman A, Nariculam J, et al. Hypomethylation of WNT5A, CRIP1 and S100P in prostate cancer. Oncogene (2007) 26:6560–5. doi:10.1038/sj.onc.1210472

41. Li JT, Wang LF, Zhao YL, Yang T, Li W, Zhao J, et al. Nuclear factor of activated T cells 5 maintained by Hotair suppression of miR-568 upregulates S100 calcium binding protein A4 to promote breast cancer metastasis. Breast Cancer Res (2014) 16:454. doi:10.1186/s13058-014-0454-2

42. Li W, Kong LB, Li JT, Guo ZY, Xue Q, Yang T, et al. MiR-568 inhibits the activation and function of CD4(+) T cells and Treg cells by targeting NFAT5. Int Immunol (2014) 26:269–81. doi:10.1093/intimm/dxt065

43. Donato R. Intracellular and extracellular roles of S100 proteins. Microsc Res Tech (2003) 60:540–51. doi:10.1002/jemt.10296

44. Edgeworth J, Gorman M, Bennett R, Freemont P, Hogg N. Identification of p8,14 as a highly abundant heterodimeric calcium binding protein complex of myeloid cells. J Biol Chem (1991) 266:7706–13.

45. Averill MM, Barnhart S, Becker L, Li X, Heinecke JW, Leboeuf RC, et al. S100A9 differentially modifies phenotypic states of neutrophils, macrophages, and dendritic cells: implications for atherosclerosis and adipose tissue inflammation. Circulation (2011) 123:1216–26. doi:10.1161/CIRCULATIONAHA.110.985523

46. Rahimi F, Hsu K, Endoh Y, Geczy CL. FGF-2, IL-1beta and TGF-beta regulate fibroblast expression of S100A8. FEBS J (2005) 272:2811–27. doi:10.1111/j.1742-4658.2005.04703.x

47. Ingersoll MA, Spanbroek R, Lottaz C, Gautier EL, Frankenberger M, Hoffmann R, et al. Comparison of gene expression profiles between human and mouse monocyte subsets. Blood (2010) 115:e10–9. doi:10.1182/blood-2009-07-235028

48. Yen T, Harrison CA, Devery JM, Leong S, Iismaa SE, Yoshimura T, et al. Induction of the S100 chemotactic protein, CP-10, in murine microvascular endothelial cells by proinflammatory stimuli. Blood (1997) 90:4812–21.

49. McCormick MM, Rahimi F, Bobryshev YV, Gaus K, Zreiqat H, Cai H, et al. S100A8 and S100A9 in human arterial wall. Implications for atherogenesis. J Biol Chem (2005) 280:41521–9. doi:10.1074/jbc.M509442200

50. Yao D, Brownlee M. Hyperglycemia-induced reactive oxygen species increase expression of the receptor for advanced glycation end products (RAGE) and RAGE ligands. Diabetes (2010) 59:249–55. doi:10.2337/db09-0801

51. Grimbaldeston MA, Geczy CL, Tedla N, Finlay-Jones JJ, Hart PH. S100A8 induction in keratinocytes by ultraviolet A irradiation is dependent on reactive oxygen intermediates. J Invest Dermatol (2003) 121:1168–74. doi:10.1046/j.1523-1747.2003.12561.x

52. Schiopu A, Cotoi OS. S100A8 and S100A9: DAMPs at the crossroads between innate immunity, traditional risk factors, and cardiovascular disease. Mediators Inflamm (2013) 2013:828354. doi:10.1155/2013/828354

53. Hsu K, Champaiboon C, Guenther BD, Sorenson BS, Khammanivong A, Ross KF, et al. Anti-infective protective properties of S100 calgranulins. Antiinflamm Antiallergy Agents Med Chem (2009) 8:290–305. doi:10.2174/187152309789838975

54. Ravasi T, Hsu K, Goyette J, Schroder K, Yang Z, Rahimi F, et al. Probing the S100 protein family through genomic and functional analysis. Genomics (2004) 84:10–22. doi:10.1016/j.ygeno.2004.02.002

55. Goebeler M, Roth J, Van Den Bos C, Ader G, Sorg C. Increase of calcium levels in epithelial cells induces translocation of calcium-binding proteins migration inhibitory factor-related protein 8 (MRP8) and MRP14 to keratin intermediate filaments. Biochem J (1995) 309(Pt 2):419–24. doi:10.1042/bj3090419

56. Stoll SW, Zhao X, Elder JT. EGF stimulates transcription of CaN19 (S100A2) in HaCaT keratinocytes. J Invest Dermatol (1998) 111:1092–7. doi:10.1046/j.1523-1747.1998.00402.x

57. Mork G, Schjerven H, Mangschau L, Soyland E, Brandtzaeg P. Proinflammatory cytokines upregulate expression of calprotectin (L1 protein, MRP-8/MRP-14) in cultured human keratinocytes. Br J Dermatol (2003) 149:484–91. doi:10.1046/j.1365-2133.2003.05536.x

58. Nukui T, Ehama R, Sakaguchi M, Sonegawa H, Katagiri C, Hibino T, et al. S100A8/A9, a key mediator for positive feedback growth stimulation of normal human keratinocytes. J Cell Biochem (2008) 104:453–64. doi:10.1002/jcb.21639

59. Bando M, Zou X, Hiroshima Y, Kataoka M, Ross KF, Shinohara Y, et al. Mechanism of interleukin-1alpha transcriptional regulation of S100A9 in a human epidermal keratinocyte cell line. Biochim Biophys Acta (2013) 1829:954–62. doi:10.1016/j.bbagrm.2013.03.010

60. Yao R, Lopez-Beltran A, Maclennan GT, Montironi R, Eble JN, Cheng L. Expression of S100 protein family members in the pathogenesis of bladder tumors. Anticancer Res (2007) 27:3051–8.

61. Nasser MW, Qamri Z, Deol YS, Ravi J, Powell CA, Trikha P, et al. S100A7 enhances mammary tumorigenesis through upregulation of inflammatory pathways. Cancer Res (2012) 72:604–15. doi:10.1158/0008-5472.CAN-11-0669

62. Glaser R, Harder J, Lange H, Bartels J, Christophers E, Schroder JM. Antimicrobial psoriasin (S100A7) protects human skin from Escherichia coli infection. Nat Immunol (2005) 6:57–64. doi:10.1038/ni1142

63. Kim JH, Oh SH, Kim EJ, Park SJ, Hong SP, Cheon JH, et al. The role of myofibroblasts in upregulation of S100A8 and S100A9 and the differentiation of myeloid cells in the colorectal cancer microenvironment. Biochem Biophys Res Commun (2012) 423:60–6. doi:10.1016/j.bbrc.2012.05.081

64. Dapunt U, Gaida MM, Meyle E, Prior B, Hansch GM. Activation of phagocytic cells by Staphylococcus epidermidis biofilms: effects of extracellular matrix proteins and the bacterial stress protein GroEL on netosis and MRP-14 release. Pathog Dis (2016) 74:ftw035. doi:10.1093/femspd/ftw035

65. Kligman D, Hilt DC. The S100 protein family. Trends Biochem Sci (1988) 13:437–43. doi:10.1016/0968-0004(88)90218-6

66. Winningham-Major F, Staecker JL, Barger SW, Coats S, Van Eldik LJ. Neurite extension and neuronal survival activities of recombinant S100 beta proteins that differ in the content and position of cysteine residues. J Cell Biol (1989) 109:3063–71. doi:10.1083/jcb.109.6.3063

67. Chan JK, Roth J, Oppenheim JJ, Tracey KJ, Vogl T, Feldmann M, et al. Alarmins: awaiting a clinical response. J Clin Invest (2012) 122:2711–9. doi:10.1172/JCI62423

68. Newton K, Dixit VM. Signaling in innate immunity and inflammation. Cold Spring Harb Perspect Biol (2012) 4:a006049. doi:10.1101/cshperspect.a006049

69. de Haan JJ, Smeets MB, Pasterkamp G, Arslan F. Danger signals in the initiation of the inflammatory response after myocardial infarction. Mediators Inflamm (2013) 2013:206039. doi:10.1155/2013/206039

70. Foell D, Wittkowski H, Vogl T, Roth J. S100 proteins expressed in phagocytes: a novel group of damage-associated molecular pattern molecules. J Leukoc Biol (2007) 81:28–37. doi:10.1189/jlb.0306170

71. Schmidt AM, Hofmann M, Taguchi A, Yan SD, Stern DM. RAGE: a multiligand receptor contributing to the cellular response in diabetic vasculopathy and inflammation. Semin Thromb Hemost (2000) 26:485–93. doi:10.1055/s-2000-13204

72. Ryckman C, Vandal K, Rouleau P, Talbot M, Tessier PA. Proinflammatory activities of S100: proteins S100A8, S100A9, and S100A8/A9 induce neutrophil chemotaxis and adhesion. J Immunol (2003) 170:3233–42. doi:10.4049/jimmunol.170.6.3233

73. Ghavami S, Eshragi M, Ande SR, Chazin WJ, Klonisch T, Halayko AJ, et al. S100A8/A9 induces autophagy and apoptosis via ROS-mediated cross-talk between mitochondria and lysosomes that involves BNIP3. Cell Res (2010) 20:314–31. doi:10.1038/cr.2009.129

74. Sunahori K, Yamamura M, Yamana J, Takasugi K, Kawashima M, Yamamoto H, et al. The S100A8/A9 heterodimer amplifies proinflammatory cytokine production by macrophages via activation of nuclear factor kappa B and p38 mitogen-activated protein kinase in rheumatoid arthritis. Arthritis Res Ther (2006) 8:R69. doi:10.1186/ar1939

75. Eue I, Pietz B, Storck J, Klempt M, Sorg C. Transendothelial migration of 27E10+ human monocytes. Int Immunol (2000) 12:1593–604. doi:10.1093/intimm/12.11.1593

76. Nisapakultorn K, Ross KF, Herzberg MC. Calprotectin expression inhibits bacterial binding to mucosal epithelial cells. Infect Immun (2001) 69:3692–6. doi:10.1128/IAI.69.6.3692-3696.2001

77. Newton RA, Hogg N. The human S100 protein MRP-14 is a novel activator of the beta 2 integrin Mac-1 on neutrophils. J Immunol (1998) 160:1427–35.

78. Manitz MP, Horst B, Seeliger S, Strey A, Skryabin BV, Gunzer M, et al. Loss of S100A9 (MRP14) results in reduced interleukin-8-induced CD11b surface expression, a polarized microfilament system, and diminished responsiveness to chemoattractants in vitro. Mol Cell Biol (2003) 23:1034–43. doi:10.1128/MCB.23.3.1034-1043.2003

79. Garrett SC, Varney KM, Weber DJ, Bresnick AR. S100A4, a mediator of metastasis. J Biol Chem (2006) 281:677–80. doi:10.1074/jbc.R500017200

80. Boye K, Maelandsmo GM. S100A4 and metastasis: a small actor playing many roles. Am J Pathol (2010) 176:528–35. doi:10.2353/ajpath.2010.090526

81. Malashkevich VN, Dulyaninova NG, Ramagopal UA, Liriano MA, Varney KM, Knight D, et al. Phenothiazines inhibit S100A4 function by inducing protein oligomerization. Proc Natl Acad Sci U S A (2010) 107:8605–10. doi:10.1073/pnas.0913660107

82. Yang Z, Yan WX, Cai H, Tedla N, Armishaw C, Di Girolamo N, et al. S100A12 provokes mast cell activation: a potential amplification pathway in asthma and innate immunity. J Allergy Clin Immunol (2007) 119:106–14. doi:10.1016/j.jaci.2006.08.021

83. Pollard JW. Tumour-educated macrophages promote tumour progression and metastasis. Nat Rev Cancer (2004) 4:71–8. doi:10.1038/nrc1256

84. Condeelis J, Pollard JW. Macrophages: obligate partners for tumor cell migration, invasion, and metastasis. Cell (2006) 124:263–6. doi:10.1016/j.cell.2006.01.007

85. Sica A, Bronte V. Altered macrophage differentiation and immune dysfunction in tumor development. J Clin Invest (2007) 117:1155–66. doi:10.1172/JCI31422

86. Dandekar RC, Kingaonkar AV, Dhabekar GS. Role of macrophages in malignancy. Ann Maxillofac Surg (2011) 1:150–4. doi:10.4103/2231-0746.92782

87. Sica A, Schioppa T, Mantovani A, Allavena P. Tumour-associated macrophages are a distinct M2 polarised population promoting tumour progression: potential targets of anti-cancer therapy. Eur J Cancer (2006) 42:717–27. doi:10.1016/j.ejca.2006.01.003

88. Lewis CE, Hughes R. Inflammation and breast cancer. Microenvironmental factors regulating macrophage function in breast tumours: hypoxia and angiopoietin-2. Breast Cancer Res (2007) 9:209. doi:10.1186/bcr1679

89. Phipps KD, Surette AP, O’Connell PA, Waisman DM. Plasminogen receptor S100A10 is essential for the migration of tumor-promoting macrophages into tumor sites. Cancer Res (2011) 71:6676–83. doi:10.1158/0008-5472.CAN-11-1748

90. O’Connell PA, Surette AP, Liwski RS, Svenningsson P, Waisman DM. S100A10 regulates plasminogen-dependent macrophage invasion. Blood (2010) 116:1136–46. doi:10.1182/blood-2010-01-264754

91. Hashimoto S, Suzuki T, Dong HY, Nagai S, Yamazaki N, Matsushima K. Serial analysis of gene expression in human monocyte-derived dendritic cells. Blood (1999) 94:845–52.

92. Cheng P, Corzo CA, Luetteke N, Yu B, Nagaraj S, Bui MM, et al. Inhibition of dendritic cell differentiation and accumulation of myeloid-derived suppressor cells in cancer is regulated by S100A9 protein. J Exp Med (2008) 205:2235–49. doi:10.1084/jem.20080132

93. Lagasse E, Weissman IL. Mouse MRP8 and MRP14, two intracellular calcium-binding proteins associated with the development of the myeloid lineage. Blood (1992) 79:1907–15.

94. Khammanivong A, Wang C, Sorenson BS, Ross KF, Herzberg MC. S100A8/A9 (calprotectin) negatively regulates G2/M cell cycle progression and growth of squamous cell carcinoma. PLoS One (2013) 8:e69395. doi:10.1371/journal.pone.0069395

95. Dapunt U, Giese T, Maurer S, Stegmaier S, Prior B, Hansch GM, et al. Neutrophil-derived MRP-14 is up-regulated in infectious osteomyelitis and stimulates osteoclast generation. J Leukoc Biol (2015) 98:575–82. doi:10.1189/jlb.3VMA1014-482R

96. Sade-Feldman M, Kanterman J, Ish-Shalom E, Elnekave M, Horwitz E, Baniyash M. Tumor necrosis factor-alpha blocks differentiation and enhances suppressive activity of immature myeloid cells during chronic inflammation. Immunity (2013) 38:541–54. doi:10.1016/j.immuni.2013.02.007

97. Goyette J, Yan WX, Yamen E, Chung YM, Lim SY, Hsu K, et al. Pleiotropic roles of S100A12 in coronary atherosclerotic plaque formation and rupture. J Immunol (2009) 183:593–603. doi:10.4049/jimmunol.0900373

98. Abbas A, Aukrust P, Dahl TB, Bjerkeli V, Sagen EB, Michelsen A, et al. High levels of S100A12 are associated with recent plaque symptomatology in patients with carotid atherosclerosis. Stroke (2012) 43:1347–53. doi:10.1161/STROKEAHA.111.642256

99. Zhao P, Wu M, Yu H, Huang Y, Wang Y, Wang W, et al. Serum S100A12 levels are correlated with the presence and severity of coronary artery disease in patients with type 2 diabetes mellitus. J Investig Med (2013) 61:861–6. doi:10.2310/JIM.0b013e318292fb1e

100. Broome AM, Ryan D, Eckert RL. S100 protein subcellular localization during epidermal differentiation and psoriasis. J Histochem Cytochem (2003) 51:675–85. doi:10.1177/002215540305100513

101. Batycka-Baran A, Hattinger E, Zwicker S, Summer B, Zack Howard OM, Thomas P, et al. Leukocyte-derived koebnerisin (S100A15) and psoriasin (S100A7) are systemic mediators of inflammation in psoriasis. J Dermatol Sci (2015) 79:214–21. doi:10.1016/j.jdermsci.2015.05.007

102. Sroussi HY, Kohler GA, Agabian N, Villines D, Palefsky JM. Substitution of methionine 63 or 83 in S100A9 and cysteine 42 in S100A8 abrogate the antifungal activities of S100A8/A9: potential role for oxidative regulation. FEMS Immunol Med Microbiol (2009) 55:55–61. doi:10.1111/j.1574-695X.2008.00498.x

103. Croce K, Gao H, Wang Y, Mooroka T, Sakuma M, Shi C, et al. Myeloid-related protein-8/14 is critical for the biological response to vascular injury. Circulation (2009) 120:427–36. doi:10.1161/CIRCULATIONAHA.108.814582

104. Wu Y, Li Y, Zhang C, A X, Wang Y, Cui W, et al. S100a8/a9 released by CD11b+Gr1+ neutrophils activates cardiac fibroblasts to initiate angiotensin II-induced cardiac inflammation and injury. Hypertension (2014) 63:1241–50. doi:10.1161/HYPERTENSIONAHA.113.02843

105. van Lent PL, Blom AB, Schelbergen RF, Sloetjes A, Lafeber FP, Lems WF, et al. Active involvement of alarmins S100A8 and S100A9 in the regulation of synovial activation and joint destruction during mouse and human osteoarthritis. Arthritis Rheum (2012) 64:1466–76. doi:10.1002/art.34315

106. Schelbergen RF, De Munter W, Van Den Bosch MH, Lafeber FP, Sloetjes A, Vogl T, et al. Alarmins S100A8/S100A9 aggravate osteophyte formation in experimental osteoarthritis and predict osteophyte progression in early human symptomatic osteoarthritis. Ann Rheum Dis (2016) 75:218–25. doi:10.1136/annrheumdis-2014-205480

107. Le Hir M, Hegyi I, Cueni-Loffing D, Loffing J, Kaissling B. Characterization of renal interstitial fibroblast-specific protein 1/S100A4-positive cells in healthy and inflamed rodent kidneys. Histochem Cell Biol (2005) 123:335–46. doi:10.1007/s00418-005-0788-z

108. Schneider M, Kostin S, Strom CC, Aplin M, Lyngbaek S, Theilade J, et al. S100A4 is upregulated in injured myocardium and promotes growth and survival of cardiac myocytes. Cardiovasc Res (2007) 75:40–50. doi:10.1016/j.cardiores.2007.03.027

109. Ma W, Lee SE, Guo J, Qu W, Hudson BI, Schmidt AM, et al. RAGE ligand upregulation of VEGF secretion in ARPE-19 cells. Invest Ophthalmol Vis Sci (2007) 48:1355–61. doi:10.1167/iovs.06-0738

110. Huttunen HJ, Kuja-Panula J, Sorci G, Agneletti AL, Donato R, Rauvala H. Coregulation of neurite outgrowth and cell survival by amphoterin and S100 proteins through receptor for advanced glycation end products (RAGE) activation. J Biol Chem (2000) 275:40096–105. doi:10.1074/jbc.M006993200

111. Shaw SS, Schmidt AM, Banes AK, Wang X, Stern DM, Marrero MB. S100B-RAGE-mediated augmentation of angiotensin II-induced activation of JAK2 in vascular smooth muscle cells is dependent on PLD2. Diabetes (2003) 52:2381–8. doi:10.2337/diabetes.52.9.2381

112. Sorci G, Skarstein F, Morand S, Hugot JP. Correlated evolution between host immunity and parasite life histories in primates and oxyurid parasites. Proc Biol Sci (2003) 270:2481–4. doi:10.1098/rspb.2003.2536

113. Riuzzi F, Sorci G, Donato R. S100B stimulates myoblast proliferation and inhibits myoblast differentiation by independently stimulating ERK1/2 and inhibiting p38 MAPK. J Cell Physiol (2006) 207:461–70. doi:10.1002/jcp.20580

114. Riuzzi F, Sorci G, Donato R. The amphoterin (HMGB1)/receptor for advanced glycation end products (RAGE) pair modulates myoblast proliferation, apoptosis, adhesiveness, migration, and invasiveness. Functional inactivation of RAGE in L6 myoblasts results in tumor formation in vivo. J Biol Chem (2006) 281:8242–53. doi:10.1074/jbc.M509436200

115. Perera C, Mcneil HP, Geczy CL. S100 calgranulins in inflammatory arthritis. Immunol Cell Biol (2010) 88:41–9. doi:10.1038/icb.2009.88

116. Odink K, Cerletti N, Bruggen J, Clerc RG, Tarcsay L, Zwadlo G, et al. Two calcium-binding proteins in infiltrate macrophages of rheumatoid arthritis. Nature (1987) 330:80–2. doi:10.1038/330080a0

117. Frosch M, Strey A, Vogl T, Wulffraat NM, Kuis W, Sunderkotter C, et al. Myeloid-related proteins 8 and 14 are specifically secreted during interaction of phagocytes and activated endothelium and are useful markers for monitoring disease activity in pauciarticular-onset juvenile rheumatoid arthritis. Arthritis Rheum (2000) 43:628–37. doi:10.1002/1529-0131(200003)43:3<628::AID-ANR20>3.0.CO;2-X

118. Zreiqat H, Belluoccio D, Smith MM, Wilson R, Rowley LA, Jones K, et al. S100A8 and S100A9 in experimental osteoarthritis. Arthritis Res Ther (2010) 12:R16. doi:10.1186/ar2917

119. van Lent PL, Grevers L, Blom AB, Sloetjes A, Mort JS, Vogl T, et al. Myeloid-related proteins S100A8/S100A9 regulate joint inflammation and cartilage destruction during antigen-induced arthritis. Ann Rheum Dis (2008) 67:1750–8. doi:10.1136/ard.2007.077800

120. Wolf R, Mirmohammadsadegh A, Walz M, Lysa B, Tartler U, Remus R, et al. Molecular cloning and characterization of alternatively spliced mRNA isoforms from psoriatic skin encoding a novel member of the S100 family. FASEB J (2003) 17:1969–71. doi:10.1096/fj.03-0148fje

121. Fessatou S, Fagerhol MK, Roth J, Stamoulakatou A, Kitra V, Hadarean M, et al. Severe anemia and neutropenia associated with hyperzincemia and hypercalprotectinemia. J Pediatr Hematol Oncol (2005) 27:477–80. doi:10.1097/01.mph.0000179958.19524.9c

122. Isidor B, Poignant S, Corradini N, Fouassier M, Quartier P, Roth J, et al. Hyperzincemia and hypercalprotectinemia: unsuccessful treatment with tacrolimus. Acta Paediatr (2009) 98:410–2. doi:10.1111/j.1651-2227.2008.01092.x

123. Vaos G, Kostakis ID, Zavras N, Chatzemichael A. The role of calprotectin in pediatric disease. Biomed Res Int (2013) 2013:542363. doi:10.1155/2013/542363

124. Moroz OV, Antson AA, Grist SJ, Maitland NJ, Dodson GG, Wilson KS, et al. Structure of the human S100A12-copper complex: implications for host-parasite defence. Acta Crystallogr D Biol Crystallogr (2003) 59:859–67. doi:10.1107/S0907444903004700

125. Realegeno S, Kelly-Scumpia KM, Dang AT, Lu J, Teles R, Liu PT, et al. S100A12 is part of the antimicrobial network against Mycobacterium leprae in human macrophages. PLoS Pathog (2016) 12:e1005705. doi:10.1371/journal.ppat.1005705

126. Haley KP, Delgado AG, Piazuelo MB, Mortensen BL, Correa P, Damo SM, et al. The human antimicrobial protein calgranulin C participates in control of Helicobacter pylori growth and regulation of virulence. Infect Immun (2015) 83:2944–56. doi:10.1128/IAI.00544-15

127. Kang JH, Hwang SM, Chung IY. S100A8, S100A9 and S100A12 activate airway epithelial cells to produce MUC5AC via extracellular signal-regulated kinase and nuclear factor-kappaB pathways. Immunology (2015) 144:79–90. doi:10.1111/imm.12352

128. Nagareddy PR, Murphy AJ, Stirzaker RA, Hu Y, Yu S, Miller RG, et al. Hyperglycemia promotes myelopoiesis and impairs the resolution of atherosclerosis. Cell Metab (2013) 17:695–708. doi:10.1016/j.cmet.2013.04.001

129. Mortensen OH, Nielsen AR, Erikstrup C, Plomgaard P, Fischer CP, Krogh-Madsen R, et al. Calprotectin – a novel marker of obesity. PLoS One (2009) 4:e7419. doi:10.1371/journal.pone.0007419

130. Catalan V, Gomez-Ambrosi J, Rodriguez A, Ramirez B, Rotellar F, Valenti V, et al. Increased levels of calprotectin in obesity are related to macrophage content: impact on inflammation and effect of weight loss. Mol Med (2011) 17:1157–67. doi:10.2119/molmed.2011.00144

131. Asakura M, Karaki F, Fujii H, Atsuda K, Itoh T, Fujiwara R. Vildagliptin and its metabolite M20.7 induce the expression of S100A8 and S100A9 in human hepatoma HepG2 and leukemia HL-60 cells. Sci Rep (2016) 6:35633. doi:10.1038/srep35633

132. van den Bos C, Roth J, Koch HG, Hartmann M, Sorg C. Phosphorylation of MRP14, an S100 protein expressed during monocytic differentiation, modulates Ca(2+)-dependent translocation from cytoplasm to membranes and cytoskeleton. J Immunol (1996) 156:1247–54.

133. Siegenthaler G, Roulin K, Chatellard-Gruaz D, Hotz R, Saurat JH, Hellman U, et al. A heterocomplex formed by the calcium-binding proteins MRP8 (S100A8) and MRP14 (S100A9) binds unsaturated fatty acids with high affinity. J Biol Chem (1997) 272:9371–7. doi:10.1074/jbc.272.14.9371

134. Yang Z, Tao T, Raftery MJ, Youssef P, Di Girolamo N, Geczy CL. Proinflammatory properties of the human S100 protein S100A12. J Leukoc Biol (2001) 69:986–94.

135. Foell D, Seeliger S, Vogl T, Koch HG, Maschek H, Harms E, et al. Expression of S100A12 (EN-RAGE) in cystic fibrosis. Thorax (2003) 58:613–7. doi:10.1136/thorax.58.7.613

136. Mocellin S, Zavagno G, Nitti D. The prognostic value of serum S100B in patients with cutaneous melanoma: a meta-analysis. Int J Cancer (2008) 123:2370–6. doi:10.1002/ijc.23794

137. Gazzolo D, Michetti F. Perinatal S100B protein assessment in human unconventional biological fluids: a minireview and new perspectives. Cardiovasc Psychiatry Neurol (2010) 2010:703563. doi:10.1155/2010/703563

138. Heizmann CW. The multifunctional S100 protein family. Methods Mol Biol (2002) 172:69–80. doi:10.1385/1-59259-183-3:069

139. Oesterle A, Bowman MA. S100A12 and the S100/calgranulins: emerging biomarkers for atherosclerosis and possibly therapeutic targets. Arterioscler Thromb Vasc Biol (2015) 35:2496–507. doi:10.1161/ATVBAHA.115.302072

140. Bogdanova MV, Rameev VV, Kozlovskaya LV, Fedorov ES, Salugina SO. [Serum calgranulin C is a highly sensitive autoinflammation activity indicator in patients with familial periodic fevers]. Ter Arkh (2016) 88:58–64. doi:10.17116/terarkh201688658-64

141. Kallinich T, Wittkowski H, Keitzer R, Roth J, Foell D. Neutrophil-derived S100A12 as novel biomarker of inflammation in familial Mediterranean fever. Ann Rheum Dis (2010) 69:677–82. doi:10.1136/ard.2009.114363

142. Geven EJ, Van Den Bosch MH, Di Ceglie I, Ascone G, Abdollahi-Roodsaz S, Sloetjes AW, et al. S100A8/A9, a potent serum and molecular imaging biomarker for synovial inflammation and joint destruction in seronegative experimental arthritis. Arthritis Res Ther (2016) 18:247. doi:10.1186/s13075-016-1121-z

143. Ionita MG, Vink A, Dijke IE, Laman JD, Peeters W, Van Der Kraak PH, et al. High levels of myeloid-related protein 14 in human atherosclerotic plaques correlate with the characteristics of rupture-prone lesions. Arterioscler Thromb Vasc Biol (2009) 29:1220–7. doi:10.1161/ATVBAHA.109.190314

144. Xia GL, Wang YK, Huang ZQ. The correlation of serum myeloid-related protein-8/14 and eosinophil cationic protein in patients with coronary artery disease. Biomed Res Int (2016) 2016:4980251. doi:10.1155/2016/4980251

145. Wittkowski H, Kuemmerle-Deschner JB, Austermann J, Holzinger D, Goldbach-Mansky R, Gramlich K, et al. MRP8 and MRP14, phagocyte-specific danger signals, are sensitive biomarkers of disease activity in cryopyrin-associated periodic syndromes. Ann Rheum Dis (2011) 70:2075–81. doi:10.1136/ard.2011.152496

146. Holzinger D, Frosch M, Kastrup A, Prince FH, Otten MH, Van Suijlekom-Smit LW, et al. The toll-like receptor 4 agonist MRP8/14 protein complex is a sensitive indicator for disease activity and predicts relapses in systemic-onset juvenile idiopathic arthritis. Ann Rheum Dis (2012) 71:974–80. doi:10.1136/annrheumdis-2011-200598

147. Hurnakova J, Zavada J, Hanova P, Hulejova H, Klein M, Mann H, et al. Serum calprotectin (S100A8/9): an independent predictor of ultrasound synovitis in patients with rheumatoid arthritis. Arthritis Res Ther (2015) 17:252. doi:10.1186/s13075-015-0764-5

148. Shakeri M, Mahdkhah A, Panahi F. S100B protein as a post-traumatic biomarker for prediction of brain death in association with patient outcomes. Arch Trauma Res (2013) 2:76–80. doi:10.5812/atr.8549

149. Pelinka LE, Kroepfl A, Leixnering M, Buchinger W, Raabe A, Redl H. GFAP versus S100B in serum after traumatic brain injury: relationship to brain damage and outcome. J Neurotrauma (2004) 21:1553–61. doi:10.1089/neu.2004.21.1553

150. Dang X, Guan L, Hu W, Du G, Li J. S100B ranks as a new marker of multiple traumas in patients and may accelerate its development by regulating endothelial cell dysfunction. Int J Clin Exp Pathol (2014) 7:3818–26.

151. Blyth BJ, Farhavar A, Gee C, Hawthorn B, He H, Nayak A, et al. Validation of serum markers for blood-brain barrier disruption in traumatic brain injury. J Neurotrauma (2009) 26:1497–507. doi:10.1089/neu.2008-0738

152. Egea-Guerrero JJ, Revuelto-Rey J, Gordillo-Escobar E, Rodriguez-Rodriguez A, Enamorado-Enamorado J, Ruiz De Azua Lopez Z, et al. Serologic behavior of S100B protein in patients who are brain dead: preliminary results. Transplant Proc (2013) 45:3569–72. doi:10.1016/j.transproceed.2013.10.021

153. Thelin EP, Jeppsson E, Frostell A, Svensson M, Mondello S, Bellander BM, et al. Utility of neuron-specific enolase in traumatic brain injury; relations to S100B levels, outcome, and extracranial injury severity. Crit Care (2016) 20:285. doi:10.1186/s13054-016-1450-y

154. Weglewski A, Ryglewicz D, Mular A, Jurynczyk J. [Changes of protein S100B serum concentration during ischemic and hemorrhagic stroke in relation to the volume of stroke lesion]. Neurol Neurochir Pol (2005) 39:310–7.

155. Kaca-Orynska M, Tomasiuk R, Friedman A. Neuron-specific enolase and S100B protein as predictors of outcome in ischaemic stroke. Neurol Neurochir Pol (2010) 44:459–63. doi:10.1016/S0028-3843(14)60136-5

156. Esposito G, Imitola J, Lu J, De Filippis D, Scuderi C, Ganesh VS, et al. Genomic and functional profiling of human Down syndrome neural progenitors implicates S100B and aquaporin 4 in cell injury. Hum Mol Genet (2008) 17:440–57. doi:10.1093/hmg/ddm322

157. Lu J, Esposito G, Scuderi C, Steardo L, Delli-Bovi LC, Hecht JL, et al. S100B and APP promote a gliocentric shift and impaired neurogenesis in Down syndrome neural progenitors. PLoS One (2011) 6:e22126. doi:10.1371/journal.pone.0022126

158. Gogas H, Eggermont AM, Hauschild A, Hersey P, Mohr P, Schadendorf D, et al. Biomarkers in melanoma. Ann Oncol (2009) 20(Suppl 6):vi8–13. doi:10.1093/annonc/mdp251

159. Shishibori T, Oyama Y, Matsushita O, Yamashita K, Furuichi H, Okabe A, et al. Three distinct anti-allergic drugs, amlexanox, cromolyn and tranilast, bind to S100A12 and S100A13 of the S100 protein family. Biochem J (1999) 338(Pt 3):583–9. doi:10.1042/bj3380583

160. Semov A, Moreno MJ, Onichtchenko A, Abulrob A, Ball M, Ekiel I, et al. Metastasis-associated protein S100A4 induces angiogenesis through interaction with Annexin II and accelerated plasmin formation. J Biol Chem (2005) 280:20833–41. doi:10.1074/jbc.M412653200

161. Faries MB, Gupta RK, Ye X, Lee C, Yee R, Leopoldo Z, et al. A comparison of 3 tumor markers (MIA, TA90IC, S100B) in stage III melanoma patients. Cancer Invest (2007) 25:285–93. doi:10.1080/07357900701208634

162. Fernandez-Fernandez MR, Veprintsev DB, Fersht AR. Proteins of the S100 family regulate the oligomerization of p53 tumor suppressor. Proc Natl Acad Sci U S A (2005) 102:4735–40. doi:10.1073/pnas.0501459102

163. Gieldon A, Mori M, Del Conte R. Theoretical study on binding of S100B protein. J Mol Model (2007) 13:1123–31. doi:10.1007/s00894-007-0231-6

164. Most P, Boerries M, Eicher C, Schweda C, Ehlermann P, Pleger ST, et al. Extracellular S100A1 protein inhibits apoptosis in ventricular cardiomyocytes via activation of the extracellular signal-regulated protein kinase 1/2 (ERK1/2). J Biol Chem (2003) 278:48404–12. doi:10.1074/jbc.M308587200

165. Pleger ST, Shan C, Ksienzyk J, Bekeredjian R, Boekstegers P, Hinkel R, et al. Cardiac AAV9-S100A1 gene therapy rescues post-ischemic heart failure in a preclinical large animal model. Sci Transl Med (2011) 3:92ra64. doi:10.1126/scitranslmed.3002097

166. Ritterhoff J, Most P. Targeting S100A1 in heart failure. Gene Ther (2012) 19:613–21. doi:10.1038/gt.2012.8

167. Tsumura H, Akimoto M, Kiyota H, Ishii Y, Ishikura H, Honma Y. Gene expression profiles in differentiating leukemia cells induced by methyl jasmonate are similar to those of cytokinins and methyl jasmonate analogs induce the differentiation of human leukemia cells in primary culture. Leukemia (2009) 23:753–60. doi:10.1038/leu.2008.347

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Xia, Braunstein, Toomey, Zhong and Rao. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

MINI REVIEW
published: 25 January 2018
doi: 10.3389/fimmu.2018.00052

Sensing Bacterial-Induced DNA Damaging Effects via Natural Killer Group 2 Member D Immune Receptor: From Dysbiosis to Autoimmunity and Carcinogenesis

J. Luis Espinoza[1]* and Mika Minami[2]

1 Department of Hematology and Rheumatology, Faculty of Medicine, Kindai University, Osakasayama, Japan

2 Faculty of Medicine, Kindai University, Higashi-osaka, Japan

OPEN ACCESS

Edited by:

Leticia A. Carneiro, Universidade Federal do Rio de Janeiro, Brazil

Reviewed by:

Marit Inngjerdingen, Oslo University Hospital, Norway
Jue Hou, Virginia Mason Medical Center, United States
Guntram A. Grassl, Hannover Medical School, Germany

*Correspondence:

J. Luis Espinoza
luis@med.kindai.ac.jp

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 06 November 2017
Accepted: 09 January 2018
Published: 25 January 2018

Citation:

Espinoza JL and Minami M (2018) Sensing Bacterial-Induced DNA Damaging Effects via Natural Killer Group 2 Member D Immune Receptor: From Dysbiosis to Autoimmunity and Carcinogenesis. Front. Immunol. 9:52. doi: 10.3389/fimmu.2018.00052

The human genome is constantly exposed to exogenous and endogenous DNA damaging factors that frequently cause DNA damages. Unless repaired, damaged DNA can result in deleterious mutations capable of causing malignant transformation. Accordingly, cells have developed an advanced and effective surveillance system, the DNA damage response (DDR) pathway, which maintains genetic integrity. In addition to well-defined outcomes, such as cell cycle arrest, apoptosis, and senescence, another consequence of DDR activation is the induction of natural killer group 2 member D ligands (NKG2D-Ls) on the surface of stressed cells. Consequently, NKG2D-Ls-expressing cells are recognized and eliminated by NKG2D receptor-expressing immune cells, including NK cells, and various subsets of T-cells. Recent pieces of evidence indicate that commensal microbial imbalance (known as dysbiosis) can trigger DDR activation in host cells, which may result in sustained inflammatory responses. Therefore, dysbiosis can be seen as an important source of DNA damage agents that may be partially responsible for the overexpression of NKG2D-Ls on intestinal epithelial cells that is frequently observed in patients with inflammatory bowel disease and other disorders associated with altered human microbiota, including the development of colorectal cancer. In this article, we discuss recent evidence that appears to link an altered human microbiota with autoimmunity and carcinogenesis via the activation of DDR signals and the induction of NKG2D-Ls in stressed cells.

Keywords: natural killer group 2 member D ligands, microbiota, dysbiosis, bacterial genotoxin, immunosurveillance, inflammatory bowel disease

INTRODUCTION

The DNA damage response (DDR) is a highly efficient network of cellular pathways that play a crucial role in maintaining DNA integrity (1, 2). This surveillance system is responsible for monitoring, detecting and repairing DNA lesions, in order to prevent the generation of potentially deleterious mutations, which otherwise may result in the irreversible damage of DNA molecules, leading to cancer and other alterations in cell behavior (3, 4). The accumulation of un-repaired DNA damages in non-replicating cells, such as most of the cells in the brains or muscles of adults, is believed to contribute to the aging process in humans (5, 6). In highly replicating cells, such as hematopoietic stem cells and epithelial cells, DNA mutations that result from unrepaired DNA damages play a crucial role in malignant transformation and cancer progression (5, 7, 8). Endogenous agents capable of harming DNA, such as reactive oxygen species (ROS), lipid peroxidation products, and reactive nitrogen species (RNS) are naturally released during cell metabolic activities or hydrolytic processes (1, 9). In addition, DDR activation can be triggered by thousands of exogenous agents, including ionizing radiation, chemotherapy, virus infections, and chronic inflammation (10–13).

DNA damage response activation is controlled by three protein kinases: ataxia telangiectasia mutated (ATM), DNA-dependent protein kinase (DNA-PK), and ATM- and Rad3-related (ATR) (7, 14). Both ATM and DNA-PK are recruited by DNA double strand breaks (DSB), however, whereas DNA-PK coordinates DSB repair via non-homologous coupling, ATM promotes homologous recombination and cell cycle arrest at various checkpoints (14). ATR is activated in response to persistent single-stranded DNA and acts at the S-phase checkpoint (14). Upon DNA damage recognition, these kinases activate various downstream mediators including p53, CHK1, CHK2, BRCA, and H2AX, which (depending on the extent of DNA damage) may lead to cell-cycle arrest, DNA repair, senescence, or apoptosis (14–16). A key mediator of ATM signal is the checkpoint kinase Chk2, which induces G1/S checkpoint via Cdk2 inactivation or can block cell cycle at G2/M by preventing cyclinB1/Cdk1 complex formation (17). On the other hand, Chk1, triggered by ATR signal, activates Cdc25A phosphatase and Treslin, which induce G2 and S phase arrest (7).

Another consequence of DDR and ATM/ATR activation is the induction of cell stress molecules that are proteins expressed on the surface of damaged cells (18, 19). These stress ligands, which are usually absent in normal cells, are specifically recognized by either, the natural killer group 2 member D (NKG2D) and the DNAX accessory molecule immunoreceptors (18–20).

Natural killer group 2 member D, also known as Klrk1, is a C-type lectin-like type II transmembrane protein constitutively expressed by NK cells, activated macrophages and various T-cell subsets, such as NKT cells, CD8+ αβ, CD4+αβ, and γδ T lymphocytes (21–23). Upon engagement of specific NKG2D ligands (NKG2D-Ls), NKG2D receptor activates downstream signaling pathways resulting in effector immune responses like cytokine releases and cellular cytotoxicity (22, 24).

Recent evidence has linked various bacterial pathogens with DDR activation caused by either the direct effect of microbe produced genotoxins (25–28) or indirectly by ROS or RNS that result from the prolonged or excessive activation of host immune cells in response to certain microbes or their metabolic end-products (29, 30). This bacterial-induced DDR is not limited to highly pathogenic bacteria, since genotoxic damage induced by certain members of the commensal bacteria community (termed the “microbiota”) have been also documented (31, 32). Notably, increased expression of NKG2D-Ls on the surface of intestinal epithelial cells and its recognition by NKG2D receptor-expressing immune cells is believed to contribute to the pathogenesis of inflammatory bowel diseases (IBDs), such as ulcerative colitis and Crohn’s disease (33–35) and dysregulated gut microbiota has been etiologically linked to IBD and colorectal cancer (CRC) (27, 36, 37). In this article, we discuss recent pieces of evidence that appear to link alterations in gut microbiota with activation of the DDR. The potential effects that perturbations in this network have on the development of autoimmunity and cancer immunosurveillance are also discussed.

NKG2D-Ls EXPRESSION

In humans, multiple families of structurally unrelated NKG2D-Ls have been identified, including the MHC class I chain-related molecules (MICA and MICB), and the UL-16 binding proteins (ULBP1, -2, -3, -4, -5, and -6) (24, 38–40). NKG2D-Ls are absent or poorly expressed on the surfaces of normal cells but they are induced under certain pathological conditions like heat shock, virus infection, oxidative stress, and malignant transformation (39, 41). The elimination of NKG2D-Ls expressing cells by NKG2D receptor-expressing immune cells is one of the underlying grounds of the concept of cancer immunosurveillance (42–44). NKG2D-Ls upregulation has been described in various human cancers, including carcinomas of the breast (45), lung, colon (46) and prostate cancer (47), as well as in melanomas (48), gliomas (49), leukemias (18), and cervix cancer (50). The expression of these molecules is tightly regulated by mechanisms that control gene transcription, mRNA stability, protein translation, and stabilization (20, 39). Intriguingly, NKG2D-Ls expression has also been documented in certain normal cells. For example, in primary bronchial epithelial cells, MICA and ULBP1-4 are detectable mainly at intracellular level, but become detectable on the cell surface when the cells are exposed to oxidative stress (51). NKG2D-Ls (mainly ULBP1) have also been detected in peripheral blood cells (52) and these proteins are particularly upregulated in activated T cells and B cells (20, 53, 54). In addition, normal gut epithelium constitutively expresses MICA, although most cells appear to express these proteins intracellularly (55). On the other hand, the aberrant expression of NKG2D-Ls has been documented in certain autoimmune diseases, especially in the damaged tissues of patients with inflammatory bowel disease (IBD) that includes Crohn’s disease (35, 56) and ulcerative colitis (33). In these disorders, NKG2D-Ls expression correlates with increased number of infiltrating NKG2D+ lymphocytes in the damaged tissues (33, 35). Consistent with these observations, a randomized controlled clinical trial recently showed that a single dose of an anti-NKG2D blocking monoclonal antibody, significantly reduced disease activity in patients with active Crohn’s disease (57). Despite the relatively small size of this study (78 patients) and the fact that patients with UC were not included, these encouraging data support the involvement of NKG2D/NKG2D-Ls axis in the pathogenesis or clinical course of IBD.

In patients with active Celiac disease, MICA is strongly expressed on the surface of intestinal epithelial cells and it is further upregulated by wheat gliadin, which triggers the activation of intraepithelial NKG2D+ lymphocytes, leading to epithelial damage and villous atrophy (55). Notably, the probiotics Lactobacillus fermentum and Bifidobacterium lactis were found to directly inhibit the toxic effects of gliadin in intestinal cells (58) and a gluten-free diet strongly downregulated NKG2D-Ls in intestinal epithelial cells and concomitantly decreased NKG2D receptor expression on infiltrating NK cells (59).

DYSBIOSIS AND NKG2D-Ls EXPRESSION

The community of commensal microorganisms living within the human intestines, known as gut microbiota, plays critical roles in maintaining immune tolerance and epithelial integrity (60–62).

Significant upregulation of NKG2D-Ls was observed in the intestinal mucosa of germ-free mice lacking commensal microbiota, as well as in commensal-depleted animals (ampicillin-treated mice), and low ligands expression level was restored when ampicillin treatment was stopped. Strikingly, the same study found low levels of NKG2D-Ls in animals treated with vancomycin, which was attributed to the selective propagation of the vancomycin-resistant bacterium Akkermansia muciniphila in mice intestines (31), indicating that NKG2D-Ls expression, at least in intestinal tissues, is largely influenced by the gut microbiota composition. Interestingly, A. muciniphila has been linked with anti-inflammatory protective properties against IBD (63).

The loss of microbial balance and the overgrowth of pathogenic bacteria (known as dysbiosis) is often associated with the development of autoimmune disorders and the development of CRC (62, 64, 65). Strikingly, direct microbe-induced NKG2D-Ls upregulation has been documented in human intestinal epithelial cells exposed to Escherichia coli strains, where the interaction between bacterial adhesin AfaE and its cellular receptor CD55 results in MICA expression (66).

Another study showed that Pseudomonas aeruginosa infection increased NKG2D-Ls (Rae1) in mouse airway epithelial cells in vivo and upregulated ULBP2 in human airway epithelial cells in vitro, although the mechanism of ligand induction by this pathogen is unknown (67).

Propionibacterium acnes was recently linked with Corpus-dominant lymphocytic gastritis (CDLG), a Helicobacter pylori negative entity and typically characterized by extensive infiltration of CD8+ T-cells in the stomach epithelium. Interestingly, P. acnes infection correlated with increased levels of IL-15 and the upregulation of NKG2D-Ls in the inflamed gastric epithelium. Although the mechanisms leading to NKG2D-L upregulation in this entity remains unclear, a microbe-derived stimuli, probably live P. acnes or microbial-derived short-chain fatty acids were proposed as triggering factors (68). Notably, CDLG frequently coexists with autoimmune disorders with altered microbiota including Celiac disease (69) and Crohn’s disease (70, 71). Moreover, propionic acid, derived from the fermentation of plant-derived dietary fiber mainly under the presence Propionibacterium, upregulated MICA/B in human cells including, activated T lymphocytes and different cancer lines (72). Mycobacterium tuberculosis (M. tuberculosis)-infected dendritic and airway epithelial cells also upregulate MICA expression in vitro and in vivo, and ligand recognition by Vγ2Vδ2 T cells expressing NKG2D receptor induces a potent inflammatory reaction (73).

Of note, albeit the above studies indicate that NKG2D-Ls upregulation is frequently observed in host cells exposed to various bacteria or their products, the molecular mechanisms of this phenomenon have not been elucidated. In addition, the exact mechanism that determines the fate of host cells exposed to dysbiosis (cell cycle arrest, apoptosis, malignant transformation or NKG2D-Ls upregulation) is currently unknown. Current data suggest that the extent of DNA damage and the resultant cellular responses determine cell fate under these stress conditions, hence in the context of dysbiosis, it is conceivable that cell fate may be dependent on the specific bacterium or group of bacteria dysregulated in the host.

As mentioned above, commensal bacteria play critical roles maintaining gut homeostasis, and this particular feature can be exploited for therapeutic purposes. The oral administration of commensal lactic acid bacteria effectively protected mice from dextran sulfate sodium-induced experimental colitis, which was attributed to the enhanced interferon-β production triggered by double-stranded RNA derived from commensal lactic acid bacteria (74). Although this study did not explore NKG2D-Ls expression on intestinal cells, it is worth mentioning that type I interferons have been shown to downregulate NKG2D-Ls expression impairing NK cells-dependent killing of target cells (38).

BACTERIAL GENOTOXINS AND DDR ACTIVATION

Various intestinal bacteria are known to release genotoxins (bacterial products capable of targeting host DNA), which together with the induction of sustained inflammation, promotes genomic instability and ultimately autoimmunity or cancer (Table 1). The first characterized bacterial genotoxin was cytolethal distending toxins (CDT), which is produced by several Gram-negative bacteria, including E. coli, Campylobacter sp., helicobacter sp., Shigella dysenteriae, and Haemophilus ducreyi. CDT induces DNA DSB in exposed host cells that may lead to transient cell cycle arrest or malignant transformation (32, 36, 75). Mouse liver cells exposed to CDT producing helicobacter develop dysplasia (76) and fibroblasts or intestinal epithelial cells chronically exposed to large concentrations of CDT, in the absence of immune cell clearance, show genomic instability, fail to activate DDR, and eventually become prone to malignant transformation (32).

TABLE 1 | Bacterial pathogens or their products that activate DNA damage response (DDR) and may induce NKG2D ligands (NKG2D-Ls) expression in host cells.

Another bacterial-derived genotoxin is colibactin produced by E. coli strains of the B2 phylogroup harboring the polyketide synthetase island (pks), which is also found in other Enterobacteriaceae members such as Proteus mirabilis and Klebsiella pneumoniae (84–86). Infection with E. coli harboring this genomic cluster generates DSB leading to DDR activation, cell cycle arrest and genomic instability (36, 85). Notably, E. coli harboring pks are frequently detectable in patients with IBD, as well as in patients with CRC, suggesting that pks is directly related to disease pathogenesis (26, 92).

Escherichia coli uropathogenic-specific protein (Usp) is another bacterial toxin that induces genotoxic stress and activates DDR in exposed cells. This genotoxin is produced by E. coli strains associated with pyelonephritis, prostatitis, and bacteremia (87, 88). Purified Usp cleaves linearized naked DNA in vitro and causes DNA fragmentation in mammalian cells (28).

Interestingly, compared with normal intestinal samples where toxin-producing bacteria constitute a minority of the commensal microbiota, human CRC tissues contain a high expression of these microorganisms (93).

Cyclo phenylalanine-proline (cFP) is other genotoxin produced by various bacteria such as Lactobacillus reuteri, Streptomyces sp. AMLK-335, Vibrio vulnificus, V. cholera, P. aeruginosa, and P. putida. Mammalian cells like INT-407, U2OS, and Huh7 cells exposed to cFP develop DSB and eventually activate ATM-CHK2 (90).

Pneumolysin, a toxin produced by Streptococcus pneumoniae and a key virulence factor against host cells, induces DSBs and ATM-mediated H2AX phosphorylation in epithelial alveolar cells. Consequently pneumolysin-exposed cells undergo cell cycle arrest and apoptosis, although the induction of NKG2D-Ls was not investigated in this study (91). Interestingly, pyruvate oxidase, another cytotoxin released by S. pneumoniae, induces DSBs and contributes to pneumolysin release (81).

Bacteria may also trigger DDR activation by inducing the expression of enzymes that enhance ROS in host cells, which lead to DNA damage or the induction of chronic inflammation (36, 94, 95). Bacterial-induced DNA damage can be further amplified by ROS released from immune cells at sites of chronic inflammation since inflammatory especially macrophages and neutrophils constitute a constant source of ROS, RNS, and cytokines that can develop in response to dysbiosis (94). Granuloma formation associated with M. tuberculosis infection was recently linked to the persistent inflammatory signals mediated by toll-like receptor (TLR) signals, which promotes macrophage polyploidy by regulating DDR signals via ATR activation (78). Although this study did not assess the expression of NKG2D-Ls in polyploidy macrophages, previous studies have shown NKG2D-Ls upregulation in macrophages exposed to bacterial-derived products via TLR signal activation (79, 80).

In addition, convincing evidence have established a link between infections with certain bacteria such as E. coli, Bacteroides fragilis, and Fusobacterium nucleatum with the development of CRC (26), indicating that dysbiosis may either cause carcinogenesis or autoimmunity. In some circumstances, bacterial can interrupt the DDR activation in host cells, thus allowing the cell cycle to progress in cells with unrepaired or in repaired with errors DNA resulting in mutations of critical genes associated with malignant transformation (26, 32). It is conceivable that the chronic exposure to genotoxin-secreting bacteria in host cells with fully functional DDR may result in NKG2D-Ls overexpression, via ATM activation, which ultimately increases the risk of autoimmunity. Conversely, in host cells with failed DDR, DSB may result in the survival of cells with unrepaired DNA and elevated risk of malignant transformation (Figure 1). This assumption is supported by the observations that epithelial cells exposed to Chlamydia trachomatis, a bacteria associated with cervical and ovarian cancer development, undergo DNA damaged but fail to activate DDR, due to bacterial-induced impaired DNA repair. Consequently, infected cells continue to proliferate in an environment favorable for malignant transformation (96).Thus, the NKG2D/NKG2D-Ls axis maintains a delicate immune equilibrium, which is crucial for cancer immunosurveillance but that, under certain conditions, it can eventually promote autoimmunity (97).

FIGURE 1 | Commensal bacteria play an important role in maintaining gut homeostasis (A). In normal condition, intestinal epithelial cells express low levels of NKG2D ligands (NKG2D-Ls) (mostly intracellular) and beneficial bacterial contribute to immune education and help to maintain immune tolerance by promoting the induction and accumulation of regulatory T cells (Treg cells). In the context of microbial imbalance (dysbiosis), pathogenic bacteria may release genotoxins that generate DNA damage in host cells. DNA damage response (DDR) is then activated, which may lead to cell cycle arrest, apoptosis, or NKG2D-Ls induction in exposed cells. These events, together with the sustained immune activation in response to dysbiosis may eventually contribute to the development of autoimmune disorders or malignant transformation. Although the precise mechanisms that determine why cells take one of these two contrasting cell fates are unclear, current data appear to suggest that (B) bacteria-induced DNA damage in cells with failed DDR (caused by mutations or by inhibitory factors secreted by bacteria) may result in the survival and proliferation of cells with unrepaired DNA, increasing the risk of malignant transformation. Transformed cells may release or shade NKG2D-Ls that impairs NKG2D receptor-mediated functions leading to failed immune surveillance and tumor growth. (C) Alternatively, the chronic exposure to genotoxin-secreting bacterial in host cells with fully functional DDR may result in NKG2D-Ls overexpression via ataxia telangiectasia mutated activation leading to increased risk of autoimmunity.

Despite the above studies have consistently shown that bacterial genotoxin can activate DDR, unfortunately, none of them explored the potential NKG2D-Ls upregulation in host cells exposed to those bacterial products; therefore, further studied are needed to elucidate the immunological impact of genotoxins.

CONCLUDING REMARKS

Cells are continuously exposed to hostile environmental stressors, including extremes of temperature, toxins, and oxygen or nutrient deprivation. As a result, cells have evolved a wide range of molecular changes and stress responses to minimize damage which, depending on the severity and duration of stress confronted, can range from the activation of survival-promoting pathways to eliciting cell senescence or programmed cell death (98). During the last few years, remarkable progress has been made in our understanding of the molecular mechanisms of DDR activation and its role in various cellular processes like aging and cancer development, and it has become apparent that the immune system constitutes an important component of the cellular response to DNA damage stressors (99). In this regard, the recognition of NKG2D-Ls overexpressed on the surface of host cells as a consequence of DDR activation constitutes one of the mechanisms by which the immune cells expressing the NKG2D receptor can sense DNA damage in host cells (100). Importantly, dysbiosis has been causally liked with DDR activation, via either the release of genotoxin or by promoting chronic inflammation (92). Several questions about the causal link of dysbiosis with NKG2D-Ls induction via DDR remain unanswered. For example, the precise mechanisms by which DDR induced by bacterial genotoxins or dysbiosis may result in malignant transformation or autoimmunity have not been fully elucidated. In addition, the specific role of dysbiosis in the upregulation of NKG2D-Ls in the intestinal epithelial cells of patients with IBD needs to be clarified. The polymorphism rs1049174 in the NKG2D gene (generating HNK and LNK genotypes) influences NKG2D receptor expression on immune cells and is implicated in individual susceptibility to certain cancers. The HNK genotype is associated with greater NK cells cytotoxic activity and lower prevalence of epithelial cells-derived malignancies, in comparison with the low cytotoxic genotype LNK (23, 50, 101, 102). It is currently unknown if HNK and LNK genotypes can affect NKG2D receptor-mediated immune responses in the context of dysbiosis and if they are implicated in the development of autoimmune disorders like IBD. Future studies on this regard are warranted as genetic variants of human genes involved in immunity and gut architecture are associated with an altered composition of the gut microbiome (103).

Compelling evidence indicate that dysbiosis is implicated in the pathogenesis of several human diseases, ranging from metabolic disorders, autoimmunity, and cancer (64, 104, 105) and various studies have shown that manipulating human microbiota, for example by using probiotics or fecal transplantation, has promising therapeutic potential (65, 106, 107). Further research is needed to uncover the specific microbes within a dysbiotic microbiota that are directly implicated in disease etiology. Designing optimal interventions aimed to remove pathogenic microorganisms or for replacing them with beneficial ones will have enormous therapeutic potential.

AUTHOR CONTRIBUTIONS

The authors contributed extensively to the work presented in this paper. JE: conceived and designed the study; created and drew figures and wrote the manuscript. MM: wrote the manuscript, searched and collected bibliography.

REFERENCES

1. Roos WP, Kaina B. DNA damage-induced apoptosis: from specific DNA lesions to the DNA damage response and apoptosis. Cancer Lett (2012) 332(2):237–48. doi:10.1016/j.canlet.2012.01.007

2. Pearl LH, Schierz AC, Ward SE, Al-Lazikani B, Pearl FM. Therapeutic opportunities within the DNA damage response. Nat Rev Cancer (2015) 15(3):166–80. doi:10.1038/nrc3891

3. Gasser S, Raulet D. The DNA damage response, immunity and cancer. Semin Cancer Biol (2006) 16(5):344–7. doi:10.1016/j.semcancer.2006.07.004

4. Cirombella R, Montrone G, Stoppacciaro A, Giglio S, Volinia S, Graziano P, et al. Fhit loss in lung preneoplasia: relation to DNA damage response checkpoint activation. Cancer Lett (2010) 291(2):230–6. doi:10.1016/j.canlet.2009.10.017

5. Iyama T, Wilson DM. DNA repair mechanisms in dividing and non-dividing cells. DNA Repair (Amst) (2013) 12(8):620–36. doi:10.1016/j.dnarep.2013.04.015

6. Vahidi Ferdousi L, Rocheteau P, Chayot R, Montagne B, Chaker Z, Flamant P, et al. More efficient repair of DNA double-strand breaks in skeletal muscle stem cells compared to their committed progeny. Stem Cell Res (2014) 13(3 Pt A):492–507. doi:10.1016/j.scr.2014.08.005

7. Delia D, Mizutani S. The DNA damage response pathway in normal hematopoiesis and malignancies. Int J Hematol (2017) 106(3):328–34. doi:10.1007/s12185-017-2300-7

8. Mo D, Zhao Y, Balajee AS. Human RecQL4 helicase plays multifaceted roles in the genomic stability of normal and cancer cells. Cancer Lett (2017) 413:1–10. doi:10.1016/j.canlet.2017.10.021

9. Lonkar P, Dedon PC. Reactive species and DNA damage in chronic inflammation: reconciling chemical mechanisms and biological fates. Int J Cancer (2011) 128(9):1999–2009. doi:10.1002/ijc.25815

10. Goldstein M, Kastan MB. The DNA damage response: implications for tumor responses to radiation and chemotherapy. Annu Rev Med (2015) 66:129–43. doi:10.1146/annurev-med-081313-121208

11. Sprung CN, Ivashkevich A, Forrester HB, Redon CE, Georgakilas A, Martin OA. Oxidative DNA damage caused by inflammation may link to stress-induced non-targeted effects. Cancer Lett (2015) 356(1):72–81. doi:10.1016/j.canlet.2013.09.008

12. Luftig MA. Viruses and the DNA damage response: activation and antagonism. Annu Rev Virol (2014) 1(1):605–25. doi:10.1146/annurev-virology-031413-085548

13. Williamson L, Saponaro M, Boeing S, East P, Mitter R, Kantidakis T, et al. UV irradiation induces a non-coding RNA that functionally opposes the protein encoded by the same gene. Cell (2017) 168(5):843–55.e13. doi:10.1016/j.cell.2017.01.019

14. Blackford AN, Jackson SP. ATM, ATR, and DNA-PK: the trinity at the heart of the DNA damage response. Mol Cell (2017) 66(6):801–17. doi:10.1016/j.molcel.2017.05.015

15. Shiloh Y, Ziv Y. The ATM protein: the importance of being active. J Cell Biol (2012) 198(3):273–5. doi:10.1083/jcb.201207063

16. Ditch S, Paull TT. The ATM protein kinase and cellular redox signaling: beyond the DNA damage response. Trends Biochem Sci (2012) 37(1):15–22. doi:10.1016/j.tibs.2011.10.002

17. Takagi M. DNA damage response and hematological malignancy. Int J Hematol (2017) 106(3):345–56. doi:10.1007/s12185-017-2226-0

18. Luis Espinoza J, Takami A, Trung LQ, Nakao S. Ataxia-telangiectasia mutated kinase-mediated upregulation of NKG2D ligands on leukemia cells by resveratrol results in enhanced natural killer cell susceptibility. Cancer Sci (2013) 104(6):657–62. doi:10.1111/cas.12141

19. Gasser S, Orsulic S, Brown EJ, Raulet DH. The DNA damage pathway regulates innate immune system ligands of the NKG2D receptor. Nature (2005) 436(7054):1186–90. doi:10.1038/nature03884

20. Cerboni C, Fionda C, Soriani A, Zingoni A, Doria M, Cippitelli M, et al. The DNA damage response: a common pathway in the regulation of NKG2D and DNAM-1 ligand expression in normal, infected, and cancer cells. Front Immunol (2014) 4:508. doi:10.3389/fimmu.2013.00508

21. Zingoni A, Ardolino M, Santoni A, Cerboni C. NKG2D and DNAM-1 activating receptors and their ligands in NK-T cell interactions: role in the NK cell-mediated negative regulation of T cell responses. Front Immunol (2012) 3:408. doi:10.3389/fimmu.2012.00408

22. Espinoza JL, Takami A, Yoshioka K, Nakata K, Sato T, Kasahara Y, et al. Human microRNA-1245 downregulates the NKG2D receptor in NK cells and impairs NKG2D-mediated functions. Haematologica (2012) 97(9):1295–303. doi:10.3324/haematol.2011.058529

23. Espinoza JL, Takami A, Onizuka M, Sao H, Akiyama H, Miyamura K, et al. NKG2D gene polymorphism has a significant impact on transplant outcomes after HLA-fully-matched unrelated bone marrow transplantation for standard risk hematologic malignancies. Haematologica (2009) 94(10):1427–34. doi:10.3324/haematol.2009.008318

24. Jelenčić V, Lenartić M, Wensveen FM, Polić B. NKG2D: a versatile player in the immune system. Immunol Lett (2017) 189:48–53. doi:10.1016/j.imlet.2017.04.006

25. Žgur-Bertok D. DNA damage repair and bacterial pathogens. PLoS Pathog (2013) 9(11):e1003711. doi:10.1371/journal.ppat.1003711

26. Gagnaire A, Nadel B, Raoult D, Neefjes J, Gorvel JP. Collateral damage: insights into bacterial mechanisms that predispose host cells to cancer. Nat Rev Microbiol (2017) 15(2):109–28. doi:10.1038/nrmicro.2016.171

27. Ray D, Kidane D. Gut microbiota imbalance and base excision repair dynamics in colon cancer. J Cancer (2016) 7(11):1421–30. doi:10.7150/jca.15480

28. Grasso F, Frisan T. Bacterial genotoxins: merging the DNA damage response into infection biology. Biomolecules (2015) 5(3):1762–82. doi:10.3390/biom5031762

29. Fahrer J, Huelsenbeck J, Jaurich H, Dörsam B, Frisan T, Eich M, et al. Cytolethal distending toxin (CDT) is a radiomimetic agent and induces persistent levels of DNA double-strand breaks in human fibroblasts. DNA Repair (Amst) (2014) 18:31–43. doi:10.1016/j.dnarep.2014.03.002

30. Kalisperati P, Spanou E, Pateras IS, Korkolopoulou P, Varvarigou A, Karavokyros I, et al. Inflammation, DNA damage, Helicobacter pylori and gastric tumorigenesis. Front Genet (2017) 8:20. doi:10.3389/fgene.2017.00020

31. Hansen CH, Holm TL, Krych Ł, Andresen L, Nielsen DS, Rune I, et al. Gut microbiota regulates NKG2D ligand expression on intestinal epithelial cells. Eur J Immunol (2013) 43(2):447–57. doi:10.1002/eji.201242462

32. Guidi R, Guerra L, Levi L, Stenerlöw B, Fox JG, Josenhans C, et al. Chronic exposure to the cytolethal distending toxins of Gram-negative bacteria promotes genomic instability and altered DNA damage response. Cell Microbiol (2013) 15(1):98–113. doi:10.1111/cmi.12034

33. La Scaleia R, Stoppacciaro A, Oliva S, Morrone S, Di Nardo G, Santoni A, et al. NKG2D/Ligand dysregulation and functional alteration of innate immunity cell populations in pediatric IBD. Inflamm Bowel Dis (2012) 18(10):1910–22. doi:10.1002/ibd.22899

34. Ito Y, Kanai T, Totsuka T, Okamoto R, Tsuchiya K, Nemoto Y, et al. Blockade of NKG2D signaling prevents the development of murine CD4+ T cell-mediated colitis. Am J Physiol Gastrointest Liver Physiol (2008) 294(1):G199–207. doi:10.1152/ajpgi.00286.2007

35. Allez M, Tieng V, Nakazawa A, Treton X, Pacault V, Dulphy N, et al. CD4+NKG2D+ T cells in Crohn’s disease mediate inflammatory and cytotoxic responses through MICA interactions. Gastroenterology (2007) 132(7):2346–58. doi:10.1053/j.gastro.2007.03.025

36. Gagnière J, Raisch J, Veziant J, Barnich N, Bonnet R, Buc E, et al. Gut microbiota imbalance and colorectal cancer. World J Gastroenterol (2016) 22(2):501–18. doi:10.3748/wjg.v22.i2.501

37. Carding S, Verbeke K, Vipond DT, Corfe BM, Owen LJ. Dysbiosis of the gut microbiota in disease. Microb Ecol Health Dis (2015) 26:26191. doi:10.3402/mehd.v26.26191

38. Zhang J, Basher F, Wu JD. NKG2D ligands in tumor immunity: two sides of a coin. Front Immunol (2015) 6:97. doi:10.3389/fimmu.2015.00097

39. Huergo-Zapico L, Acebes-Huerta A, López-Soto A, Villa-Álvarez M, Gonzalez-Rodriguez AP, Gonzalez S. Molecular bases for the regulation of NKG2D ligands in cancer. Front Immunol (2014) 5:106. doi:10.3389/fimmu.2014.00106

40. Lanier LL. NKG2D receptor and its ligands in host defense. Cancer Immunol Res (2015) 3(6):575–82. doi:10.1158/2326-6066.CIR-15-0098

41. Ardolino M, Zingoni A, Cerboni C, Cecere F, Soriani A, Iannitto ML, et al. DNAM-1 ligand expression on Ag-stimulated T lymphocytes is mediated by ROS-dependent activation of DNA-damage response: relevance for NK-T cell interaction. Blood (2011) 117(18):4778–86. doi:10.1182/blood-2010-08-300954

42. Sagiv A, Burton DG, Moshayev Z, Vadai E, Wensveen F, Ben-Dor S, et al. NKG2D ligands mediate immunosurveillance of senescent cells. Aging (Albany NY) (2016) 8(2):328–44. doi:10.18632/aging.100897

43. Weil S, Memmer S, Lechner A, Huppert V, Giannattasio A, Becker T, et al. Natural killer group 2D ligand depletion reconstitutes natural killer cell immunosurveillance of head and neck squamous cell carcinoma. Front Immunol (2017) 8:387. doi:10.3389/fimmu.2017.00387

44. Guerra N, Tan Y, Joncker N, Choy A, Gallardo F, Xiong N, et al. NKG2D-deficient mice are defective in tumor surveillance in models of spontaneous malignancy. Immunity (2008) 28(4):571–80. doi:10.1016/j.immuni.2008.02.016

45. Breunig C, Pahl J, Küblbeck M, Miller M, Antonelli D, Erdem N, et al. MicroRNA-519a-3p mediates apoptosis resistance in breast cancer cells and their escape from recognition by natural killer cells. Cell Death Dis (2017) 8(8):e2973. doi:10.1038/cddis.2017.364

46. Kim GR, Ha GH, Bae JH, Oh SO, Kim SH, Kang CD. Metastatic colon cancer cell populations contain more cancer stem-like cells with a higher susceptibility to natural killer cell-mediated lysis compared with primary colon cancer cells. Oncol Lett (2015) 9(4):1641–6. doi:10.3892/ol.2015.2918

47. Chávez-Blanco A, De la Cruz-Hernández E, Domínguez GI, Rodríguez-Cortez O, Alatorre B, Pérez-Cárdenas E, et al. Upregulation of NKG2D ligands and enhanced natural killer cell cytotoxicity by hydralazine and valproate. Int J Oncol (2011) 39(6):1491–9. doi:10.3892/ijo.2011.1144

48. Paschen A, Baingo J, Schadendorf D. Expression of stress ligands of the immunoreceptor NKG2D in melanoma: regulation and clinical significance. Eur J Cell Biol (2014) 93(1–2):49–54. doi:10.1016/j.ejcb.2014.01.009

49. Zhang X, Rao A, Sette P, Deibert C, Pomerantz A, Kim WJ, et al. IDH mutant gliomas escape natural killer cell immune surveillance by downregulation of NKG2D ligand expression. Neuro Oncol (2016) 18(10):1402–12. doi:10.1093/neuonc/now061

50. Espinoza JL, Nguyen VH, Ichimura H, Pham TT, Nguyen CH, Pham TV, et al. A functional polymorphism in the NKG2D gene modulates NK-cell cytotoxicity and is associated with susceptibility to human papilloma virus-related cancers. Sci Rep (2016) 6:39231. doi:10.1038/srep39231

51. Borchers MT, Harris NL, Wesselkamper SC, Vitucci M, Cosman D. NKG2D ligands are expressed on stressed human airway epithelial cells. Am J Physiol Lung Cell Mol Physiol (2006) 291(2):L222–31. doi:10.1152/ajplung.00327.2005

52. Nowbakht P, Ionescu MC, Rohner A, Kalberer CP, Rossy E, Mori L, et al. Ligands for natural killer cell-activating receptors are expressed upon the maturation of normal myelomonocytic cells but at low levels in acute myeloid leukemias. Blood (2005) 105(9):3615–22. doi:10.1182/blood-2004-07-2585

53. Raulet DH, Gasser S, Gowen BG, Deng W, Jung H. Regulation of ligands for the NKG2D activating receptor. Annu Rev Immunol (2013) 31:413–41. doi:10.1146/annurev-immunol-032712-095951

54. Cerboni C, Zingoni A, Cippitelli M, Piccoli M, Frati L, Santoni A. Antigen-activated human T lymphocytes express cell-surface NKG2D ligands via an ATM/ATR-dependent mechanism and become susceptible to autologous NK-cell lysis. Blood (2007) 110(2):606–15. doi:10.1182/blood-2006-10-052720

55. Hüe S, Mention JJ, Monteiro RC, Zhang S, Cellier C, Schmitz J, et al. A direct role for NKG2D/MICA interaction in villous atrophy during celiac disease. Immunity (2004) 21(3):367–77. doi:10.1016/j.immuni.2004.06.018

56. Vadstrup K, Galsgaard ED, Jensen H, Lanier LL, Ryan JC, Chen SY, et al. NKG2D ligand expression in Crohn’s disease and NKG2D-dependent stimulation of CD8(+) T cell migration. Exp Mol Pathol (2017) 103(1):56–70. doi:10.1016/j.yexmp.2017.06.010

57. Allez M, Skolnick BE, Wisniewska-Jarosinska M, Petryka R, Overgaard RV. Anti-NKG2D monoclonal antibody (NNC0142-0002) in active Crohn’s disease: a randomised controlled trial. Gut (2017) 66(11):1918–25. doi:10.1136/gutjnl-2016-311824

58. Lindfors K, Blomqvist T, Juuti-Uusitalo K, Stenman S, Venäläinen J, Mäki M, et al. Live probiotic Bifidobacterium lactis bacteria inhibit the toxic effects induced by wheat gliadin in epithelial cell culture. Clin Exp Immunol (2008) 152(3):552–8. doi:10.1111/j.1365-2249.2008.03635.x

59. Adlercreutz EH, Weile C, Larsen J, Engkilde K, Agardh D, Buschard K, et al. A gluten-free diet lowers NKG2D and ligand expression in BALB/c and non-obese diabetic (NOD) mice. Clin Exp Immunol (2014) 177(2):391–403. doi:10.1111/cei.12340

60. Espinoza JL, Elbadry MI, Nakao S. An altered gut microbiota may trigger autoimmune-mediated acquired bone marrow failure syndromes. Clin Immunol (2016) 171:62–4. doi:10.1016/j.clim.2016.08.008

61. Green ED, Watson JD, Collins FS. Human genome project: twenty-five years of big biology. Nature (2015) 526(7571):29–31. doi:10.1038/526029a

62. Espinoza JL, Kotecha R, Nakao S. Microbe-induced inflammatory signals triggering acquired bone marrow failure syndromes. Front Immunol (2017) 8:186. doi:10.3389/fimmu.2017.00186

63. Png CW, Lindén SK, Gilshenan KS, Zoetendal EG, McSweeney CS, Sly LI, et al. Mucolytic bacteria with increased prevalence in IBD mucosa augment in vitro utilization of mucin by other bacteria. Am J Gastroenterol (2010) 105(11):2420–8. doi:10.1038/ajg.2010.281

64. Lerner A, Aminov R, Matthias T. Dysbiosis may trigger autoimmune diseases via inappropriate post-translational modification of host proteins. Front Microbiol (2016) 7:84. doi:10.3389/fmicb.2016.00084

65. Belizário JE, Napolitano M. Human microbiomes and their roles in dysbiosis, common diseases, and novel therapeutic approaches. Front Microbiol (2015) 6:1050. doi:10.3389/fmicb.2015.01050

66. Tieng V, Le Bouguénec C, du Merle L, Bertheau P, Desreumaux P, Janin A, et al. Binding of Escherichia coli adhesin AfaE to CD55 triggers cell-surface expression of the MHC class I-related molecule MICA. Proc Natl Acad Sci U S A (2002) 99(5):2977–82. doi:10.1073/pnas.032668099

67. Borchers MT, Harris NL, Wesselkamper SC, Zhang S, Chen Y, Young L, et al. The NKG2D-activating receptor mediates pulmonary clearance of Pseudomonas aeruginosa. Infect Immun (2006) 74(5):2578–86. doi:10.1128/IAI.74.5.2578-2586.2006

68. Montalban-Arques A, Wurm P, Trajanoski S, Schauer S, Kienesberger S, Halwachs B, et al. Propionibacterium acnes overabundance and natural killer group 2 member D system activation in corpus-dominant lymphocytic gastritis. J Pathol (2016) 240(4):425–36. doi:10.1002/path.4782

69. Nielsen JA, Roberts CA, Lager DJ, Putcha RV, Jain R, Lewin M. Lymphocytic gastritis is not associated with active Helicobacter pylori infection. Helicobacter (2014) 19(5):349–55. doi:10.1111/hel.12139

70. Wu TT, Hamilton SR. Lymphocytic gastritis: association with etiology and topology. Am J Surg Pathol (1999) 23(2):153–8. doi:10.1097/00000478-199902000-00003

71. Polydorides AD. Pathology and differential diagnosis of chronic, noninfectious gastritis. Semin Diagn Pathol (2014) 31(2):114–23. doi:10.1053/j.semdp.2014.02.008

72. Andresen L, Hansen K, Jensen H, Pedersen S, Stougaard P, Hansen H, et al. Propionic acid secreted from propionibacteria induces NKG2D ligand expression on human-activated T lymphocytes and cancer cells. J Immunol (2009) 183(2):897–906. doi:10.4049/jimmunol.0803014

73. Das H, Groh V, Kuijl C, Sugita M, Morita CT, Spies T, et al. MICA engagement by human Vgamma2Vdelta2 T cells enhances their antigen-dependent effector function. Immunity (2001) 15(1):83–93. doi:10.1016/S1074-7613(01)00168-6

74. Kawashima T, Kosaka A, Yan H, Guo Z, Uchiyama R, Fukui R, et al. Double-stranded RNA of intestinal commensal but not pathogenic bacteria triggers production of protective interferon-β. Immunity (2013) 38(6):1187–97. doi:10.1016/j.immuni.2013.02.024

75. Frisan T, Cortes-Bratti X, Chaves-Olarte E, Stenerlöw B, Thelestam M. The Haemophilus ducreyi cytolethal distending toxin induces DNA double-strand breaks and promotes ATM-dependent activation of RhoA. Cell Microbiol (2003) 5(10):695–707. doi:10.1046/j.1462-5822.2003.00311.x

76. Fox JG, Rogers AB, Whary MT, Ge Z, Taylor NS, Xu S, et al. Gastroenteritis in NF-kappaB-deficient mice is produced with wild-type Campylobacter jejuni but not with C. jejuni lacking cytolethal distending toxin despite persistent colonization with both strains. Infect Immun (2004) 72(2):1116–25. doi:10.1128/IAI.72.2.1116-1125.2004

77. Saliba AM, de Assis MC, Nishi R, Raymond B, Marques EA, Lopes UG, et al. Implications of oxidative stress in the cytotoxicity of Pseudomonas aeruginosa ExoU. Microbes Infect (2006) 8(2):450–9. doi:10.1016/j.micinf.2005.07.011

78. Herrtwich L, Nanda I, Evangelou K, Nikolova T, Horn V, Sagar, et al. DNA damage signaling instructs polyploid macrophage fate in granulomas. Cell (2016) 167(5):1264–80.e18. doi:10.1016/j.cell.2016.09.054

79. Hamerman JA, Ogasawara K, Lanier LL. Cutting edge: toll-like receptor signaling in macrophages induces ligands for the NKG2D receptor. J Immunol (2004) 172(4):2001–5. doi:10.4049/jimmunol.172.4.2001

80. Antonangeli F, Soriani A, Cerboni C, Sciumè G, Santoni A. How mucosal epithelia deal with stress: role of NKG2D/NKG2D ligands during inflammation. Front Immunol (2017) 8:1583. doi:10.3389/fimmu.2017.01583

81. Bryant JC, Dabbs RC, Oswalt KL, Brown LR, Rosch JW, Seo KS, et al. Pyruvate oxidase of Streptococcus pneumoniae contributes to pneumolysin release. BMC Microbiol (2016) 16(1):271. doi:10.1186/s12866-016-0881-6

82. Rai P, Parrish M, Tay IJ, Li N, Ackerman S, He F, et al. Streptococcus pneumoniae secretes hydrogen peroxide leading to DNA damage and apoptosis in lung cells. Proc Natl Acad Sci U S A (2015) 112(26):E3421–30. doi:10.1073/pnas.1424144112

83. Li Z, Zhang C, Zhou Z, Zhang J, Tian Z. Small intestinal intraepithelial lymphocytes expressing CD8 and T cell receptor γδ are involved in bacterial clearance during Salmonella enterica serovar Typhimurium infection. Infect Immun (2012) 80(2):565–74. doi:10.1128/IAI.05078-11

84. Buc E, Dubois D, Sauvanet P, Raisch J, Delmas J, Darfeuille-Michaud A, et al. High prevalence of mucosa-associated E. coli producing cyclomodulin and genotoxin in colon cancer. PLoS One (2013) 8(2):e56964. doi:10.1371/journal.pone.0056964

85. Nougayrède JP, Homburg S, Taieb F, Boury M, Brzuszkiewicz E, Gottschalk G, et al. Escherichia coli induces DNA double-strand breaks in eukaryotic cells. Science (2006) 313(5788):848–51. doi:10.1126/science.1127059

86. Putze J, Hennequin C, Nougayrède JP, Zhang W, Homburg S, Karch H, et al. Genetic structure and distribution of the colibactin genomic island among members of the family Enterobacteriaceae. Infect Immun (2009) 77(11):4696–703. doi:10.1128/IAI.00522-09

87. Crnigoj M, Podlesek Z, Budič M, Zgur-Bertok D. The Escherichia coli uropathogenic-specific-protein-associated immunity protein 3 (Imu3) has nucleic acid-binding activity. BMC Microbiol (2014) 14:16. doi:10.1186/1471-2180-14-16

88. Zaw MT, Yamasaki E, Yamamoto S, Nair GB, Kawamoto K, Kurazono H. Uropathogenic specific protein gene, highly distributed in extraintestinal uropathogenic Escherichia coli, encodes a new member of H-N-H nuclease superfamily. Gut Pathog (2013) 5(1):13. doi:10.1186/1757-4749-5-13

89. Nipič D, Podlesek Z, Budič M, Črnigoj M, Žgur-Bertok D. Escherichia coli uropathogenic-specific protein, Usp, is a bacteriocin-like genotoxin. J Infect Dis (2013) 208(10):1545–52. doi:10.1093/infdis/jit480

90. Lee K, Jeong JE, Kim IH, Kim KS, Ju BG. Cyclo(phenylalanine-proline) induces DNA damage in mammalian cells via reactive oxygen species. J Cell Mol Med (2015) 19(12):2851–64. doi:10.1111/jcmm.12678

91. Rai P, He F, Kwang J, Engelward BP, Chow VT. Pneumococcal pneumolysin induces DNA damage and cell cycle arrest. Sci Rep (2016) 6:22972. doi:10.1038/srep22972

92. Kidane D, Chae WJ, Czochor J, Eckert KA, Glazer PM, Bothwell AL, et al. Interplay between DNA repair and inflammation, and the link to cancer. Crit Rev Biochem Mol Biol (2014) 49(2):116–39. doi:10.3109/10409238.2013.875514

93. Dutilh BE, Backus L, van Hijum SA, Tjalsma H. Screening metatranscriptomes for toxin genes as functional drivers of human colorectal cancer. Best Pract Res Clin Gastroenterol (2013) 27(1):85–99. doi:10.1016/j.bpg.2013.03.008

94. Bhatt AP, Redinbo MR, Bultman SJ. The role of the microbiome in cancer development and therapy. CA Cancer J Clin (2017) 67(4):326–44. doi:10.3322/caac.21398

95. Wroblewski LE, Peek RM, Coburn LA. The role of the microbiome in gastrointestinal cancer. Gastroenterol Clin North Am (2016) 45(3):543–56. doi:10.1016/j.gtc.2016.04.010

96. Chumduri C, Gurumurthy RK, Zadora PK, Mi Y, Meyer TF. Chlamydia infection promotes host DNA damage and proliferation but impairs the DNA damage response. Cell Host Microbe (2013) 13(6):746–58. doi:10.1016/j.chom.2013.05.010

97. Guerra N, Pestal K, Juarez T, Beck J, Tkach K, Wang L, et al. A selective role of NKG2D in inflammatory and autoimmune diseases. Clin Immunol (2013) 149(3):432–9. doi:10.1016/j.clim.2013.09.003

98. Chen F, Evans A, Pham J, Plosky B. Cellular stress responses: a balancing act. Mol Cell (2010) 40(2):175. doi:10.1016/j.molcel.2010.10.008

99. Nakad R, Schumacher B. DNA damage response and immune defense: links and mechanisms. Front Genet (2016) 7:147. doi:10.3389/fgene.2016.00147

100. Yin X, Lu X, Xiuwen Z, Min Z, Xiao R, Mao Z, et al. Role of NKG2D in cytokine-induced killer cells against lung cancer. Oncol Lett (2017) 13(5):3139–43. doi:10.3892/ol.2017.5800

101. Hayashi T, Imai K, Morishita Y, Hayashi I, Kusunoki Y, Nakachi K. Identification of the NKG2D haplotypes associated with natural cytotoxic activity of peripheral blood lymphocytes and cancer immunosurveillance. Cancer Res (2006) 66(1):563–70. doi:10.1158/0008-5472.CAN-05-2776

102. Imai K, Matsuyama S, Miyake S, Suga K, Nakachi K. Natural cytotoxic activity of peripheral-blood lymphocytes and cancer incidence: an 11-year follow-up study of a general population. Lancet (2000) 356(9244):1795–9. doi:10.1016/S0140-6736(00)03231-1

103. Hall AB, Tolonen AC, Xavier RJ. Human genetic variation and the gut microbiome in disease. Nat Rev Genet (2017) 18(11):690–9. doi:10.1038/nrg.2017.63

104. Castanys-Muñoz E, Martin MJ, Vazquez E. Building a beneficial microbiome from birth. Adv Nutr (2016) 7(2):323–30. doi:10.3945/an.115.010694

105. Ruff WE, Kriegel MA. Autoimmune host-microbiota interactions at barrier sites and beyond. Trends Mol Med (2015) 21(4):233–44. doi:10.1016/j.molmed.2015.02.006

106. Blaser MJ. The microbiome revolution. J Clin Invest (2014) 124(10):4162–5. doi:10.1172/JCI78366

107. Neemann K, Eichele DD, Smith PW, Bociek R, Akhtari M, Freifeld A. Fecal microbiota transplantation for fulminant Clostridium difficile infection in an allogeneic stem cell transplant patient. Transpl Infect Dis (2012) 14(6):E161–5. doi:10.1111/tid.12017

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Espinoza and Minami. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

REVIEW
published: 05 March 2018
doi: 10.3389/fimmu.2018.00422

Regulation of Cytokine Production by the Unfolded Protein Response; Implications for Infection and Autoimmunity

Judith A. Smith1,2*

1 Department of Pediatrics, University of Wisconsin-Madison School of Medicine and Public Health, Madison, WI, United States

2 Department of Medical Microbiology and Immunology, University of Wisconsin-Madison School of Medicine and Public Health, Madison, WI, United States

OPEN ACCESS

Edited by:

Renata Meirelles Pereira, Universidade Federal do Rio de Janeiro, Brazil

Reviewed by:

Marco A. Cassatella, University of Verona, Italy
Robert Adam Harris, Karolinska Institute (KI), Sweden

*Correspondence:

Judith A. Smith
jsmith27@wisc.edu

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 21 December 2017
Accepted: 16 February 2018
Published: 05 March 2018

Citation:

Smith JA (2018) Regulation of Cytokine Production by the Unfolded Protein Response; Implications for Infection and Autoimmunity. Front. Immunol. 9:422. doi: 10.3389/fimmu.2018.00422

Protein folding in the endoplasmic reticulum (ER) is an essential cell function. To safeguard this process in the face of environmental threats and internal stressors, cells mount an evolutionarily conserved response known as the unfolded protein response (UPR). Invading pathogens induce cellular stress that impacts protein folding, thus the UPR is well situated to sense danger and contribute to immune responses. Cytokines (inflammatory cytokines and interferons) critically mediate host defense against pathogens, but when aberrantly produced, may also drive pathologic inflammation. The UPR influences cytokine production on multiple levels, from stimulation of pattern recognition receptors, to modulation of inflammatory signaling pathways, and the regulation of cytokine transcription factors. This review will focus on the mechanisms underlying cytokine regulation by the UPR, and the repercussions of this relationship for infection and autoimmune/autoinflammatory diseases. Interrogation of viral and bacterial infections has revealed increasing numbers of examples where pathogens induce or modulate the UPR and implicated UPR-modulated cytokines in host response. The flip side of this coin, the UPR/ER stress responses have been increasingly recognized in a variety of autoimmune and inflammatory diseases. Examples include monogenic disorders of ER function, diseases linked to misfolding protein (HLA-B27 and spondyloarthritis), diseases directly implicating UPR and autophagy genes (inflammatory bowel disease), and autoimmune diseases targeting highly secretory cells (e.g., diabetes). Given the burgeoning interest in pharmacologically targeting the UPR, greater discernment is needed regarding how the UPR regulates cytokine production during specific infections and autoimmune processes, and the relative place of this interaction in pathogenesis.

Keywords: unfolded protein response, endoplasmic reticulum stress, infection, virus, bacteria, autoimmunity, cytokine regulation, autoinflammatory disease

INTRODUCTION: IMMUNE SENSING OF DANGER AND ENDOPLASMIC RETICULUM (ER) STRESS

How does the immune system sense pathogenic threats and respond appropriately? Cells in the immune system “see” the environment in little snippets: adaptive immune cells such as T cells bear surface receptors triggered by major histocompatibility complexes (MHC) loaded with peptides 8–20 amino acids in length (1). Even within these short stretches, the T cell receptor may physically interact with only five amino acids (2, 3). Antibodies, constituting the B cell receptors, also recognize similarly small molecules, averaging 18–19 contact residues (up to 5 contiguous) (4). Innate immune cells, the first responders on the scene of infection, including neutrophils, macrophages, and dendritic cells, express pathogen-sensing receptors on their surfaces, inside their endosomes and cytosol collectively referred to as pattern recognition receptors (PRRs). These PRRs recognize conserved molecular arrays on pathogens, or pathogen-associated molecular patterns (PAMPs) such as bacterial lipopolysaccharide (LPS), flagellin and lipoproteins, β-glucans and mannans on yeast, and nucleic acids from viruses. The nucleic acid sensors detect types of nucleic acids that are not normally produced (e.g., dsRNA) or located in unusual settings, such as dsDNA in the cytosol or single stranded RNA in endosomes. Classes of PRRs include the toll-like receptors (TLRs), C-type lectin receptors, nucleotide-binding domain and leucine-rich repeat containing receptors (NLRs), retinoic acid inducible gene I (RIG-I) family helicases, and other cytosolic nucleic acid sensors (5–8).

When the outside world is observed in small pieces, the specificity of immune receptors becomes problematic. For instance, pathogens may express peptides with identical or functionally analogous amino acids stretches as endogenous peptides, a phenomenon referred to as “molecular mimicry” (2, 9). The classic example is the antibody cross-reactivity between streptococcal N-acetyl-beta-D-glucosamine and the cardiac myosin protein (10). This specificity issue, complicating the discrimination of self and infectious non-self, led to the “danger theory” put forth by Polly Matzinger in 1994, and then later refined over the years, that the immune system responds to challenges in accordance with contextual clues from damaged tissues (11–13). These damage-associated signals have been termed “danger-associated molecule patterns” (DAMPs). Different types of DAMPs have been reviewed recently in Ref. (14). When tissue is damaged, and cells destroyed by necrosis rather than apoptosis, specific molecules are released into the surrounding milieu. Examples of released products include dramatic increases in extracellular ATP, extracellular nucleic acids such as double-stranded DNA, mitochondrial DNA, chaperones such as high-mobility group box 1 (HMGB1) and heat shock proteins, interleukins IL-1α and IL-33, and uric acid (15, 16). Even in the absence of actual cellular destruction, infection or stress-triggered calcium signaling, and the generation of reactive oxygen species (ROS) may be considered DAMPs. During infections, the generation of multiple DAMPs provides the context to signal significant organismal insult.

The “danger” hypothesis was initially conceived to address issues with adaptive immune (T and B cell) self-non-self-discrimination. However, this same conceptual requirement for damage that provides context for dendritic cell activation and T cell stimulation may also help with several other specificity issues in innate immunity. Consider the microbiome: humans are widely covered on external and internal surfaces with trillions of microbes that constitute our natural microflora. Microbial-associated molecular patterns (MAMPs) also stimulate PRRs. For instance, theoretically, the same TLR4 that recognizes LPS on an invading pathogen could also be triggered by gut gram-negative bacteria. However, in the absence of tissue damage or stress, the healthy steady-state microbiome does not normally trigger inflammatory responses.

Many damage-generated endogenous products, such as extracellular matrix proteins, also stimulate PRRs (17). Indeed, the same PRRs poised to recognize PAMPs/MAMPs do “double duty” and respond to DAMPs, a testament to natural efficiency and repurposing (14). Alternatively, it has been suggested that pathogens have evolved to take advantage of PRRs evolutionarily aimed at wound repair (12). As an example of endogenous product recognition, the nucleic acids released by dying cells that are taken up into endosomes stimulate endosomal TLRs. The same non-specificity inherent in TLR4 that enables recognition of a broad variety of LPS structures may also allow TLR4 to respond to endogenous products such as fibrinogen or HMGB1 (18). The dual recognition of endogenous products and pathogens by the same receptors again poses a problem of specificity, as non-infectious damage (that releases DAMPs) may not merit an anti-pathogen response. How does the immune system determine whether to mount a wound healing response or an inflammatory response? Is there a titration by numbers or types of DAMPs (and PAMPs) and does this discrimination occasionally fail? Even in the absence of pathogens, “sterile” damage may liberate significant endogenous ligands for PRRs. One example of an over-exuberant inflammatory response in the face of sterile damage is the post-traumatic inflammatory response syndrome that occurs in the absence of inciting infection (16). Aberrant recognition of endogenous products may also drive non-resolving wound responses that lead to fibrosis (19).

Vance et al. have proposed that pathogenic organisms provide extra contextual clues that alert the immune system (20). The immune system recognizes certain bacterial products produced only by living (rather than dead), invasive pathogenic bacteria, so-called “vita PAMPs”: for instance, live bacteria produce cyclic-di-nucleotides second messengers that activate the host cytosolic stimulator of interferon gene (STING) (8, 21). Access to the cytosol may be the factor that provides the key information. As an example, the lysteriolysin O that enables Listeria release into the cytosol is required for immunogenicity (20). Other bacterial pathogens contain secretion systems that provide a conduit between vacuoles and host cytosol. Release of products via this route (e.g., flagellin) may then trigger cytosolic inflammasome sensors (22, 23). Disruption of the cytoskeleton may also be directly sensed by the host cell. The mechanisms by which this occurs remain unclear, but may involve co-localization of PRRs (NOD proteins and inflammasome components) with the actin cytoskeleton (24, 25).

Disruption of fundamental cellular processes such as protein production, may also contribute to immune calibration, titrating up the threat level either appropriately, as in the case of infection, or inappropriately in autoimmunity. All cells must make protein to survive. Secreted and transmembrane proteins are manufactured in the ER. Amazingly, the ER accomplishes protein folding in a very crowded environment, estimated at 100 mg/ml, a concentration that could theoretically promote aggregation (26). The ER is also the site of sterol and phospholipid synthesis and the major cellular store for calcium. Indeed, many of the protein folding chaperones, including the carbohydrate-binding calnexin and calreticulin, immunoglobulin heavy chain binding protein (BiP/Grp78), and protein disulfide isomerases require high concentrations of calcium for their function (27). The formation of intermolecular and intramolecular disulfide bonds during protein folding generates ROS. Thus, to maintain redox equilibrium, the ER contains buffering anti-oxidant enzymes. Related to the exigencies of the folding process, a broad variety of environmental stressors may adversely impact protein folding, such as decreased glucose or amino acids, hypoxia, decreases in ER calcium, excessive reactive oxygen radicals, increased demands in protein production, as well as the presence of misfolding proteins. To safeguard protein production and ensure quality control, ER-stress triggers the activation of several biochemical pathways collectively referred to as the unfolded protein response (UPR). The UPR restores proteostasis equilibrium by increasing capacity (ER size and chaperone production) as well as decreasing protein client load, through translational inhibition and the process of ER-associated peptide degradation (ERAD). If ER stress becomes irremediable (excessively severe or prolonged), the UPR initiates apoptosis. One could envision how infections result in the multiple ER stresses listed above: viral infections dramatically increase protein production; bacteria consume nutrient resources and stimulate oxygen radical production. Because of the universal need for protein production, and the sensitivity to a wide variety of environmental or internal stressors, the UPR is well poised to sense pathogenic danger and transduce the stress signal into a heightened immune response (Figure 1).

FIGURE 1 | Amplification of pathogen immune responses via endoplasmic reticulum (ER) stress and the unfolded protein response (UPR). Pathogens cause tissue damage, intracellular host stress, and stimulate pattern recognition receptors (PRRs) which then induce cytokines. Multiple pathogen-triggered cellular insults cause stress in the ER that impacts protein folding and thus induces the UPR. The UPR, PRR activation, and cytokine production intersects on multiple levels (see main text and Figure 3), with interactions going in both directions (blue two-sided arrows). This amplification mechanism generates an immune response commensurate with the degree of pathogenic threat.

The UPR plays a physiologic role by enabling the function of highly secretory cells such as hepatocytes, plasma cells, and acinar or islet pancreatic cells. For example, mice deficient in key UPR components die early of pancreatic insufficiency and diabetes (28, 29). The UPR also supports the development of specific cells in the immune system. Even before B cells ramp up antibody production to become plasma cells, the UPR is engaged by the plasma cell differentiation program (30–32). Similarly, optimal development and survival of dendritic cells, and differentiation of eosinophils requires the UPR (33, 34). Not only does the UPR support the development of specific immune cells, but it also globally shapes the immune responses in many cell types (32, 35).

Over the past 10–15 years, it has become apparent that one way in which the UPR tunes immune responses is through the modulation of cytokine production (35). Cells of the immune system communicate via cytokines, which are soluble secreted proteins encompassing the families of interleukins, interferons (IFNs), and tumor necrosis factor (TNF) family members among other mediators (36). Both the magnitude and types of cytokines produced program the immune response to respond appropriately to different types of threats. For instance, type I IFN-α/β induce hundreds of target genes aimed at containing and eliminating viral invasion. Cytokines such as TNF-α, IFN-γ, and IL-6 promote inflammatory innate responses that enhance anti-bacterial activities. IL-4, IL-5, and IL-13 heighten anti-parasitic immunity. On the other hand, IL-10 and transforming growth factor β (TGF-β) limit immune destructiveness and collateral damage to the host by toning down innate and adaptive immune responses. Following the sensing of danger signals via PRRs or adaptive immune leukocyte receptors, inflammatory pathways are set in motion that culminate in the activation of cytokine-regulatory transcription factors. Intriguingly, UPR pathways interweave through all levels of cytokine regulation: the UPR impacts the PRRs that sense pathogenic molecules, downstream inflammatory signaling pathways, and ultimately, the activation of cytokine-regulatory transcription factors, such as nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB), activator protein 1 (AP-1), and the interferon regulatory factors (IRFs). This interaction between the UPR and inflammation is a “two-way street”; in certain tissues (e.g., the liver), inflammatory cytokines induce ER stress, setting up the potential for a positive feedback loop (37). The modulation of cytokine production by intracellular stress during infection has implications for how the immune system detects and responds appropriately to pathogens. The drawback to anti-pathogen cytokine augmentation is the potential for inappropriate boosting of immune responses resulting in autoimmunity. Below, the variety of mechanisms linking the UPR with cytokine production and the implications of this interaction for infection and autoimmunity will be addressed.

THE UPR

The metazoan UPR comprises three primary signaling pathways stemming from the activation of ER-stress sensors inositol requiring enzyme 1 (IRE1), activating transcription factor 6 (ATF6), and protein kinase R (PKR) like endoplasmic reticulum kinase (PERK) (27, 38). These three sensors reside in the membrane of the ER, poised to respond to stressors that increase the abundance of unfolded proteins. In their inactive state, the stress sensors associate with the folding chaperone BiP. When ER luminal load of unfolded proteins increases, BiP releases the sensors to preferentially bind hydrophobic patches on misfolded protein, thus resulting in the activation of the three pathways. In addition to this BiP “titration” model, alternative mechanisms of activation have been described: crystallographic resolution of the yeast IRE1 revealed an ER luminal structure that forms an MHC-like peptide-binding groove upon dimerization, thus potentially allowing direct sensing of unfolded peptides (39). The mammalian pocket is too narrow to accommodate peptides, but may undergo a conformational change upon activation by peptide binding (40, 41). Although the PERK luminal domain has high-structural homology with IRE1, direct peptide binding has not been described for PERK. On the other hand, significant alterations in lipid content of the ER (e.g., increased acyl chain saturation) may also directly activate IRE1 and PERK, independently of their ER luminal domains (42).

Inositol requiring enzyme 1 is the most evolutionarily conserved ER stress sensor, and the only UPR pathway present in single cell organisms such as yeast. In mammals, the IRE1α (ERN1) isoform is ubiquitously expressed, whereas IRE1β (ERN2) is restricted to mucosal epithelial surfaces such as the lung and gut (43, 44). The cytosolic portion of IRE1 contains two functional domains: a kinase domain and an endonuclease domain. Upon sensing unfolded protein, IRE1 dimerizes and auto-trans phosphorylates, a prerequisite for activation. Intriguingly, the IRE1 endonuclease has only one specific mRNA target, known as Hac-1 in yeast and XBP1 in higher eukaryotes. IRE1 cleaves a 26 base pair loop out of the XBP1 mRNA, causing a frame shift mutation that removes a premature stop codon. The “unspliced” XBP1 mRNA encodes a shorter unstable protein with DNA binding domain only, but the longer “spliced” XBP1 mRNA encodes the full length transcription factor with DNA binding and transcriptional transactivating domains (45). XBP1 increases the production of folding chaperones (e.g., ERdj4), components involved in ERAD and increases phospholipid synthesis and ER size (31, 46, 47). By increasing ER capacity and decreasing ER client load, XBP1 is considered a largely “adaptive” pro-life response (48, 49). In addition to splicing XBP1, upon prolonged or severe stress, IRE1 may non-specifically degrade mRNAs in proximity to the ER in a process termed regulated IRE1-dependent decay (RIDD) of mRNA (50, 51). This non-specific endonuclease process is thought to decrease ER protein client load, as many of the degraded mRNAs encode proteins in the secretory pathway. XBP1 splicing and the RIDD functions of IRE1 may be experimentally dissociated, but the precise mechanisms governing the switch between these activities remain elusive (52). Degree of IRE1 oligomerization may regulate RNase substrate preference (53). The IRE1 kinase domain associates with other molecules in a multi-molecular complex referred to as the “UPRosome” (45, 54). Through association with TNF-receptor-associated factor 2 (TRAF2) and apoptosis signal-regulating kinase 1 (ASK1), IRE1 phosphorylates c-Jun N-terminal kinase (JNK), thus linking ER stress with autophagy, apoptosis, and inflammatory signaling (described more below) (55). Intriguingly, IRE1 also associates with the pro-apoptotic B cell lymphoma 2 (Bcl2)-family members Bcl2-antagonist/killer 1 (Bak) and Bcl2-associated X protein (Bax), which, through unknown mechanisms, enhance IRE1 kinase activity (56).

Protein kinase R like endoplasmic reticulum kinase oligomerizes and trans phosphorylates early during the UPR. PERK phosphorylates eukaryotic initiation factor 2α (eIF2α) on serine 51, thus inhibiting the guanine nucleotide-exchange activity of eIF2B required for recycling eIF2α to its GTP-bound form (57). By this mechanism, PERK inhibits ribosomal function and globally diminishes protein translation of capped mRNAs. This decrease in protein production is essential for stress adaptation, in that interference with eIF2α phosphorylation leads to proteotoxicity during ER stress (58). Certain mRNAs with inhibitory upstream short open reading frames such as the mRNA encoding the transcription factor ATF4 are preferentially translated when eIF2α is phosphorylated (59, 60). ATF4 stimulates the production of a pro-apoptotic transcription factor C/EBP homologous protein (CHOP). Together CHOP and ATF4 achieve most of the transcriptional program stemming from PERK activation, which includes the induction of proteins involved in amino acid transport, autophagy, folding chaperones, and redox regulatory proteins in addition to pro-apoptotic molecules (61, 62). ATF4 also initiates relief of the translational blockade through induction of growth arrest and DNA damage-inducible 34 (GADD34). GADD34 forms a complex with, and activates protein phosphatase 1, which dephosphorylates eIF2α (63). Thus, the global translational decrease is transient. Other molecules also impact eIF2α phosphorylation status: as an example of cross-talk between UPR pathways, XBP1 regulates p58IPK which binds and inhibits PERK, thus promoting eIF2α dephosphorylation (46, 64, 65). During the non-stressed state, constitutive repressor of eIF2α phosphorylation maintains eIF2α dephosphorylation (66). Interestingly, other molecules with PERK homology, such as PKR, general control non-derepressible 2 (GCN2), and Heme-regulated eIF2α kinase also phosphorylate eIF2α, in response to dsRNA, amino acid deprivation, and heme deficiency, respectively, thus broadening the scope of stressors utilizing this response pathway. For this reason, the eIF2α pathway has also been referred to as the “Integrated Stress Response” pathway (61, 67). In addition to eIF2α, PERK also phosphorylates nuclear factor erythroid 2 (Nrf2), freeing it from the Kelch-like ECH associated protein 1 inhibitory protein. PERK thus enables Nrf2 nuclear translocation and an increase in anti-oxidant protein production (68).

In the third major UPR pathway, release of BiP from ATF6 uncovers a Golgi localization signal, enabling translocation of ATF6 from ER to Golgi (69). In the Golgi, Site-1 and Site-2 proteases cleave ATF6, liberating the active transcription factor. ATF6 also has two isoforms, ATF6α, and ATF6β. Most of the UPR-related activity is dependent upon ATF6α, but there is some redundancy required for development as deletion of both in mice is embryonic lethal (70). ATF6 binds ER stress element sites by itself or as a heterodimer with XBP1; thus, there is some overlap in function. ATF6 also upregulates XBP1 message, another instance of UPR pathway cross-talk (71). Certain UPR target gene chaperones, such as glucose-regulated protein 94 and BiP itself are primarily ATF6-dependent (70). Besides ATF6, in specific cell types, ER stress regulates Site-1 cleavage of other basic leucine zipper transcription factor proteins [e.g., Cyclic AMP-responsive element-binding protein H (CREBH), old astrocyte specifically induced substance, CREB4] (72). In the liver, CREBH participates in inflammatory responses by activating the production of C-reactive protein and serum amyloid P components of the acute phase response (37). For a basic summary of the three UPR pathways, see Figure 2.

FIGURE 2 | Three pathways of the unfolded protein response (UPR). (1) inositol requiring enzyme 1 (IRE1) pathway (left, green), a dual endonuclease and kinase, binds the chaperone binding protein (BiP) in its monomeric state. On sensing unfolded/misfolded protein IRE1 oligomerizes and auto-trans phosphorylates (red Ps). Activation of the endonuclease specifically splices 26 nucleotides out of the XBP1 mRNA, causing a frameshift mutation that removes a premature stop codon, thus enabling translation of the full length transcription factor. With increased stress, the non-specific endonuclease function cleaves endoplasmic reticulum (ER)-associated mRNAs in a process called regulated IRE1-dependent decay (RIDD). The IRE1 kinase domain associates with other signaling partners that phosphorylate Jun N-terminal kinase (JNK). ERAD, ER-associated degradation. (2) Activating transcription factor 6 (ATF6) pathway (middle, blue): ATF6 release of BiP uncovers a Golgi localization signal (GLS) enabling translocation to the Golgi. There it is cleaved by Site-1 and Site-2 proteases (scissors), liberating the ATF6 transcription factor. (3) Protein kinase R like endoplasmic reticulum kinase (PERK) pathway (right, pink): in the presence of phosphorylated protein, PERK also oligomerizes and transphosphorylates, activating its kinase activity. PERK in turn phosphorylates eIF2α, resulting in transient global translational inhibition apart from a few specific mRNAs such as ATF4. ATF4 promotes transcription of the apoptosis-inducing transcription factor C/EBP homologous protein (CHOP). Cellular processes altered by the UPR pathways and key gene targets that are UPR components are in boxes.

MECHANISMS OF CYTOKINE REGULATION BY ER STRESS

Activation of the UPR is sufficient to induce low levels of inflammatory cytokine production, even in the absence of ostensible infectious stimuli or PRR ligation. In one of the earliest studies to note this phenomenon, a 2005 study by Li et al., free cholesterol loading of macrophages induced ER stress-dependent mitogen-activated protein (MAP) kinase signaling and NF-κB activation, resulting in production of IL-6 and TNF-α (73). Subsequently, studies using classic pharmacologic UPR inducers such as tunicamycin and thapsigargin have also noted low level “sterile” inflammatory cytokine production (74, 75). ER stress induces inflammatory cytokines by modulating inflammatory signaling cascades, activating “canonical” cytokine-regulatory transcription factors, as well as via the actions of the UPR-activated transcription factors themselves.

All three UPR pathways impact the activation of NF-κB. In the quiescent state, NF-κB family members (p50, p52, p65, RelB, and c-Rel) reside in the cytoplasm, bound to inhibitory factor κB (IκB). Upon immune signaling, IκB kinase (IKK) phosphorylates IκB, targeting it for ubiquitination and proteolytic destruction. The degradation of IκB permits NF-κB to translocate into the nucleus where it induces inflammatory cytokines such as IL-6 and TNF-α (76). IRE1 increases basal IKK activity via TRAF2, promoting NF-κB translocation (77–79). IRE1 may also promote NF-κB activation indirectly via regulation of glycogen synthase kinase 3 (80, 81). The IκBα protein has a shorter half-life compared with NF-κB, thus the PERK-dependent global translational shutdown preferentially affects IκB expression levels over NF-κB, leaving NF-κB free to translocate (82). Downstream of PERK, CHOP also enhances NF-κB signaling via transcriptional repression of the negative regulator peroxisome proliferator-activated receptor (83). ATF6 impacts NF-κB activation through a pathway involving mammalian target of rapamycin signaling and protein kinase B (Akt) dephosphorylation (84, 85). Finally, the calcium dysregulation and ROS generated during ER stress may contribute to NF-κB activation, either by enhancing induction of UPR pathways or other mechanisms (86). NF-κB regulates cytokine production in conjunction with other transcription factors such as the AP-1 heterodimer of Fos and Jun family transcription factors. MAP kinases [e.g., p38, extracellular regulated kinase (ERK) and JNK] regulate the activation of AP-1 factors (87). ER stress intersects with MAP kinase signaling in multiple ways [reviewed in Ref. (88)]: IRE1 promotes the activation of AP-1 family members via ASK-1 mediated JNK and p38 phosphorylation (55, 89). ERK phosphorylation during ER stress is also partially IRE1-dependent (90). In bronchial epithelial cells, PERK and ATF6 promote ERK and p38 signaling, and in cholesterol loaded macrophages, CHOP was required for ERK activation (73, 91). P38 positively feeds back on the UPR, phosphorylating CHOP and ATF6, and thus increasing their activities (92–94).

In addition to the classic pro-inflamamtory cytokines, the UPR regulates type I IFN. IFN-β is one of the earliest IFNs produced in response to viral infection and PRR engagement, and by binding the type I IFN receptor (IFNAR) and upregulating IRF7, promotes the production of multiple IFN-α species and induction of the full anti-viral interferon program (95). In the ifnb1 promoter “enhanceosome” region, NF-κB, AP-1, and IRF3 bind cooperatively to initiate transcription (96–98). Like NF-κB, unactuated IRF3 remains cytoplasmic. Upon phosphorylation on multiple serines and threonines, IRF3 dimerizes and translocates into the nucleus where it binds its gene targets (99). IRF3 phosphorylation also enables association with the transcriptional co-activator CREB-binding protein (CBP/p300) (100). ER stress induces IRF3 phosphorylation and nuclear translocation, although the precise mechanisms are not yet clear and may depend upon the type of stress. Calcium disruption (as through the SERCA pump inhibitor thapsigargin) and oxygen glucose deprivation activate IRF3 through a STING-dependent mechanism, whereas agents that disrupt N-linked glycosylation (e.g., tunicamycin) appear to utilize a STING-independent, but Site-1/Site-2 protease (ATF6?)-dependent pathway (101).

In addition to the activation of canonical inflammatory transcription factors and IRFs, the classic UPR transcription factors which orchestrate the UPR bind directly to genetic cytokine-regulatory elements. Through chromatin precipitation analyses, XBP1 was detected at the promoters of the IL-6, and TNF-α encoding genes, a tnf enhancer, as well as an enhancer element downstream of the ifnb1 gene (74, 102). In response to short chain fatty acids, ATF4 (downstream of PERK and the integrated stress response) binds the cAMP response element in the Il6 promoter (103). CHOP binds the IL-23p19 (Il23A) promoter in dendritic cells in response to LPS, ER stress, and most fully to the combination of LPS and ER stress (104). On the other hand, certain UPR-regulated transcription factors such as ATF3 have anti-inflammatory effects, and may play a role in regulating pathogen responses, ischemic preconditioning, and cancer (105–109).

More recently, evidence has suggested that beyond directly regulating transcription factors or cytokine promoters, ER stress also impacts the activation of upstream PRRs. For example, ER stress activates the inflammasome, thus promoting IL-1β production and potentially programmed cell death. IRE1 activation, possibly via RIDD, inhibits a micro-RNA, miR-17, that down-regulates the production of thioredoxin-interacting protein (TXNIP) (110, 111). Thus, ER stress rapidly increases TXNIP expression (111, 112). PERK also increases TXNIP expression via the induction of transcription factors carbohydrate-responsive element-binding protein and ATF5 (112). TXNIP associates with and activates the NLRP3 inflammasome at the mitochondria. NLRP3 in turn, in a caspase-2 and BH3 domain interacting agonist (Bid)-dependent mechanism, causes mitochondrial damage, cytochrome C release, and production of oxygen radicals that further stimulates inflammasome production of IL-1β (113). The IRE1-RIDD function has also been implicated in the generation of small RNAs that trigger RIG-I-dependent NF-κB activation (114). UPR-dependent mitochondrial damage and mitochondrial DNA release may also play a role in the activation of another cytosolic sensor STING: mitochondrial DNA triggers the molecule cGAS, which in turn generates a cGAMP ligand that stimulates STING (115). As noted, certain types of ER stress mobilize STING translocation and STING-dependent IFN production (101). However, the link between ER stress-dependent mitochondrial damage and STING activation remains speculative. ER stress is well poised to initiate mitochondrial ROS-dependent events that activate and amplify innate immune signaling: protein folding is an oxidative process (116). The UPR and ROS trigger calcium release from the ER through activation of the inositol-1,4,5-triphosphate (IP3) receptor and ryanodine receptor ER channels. ER and mitochondria are spatially juxtaposed at the mitochondria-associated ER membranes, where ER IP3R channels are linked via chaperones to mitochondrial voltage-dependent anion channels (117). Increased cytosolic calcium thus triggers ROS release from mitochondria, which induces increased levels of ER stress, resulting in a relentless feed-forward loop (116). Finally, the UPR also interacts with the cytosolic peptidoglycan receptors NOD1 and NOD2 to induce production of the pro-inflammatory cytokine IL-6. Activation of this NOD1/2-dependent pathway by thapsigargin or infection with Brucella abortus was suppressed by the general UPR inhibitor tauroursodeoxycholic acid (TUDCA) and the IRE1 kinase inhibitor KIRA6 (118). The proposed mechanism involves IRE1 kinase activation and recruitment of NOD-interacting proteins TRAF2 and receptor-interacting serine/threonine-protein kinase 2 (119). For a summary highlighting mechanisms at the intersection of UPR and cytokine induction, see Figure 3.

FIGURE 3 | Intersections of endoplasmic reticulum (ER) stress/unfolded protein response (UPR) and immune signaling. ER stress and the UPR impact innate immune signaling and cytokine production on many levels between pathogen-sensing pattern recognition receptors (PRRs) and ultimate cytokine production: (1) PRR activation: ER stress and the UPR activate multiple PRRs (purple) including stimulator of interferon gene (STING), NLRP3, and other inflammasomes via thioredoxin-interacting protein (TXNIP) upregulation and reactive oxygen species (ROS), and the NOD1/2 receptors. Much of this interaction occurs at the mitochondrial-ER interface, where released calcium (Ca2+) and ROS feed into PRR activation. (2) The UPR enhances inflammatory signaling pathways leading to mitogen-activated protein kinase activation [IRE1 shown here, but protein kinase R-like endoplasmic reticulum kinase (PERK) and activating transcription factor 6 (ATF6) also impact ERK and p38 activation], and inhibitory factor κB (IκB) phosphorylation and degradation. (3) Transcription factors: the UPR activates canonical pro-inflammatory and IFN-regulatory transcription factors activator protein 1 (AP-1), nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB), and IRF3. Core UPR generated transcription factors such as XBP1 and C/EBP homologous protein (CHOP) also directly stimulate cytokine production by binding cytokine promoter and enhancer elements.

Sterile ER stress results in relatively low levels of cytokine production, particularly compared with PRR stimulation (74, 120). In the case of IFN-β, this is perhaps surprising, as ER stress activates the critical enhanceosome components NF-κB, AP-1, and IRF3. One possible explanation is that another signal is required (e.g., PRR ligation) for full phosphorylation of IRF3 at multiple sites (101). Alternatively, although multiple UPR pathways activate NF-κB, it may still be at a low level compared with that induced by PRR ligation. Another possibility, extending to other cytokines, is that PRR ligation may be required to generate certain transcriptional co-factors or epigenetic modifiers.

In contrast to situations involving either sterile ER stress or isolated PRR stimulation, subsequent PRR ligation of cells undergoing ER stress has profound consequences for inflammation. Specifically, induction of ER stress has the capacity to render cells exquisitely sensitive to PRR stimulation, resulting in dramatically synergistic production of certain cytokines. This synergism has been demonstrated using pharmacologic UPR inducers, XBP1 overexpression, and misfolding proteins (74, 104, 120–122). Prominently increased cytokines include IL-6, TNF-α, IL-23, and IFN-β. In the cases of IL-6, TNF-α, and IFN-β, synergy appears to be XBP1 dependent but for IL-23, it is CHOP-dependent, consistent with their detected binding to specific cytokine gene regulatory elements (74, 102, 104). ER stress may also enable cells to produce IL-1β in response to TLR4 ligation in a TRIF (TIR domain containing adaptor protein inducing interferon beta)-dependent and caspase 8-dependent, but XBP1 and CHOP independent manner (123). Synergy is not the invariable outcome of PRR stimulation of stressed cells but may depend upon the context. In ischemic preconditioning, which induces ER stress, inflammatory cytokine production is blunted, possibly via ATF3 induction or decreased NF-κB activity (106, 124).

Direct ligation of PRRs on the other hand, in the absence of a specific ER stressor, appears to partially activate UPR signaling pathways and selectively suppress others. Woo et al. reported that TLR3 or TLR4 stimulation suppressed subsequent ER stress-induced ATF4 and CHOP activation (but not upstream PERK or eIF2α phosphorylation) in a TRIF-dependent manner (125). LPS suppression of CHOP limited apoptosis (126). Stimulation of TLR2 and TLR4 activates IRE1 sufficiently to induce XBP1 mRNA splicing and binding of XBP1 to cytokine promoters. Interestingly, in this setting the nominal XBP1 UPR targets genes (e.g., ERdj4) were not transcribed. TLR signaling did not trigger the other two UPR pathways, as assessed by PERK phosphorylation and ATF6 cleavage, and inhibited tunicamycin-dependent upregulation of CHOP and the ATF6 target BiP. Canonical TLR signaling pathways and ROS appear to be involved in TLR-induced XBP1 splicing, as NOX2, TRAF6, and TLR adaptors myeloid differentiation primary response 88 (MyD88) and/or TRIF were all required (74). As another example of partial UPR activation and modification, viral infections that release dsRNA stimulate PKR, eIF2α phosphorylation, and GADD34 induction, in a TRIF-dependent manner. Interestingly, in the setting of virus/dsRNA, GADD34 relieves the translational inhibition of IFN-β and IL-6, but not global translation (127, 128). The basis of this specificity, or the resistance of global translational reversal remains unclear. Likewise, it is not yet understood why TLR4 induced XBP1 would promote the production of cytokines, but not its nominal chaperone targets. This phenomenon of partial UPR signaling and modulation in response to PRR ligation has been termed the “microbial stress response” pathway (129). As a net result, PRR adaptation of the UPR machinery potentially boosts cytokine production while avoiding the apoptotic sequelae of a fully engaged UPR.

One other mode of UPR-cytokine cross-talk occurs between cells rather than within individual cells. Surface translocation of calreticulin in cancer cells due to ER stress enhances immunogenicity and phagocytic uptake by dendritic cells—an immunostimulatory “eat me” signal (130). In a subsequent study, thapsigargin but not tunicamycin treatment of fibroblasts increased surface calreticulin expression and phagocytic uptake by co-incubated dendritic cells, suggesting the type of ER stress may be important. Interestingly, incubation of these thapsigargin-treated fibroblasts with bone marrow cells augmented LPS-induced production of pro-inflammatory cytokines such as IL-1β, IL-12, IL-23, and TNF-α (75).

IMPLICATIONS FOR VIRAL INFECTIONS

Intracellular infections provide a stage where ER stress interacts with signals from multiple PAMPs and DAMPs. The impact of the UPR on host-pathogen interaction has been increasingly recognized in viral, bacterial, and parasitic infections. UPR in the setting of parasitic infections has been reviewed recently and will not be discussed below (131).

The dramatic synergy observed by multiple groups between UPR and PRR signaling in the induction of type I IFN has particularly compelling implications for viral infection where the IFN response forms the capstone of host resistance. Viruses notoriously sabotage IFN production in a variety of ways. Several viruses interfere with the signaling leading to IRF3 activation or association with CBP/p300 (132–134). For instance, Dengue virus infection cleaves STING and also targets its upstream ligand-generator cGAS (135, 136). Other viruses target the type I IFN receptor IFNAR for proteolytic degradation (137). Paramyxovirus V proteins target STAT1 and STAT2 for proteolytic degradation (138). Therefore, one could speculate, that given all the viral obstacles to mounting an effective IFN response, even a partial UPR with XBP1 splicing or GADD34 induction to promote IFN-β transcription and translation might improve the odds.

Viruses induce ER stress through multiple mechanisms: during viral infection, cells dramatically increase protein production to manufacture new progeny virus. Some viruses reorganize the ER to develop replication platforms (e.g., Hepatitis C virus, coronavirus), and disrupt ER-Golgi trafficking (e.g., Picornavirus) (139–142). Viral infection also generate ROS. Beyond the host’s direct response to ER stress, the catalog of viral proteins that induce or manipulate UPR pathways has grown exponentially. One could envisage how the UPR could be both helpful and harmful to viral infection, even aside from any effects on the anti-viral IFN program. On the one hand, adaptive pathways within the UPR could enable host cells to survive the inordinate stress of significantly increased viral protein production by significantly increasing ER capacity. However, both translational inhibition and ERAD could diminish viral protein production. Premature UPR-related apoptosis could also limit viral replication and spread.

In recent reviews, 35 animal viruses and several plant viruses have been reported to provoke ER stress and/or UPR induction (143, 144). Viruses vary greatly in their capacity to both induce and inhibit individual UPR pathways. Multiple RNA viruses (e.g., Dengue virus, Hepatitis C, Coxsackie B3, and SARS coronavirus) and DNA viruses (Ebstein Barr virus, Hepatitis B) induce all three UPR signaling axes (65, 114, 143, 145–150). Several viruses have been reported to induce two UPR axes, for instance IRE1 and PERK (Sindbis) or IRE1 and ATF6 (Influenza A, Chikungunya), whereas some may induce only one arm [e.g., ATF6, lymphocytic choriomeningitis virus (LCMV)] (151–153). Different aspects of the UPR may also prevail at specific times during the viral life cycle (145). The basis for this selective activation is not well understood but may depend upon specific viral factors and intracellular lifestyle.

Viruses have co-evolved multiple mechanisms to manipulate specific UPR pathways to avoid some of the potentially detrimental effects of UPR induction. For instance, several viruses encode GADD34 homologs: the Herpes Simplex Virus 1 product γ_134.5 forms a complex with protein phosphatase 1, which dephosphorylates eIF2α, thus limiting translational inhibition (154). Further, γ_134.5 contributes to viral resistance to IFN-α/β (155). The African swine fever virus DP71L functions similarly, inhibiting induction of ATF4 and CHOP (156). Japanese encephalitis virus induces RIDD to enhance replication, but intriguingly appears resistant to the RNAse activity (157). Herpes Simplex Virus UL41 protein suppresses XBP1 mRNA induction and splicing, possibly to decrease ERAD (158). There are also examples of viruses (e.g., Hepatitis C) that are permissive for XBP1 splicing, but prevent induction of XBP1’s nominal UPR gene targets, which would include ERAD proteins such as ER degradation-enhancing α-mannosidase-like protein EDEM (159). This separation of XBP1 splicing and UPR target induction is reminiscent of the TLR-induced XBP1 disjunction. In these cases, it would be interesting to determine if the “blocked” XBP1s could still synergize in promoting IFN or pro-inflammatory cytokine production.

Modulation of the UPR appears to have varying effects, depending upon the virus and the type of UPR inhibition used. Viruses often induce pathways that enhance their replication. For instance, LCMV induces ATF6 activation, and cells defective in Site-2 protease produce lower titers of infectious virus (153). Likewise, blocking IRE1 with pharmacologic agents inhibits Influenza A replication (160). There are also multiple examples of viruses where the UPR appears to limit replication, suggesting a contribution to host defense. For instance, PERK is required for control of Dengue replication and pharmacologic eIF2α phosphorylators exhibit anti-viral activity (145, 161). Similarly, West Nile virus replicates at much greater titers in the absence of pro-apoptotic CHOP (162). Together, these studies support a general, but not universal concept that the IRE1 and ATF6 pathways are more likely to benefit virus, but the PERK pathway supports host defense. As an example where integrated stress response benefits virus, HIV induced ATF4 directly promotes HIV transcription through its long terminal repeat (163).

Although the UPR limits some viral infections, direct evidence for the role of the UPR in promoting type I IFN or other inflammatory cytokines during viral infection has been limited. It can also be difficult to tease apart cytokine vs. other effects of UPR modulation. For instance, in a Dengue model, induction of the UPR with a BiP inhibitor increased activation of IRF3 and NF-κB. However, it was not clear if these transcription factor effects contributed to anti-Dengue activity (164). There is some evidence viruses induce collateral damaging inflammation via UPR activation. For instance, the Hepatitis B protein HBx induced inflammatory cyclooxygenase 2 via an eIF2α-ATF4 pathway (114). Dengue virus-induced PERK/Nrf2 activation enhanced TNF-α production via increases in c-type lectin domain family 5, member A (CLELC5A), thus exacerbating pathology in a mouse model (165). Regarding IFN, in dendritic cells, XBP1 overexpression enhanced IFN-β production and markedly suppressed Vesicular stomatitis virus replication (122). In murine embryonic fibroblasts, GADD34 was required for dsRNA induced IFN-β and IL-6 production and resistance to Chikungunya virus. In vivo, IFN-dependence upon GADD34 appeared age-specific: adult mice were resistant to Chikungunya. However GADD34−/− neonates produced greatly diminished IFN-β in response to infection and rapidly succumbed (127). These two studies support a role for the UPR or microbial stress response pathways in supporting IFN and anti-viral immunity. However, viruses can also manipulate the UPR to limit IFN production. For instance, vesicular stomatitis virus and hepatitis C virus target IFNAR for proteolytic degradation via a PERK-dependent pathway, and this pathway appeared to enhance viral infection (137). Hepatitis C activation of UPR-autophagy pathways, including induction of CHOP and autophagy protein 5, also limited IFN-β production (166). Overall, the precise role of UPR pathways in supporting or limiting IFN or other cytokine production during viral infection, and the ultimate effect on pathogenesis remain important areas for further investigation.

IMPLICATIONS FOR BACTERIAL INFECTIONS

The study of the UPR in bacterial infections is much younger and less well developed than for viral infection, but the complexity of bacterial lifestyles promises many interesting variations on the interactions between host UPR and immunity. The list of bacteria inducing UPR pathways through their intracellular lifecycles or elaboration of bacterial products is steadily growing. Regarding bacterial products, Subtilase toxin, produced by Shiga endotoxic Escherichia coli, cleaves BiP, thus initiating all three arms of the UPR (167, 168). Interestingly, this UPR activation may either promote apoptosis, or dampen NF-κB responses and endotoxic pathology at subcytotoxic doses (169, 170). Listeriolysin O, produced by Listeria monocytogenes, also induces all three axes of the UPR (171). The current mechanism is unknown, but may involve depletion of intracellular calcium stores (172). Cholera toxin selectively binds IRE1, activating its RIDD activity (173). Brucella abortus secretes a factor VceC via its type IV secretion system that binds BiP and selectively induced IRE1activation (174). Interestingly, when ectopically expressed, several other Brucella type IV secretion system substrates also appear to accumulate in the ER, inhibit protein secretion and induce varying amounts of ER stress (175).

Beyond secretion of ER/UPR modifying factors, several pathogens form intimate spatial relationships with the ER during their intracellular lifecycle. For instance, Legionella and Brucella traffic in the endosomal pathway, preventing full phagosome-lysosome fusion, and establish replicative vacuoles within ER-derived compartments (176, 177). Chlamydia containing inclusion compartments also contact the ER (178). Intriguingly, reports of the interactions of these three ER-localized pathogens with ER stress responses have varied. One group reported that persistent (non-productive) Chlamydia infection induced transient BiP upregulation and eIF2α phosphorylation but not ATF6 cleavage or XBP1 splicing (179). However, in another study, Chlamydia stimulated “robust” IRE1 activation and XBP1 splicing, and induced CHOP in a GCN2-dependent manner (180). Legionella actively inhibited XBP1 splicing via bacterial translation elongation inhibitors (181). Brucella infection induces pronounced activation of UPR pathways. Within 24–48 h of infection, Brucella causes massive restructuring of the ER marked by condensation, fragmentation, and vacuolization (176, 182). This restructuring is mediated, at least in part via a microtubule stabilizing factor produced by Brucella, TcpB, which also has UPR-inducing properties (182, 183). Although the UPR induced by B. melitensis involves all three axes, with prominent CHOP induction, the B. abortus triggered UPR appears more targeted in scope (174, 182). Interestingly, the UPR appears to benefit B. melitensis replication in that targeting IRE1 with RNAi in a Drosophila S2 cell line or in IRE1−/− fibroblasts, or treatment of macrophages with the general UPR inhibitor TUDCA all diminished replication (182, 184). The UPR may help the host cell to survive the tremendous structural insult to its protein producing factory through its adaptive pro-survival ER stress coping mechanisms. The UPR also induces autophagy through multiple pathways, thus providing increased nutrients to “feed” the bacteria (185). Autophagy may also promote cell–cell spread (186). In contrast to B. melitensis, B. abortus replication was not affected by TUDCA (118). The basis for this species difference in UPR induction and consequence is not clear.

Several lines of evidence support a role for the UPR in innate immune sensing of bacterial infection and control of infection or collateral inflammation. The cytokine response to Chlamydia involves multiple ER stress pathways: CHOP critically contributed to Chlamydia-induced IL-23 production (104). Chlamydia also induced PKR-dependent IFN-β through a mechanism requiring TLR4 and IRE1 RNase activity. Interestingly, this TLR4 activity may limit CHOP induction, stressing the importance of the multiple innate immune and ER stress inputs that impact cytokine production during infection (180). XBP1 deficiency significantly decreased TLR2-dependent TNF-α and IL-6 responses to Francisella in vitro. Furthermore, XBP1 conditional knockout mice infected with F. tularensis exhibited greater organ disease burden (74). UPR augmentation of cytokine production may be particularly important in Brucella infection because of the unusually low endotoxicity of its LPS, as well as the sabotage of TLR signaling by TIR-domain analog-containing bacterial factors (e.g., TcpB) (187, 188). In B. abortus infected macrophages, VceC and IRE1 was required for optimal IL-6 responses in vitro (174). In a subsequent study, this same group implicated the NOD1/NOD2 PRRs downstream of ER stress in Brucella-stimulated IL-6 production (118). In vivo, the VceC mutant stimulated much less splenic IL-6 production, despite similar bacterial burden. Furthermore, in an inflammatory abortion model, the VceC mutant, TUDCA treatment, or NOD1/2 deficiency all decreased placentitis, placenta IL-6 expression, and increased mouse pup survival (118). Thus, ultimately, the net benefit of UPR-supported inflammatory responses during infection may represent a balance between augmented host sensing of infection, containment, and collateral inflammatory damage.

IMPLICATIONS FOR AUTOIMMUNITY AND AUTOINFLAMMATORY DISEASES

The UPR potentially enhances host responses to invading pathogens by boosting PRR signals. However, the down side to immune augmentation is the capacity to cause pathologic cytokine production, even in the absence of infection. Aberrant cytokine production plays a critical role in fomenting inflammatory disease, as attested to by the tremendous clinical utility of cytokine blocking antibody therapies. Cytokine targeting therapy has been remarkably effective in both autoimmune disease [e.g., rheumatoid arthritis (RA)], where “self” autoantigens play key roles in disease pathogenesis, as well as autoinflammatory diseases, which are driven primarily by abnormalities in cytokine production [e.g., TNF-receptor-associated periodic fever syndrome (TRAPs) and cryopyrinopathies] (189). Some of the diseases discussed below [inflammatory bowel disease (IBD), spondyloarthritis (SpA)], although not a result of a monogenic cytokine dysregulation, also have prominent autoinflammatory features. For instance, in mouse models, exogenous expression of IL-23 (generated by genetic minicircle infusion) reproduces many of the clinical features of SpA, including sacroiliitis, enthesitis, and inflammatory skin disease (190). General overexpression of human TNF in mice phenocopies RA, whereas a stabilized TNF-α in mice (TNFΔARE) produces aggressive widespread (polyarticular) joint disease and Crohn’s like IBD, with arthritis occurring independently of T or B cells (191–193). In humans, genome wide association studies in polygenic autoimmune and autoinflammatory disorders have identified numerous associations with polymorphisms in cytokine or cytokine-regulatory genes (194–196). Thus, given the centrality of cytokine production in these inflammatory diseases, as indicated by clinical data, mouse models, and genetic studies, ER stress could theoretically have a major impact on disease pathogenesis. Indeed, the UPR has been implicated in an increasingly greater number of inflammatory diseases. A few themes will be highlighted below, including linkage of UPR components to polygenic autoimmune diseases, diseases of altered ER function, misfolding protein diseases, and autoimmunity in highly secretory cells.

Inflammatory bowel disease results from the aberrant, over-exuberant response to endogenous gut flora (197). Further, the association with NOD2, the first major gene linked to IBD, implicates innate immunity in the abnormal gut inflammation (198, 199). IBD is also one of the first polygenic disease to be genetically linked to UPR components (200). Specifically, a hypomorphic allele of XBP1 increases risk of developing IBD. XBP1ΔIEX mice, lacking XP1 in intestinal epithelial cells, develop spontaneous mild enteritis and are more susceptible to Dextran sodium sulfate-induced colitis (an experimental IBD model) (200). Autophagy or the process of “self-eating” interacts with the UPR on multiple levels, in that the UPR induces autophagic pathways and autophagy may limit the UPR (185). Interestingly, in the case of IBD, ATG16L1, encoding a core autophagy effector, also associates with IBD in human genetic screens, and ATG16L1ΔIEX mice develop spontaneous Crohn’s like ileitis (201–203). ATG16L1ΔIEC and XBP1ΔIEC double knockout mice develop very severe colitis, suggesting a functional synergy between defective autophagy and UPR in predisposing to colitis (202). Part of the role of the UPR in colitis appears to be in support of gut-protective secretion: XBP1 supports Paneth cell development and function (200). However, there is also a more direct inflammatory consequence of XBP1 deletion. Through an unclear mechanism, XBP1 deficiency results in hyperactivation of IRE1. ATG16L1 deficiency in gut intraepithelial cells independently results in increased IRE1, related to defective IRE1 clearance by autophagy (203). Increased IRE1 kinase activity induces augmented NF-κB activation and thus pro-inflammatory cytokine production. Indeed, gut deficiency of IRE1 or TNFR1 relieves the XBP1ΔIEC inflammatory phenotype (202). Mucin production maintains the barrier between gut flora and epithelial cells; “Winnie” and “Eyeore” mice expressing misfolding Mucin2 have a deficiency in mucin production, exhibit increased gut permeability and strong UPR induction, and develop gut inflammation (innate immune and Th17) (204, 205). Two other ER genes have also been linked to IBD in humans, anterior gradient 2 (AGR2), encoding a protein disulfide isomerase, and Orosomucoid-like 3 (ORMDL3), which regulates ER calcium and induces UPR pathways (206–209). Agr2−/− mice develop severe spontaneous ileocolitis associated with defective mucin folding and ER stress (210). At this time, it is not clear how ORMDL3 regulates gut inflammation. Together these studies suggest that the UPR-autophagy interaction regulates the extent of inflammatory responses to gut flora and that defects in this axis predispose to IBD.

More recently, protein mishandling/altered trafficking at the ER has been identified as a monogenic cause of an autoimmune syndrome. Patients with mutations in COPA develop inflammatory interstitial lung disease with pulmonary hemorrhages, arthritis, autoantibody production, and renal disease (211, 212). COPA encodes a component of the COP I complex responsible for Golgi-ER retrograde transit and the syndromic mutations in this gene appear to disturb protein cargo recognition. COPA mutant cells display signs of ER stress with increased BiP, ATF4, and CHOP expression, although the precise mechanism linking this defect in retrograde transit with ER stress are not yet clear. The ER stress correlates with increased expression of IL-1β, IL-6, and IL-23 in immortalized B cells from these subjects, previously noted ER stress augmented cytokines (74, 104, 123). Perhaps as a result of increases in these specific cytokines, patients also exhibit an expansion of T helper 17 CD4+ T cells, a cell type implicated in autoimmunity (213). Interestingly, a number of these patients also have evidence for a type I IFN-regulated gene signature in their peripheral blood (214).

TNFR1-associated periodic fever syndrome is an autosomal dominant monogenic autoinflammatory disease that manifests with episodes of prolonged high fever, rash, abdominal pain, peri-orbital edema, and myalgia (189). Defective surface shedding of TNF receptors (and thus prolonged TNF signaling) was initially postulated as a pathogenic mechanism; however, several studies have shown that TNFR1-associated mutants form oligomers and aggregates in the ER, resulting in ER retention (215). Interestingly, these mutations were also associated with defective autophagy, and increasing autophagy with geldanamycin decreased IL-β production (216). Patient peripheral blood mononuclear cells (PBMC) expressed elevated levels of phosphor PERK and spliced XBP1 mRNA, but not increases in other UPR-associated transcripts. Their monocytes had increased ROS as well (217). Transfection of cells with mutant TNFR1 did not induce BiP or CHOP expression, suggesting the UPR is not a direct contributor (215). However, cells from patients expressing mutant TNFR1 displayed increased mitochondrial ROS production, which promotes inflammatory cytokine production (218). Thus, ER stress may link misfolding TNFR1 to inflammation via ROS.

Spondyloarthritis encompasses a group of genetically and pathologically related inflammatory diseases which manifest with axial (spinal) arthritis, enthesitis, uveitis, gut inflammation, and psoriasis (219). SpA is highly linked to an MHC protein HLA-B27 that misfolds during biogenesis: in patients with the prototypic SpA, ankylosing spondylitis, 80–90% of subjects are HLA-B27 positive vs 6% of the United States population (220, 221). Although ankylosing spondylitis is a polygenic disease, the presence of HLA-B27 accounts for the preponderance (67%) of the currently identified heritability, conferring an odds ratio of >50 (222, 223). This misfolding propensity and prolonged association with BiP in the ER results from specific amino acids in its peptide-binding B pocket and unpaired cysteines (224–226). The subtypes of B27 with differential disease association also exhibit variance in biochemical features including thermos-stability, folding rates, and intracellular aggregation (227–229).

Transgenic HLA-B27 expression alone is sufficient to drive an inflammatory disease analogous to SpA in susceptible rat strains, although disease requires very high-transgene numbers (230, 231). Interestingly, disease does not occur in germ-free rats, but requires microbiota (232). Although there are many reasons why this might be the case, in light of the current discussion, one could speculate that microbiota may also be required to provide PRR signals that synergize with ER stress. Interestingly, CD8+ T cells are dispensable for disease development in rats, suggesting another property of HLA-B27 besides its antigen-presenting capacity may be important in driving disease (233). Bone marrow-derived macrophages from HLA-B27, but not HLA-B7 transgenic rats showed evidence for a UPR gene signature, particularly when class I MHC was acutely upregulated by cytokines such as TNF-α and/or IFN (234). These ER stressed macrophages responded to TLR agonists with greatly increased type I IFN in vitro (120). Interestingly, the bone marrow macrophages from the diseased B27 transgenic animals displaying a UPR gene signature also exhibited a very prominent IFN signature (234). However, the role of IFN, if any, in SpA has not been established. The inflamed colons in diseased animals exhibited upregulation of UPR target genes, along with increased IL-23, IL-17, IFN-γ expression, and expansion of Th17 cells (121). In an effort to more directly address the role of the UPR in these rats, one study interbred HLA-B27 transgenic rats with human beta-2 microglobulin overexpressing rats to stabilize and aid in HLA-B27 folding. This breeding did indeed reduce misfolding in Con-A stimulated splenocytes, although macrophages and tissue UPR were not assessed (235, 236). Surprisingly, these animals developed more severe arthritis, without changes to their colitis. This study suggests the role of HLA-B27-linked UPR may be discordant in the joints and the gut during SpA and raises further questions regarding HLA-B27 misfolding, UPR, and disease pathogenesis.

Although HLA-B27 can induce a UPR, it is not clear this property is the culprit in human subjects expressing at most two copies of the MHC molecule. HLA-B27 also forms surface dimers that can stimulate IL-17 producing cells, providing an alternative mechanism (237). Studies examining UPR in human subjects have yielded inconsistent results: increased BiP has been observed in knee fluid macrophages from ankylosing spondylitis patients (238). PBMC monocytes have been reported to express higher levels of UPR target genes, although other groups have reported a lack of UPR in PBMC and synovium (239, 240). Blood-derived macrophages from ankylosing spondylitis patients produce increased IL-23 in response to LPS without increased UPR target gene expression (241). Misfolded HLA-B27 has been detected in gut biopsies from SpA patients, but associated with activation of autophagy rather than UPR (242). Also, not all SpA (or even ankylosing spondylitis) patients are HLA-B27 positive. Interestingly, in a mouse model with altered autoreactive T cell repertoire, curdlan or zymosan treatment induces an SpA-like disease with enteritis, sacroiliitis, enthesitis, and psoriatic skin inflammation (243, 244). This disease model is also cytokine (IL-23 in particular) and gut microbiome-dependent (245). Interestingly, the inflamed colons from these animals showed evidence of UPR target gene induction (243). Thus, misfolding HLA is not an absolute prerequisite for UPR induction in SpA pathogenesis. These observations also raise the possibility that the UPR may be an integral part of the developing inflammatory process and not just the inciting event.

Myositis is another rheumatologic entity linking aberrant MHC, a type I IFN signature and the UPR. This group of diseases includes dermatomyositis, inclusion body myositis and dermatomyositis. Muscle biopsies from these patients exhibit either CD4+ or CD8+ T cell infiltrate, along with macrophages, and dendritic cells, implicating adaptive and innate immunity (246). Both peripheral blood (dermatomyostis and polymyositis) and muscle biopsies (dermatomyositis) showed evidence for a type I IFN signature and the blood signature correlated with disease activity (247–249). Muscle biopsies from autoimmune myositis patients and inclusion body myositis patients also showed evidence for UPR activation, supported by increased expression of BiP, PERK, GADD 153, ATF3, and chaperones such as grp94, calnexin, calreticulin, and ERp72 (250, 251). Myocytes do not typically express abundant MHC class I, but class I molecules are highly expressed in muscle from these patients, in conjunction with elevated ER stress markers and NF-κB activation (250, 252). Although the link between aberrant MHC expression and ER stress driven inflammation in human cells is mainly correlative, in mice, transgenic overexpression of H-2Kb in skeletal muscle drives an inflammatory myositis phenotype associated with autoantibodies and ER stress (253, 254). Myositis was particularly severe in young mice compared with adults (254).

Besides myositis, an increasing number of rheumatologic conditions appear to be associated with a type I IFN gene signature. This list prominently includes systemic lupus erythematosus (SLE), Sjogren’s disease and systemic sclerosis (255). Moreover, in SLE, the gene signature also correlates with disease activity (256). Outside of plasma B cell development, current evidence for UPR activation in SLE is meager: lupus PBMC showed increased XBP1s but decreased expression of IRE1, PERK, and CHOP (257). T lymphocytes from SLE patients may be more susceptible to ER stress-induced apoptosis, related to defective BiP and autophagy (258). On the other hand, anti-double-stranded DNA antibodies, which are characteristic of lupus, induced both ER stress and cytokine production from human kidney mesangial cells (259). In systemic sclerosis, PBMC from patients showed upregulation of BiP, ATF4, ATF6, XBP1s, along with increased DNAJB1 and IFN-related genes. Furthermore, ER stress markers correlated with disease severity (the presence of pulmonary arterial hypertension) and IL-6 levels (260). Systemic sclerosis involves overproduction of pro-fibrotic cytokines, such as TGF-β, aberrant tissue deposition of collagen, and differentiation of fibroblasts and epithelial cells into myofibroblasts (261). TGF-β increased ER stress in lung fibroblasts, as evident by BiP, ATF6, and XBP1s induction, and also increased expression of α-smooth muscle actin and collagen. Indeed, ER stress may mediate the induction of the myofibroblast proteins, as the chemical chaperone 4-PBA inhibited TGF-β induced α-smooth muscle actin and collagen induction (262). The IRE1α endonuclease pathway also regulated TGF-β driven myofibroblast differentiation in human cells (263).

Finally, autoimmunity frequently targets physiologically highly secretory cells. Autoimmune thyroid diseases are the most prevalent autoimmune conditions and thyrocytes pump out abundant thyroglobulin (264). Melanocytes mount a UPR to cope with melanin production and become targets in vitiligo (265). In the pancreas, β-cells are insulin-producing factories that increase production up to 25-fold in response to glucose (266). In vitiligo and diabetes, CD8+ T cells appear to kill their cellular targets very specifically, without damage to neighboring tissue (265). However, although the autoimmune destruction is carried out by adaptive immune cells, pro-inflammatory cytokine production plays a critical inciting role. The T cell recruiting IFN-regulated chemokine CXCL10 is critical for the development and maintenance of vitiligo (267). In diabetes, IL-1β and IFN-γ induce β-cell apoptosis by stimulating reactive oxygen and nitrogen species (268). Beta-cell death generates autoantigen. Beta-cells also secrete chemokines CXCL10 and CXCL9 that recruit T lymphocytes to the islets (269).

Endoplasmic reticulum stress and the UPR interweave through diabetes pathogenesis on multiple levels. The UPR is absolutely required for basal pancreatic function; PERK−/− mice die early from diabetes and exocrine pancreas failure (28). IRE1/XBP1s activity was also required for glucose-stimulated increases in insulin production and protection from oxidative stress (270). Islets from both diabetes-prone non-obese diabetic (NOD) mice and early human diabetes patients exhibited signs of chronic ER stress with increased CHOP expression and decreased pro-adaptive XBP1 and ATF6 (271, 272). Furthermore, treatment of the NOD mice with TUDCA restored UPR function and markedly protected NOD mice from the development of diabetes (271). Pro-inflammatory cytokines, particularly TNF-α, IFN-γ, and IL-1β induced ER stress (particularly CHOP upregulation) in β-cells (273, 274). TUDCA also protected islet cells from cytokine-induced JNK activation and apoptosis (274). The pro-insulin molecule is prone to misfolding, and human mutations that increase misfolding cause infantile diabetes (275, 276). In the Akita mouse model of diabetes, a mutation in the Ins2 gene that prevents proper proinsulin folding (C96Y) leads to early onset diabetes associated with ER stress. CHOP deficiency delayed diabetes onset in this model by 8–10 weeks (277). In β-cells, activation of IRE1 promoted increased TXNIP expression via miR-17 degradation. TXNIP induction also depended upon PERK. ER stress-induced IL-1β and TXNIP-dependent apoptosis in islets. In THP-1 monocytes, induction of IL-1β depended upon TXNIP and NLRP3 (111). Other studies have also linked NLRP3 and islet IL-1β in type 2 diabetes (278). In the Akita model, deletion of TXNIP protected against β-cell apoptosis and ameliorates diabetes severity (111). Interestingly, NLRP3 deficiency did not prevent diabetes in Akita mice, suggesting other inflammasomes or TXNIP activities may play a role (279). IRE1α has also been linked to the development of diabetes in the NOD mice: targeting the ABL kinases that hyperactivate IRE1 (and thus decreasing IRE1 activity) reversed diabetes in NOD mice (280). These studies provide tantalizing clues that link diabetes and IRE1 activation; however, the connection between ER stress and early cytokine production and apoptosis in these autoimmune conditions remains an open area of investigation. For a summary of the autoimmune and autoinflammatory disorders highlighted above, see Figure 4.

FIGURE 4 | Autoimmune and autoinflammatory diseases involving the unfolded protein response (UPR) and endoplasmic reticulum (ER) stress. Aberrant or excess cytokine production plays a key role in driving autoimmune and autoinflammatory disorders. Interestingly, ER stress and/or the UPR has been increasingly implicated in these same diseases. Thus, the multiple mechanisms by which the UPR interacts with cytokines (both cytokines inducing ER stress and UPR regulating cytokine production) have repercussions for the pathogenesis of inflammatory diseases. Several of the diseases highlighted in this review and the prominent features surrounding ER stress and cytokine induction are in boxes. More autoinflammatory disorders are to the left and autoimmune on the right.

CONCLUSION AND PERSPECTIVES

In summary, the ER plays an indispensable role in cell function and is sensitive to many types of stress; the ER is thus perhaps uniquely poised to transmute significant threats to cell function into amplified immune responses. Because of this role in sensing threats that perturb proteostasis, ER stress has been referred to as a “dyshomeostatic DAMP” (14). From an evolutionary perspective, it may not be mere serendipity that UPR molecules exhibit homology with ancient cytosolic anti-viral proteins, PERK with PKR and IRE1 with RNaseL (281, 282). Numerous pathways interweave the UPR and inflammation, making the ER an effective nidus for promoting sterile inflammation or dramatically amplifying PRR responses. Specifically, the UPR regulates cytokine production through a variety of mechanisms extending from PRR sensing to inflammatory signaling and cytokine transcription factor activation. During infection, the UPR may enable cells to titer the degree of threat, providing greater cytokine responses for threats that impact cell function vs. those that merely stimulate PRRs. The UPR may also enable infected cells to sense invasion by pathogens that otherwise sabotage PRR signaling. Perhaps one of the costs of this inflammatory amplification is the potential for inappropriate activation in the absence of pathogens. The UPR has been increasingly implicated in the pathogenesis of a number of autoimmune and autoinflammatory conditions where cytokines play a central role. However, at this point, much of this is guilt by associations. Although the pieces are there (evidence for UPR, aberrant cytokine production), the exact causative relationships await further definition.

The material presented above raise a number of questions, ranging from mechanistic to teleological. Several questions surround the regulation of the different modes of IRE1 (kinase, RIDD, and XBP1 splicing) activation. Is degree of oligomerization critical or association with co-factors? Does XBP1 directly or indirectly limit kinase activity? Is this occurring via ERAD of IRE1? During TLR4 ligation how does XBP1 promote cytokine production but not its other UPR gene targets? Is this also related to co-factor or heterotypic binding? During viral infections, how does GADD34 promote IFN production and not translation of other targets? For that matter, how does Japanese encephalitis virus trigger RIDD but specifically avoid degradation? The relationship between the proposed microbial stress response and UPR also requires further clarification. Drawbacks to the TLR-mediated suppression of ATF6 and PERK include inhibiting cytokine promotion by these pathways (e.g., NF-κB activation) or adaptive pathways that enable cells to survive stress or commit apoptosis when infected. Infections may induce both ER stress and stimulate multiple PRRs. Perhaps the relative balance of PRR stimulation vs degree of ER stress sways the cell toward either UPR or microbial stress response.

Although the UPR can regulate cytokines, how much of a role does the UPR actually play in cytokine induction during infection and autoimmunity? Moving from the relatively clean results obtained with selective pharmacologic UPR agonists or PRR agonists to the “messy” reality of an intracellular infection or autoimmune disease has been challenging, related to the tremendous increase in complexity. Beyond cytokine regulation, the UPR heavily influences autophagy, nutrient mobilization, and cell death. These other effects of the UPR make it difficult to assign particular responsibility to its effects on cytokines. For instance, it is challenging to tease apart the direct effect of the UPR on viral replication vs. augmented IFN production. In autoimmunity, the UPR may critically regulate autoantigen generation (and presentation) or the basal function of immune type cells. This may be a deus ex machina concept, but perhaps the sheer number of intersections between cytokine regulation and the UPR and the magnitude of effect (e.g., log-fold for IFN) provide support for their significance in disease pathogenesis.

The availability of small molecule inhibitors or agonists for different UPR pathways has grown exponentially, driven by the interest in developing novel therapeutic approaches to cancer and autoimmunity. As an example of repurposed cancer drugs, proteosome inhibitors, which affect proteostasis (and thus ER function) and cytokine production, have demonstrated efficacy in murine lupus models (283, 284). UPR modulating agents may also be useful for intractable infectious diseases or vaccine development. Some of these UPR drugs have already moved to clinical trials. For instance, Celgosivir, which inhibits N-linked glycosylation, is undergoing a phase II trial for Dengue (285). Better understanding of the role of the UPR in specific settings will be critical for the judicious trial of these new therapies; given the complexity of UPR-immune interactions, carefully conceived pre-clinical studies may be necessary to gage the net effect of individual UPR modulating agents on specific infectious or inflammatory conditions. It will be important not to generalize, as the role of the UPR is likely to be highly context specific, even between species of pathogen. An example described above, B. melitensis and B. abortus have been reported to induce different degrees of UPR activation and blockade with TUDCA appears to have different effects on replication (118, 182). Also, it will be important to balance the anti-pathogenic effects of UPR modulation against the potential of disturbing physiologic UPR responses. Given the exciting clinical potential for UPR modulation, clarification of these issues has become a compelling mandate.

AUTHOR CONTRIBUTIONS

The author confirms being the sole contributor of this work and approved it for publication.

FUNDING

This work was supported by the NIH: R01AI073558, R01 AI116453-01 and R21AI121808-01.

REFERENCES

1. O’Brien C, Flower DR, Feighery C. Peptide length significantly influences in vitro affinity for MHC class II molecules. Immunome Res (2008) 4:6. doi:10.1186/1745-7580-4-6

2. Calis JJ, de Boer RJ, Kesmir C. Degenerate T-cell recognition of peptides on MHC molecules creates large holes in the T-cell repertoire. PLoS Comput Biol (2012) 8(3):e1002412. doi:10.1371/journal.pcbi.1002412

3. Bremel RD, Homan EJ. Frequency patterns of T-cell exposed amino acid motifs in immunoglobulin heavy chain peptides presented by MHCs. Front Immunol (2014) 5:541. doi:10.3389/fimmu.2014.00541

4. Stave JW, Lindpaintner K. Antibody and antigen contact residues define epitope and paratope size and structure. J Immunol (2013) 191(3):1428–35. doi:10.4049/jimmunol.1203198

5. Akira S, Uematsu S, Takeuchi O. Pathogen recognition and innate immunity. Cell (2006) 124(4):783–801. doi:10.1016/j.cell.2006.02.015

6. van Vliet SJ, Garcia-Vallejo JJ, van Kooyk Y. Dendritic cells and C-type lectin receptors: coupling innate to adaptive immune responses. Immunol Cell Biol (2008) 86(7):580–7. doi:10.1038/icb.2008.55

7. Place DE, Kanneganti TD. Recent advances in inflammasome biology. Curr Opin Immunol (2017) 50:32–8. doi:10.1016/j.coi.2017.10.011

8. Barber GN. STING-dependent cytosolic DNA sensing pathways. Trends Immunol (2014) 35(2):88–93. doi:10.1016/j.it.2013.10.010

9. Rose NR. Negative selection, epitope mimicry and autoimmunity. Curr Opin Immunol (2017) 49:51–5. doi:10.1016/j.coi.2017.08.014

10. Cunningham MW. Post-streptococcal autoimmune sequelae: rheumatic fever and beyond. In: Ferretti JJ, Stevens DL, Fischetti VA, editors. Streptococcus pyogenes: Basic Biology to Clinical Manifestations [Internet]. Oklahoma: University of Oklahoma Health Sciences Center (2016).

11. Matzinger P. Tolerance, danger, and the extended family. Annu Rev Immunol (1994) 12:991–1045. doi:10.1146/annurev.iy.12.040194.005015

12. Matzinger P. The danger model: a renewed sense of self. Science (2002) 296(5566):301–5. doi:10.1126/science.1071059

13. Pradeu T, Cooper EL. The danger theory: 20 years later. Front Immunol (2012) 3:287. doi:10.3389/fimmu.2012.00287

14. Land WG, Agostinis P, Gasser S, Garg AD, Linkermann A. Transplantation and damage-associated molecular patterns (DAMPs). Am J Transplant (2016) 16(12):3338–61. doi:10.1111/ajt.13963

15. Kono H, Rock KL. How dying cells alert the immune system to danger. Nat Rev Immunol (2008) 8(4):279–89. doi:10.1038/nri2215

16. Hirsiger S, Simmen HP, Werner CM, Wanner GA, Rittirsch D. Danger signals activating the immune response after trauma. Mediators Inflamm (2012) 2012:315941. doi:10.1155/2012/315941

17. Yu L, Wang L, Chen S. Endogenous toll-like receptor ligands and their biological significance. J Cell Mol Med (2010) 14(11):2592–603. doi:10.1111/j.1582-4934.2010.01127.x

18. Erridge C. Endogenous ligands of TLR2 and TLR4: agonists or assistants? J Leukoc Biol (2010) 87(6):989–99. doi:10.1189/jlb.1209775

19. Bhattacharyya S, Varga J. Endogenous ligands of TLR4 promote unresolving tissue fibrosis: implications for systemic sclerosis and its targeted therapy. Immunol Lett (2018) 195:9–17. doi:10.1016/j.imlet.2017.09.011

20. Vance RE, Isberg RR, Portnoy DA. Patterns of pathogenesis: discrimination of pathogenic and nonpathogenic microbes by the innate immune system. Cell Host Microbe (2009) 6(1):10–21. doi:10.1016/j.chom.2009.06.007

21. Karaolis DK, Means TK, Yang D, Takahashi M, Yoshimura T, Muraille E, et al. Bacterial c-di-GMP is an immunostimulatory molecule. J Immunol (2007) 178(4):2171–81. doi:10.4049/jimmunol.178.4.2171

22. Sun YH, Rolan HG, Tsolis RM. Injection of flagellin into the host cell cytosol by Salmonella enterica serotype typhimurium. J Biol Chem (2007) 282(47):33897–901. doi:10.1074/jbc.C700181200

23. Lightfield KL, Persson J, Brubaker SW, Witte CE, von Moltke J, Dunipace EA, et al. Critical function for Naip5 in inflammasome activation by a conserved carboxy-terminal domain of flagellin. Nat Immunol (2008) 9(10):1171–8. doi:10.1038/ni.1646

24. Legrand-Poels S, Kustermans G, Bex F, Kremmer E, Kufer TA, Piette J. Modulation of Nod2-dependent NF-kappaB signaling by the actin cytoskeleton. J Cell Sci (2007) 120(Pt 7):1299–310. doi:10.1242/jcs.03424

25. Waite AL, Schaner P, Hu C, Richards N, Balci-Peynircioglu B, Hong A, et al. Pyrin and ASC co-localize to cellular sites that are rich in polymerizing actin. Exp Biol Med (Maywood) (2009) 234(1):40–52. doi:10.3181/0806-RM-184

26. Stevens FJ, Argon Y. Protein folding in the ER. Semin Cell Dev Biol (1999) 10(5):443–54. doi:10.1006/scdb.1999.0315

27. Schroder M, Kaufman RJ. The mammalian unfolded protein response. Annu Rev Biochem (2005) 74:739–89. doi:10.1146/annurev.biochem.73.011303.074134

28. Harding HP, Zeng H, Zhang Y, Jungries R, Chung P, Plesken H, et al. Diabetes mellitus and exocrine pancreatic dysfunction in perk-/- mice reveals a role for translational control in secretory cell survival. Mol Cell (2001) 7(6):1153–63. doi:10.1016/S1097-2765(01)00264-7

29. Lee AH, Chu GC, Iwakoshi NN, Glimcher LH. XBP-1 is required for biogenesis of cellular secretory machinery of exocrine glands. EMBO J (2005) 24(24):4368–80. doi:10.1038/sj.emboj.7600903

30. Iwakoshi NN, Lee AH, Vallabhajosyula P, Otipoby KL, Rajewsky K, Glimcher LH. Plasma cell differentiation and the unfolded protein response intersect at the transcription factor XBP-1. Nat Immunol (2003) 4(4):321–9. doi:10.1038/ni907

31. Shaffer AL, Shapiro-Shelef M, Iwakoshi NN, Lee AH, Qian SB, Zhao H, et al. XBP1, downstream of Blimp-1, expands the secretory apparatus and other organelles, and increases protein synthesis in plasma cell differentiation. Immunity (2004) 21(1):81–93. doi:10.1016/j.immuni.2004.06.010

32. Todd DJ, Lee AH, Glimcher LH. The endoplasmic reticulum stress response in immunity and autoimmunity. Nat Rev Immunol (2008) 8(9):663–74. doi:10.1038/nri2359

33. Iwakoshi NN, Pypaert M, Glimcher LH. The transcription factor XBP-1 is essential for the development and survival of dendritic cells. J Exp Med (2007) 204(10):2267–75. doi:10.1084/jem.20070525

34. Bettigole SE, Lis R, Adoro S, Lee AH, Spencer LA, Weller PF, et al. The transcription factor XBP1 is selectively required for eosinophil differentiation. Nat Immunol (2015) 16(8):829–37. doi:10.1038/ni.3225

35. Grootjans J, Kaser A, Kaufman RJ, Blumberg RS. The unfolded protein response in immunity and inflammation. Nat Rev Immunol (2016) 16(8):469–84. doi:10.1038/nri.2016.62

36. Oppenheim JJ. Cytokines: past, present, and future. Int J Hematol (2001) 74(1):3–8. doi:10.1007/BF02982543

37. Zhang K, Shen X, Wu J, Sakaki K, Saunders T, Rutkowski DT, et al. Endoplasmic reticulum stress activates cleavage of CREBH to induce a systemic inflammatory response. Cell (2006) 124(3):587–99. doi:10.1016/j.cell.2005.11.040

38. Walter P, Ron D. The unfolded protein response: from stress pathway to homeostatic regulation. Science (2011) 334(6059):1081–6. doi:10.1126/science.1209038

39. Credle JJ, Finer-Moore JS, Papa FR, Stroud RM, Walter P. On the mechanism of sensing unfolded protein in the endoplasmic reticulum. Proc Natl Acad Sci U S A (2005) 102(52):18773–84. doi:10.1073/pnas.0509487102

40. Zhou J, Liu CY, Back SH, Clark RL, Peisach D, Xu Z, et al. The crystal structure of human IRE1 luminal domain reveals a conserved dimerization interface required for activation of the unfolded protein response. Proc Natl Acad Sci U S A (2006) 103(39):14343–8. doi:10.1073/pnas.0606480103

41. Karagöz GE, Acosta-Alvear D, Nguyen HT, Lee CP, Chu F, Walter P. An unfolded protein-induced conformational switch activates mammalian IRE1. Elife (2017) 6:e30700. doi:10.7554/eLife.30700

42. Volmer R, van der Ploeg K, Ron D. Membrane lipid saturation activates endoplasmic reticulum unfolded protein response transducers through their transmembrane domains. Proc Natl Acad Sci U S A (2013) 110(12):4628–33. doi:10.1073/pnas.1217611110

43. Bertolotti A, Wang X, Novoa I, Jungreis R, Schlessinger K, Cho JH, et al. Increased sensitivity to dextran sodium sulfate colitis in IRE1beta-deficient mice. J Clin Invest (2001) 107(5):585–93. doi:10.1172/JCI11476

44. Martino MB, Jones L, Brighton B, Ehre C, Abdulah L, Davis CW, et al. The ER stress transducer IRE1beta is required for airway epithelial mucin production. Mucosal Immunol (2013) 6(3):639–54. doi:10.1038/mi.2012.105

45. Hetz C, Martinon F, Rodriguez D, Glimcher LH. The unfolded protein response: integrating stress signals through the stress sensor IRE1alpha. Physiol Rev (2011) 91(4):1219–43. doi:10.1152/physrev.00001.2011

46. Lee AH, Iwakoshi NN, Glimcher LH. XBP-1 regulates a subset of endoplasmic reticulum resident chaperone genes in the unfolded protein response. Mol Cell Biol (2003) 23(21):7448–59. doi:10.1128/MCB.23.21.7448-7459.2003

47. Sriburi R, Bommiasamy H, Buldak GL, Robbins GR, Frank M, Jackowski S, et al. Coordinate regulation of phospholipid biosynthesis and secretory pathway gene expression in XBP-1(S)-induced endoplasmic reticulum biogenesis. J Biol Chem (2007) 282(10):7024–34. doi:10.1074/jbc.M609490200

48. Jäger R, Bertrand MJ, Gorman AM, Vandenabeele P, Samali A. The unfolded protein response at the crossroads of cellular life and death during endoplasmic reticulum stress. Biol Cell (2012) 104(5):259–70. doi:10.1111/boc.201100055

49. Lin JH, Li H, Yasumura D, Cohen HR, Zhang C, Panning B, et al. IRE1 signaling affects cell fate during the unfolded protein response. Science (2007) 318(5852):944–9. doi:10.1126/science.1146361

50. Hollien J, Weissman JS. Decay of endoplasmic reticulum-localized mRNAs during the unfolded protein response. Science (2006) 313(5783):104–7. doi:10.1126/science.1129631

51. Hollien J, Lin JH, Li H, Stevens N, Walter P, Weissman JS. Regulated Ire1-dependent decay of messenger RNAs in mammalian cells. J Cell Biol (2009) 186(3):323–31. doi:10.1083/jcb.200903014

52. Han D, Lerner AG, Vande Walle L, Upton JP, Xu W, Hagen A, et al. IRE1alpha kinase activation modes control alternate endoribonuclease outputs to determine divergent cell fates. Cell (2009) 138(3):562–75. doi:10.1016/j.cell.2009.07.017

53. Ghosh R, Wang L, Wang ES, Perera BG, Igbaria A, Morita S, et al. Allosteric inhibition of the IRE1alpha RNase preserves cell viability and function during endoplasmic reticulum stress. Cell (2014) 158(3):534–48. doi:10.1016/j.cell.2014.07.002

54. Hetz C, Glimcher LH. Fine-tuning of the unfolded protein response: assembling the IRE1alpha interactome. Mol Cell (2009) 35(5):551–61. doi:10.1016/j.molcel.2009.08.021

55. Nishitoh H, Matsuzawa A, Tobiume K, Saegusa K, Takeda K, Inoue K, et al. ASK1 is essential for endoplasmic reticulum stress-induced neuronal cell death triggered by expanded polyglutamine repeats. Genes Dev (2002) 16(11):1345–55. doi:10.1101/gad.992302

56. Hetz C, Bernasconi P, Fisher J, Lee AH, Bassik MC, Antonsson B, et al. Proapoptotic BAX and BAK modulate the unfolded protein response by a direct interaction with IRE1alpha. Science (2006) 312(5773):572–6. doi:10.1126/science.1123480

57. Young SK, Palam LR, Wu C, Sachs MS, Wek RC. Ribosome elongation stall directs gene-specific translation in the integrated stress response. J Biol Chem (2016) 291(12):6546–58. doi:10.1074/jbc.M115.705640

58. Harding HP, Zhang Y, Bertolotti A, Zeng H, Ron D. Perk is essential for translational regulation and cell survival during the unfolded protein response. Mol Cell (2000) 5(5):897–904. doi:10.1016/S1097-2765(00)80330-5

59. Lu PD, Harding HP, Ron D. Translation reinitiation at alternative open reading frames regulates gene expression in an integrated stress response. J Cell Biol (2004) 167(1):27–33. doi:10.1083/jcb.200408003

60. Vattem KM, Wek RC. Reinitiation involving upstream ORFs regulates ATF4 mRNA translation in mammalian cells. Proc Natl Acad Sci U S A (2004) 101(31):11269–74. doi:10.1073/pnas.0400541101

61. Harding HP, Zhang Y, Zeng H, Novoa I, Lu PD, Calfon M, et al. An integrated stress response regulates amino acid metabolism and resistance to oxidative stress. Mol Cell (2003) 11(3):619–33. doi:10.1016/S1097-2765(03)00105-9

62. B’Chir W, Maurin AC, Carraro V, Averous J, Jousse C, Muranishi Y, et al. The eIF2alpha/ATF4 pathway is essential for stress-induced autophagy gene expression. Nucleic Acids Res (2013) 41(16):7683–99. doi:10.1093/nar/gkt563

63. Novoa I, Zeng H, Harding HP, Ron D. Feedback inhibition of the unfolded protein response by GADD34-mediated dephosphorylation of eIF2alpha. J Cell Biol (2001) 153(5):1011–22. doi:10.1083/jcb.153.5.1011

64. Yan W, Frank CL, Korth MJ, Sopher BL, Novoa I, Ron D, et al. Control of PERK eIF2alpha kinase activity by the endoplasmic reticulum stress-induced molecular chaperone P58IPK. Proc Natl Acad Sci U S A (2002) 99(25):15920–5. doi:10.1073/pnas.252341799

65. Zhang HM, Ye X, Su Y, Yuan J, Liu Z, Stein DA, et al. Coxsackievirus B3 infection activates the unfolded protein response and induces apoptosis through downregulation of p58IPK and activation of CHOP and SREBP1. J Virol (2010) 84(17):8446–59. doi:10.1128/JVI.01416-09

66. Jousse C, Oyadomari S, Novoa I, Lu P, Zhang Y, Harding HP, et al. Inhibition of a constitutive translation initiation factor 2alpha phosphatase, CReP, promotes survival of stressed cells. J Cell Biol (2003) 163(4):767–75. doi:10.1083/jcb.200308075

67. Pakos-Zebrucka K, Koryga I, Mnich K, Ljujic M, Samali A, Gorman AM. The integrated stress response. EMBO Rep (2016) 17(10):1374–95. doi:10.15252/embr.201642195

68. Cullinan SB, Zhang D, Hannink M, Arvisais E, Kaufman RJ, Diehl JA. Nrf2 is a direct PERK substrate and effector of PERK-dependent cell survival. Mol Cell Biol (2003) 23(20):7198–209. doi:10.1128/MCB.23.20.7198-7209.2003

69. Shen J, Chen X, Hendershot L, Prywes R. ER stress regulation of ATF6 localization by dissociation of BiP/GRP78 binding and unmasking of Golgi localization signals. Dev Cell (2002) 3(1):99–111. doi:10.1016/S1534-5807(02)00203-4

70. Yamamoto K, Sato T, Matsui T, Sato M, Okada T, Yoshida H, et al. Transcriptional induction of mammalian ER quality control proteins is mediated by single or combined action of ATF6alpha and XBP1. Dev Cell (2007) 13(3):365–76. doi:10.1016/j.devcel.2007.07.018

71. Lee K, Tirasophon W, Shen X, Michalak M, Prywes R, Okada T, et al. IRE1-mediated unconventional mRNA splicing and S2P-mediated ATF6 cleavage merge to regulate XBP1 in signaling the unfolded protein response. Genes Dev (2002) 16(4):452–66. doi:10.1101/gad.964702

72. Asada R, Kanemoto S, Kondo S, Saito A, Imaizumi K. The signalling from endoplasmic reticulum-resident bZIP transcription factors involved in diverse cellular physiology. J Biochem (2011) 149(5):507–18. doi:10.1093/jb/mvr041

73. Li Y, Schwabe RF, DeVries-Seimon T, Yao PM, Gerbod-Giannone MC, Tall AR, et al. Free cholesterol-loaded macrophages are an abundant source of tumor necrosis factor-alpha and interleukin-6: model of NF-kappaB- and map kinase-dependent inflammation in advanced atherosclerosis. J Biol Chem (2005) 280(23):21763–72. doi:10.1074/jbc.M501759200

74. Martinon F, Chen X, Lee AH, Glimcher LH. TLR activation of the transcription factor XBP1 regulates innate immune responses in macrophages. Nat Immunol (2010) 11(5):411–8. doi:10.1038/ni.1857

75. Peters LR, Raghavan M. Endoplasmic reticulum calcium depletion impacts chaperone secretion, innate immunity, and phagocytic uptake of cells. J Immunol (2011) 187(2):919–31. doi:10.4049/jimmunol.1100690

76. Hayden MS, Ghosh S. Shared principles in NF-kappaB signaling. Cell (2008) 132(3):344–62. doi:10.1016/j.cell.2008.01.020

77. Hu P, Han Z, Couvillon AD, Kaufman RJ, Exton JH. Autocrine tumor necrosis factor alpha links endoplasmic reticulum stress to the membrane death receptor pathway through IRE1alpha-mediated NF-kappaB activation and down-regulation of TRAF2 expression. Mol Cell Biol (2006) 26(8):3071–84. doi:10.1128/MCB.26.8.3071-3084.2006

78. Kaneko M, Niinuma Y, Nomura Y. Activation signal of nuclear factor-kappa B in response to endoplasmic reticulum stress is transduced via IRE1 and tumor necrosis factor receptor-associated factor 2. Biol Pharm Bull (2003) 26(7):931–5. doi:10.1248/bpb.26.931

79. Tam AB, Mercado EL, Hoffmann A, Niwa M. ER stress activates NF-kappaB by integrating functions of basal IKK activity, IRE1 and PERK. PLoS One (2012) 7(10):e45078. doi:10.1371/journal.pone.0045078

80. Kim S, Joe Y, Kim HJ, Kim YS, Jeong SO, Pae HO, et al. Endoplasmic reticulum stress-induced IRE1alpha activation mediates cross-talk of GSK-3beta and XBP-1 to regulate inflammatory cytokine production. J Immunol (2015) 194(9):4498–506. doi:10.4049/jimmunol.1401399

81. Demarchi F, Bertoli C, Sandy P, Schneider C. Glycogen synthase kinase-3 beta regulates NF-kappa B1/p105 stability. J Biol Chem (2003) 278(41):39583–90. doi:10.1074/jbc.M305676200

82. Deng J, Lu PD, Zhang Y, Scheuner D, Kaufman RJ, Sonenberg N, et al. Translational repression mediates activation of nuclear factor kappa B by phosphorylated translation initiation factor 2. Mol Cell Biol (2004) 24(23):10161–8. doi:10.1128/MCB.24.23.10161-10168.2004

83. Park SH, Choi HJ, Yang H, Do KH, Kim J, Lee DW, et al. Endoplasmic reticulum stress-activated C/EBP homologous protein enhances nuclear factor-kappaB signals via repression of peroxisome proliferator-activated receptor gamma. J Biol Chem (2010) 285(46):35330–9. doi:10.1074/jbc.M110.136259

84. Yamazaki H, Hiramatsu N, Hayakawa K, Tagawa Y, Okamura M, Ogata R, et al. Activation of the Akt-NF-kappaB pathway by subtilase cytotoxin through the ATF6 branch of the unfolded protein response. J Immunol (2009) 183(2):1480–7. doi:10.4049/jimmunol.0900017

85. Nakajima S, Hiramatsu N, Hayakawa K, Saito Y, Kato H, Huang T, et al. Selective abrogation of BiP/GRP78 blunts activation of NF-kappaB through the ATF6 branch of the UPR: involvement of C/EBPbeta and mTOR-dependent dephosphorylation of Akt. Mol Cell Biol (2011) 31(8):1710–8. doi:10.1128/MCB.00939-10

86. Pahl HL, Baeuerle PA. Activation of NF-kappa B by ER stress requires both Ca2+ and reactive oxygen intermediates as messengers. FEBS Lett (1996) 392(2):129–36. doi:10.1016/0014-5793(96)00800-9

87. Whitmarsh AJ, Davis RJ. Transcription factor AP-1 regulation by mitogen-activated protein kinase signal transduction pathways. J Mol Med (Berl) (1996) 74(10):589–607. doi:10.1007/s001090050063

88. Darling NJ, Cook SJ. The role of MAPK signalling pathways in the response to endoplasmic reticulum stress. Biochim Biophys Acta (2014) 1843(10):2150–63. doi:10.1016/j.bbamcr.2014.01.009

89. Ichijo H, Nishida E, Irie K, ten Dijke P, Saitoh M, Moriguchi T, et al. Induction of apoptosis by ASK1, a mammalian MAPKKK that activates SAPK/JNK and p38 signaling pathways. Science (1997) 275(5296):90–4. doi:10.1126/science.275.5296.90

90. Nguyên DT, Kebache S, Fazel A, Wong HN, Jenna S, Emadali A, et al. Nck-dependent activation of extracellular signal-regulated kinase-1 and regulation of cell survival during endoplasmic reticulum stress. Mol Biol Cell (2004) 15(9):4248–60. doi:10.1091/mbc.E03-11-0851

91. Mijošek V, Lasitschka F, Warth A, Zabeck H, Dalpke AH, Weitnauer M. Endoplasmic reticulum stress is a danger signal promoting innate inflammatory responses in bronchial epithelial cells. J Innate Immun (2016) 8(5):464–78. doi:10.1159/000447668

92. Wang XZ, Ron D. Stress-induced phosphorylation and activation of the transcription factor CHOP (GADD153) by p38 MAP Kinase. Science (1996) 272(5266):1347–9. doi:10.1126/science.272.5266.1347

93. Luo S, Lee AS. Requirement of the p38 mitogen-activated protein kinase signalling pathway for the induction of the 78 kDa glucose-regulated protein/immunoglobulin heavy-chain binding protein by azetidine stress: activating transcription factor 6 as a target for stress-induced phosphorylation. Biochem J (2002) 366(Pt 3):787–95. doi:10.1042/BJ20011802

94. Thuerauf DJ, Arnold ND, Zechner D, Hanford DS, DeMartin KM, McDonough PM, et al. p38 Mitogen-activated protein kinase mediates the transcriptional induction of the atrial natriuretic factor gene through a serum response element. A potential role for the transcription factor ATF6. J Biol Chem (1998) 273(32):20636–43. doi:10.1074/jbc.273.32.20636

95. Sato M, Suemori H, Hata N, Asagiri M, Ogasawara K, Nakao K, et al. Distinct and essential roles of transcription factors IRF-3 and IRF-7 in response to viruses for IFN-alpha/beta gene induction. Immunity (2000) 13(4):539–48. doi:10.1016/S1074-7613(00)00053-4

96. Agalioti T, Lomvardas S, Parekh B, Yie J, Maniatis T, Thanos D. Ordered recruitment of chromatin modifying and general transcription factors to the IFN-beta promoter. Cell (2000) 103(4):667–78. doi:10.1016/S0092-8674(00)00169-0

97. Merika M, Thanos D. Enhanceosomes. Curr Opin Genet Dev (2001) 11(2):205–8. doi:10.1016/S0959-437X(00)00180-5

98. Panne D, Maniatis T, Harrison SC. An atomic model of the interferon-beta enhanceosome. Cell (2007) 129(6):1111–23. doi:10.1016/j.cell.2007.05.019

99. Hiscott J. Triggering the innate antiviral response through IRF-3 activation. J Biol Chem (2007) 282(21):15325–9. doi:10.1074/jbc.R700002200

100. Chen W, Srinath H, Lam SS, Schiffer CA, Royer WE Jr, Lin K. Contribution of Ser386 and Ser396 to activation of interferon regulatory factor 3. J Mol Biol (2008) 379(2):251–60. doi:10.1016/j.jmb.2008.03.050

101. Liu YP, Zeng L, Tian A, Bomkamp A, Rivera D, Gutman D, et al. Endoplasmic reticulum stress regulates the innate immunity critical transcription factor IRF3. J Immunol (2012) 189(9):4630–9. doi:10.4049/jimmunol.1102737

102. Zeng L, Liu YP, Sha H, Chen H, Qi L, Smith JA. XBP-1 couples endoplasmic reticulum stress to augmented IFN-beta induction via a cis-acting enhancer in macrophages. J Immunol (2010) 185(4):2324–30. doi:10.4049/jimmunol.0903052

103. Iwasaki Y, Suganami T, Hachiya R, Shirakawa I, Kim-Saijo M, Tanaka M, et al. Activating transcription factor 4 links metabolic stress to interleukin-6 expression in macrophages. Diabetes (2014) 63(1):152–61. doi:10.2337/db13-0757

104. Goodall JC, Wu C, Zhang Y, McNeill L, Ellis L, Saudek V, et al. Endoplasmic reticulum stress-induced transcription factor, CHOP, is crucial for dendritic cell IL-23 expression. Proc Natl Acad Sci U S A (2010) 107(41):17698–703. doi:10.1073/pnas.1011736107

105. Jiang HY, Wek SA, McGrath BC, Lu D, Hai T, Harding HP, et al. Activating transcription factor 3 is integral to the eukaryotic initiation factor 2 kinase stress response. Mol Cell Biol (2004) 24(3):1365–77. doi:10.1128/MCB.24.3.1365-1377.2004

106. Brooks AC, Guo Y, Singh M, McCracken J, Xuan YT, Srivastava S, et al. Endoplasmic reticulum stress-dependent activation of ATF3 mediates the late phase of ischemic preconditioning. J Mol Cell Cardiol (2014) 76:138–47. doi:10.1016/j.yjmcc.2014.08.011

107. Gilchrist M, Thorsson V, Li B, Rust AG, Korb M, Roach JC, et al. Systems biology approaches identify ATF3 as a negative regulator of Toll-like receptor 4. Nature (2006) 441(7090):173–8. doi:10.1038/nature04768

108. Whitmore MM, Iparraguirre A, Kubelka L, Weninger W, Hai T, Williams BR. Negative regulation of TLR-signaling pathways by activating transcription factor-3. J Immunol (2007) 179(6):3622–30. doi:10.4049/jimmunol.179.6.3622

109. Thompson MR, Xu D, Williams BR. ATF3 transcription factor and its emerging roles in immunity and cancer. J Mol Med (Berl) (2009) 87(11):1053–60. doi:10.1007/s00109-009-0520-x

110. Dong D, Fu N, Yang P. MiR-17 downregulation by high glucose stabilizes thioredoxin-interacting protein and removes thioredoxin inhibition on ASK1 leading to apoptosis. Toxicol Sci (2016) 150(1):84–96. doi:10.1093/toxsci/kfv313

111. Lerner AG, Upton JP, Praveen PV, Ghosh R, Nakagawa Y, Igbaria A, et al. IRE1alpha induces thioredoxin-interacting protein to activate the NLRP3 inflammasome and promote programmed cell death under irremediable ER stress. Cell Metab (2012) 16(2):250–64. doi:10.1016/j.cmet.2012.07.007

112. Oslowski CM, Hara T, O’Sullivan-Murphy B, Kanekura K, Lu S, Hara M, et al. Thioredoxin-interacting protein mediates ER stress-induced beta cell death through initiation of the inflammasome. Cell Metab (2012) 16(2):265–73. doi:10.1016/j.cmet.2012.07.005

113. Bronner DN, Abuaita BH, Chen X, Fitzgerald KA, Nuñez G, He Y, et al. Endoplasmic reticulum stress activates the inflammasome via NLRP3- and caspase-2-driven mitochondrial damage. Immunity (2015) 43(3):451–62. doi:10.1016/j.immuni.2015.08.008

114. Cho HK, Cheong KJ, Kim HY, Cheong J. Endoplasmic reticulum stress induced by hepatitis B virus X protein enhances cyclo-oxygenase 2 expression via activating transcription factor 4. Biochem J (2011) 435(2):431–9. doi:10.1042/BJ20102071

115. West AP, Khoury-Hanold W, Staron M, Tal MC, Pineda CM, Lang SM, et al. Mitochondrial DNA stress primes the antiviral innate immune response. Nature (2015) 520(7548):553–7. doi:10.1038/nature14156

116. Cao SS, Kaufman RJ. Endoplasmic reticulum stress and oxidative stress in cell fate decision and human disease. Antioxid Redox Signal (2014) 21(3):396–413. doi:10.1089/ars.2014.5851

117. Szabadkai G, Bianchi K, Várnai P, De Stefani D, Wieckowski MR, Cavagna D, et al. Chaperone-mediated coupling of endoplasmic reticulum and mitochondrial Ca2+ channels. J Cell Biol (2006) 175(6):901–11. doi:10.1083/jcb.200608073

118. Keestra-Gounder AM, Byndloss MX, Seyffert N, Young BM, Chávez-Arroyo A, Tsai AY, et al. NOD1 and NOD2 signalling links ER stress with inflammation. Nature (2016) 532(7599):394–7. doi:10.1038/nature17631

119. Xie P. TRAF molecules in cell signaling and in human diseases. J Mol Signal (2013) 8(1):7. doi:10.1186/1750-2187-8-7

120. Smith JA, Turner MJ, DeLay ML, Klenk EI, Sowders DP, Colbert RA. Endoplasmic reticulum stress and the unfolded protein response are linked to synergistic IFN-beta induction via X-box binding protein 1. Eur J Immunol (2008) 38(5):1194–203. doi:10.1002/eji.200737882

121. DeLay ML, Turner MJ, Klenk EI, Smith JA, Sowders DP, Colbert RA. HLA-B27 misfolding and the unfolded protein response augment interleukin-23 production and are associated with Th17 activation in transgenic rats. Arthritis Rheum (2009) 60(9):2633–43. doi:10.1002/art.24763

122. Hu F, Yu X, Wang H, Zuo D, Guo C, Yi H, et al. ER stress and its regulator X-box-binding protein-1 enhance polyIC-induced innate immune response in dendritic cells. Eur J Immunol (2011) 41(4):1086–97. doi:10.1002/eji.201040831

123. Shenderov K, Riteau N, Yip R, Mayer-Barber KD, Oland S, Hieny S, et al. Cutting edge: endoplasmic reticulum stress licenses macrophages to produce mature IL-1beta in response to TLR4 stimulation through a caspase-8- and TRIF-dependent pathway. J Immunol (2014) 192(5):2029–33. doi:10.4049/jimmunol.1302549

124. Leonard A, Paton AW, El-Quadi M, Paton JC, Fazal F. Preconditioning with endoplasmic reticulum stress ameliorates endothelial cell inflammation. PLoS One (2014) 9(10):e110949. doi:10.1371/journal.pone.0110949

125. Woo CW, Cui D, Arellano J, Dorweiler B, Harding H, Fitzgerald KA, et al. Adaptive suppression of the ATF4-CHOP branch of the unfolded protein response by toll-like receptor signalling. Nat Cell Biol (2009) 11(12):1473–80. doi:10.1038/ncb1996

126. Nakayama Y, Endo M, Tsukano H, Mori M, Oike Y, Gotoh T. Molecular mechanisms of the LPS-induced non-apoptotic ER stress-CHOP pathway. J Biochem (2010) 147(4):471–83. doi:10.1093/jb/mvp189

127. Clavarino G, Cláudio N, Couderc T, Dalet A, Judith D, Camosseto V, et al. Induction of GADD34 is necessary for dsRNA-dependent interferon-beta production and participates in the control of chikungunya virus infection. PLoS Pathog (2012) 8(5):e1002708. doi:10.1371/journal.ppat.1002708

128. Clavarino G, Cláudio N, Dalet A, Terawaki S, Couderc T, Chasson L, et al. Protein phosphatase 1 subunit Ppp1r15a/GADD34 regulates cytokine production in polyinosinic:polycytidylic acid-stimulated dendritic cells. Proc Natl Acad Sci U S A (2012) 109(8):3006–11. doi:10.1073/pnas.1104491109

129. Cláudio N, Dalet A, Gatti E, Pierre P. Mapping the crossroads of immune activation and cellular stress response pathways. EMBO J (2013) 32(9):1214–24. doi:10.1038/emboj.2013.80

130. Obeid M, Tesniere A, Ghiringhelli F, Fimia GM, Apetoh L, Perfettini JL, et al. Calreticulin exposure dictates the immunogenicity of cancer cell death. Nat Med (2007) 13(1):54–61. doi:10.1038/nm1523

131. Galluzzi L, Diotallevi A, Magnani M. Endoplasmic reticulum stress and unfolded protein response in infection by intracellular parasites. Future Sci OA (2017) 3(3):FSO198. doi:10.4155/fsoa-2017-0020

132. Ren J, Liu T, Pang L, Li K, Garofalo RP, Casola A, et al. A novel mechanism for the inhibition of interferon regulatory factor-3-dependent gene expression by human respiratory syncytial virus NS1 protein. J Gen Virol (2011) 92(Pt 9):2153–9. doi:10.1099/vir.0.032987-0

133. Nitta S, Sakamoto N, Nakagawa M, Kakinuma S, Mishima K, Kusano-Kitazume A, et al. Hepatitis C virus NS4B protein targets STING and abrogates RIG-I-mediated type I interferon-dependent innate immunity. Hepatology (2013) 57(1):46–58. doi:10.1002/hep.26017

134. Sun L, Xing Y, Chen X, Zheng Y, Yang Y, Nichols DB, et al. Coronavirus papain-like proteases negatively regulate antiviral innate immune response through disruption of STING-mediated signaling. PLoS One (2012) 7(2):e30802. doi:10.1371/journal.pone.0030802

135. Aguirre S, Maestre AM, Pagni S, Patel JR, Savage T, Gutman D, et al. DENV inhibits type I IFN production in infected cells by cleaving human STING. PLoS Pathog (2012) 8(10):e1002934. doi:10.1371/journal.ppat.1002934

136. Aguirre S, Luthra P, Sanchez-Aparicio MT, Maestre AM, Patel J, Lamothe F, et al. Dengue virus NS2B protein targets cGAS for degradation and prevents mitochondrial DNA sensing during infection. Nat Microbiol (2017) 2:17037. doi:10.1038/nmicrobiol.2017.37

137. Liu J, HuangFu WC, Kumar KG, Qian J, Casey JP, Hamanaka RB, et al. Virus-induced unfolded protein response attenuates antiviral defenses via phosphorylation-dependent degradation of the type I interferon receptor. Cell Host Microbe (2009) 5(1):72–83. doi:10.1016/j.chom.2008.11.008

138. Horvath CM. Weapons of STAT destruction. Interferon evasion by paramyxovirus V protein. Eur J Biochem (2004) 271(23–24):4621–8. doi:10.1111/j.1432-1033.2004.04425.x

139. Choe SS, Dodd DA, Kirkegaard K. Inhibition of cellular protein secretion by picornaviral 3A proteins. Virology (2005) 337(1):18–29. doi:10.1016/j.virol.2005.03.036

140. Moradpour D, Penin F, Rice CM. Replication of hepatitis C virus. Nat Rev Microbiol (2007) 5(6):453–63. doi:10.1038/nrmicro1645

141. Romero-Brey I, Merz A, Chiramel A, Lee JY, Chlanda P, Haselman U, et al. Three-dimensional architecture and biogenesis of membrane structures associated with hepatitis C virus replication. PLoS Pathog (2012) 8(12):e1003056. doi:10.1371/journal.ppat.1003056

142. Knoops K, Kikkert M, Worm SH, Zevenhoven-Dobbe JC, van der Meer Y, Koster AJ, et al. SARS-coronavirus replication is supported by a reticulovesicular network of modified endoplasmic reticulum. PLoS Biol (2008) 6(9):e226. doi:10.1371/journal.pbio.0060226

143. Li S, Kong L, Yu X. The expanding roles of endoplasmic reticulum stress in virus replication and pathogenesis. Crit Rev Microbiol (2015) 41(2):150–64. doi:10.3109/1040841X.2013.813899

144. Zhang L, Wang A. Virus-induced ER stress and the unfolded protein response. Front Plant Sci (2012) 3:293. doi:10.3389/fpls.2012.00293

145. Pena J, Harris E. Dengue virus modulates the unfolded protein response in a time-dependent manner. J Biol Chem (2011) 286(16):14226–36. doi:10.1074/jbc.M111.222703

146. Zheng Y, Gao B, Ye L, Kong L, Jing W, Yang X, et al. Hepatitis C virus non-structural protein NS4B can modulate an unfolded protein response. J Microbiol (2005) 43(6):529–36.

147. Chan SW, Egan PA. Hepatitis C virus envelope proteins regulate CHOP via induction of the unfolded protein response. FASEB J (2005) 19(11):1510–2. doi:10.1096/fj.04-3455fje

148. Lee DY, Sugden B. The LMP1 oncogene of EBV activates PERK and the unfolded protein response to drive its own synthesis. Blood (2008) 111(4):2280–9. doi:10.1182/blood-2007-07-100032

149. Li B, Gao B, Ye L, Han X, Wang W, Kong L, et al. Hepatitis B virus X protein (HBx) activates ATF6 and IRE1-XBP1 pathways of unfolded protein response. Virus Res (2007) 124(1–2):44–9. doi:10.1016/j.virusres.2006.09.011

150. Chan CP, Siu KL, Chin KT, Yuen KY, Zheng B, Jin DY. Modulation of the unfolded protein response by the severe acute respiratory syndrome coronavirus spike protein. J Virol (2006) 80(18):9279–87. doi:10.1128/JVI.80.7.3225-3237.2006

151. Roberson EC, Tully JE, Guala AS, Reiss JN, Godburn KE, Pociask DA, et al. Influenza induces endoplasmic reticulum stress, caspase-12-dependent apoptosis, and c-Jun N-terminal kinase-mediated transforming growth factor-beta release in lung epithelial cells. Am J Respir Cell Mol Biol (2012) 46(5):573–81. doi:10.1165/rcmb.2010-0460OC

152. Rathore AP, Ng ML, Vasudevan SG. Differential unfolded protein response during chikungunya and sindbis virus infection: CHIKV nsP4 suppresses eIF2alpha phosphorylation. Virol J (2013) 10:36. doi:10.1186/1743-422X-10-36

153. Pasqual G, Burri DJ, Pasquato A, de la Torre JC, Kunz S. Role of the host cell’s unfolded protein response in arenavirus infection. J Virol (2011) 85(4):1662–70. doi:10.1128/JVI.01782-10

154. He B, Gross M, Roizman B. The gamma(1)34.5 protein of herpes simplex virus 1 complexes with protein phosphatase 1alpha to dephosphorylate the alpha subunit of the eukaryotic translation initiation factor 2 and preclude the shutoff of protein synthesis by double-stranded RNA-activated protein kinase. Proc Natl Acad Sci U S A (1997) 94(3):843–8.

155. Cheng G, Brett ME, He B. Val193 and Phe195 of the gamma 1 34.5 protein of herpes simplex virus 1 are required for viral resistance to interferon-alpha/beta. Virology (2001) 290(1):115–20. doi:10.1006/viro.2001.1148

156. Zhang F, Moon A, Childs K, Goodbourn S, Dixon LK. The African swine fever virus DP71L protein recruits the protein phosphatase 1 catalytic subunit to dephosphorylate eIF2alpha and inhibits CHOP induction but is dispensable for these activities during virus infection. J Virol (2010) 84(20):10681–9. doi:10.1128/JVI.01027-10

157. Bhattacharyya S, Sen U, Vrati S. Regulated IRE1-dependent decay pathway is activated during Japanese encephalitis virus-induced unfolded protein response and benefits viral replication. J Gen Virol (2014) 95(Pt 1):71–9. doi:10.1099/vir.0.057265-0

158. Zhang P, Su C, Jiang Z, Zheng C. Herpes simplex virus 1 UL41 protein suppresses the IRE1/XBP1 signal pathway of the unfolded protein response via Its RNase activity. J Virol (2017) 91(4):e2056–2016. doi:10.1128/JVI.02056-16

159. Tardif KD, Mori K, Kaufman RJ, Siddiqui A. Hepatitis C virus suppresses the IRE1-XBP1 pathway of the unfolded protein response. J Biol Chem (2004) 279(17):17158–64. doi:10.1074/jbc.M312144200

160. Hassan IH, Zhang MS, Powers LS, Shao JQ, Baltrusaitis J, Rutkowski DT, et al. Influenza A viral replication is blocked by inhibition of the inositol-requiring enzyme 1 (IRE1) stress pathway. J Biol Chem (2012) 287(7):4679–89. doi:10.1074/jbc.M111.284695

161. Fraser JE, Wang C, Chan KW, Vasudevan SG, Jans DA. Novel dengue virus inhibitor 4-HPR activates ATF4 independent of protein kinase R-like endoplasmic reticulum kinase and elevates levels of eIF2alpha phosphorylation in virus infected cells. Antiviral Res (2016) 130:1–6. doi:10.1016/j.antiviral.2016.03.006

162. Medigeshi GR, Lancaster AM, Hirsch AJ, Briese T, Lipkin WI, Defilippis V, et al. West nile virus infection activates the unfolded protein response, leading to CHOP induction and apoptosis. J Virol (2007) 81(20):10849–60. doi:10.1128/JVI.01151-07

163. Jiang G, Santos Rocha C, Hirao LA, Mendes EA, Tang Y, Thompson GR III, et al. HIV exploits antiviral host innate GCN2-ATF4 signaling for establishing viral replication early in infection. MBio (2017) 8(3):e1518–1516. doi:10.1128/mBio.01518-16

164. Diwaker D, Mishra KP, Ganju L. Effect of modulation of unfolded protein response pathway on dengue virus infection. Acta Biochim Biophys Sin (Shanghai) (2015) 47(12):960–8. doi:10.1093/abbs/gmv108

165. Cheng YL, Lin YS, Chen CL, Tsai TT, Tsai CC, Wu YW, et al. Activation of Nrf2 by the dengue virus causes an increase in CLEC5A, which enhances TNF-alpha production by mononuclear phagocytes. Sci Rep (2016) 6:32000. doi:10.1038/srep32000

166. Ke PY, Chen SS. Activation of the unfolded protein response and autophagy after hepatitis C virus infection suppresses innate antiviral immunity in vitro. J Clin Invest (2011) 121(1):37–56. doi:10.1172/JCI41474

167. Wolfson JJ, May KL, Thorpe CM, Jandhyala DM, Paton JC, Paton AW. Subtilase cytotoxin activates PERK, IRE1 and ATF6 endoplasmic reticulum stress-signalling pathways. Cell Microbiol (2008) 10(9):1775–86. doi:10.1111/j.1462-5822.2008.01164.x

168. Paton AW, Srimanote P, Talbot UM, Wang H, Paton JC. A new family of potent AB(5) cytotoxins produced by Shiga toxigenic Escherichia coli. J Exp Med (2004) 200(1):35–46. doi:10.1084/jem.20040392

169. Harama D, Koyama K, Mukai M, Shimokawa N, Miyata M, Nakamura Y, et al. A subcytotoxic dose of subtilase cytotoxin prevents lipopolysaccharide-induced inflammatory responses, depending on its capacity to induce the unfolded protein response. J Immunol (2009) 183(2):1368–74. doi:10.4049/jimmunol.0804066

170. Lee SY, Lee MS, Cherla RP, Tesh VL. Shiga toxin 1 induces apoptosis through the endoplasmic reticulum stress response in human monocytic cells. Cell Microbiol (2008) 10(3):770–80. doi:10.1111/j.1462-5822.2007.01083.x

171. Pillich H, Loose M, Zimmer KP, Chakraborty T. Activation of the unfolded protein response by Listeria monocytogenes. Cell Microbiol (2012) 14(6):949–64. doi:10.1111/j.1462-5822.2012.01769.x

172. Gekara NO, Groebe L, Viegas N, Weiss S. Listeria monocytogenes desensitizes immune cells to subsequent Ca2+ signaling via listeriolysin O-induced depletion of intracellular Ca2+ stores. Infect Immun (2008) 76(2):857–62. doi:10.1128/IAI.00622-07

173. Cho JA, Lee AH, Platzer B, Cross BC, Gardner BM, De Luca H, et al. The unfolded protein response element IRE1alpha senses bacterial proteins invading the ER to activate RIG-I and innate immune signaling. Cell Host Microbe (2013) 13(5):558–69. doi:10.1016/j.chom.2013.03.011

174. de Jong MF, Starr T, Winter MG, den Hartigh AB, Child R, Knodler LA, et al. Sensing of bacterial type IV secretion via the unfolded protein response. MBio (2013) 4(1):e418–412. doi:10.1128/mBio.00418-12

175. Myeni S, Child R, Ng TW, Kupko JJ III, Wehrly TD, Porcella SF, et al. Brucella modulates secretory trafficking via multiple type IV secretion effector proteins. PLoS Pathog (2013) 9(8):e1003556. doi:10.1371/journal.ppat.1003556

176. Celli J, de Chastellier C, Franchini DM, Pizarro-Cerda J, Moreno E, Gorvel JP. Brucella evades macrophage killing via VirB-dependent sustained interactions with the endoplasmic reticulum. J Exp Med (2003) 198(4):545–56. doi:10.1084/jem.20030088

177. Tilney LG, Harb OS, Connelly PS, Robinson CG, Roy CR. How the parasitic bacterium Legionella pneumophila modifies its phagosome and transforms it into rough ER: implications for conversion of plasma membrane to the ER membrane. J Cell Sci (2001) 114(Pt 24):4637–50.

178. Derre I, Swiss R, Agaisse H. The lipid transfer protein CERT interacts with the Chlamydia inclusion protein IncD and participates to ER-Chlamydia inclusion membrane contact sites. PLoS Pathog (2011) 7(6):e1002092. doi:10.1371/journal.ppat.1002092

179. Shima K, Klinger M, Schütze S, Kaufhold I, Solbach W, Reiling N, et al. The role of endoplasmic reticulum-related BiP/GRP78 in interferon gamma-induced persistent Chlamydia pneumoniae infection. Cell Microbiol (2015) 17(7):923–34. doi:10.1111/cmi.12416

180. Webster SJ, Ellis L, O’Brien LM, Tyrrell B, Fitzmaurice TJ, Elder MJ, et al. IRE1alpha mediates PKR activation in response to Chlamydia trachomatis infection. Microbes Infect (2016) 18(7–8):472–83. doi:10.1016/j.micinf.2016.03.010

181. Hempstead AD, Isberg RR. Inhibition of host cell translation elongation by Legionella pneumophila blocks the host cell unfolded protein response. Proc Natl Acad Sci U S A (2015) 112(49):E6790–7. doi:10.1073/pnas.1508716112

182. Smith JA, Khan M, Magnani DD, Harms JS, Durward M, Radhakrishnan GK, et al. Brucella induces an unfolded protein response via TcpB that supports intracellular replication in macrophages. PLoS Pathog (2013) 9(12):e1003785. doi:10.1371/journal.ppat.1003785

183. Radhakrishnan GK, Harms JS, Splitter GA. Modulation of microtubule dynamics by a TIR domain protein from the intracellular pathogen Brucella melitensis. Biochem J (2011) 439(1):79–83. doi:10.1042/BJ20110577

184. Qin QM, Pei J, Ancona V, Shaw BD, Ficht TA, de Figueiredo P. RNAi screen of endoplasmic reticulum-associated host factors reveals a role for IRE1alpha in supporting Brucella replication. PLoS Pathog (2008) 4(7):e1000110. doi:10.1371/journal.ppat.1000110

185. Deegan S, Saveljeva S, Gorman AM, Samali A. Stress-induced self-cannibalism: on the regulation of autophagy by endoplasmic reticulum stress. Cell Mol Life Sci (2013) 70(14):2425–41. doi:10.1007/s00018-012-1173-4

186. Starr T, Child R, Wehrly TD, Hansen B, Hwang S, López-Otin C, et al. Selective subversion of autophagy complexes facilitates completion of the Brucella intracellular cycle. Cell Host Microbe (2012) 11(1):33–45. doi:10.1016/j.chom.2011.12.002

187. Byndloss MX, Tsolis RM. Brucella spp. Virulence factors and immunity. Annu Rev Anim Biosci (2016) 4:111–27. doi:10.1146/annurev-animal-021815-111326

188. Radhakrishnan GK, Yu Q, Harms JS, Splitter GA. Brucella TIR domain-containing protein mimics properties of the toll-like receptor adaptor protein TIRAP. J Biol Chem (2009) 284(15):9892–8. doi:10.1074/jbc.M805458200

189. Sag E, Bilginer Y, Ozen S. Autoinflammatory diseases with periodic fevers. Curr Rheumatol Rep (2017) 19(7):41. doi:10.1007/s11926-017-0670-8

190. Sherlock JP, Joyce-Shaikh B, Turner SP, Chao CC, Sathe M, Grein J, et al. IL-23 induces spondyloarthropathy by acting on ROR-gammat+ CD3+CD4-CD8- entheseal resident T cells. Nat Med (2012) 18(7):1069–76. doi:10.1038/nm.2817

191. Kontoyiannis D, Pasparakis M, Pizarro TT, Cominelli F, Kollias G. Impaired on/off regulation of TNF biosynthesis in mice lacking TNF AU-rich elements: implications for joint and gut-associated immunopathologies. Immunity (1999) 10(3):387–98. doi:10.1016/S1074-7613(00)80038-2

192. Hreggvidsdottir HS, Noordenbos T, Baeten DL. Inflammatory pathways in spondyloarthritis. Mol Immunol (2014) 57(1):28–37. doi:10.1016/j.molimm.2013.07.016

193. Keffer J, Probert L, Cazlaris H, Georgopoulos S, Kaslaris E, Kioussis D, et al. Transgenic mice expressing human tumour necrosis factor: a predictive genetic model of arthritis. EMBO J (1991) 10(13):4025–31.

194. Smith JA. Update on ankylosing spondylitis: current concepts in pathogenesis. Curr Allergy Asthma Rep (2015) 15(1):489. doi:10.1007/s11882-014-0489-6

195. Lees CW, Barrett JC, Parkes M, Satsangi J. New IBD genetics: common pathways with other diseases. Gut (2011) 60(12):1739–53. doi:10.1136/gut.2009.199679

196. Ramos PS, Criswell LA, Moser KL, Comeau ME, Williams AH, Pajewski NM, et al. A comprehensive analysis of shared loci between systemic lupus erythematosus (SLE) and sixteen autoimmune diseases reveals limited genetic overlap. PLoS Genet (2011) 7(12):e1002406. doi:10.1371/journal.pgen.1002406

197. Xavier RJ, Podolsky DK. Unravelling the pathogenesis of inflammatory bowel disease. Nature (2007) 448(7152):427–34. doi:10.1038/nature06005

198. Hugot JP, Chamaillard M, Zouali H, Lesage S, Cézard JP, Belaiche J, et al. Association of NOD2 leucine-rich repeat variants with susceptibility to Crohn’s disease. Nature (2001) 411(6837):599–603. doi:10.1038/35079107

199. Ogura Y, Bonen DK, Inohara N, Nicolae DL, Chen FF, Ramos R, et al. A frameshift mutation in NOD2 associated with susceptibility to Crohn’s disease. Nature (2001) 411(6837):603–6. doi:10.1038/35079114

200. Kaser A, Lee AH, Franke A, Glickman JN, Zeissig S, Tilg H, et al. XBP1 links ER stress to intestinal inflammation and confers genetic risk for human inflammatory bowel disease. Cell (2008) 134(5):743–56. doi:10.1016/j.cell.2008.07.021

201. Rioux JD, Xavier RJ, Taylor KD, Silverberg MS, Goyette P, Huett A, et al. Genome-wide association study identifies new susceptibility loci for Crohn’s disease and implicates autophagy in disease pathogenesis. Nat Genet (2007) 39(5):596–604. doi:10.1038/ng2032

202. Adolph TE, Tomczak MF, Niederreiter L, Ko HJ, Böck J, Martinez-Naves E, et al. Paneth cells as a site of origin for intestinal inflammation. Nature (2013) 503(7475):272–6. doi:10.1038/nature12599

203. Tschurtschenthaler M, Adolph TE, Ashcroft JW, Niederreiter L, Bharti R, Saveljeva S, et al. Defective ATG16L1-mediated removal of IRE1alpha drives Crohn’s disease-like ileitis. J Exp Med (2017) 214(2):401–22. doi:10.1084/jem.20160791

204. Heazlewood CK, Cook MC, Eri R, Price GR, Tauro SB, Taupin D, et al. Aberrant mucin assembly in mice causes endoplasmic reticulum stress and spontaneous inflammation resembling ulcerative colitis. PLoS Med (2008) 5(3):e54. doi:10.1371/journal.pmed.0050054

205. Wenzel UA, Jonstrand C, Hansson GC, Wick MJ. CD103+ CD11b+ dendritic cells induce Th17 T cells in Muc2-deficient mice with extensively spread colitis. PLoS One (2015) 10(6):e0130750. doi:10.1371/journal.pone.0130750

206. Barrett JC, Hansoul S, Nicolae DL, Cho JH, Duerr RH, Rioux JD, et al. Genome-wide association defines more than 30 distinct susceptibility loci for Crohn’s disease. Nat Genet (2008) 40(8):955–62. doi:10.1038/ng.175

207. Miller M, Tam AB, Cho JY, Doherty TA, Pham A, Khorram N, et al. ORMDL3 is an inducible lung epithelial gene regulating metalloproteases, chemokines, OAS, and ATF6. Proc Natl Acad Sci U S A (2012) 109(41):16648–53. doi:10.1073/pnas.1204151109

208. Cantero-Recasens G, Fandos C, Rubio-Moscardo F, Valverde MA, Vicente R. The asthma-associated ORMDL3 gene product regulates endoplasmic reticulum-mediated calcium signaling and cellular stress. Hum Mol Genet (2010) 19(1):111–21. doi:10.1093/hmg/ddp471

209. Zheng W, Rosenstiel P, Huse K, Sina C, Valentonyte R, Mah N, et al. Evaluation of AGR2 and AGR3 as candidate genes for inflammatory bowel disease. Genes Immun (2006) 7(1):11–8. doi:10.1038/sj.gene.6364263

210. Zhao F, Edwards R, Dizon D, Afrasiabi K, Mastroianni JR, Geyfman M, et al. Disruption of paneth and goblet cell homeostasis and increased endoplasmic reticulum stress in Agr2-/- mice. Dev Biol (2010) 338(2):270–9. doi:10.1016/j.ydbio.2009.12.008

211. Watkin LB, Jessen B, Wiszniewski W, Vece TJ, Jan M, Sha Y, et al. COPA mutations impair ER-Golgi transport and cause hereditary autoimmune-mediated lung disease and arthritis. Nat Genet (2015) 47(6):654–60. doi:10.1038/ng.3279

212. Vece TJ, Watkin LB, Nicholas S, Canter D, Braun MC, Guillerman RP, et al. Copa syndrome: a novel autosomal dominant immune dysregulatory disease. J Clin Immunol (2016) 36(4):377–87. doi:10.1007/s10875-016-0271-8

213. Patel DD, Kuchroo VK. Th17 cell pathway in human immunity: lessons from genetics and therapeutic interventions. Immunity (2015) 43(6):1040–51. doi:10.1016/j.immuni.2015.12.003

214. Volpi S, Tsui J, Mariani M, Pastorino C, Caorsi R, Sacco O, et al. Type I interferon pathway activation in COPA syndrome. Clin Immunol (2017). doi:10.1016/j.clim.2017.10.001

215. Lobito AA, Kimberley FC, Muppidi JR, Komarow H, Jackson AJ, Hull KM, et al. Abnormal disulfide-linked oligomerization results in ER retention and altered signaling by TNFR1 mutants in TNFR1-associated periodic fever syndrome (TRAPS). Blood (2006) 108(4):1320–7. doi:10.1182/blood-2005-11-006783

216. Bachetti T, Chiesa S, Castagnola P, Bani D, Di Zanni E, Omenetti A, et al. Autophagy contributes to inflammation in patients with TNFR-associated periodic syndrome (TRAPS). Ann Rheum Dis (2013) 72(6):1044–52. doi:10.1136/annrheumdis-2012-201952

217. Dickie LJ, Aziz AM, Savic S, Lucherini OM, Cantarini L, Geiler J, et al. Involvement of X-box binding protein 1 and reactive oxygen species pathways in the pathogenesis of tumour necrosis factor receptor-associated periodic syndrome. Ann Rheum Dis (2012) 71(12):2035–43. doi:10.1136/annrheumdis-2011-201197

218. Bulua AC, Simon A, Maddipati R, Pelletier M, Park H, Kim KY, et al. Mitochondrial reactive oxygen species promote production of proinflammatory cytokines and are elevated in TNFR1-associated periodic syndrome (TRAPS). J Exp Med (2011) 208(3):519–33. doi:10.1084/jem.20102049

219. Dougados M, Baeten D. Spondyloarthritis. Lancet (2011) 377(9783):2127–37. doi:10.1016/S0140-6736(11)60071-8

220. Reveille JD, Weisman MH. The epidemiology of back pain, axial spondyloarthritis and HLA-B27 in the United States. Am J Med Sci (2013) 345(6):431–6. doi:10.1097/MAJ.0b013e318294457f

221. Colbert RA, Tran TM, Layh-Schmitt G. HLA-B27 misfolding and ankylosing spondylitis. Mol Immunol (2014) 57(1):44–51. doi:10.1016/j.molimm.2013.07.013

222. Hanson A, Brown MA. Genetics and the causes of ankylosing spondylitis. Rheum Dis Clin North Am (2017) 43(3):401–14. doi:10.1016/j.rdc.2017.04.006

223. Robinson PC, Brown MA. Genetics of ankylosing spondylitis. Mol Immunol (2014) 57(1):2–11. doi:10.1016/j.molimm.2013.06.013

224. Mear JP, Schreiber KL, Münz C, Zhu X, Stevanović S, Rammensee HG, et al. Misfolding of HLA-B27 as a result of its B pocket suggests a novel mechanism for its role in susceptibility to spondyloarthropathies. J Immunol (1999) 163(12):6665–70.

225. Dangoria NS, DeLay ML, Kingsbury DJ, Mear JP, Uchanska-Ziegler B, Ziegler A, et al. HLA-B27 misfolding is associated with aberrant intermolecular disulfide bond formation (dimerization) in the endoplasmic reticulum. J Biol Chem (2002) 277(26):23459–68. doi:10.1074/jbc.M110336200

226. Tran TM, Satumtira N, Dorris ML, May E, Wang A, Furuta E, et al. HLA-B27 in transgenic rats forms disulfide-linked heavy chain oligomers and multimers that bind to the chaperone BiP. J Immunol (2004) 172(8):5110–9. doi:10.4049/jimmunol.172.8.5110

227. Guiliano DB, North H, Panayoitou E, Campbell EC, McHugh K, Cooke FG, et al. Polymorphisms in the F pocket of HLA-B27 subtypes strongly affect assembly, chaperone interactions, and heavy-chain misfolding. Arthritis Rheumatol (2017) 69(3):610–21. doi:10.1002/art.39948

228. García-Medel N, Sanz-Bravo A, Alvarez-Navarro C, Gómez-Molina P, Barnea E, Marcilla M, et al. Peptide handling by HLA-B27 subtypes influences their biological behavior, association with ankylosing spondylitis and susceptibility to endoplasmic reticulum aminopeptidase 1 (ERAP1). Mol Cell Proteomics (2014) 13(12):3367–80. doi:10.1074/mcp.M114.039214

229. Jeanty C, Sourisce A, Noteuil A, Jah N, Wielgosik A, Fert I, et al. HLA-B27 Subtype oligomerization and intracellular accumulation patterns correlate with predisposition to spondyloarthritis. Arthritis Rheumatol (2014) 66(8):2113–23. doi:10.1002/art.38644

230. Hammer RE, Maika SD, Richardson JA, Tang JP, Taurog JD. Spontaneous inflammatory disease in transgenic rats expressing HLA-B27 and human beta 2m: an animal model of HLA-B27-associated human disorders. Cell (1990) 63(5):1099–112. doi:10.1016/0092-8674(90)90512-D

231. Taurog JD, Maika SD, Simmons WA, Breban M, Hammer RE. Susceptibility to inflammatory disease in HLA-B27 transgenic rat lines correlates with the level of B27 expression. J Immunol (1993) 150(9):4168–78.

232. Taurog JD, Richardson JA, Croft JT, Simmons WA, Zhou M, Fernández-Sueiro JL, et al. The germfree state prevents development of gut and joint inflammatory disease in HLA-B27 transgenic rats. J Exp Med (1994) 180(6):2359–64. doi:10.1084/jem.180.6.2359

233. May E, Dorris ML, Satumtira N, Iqbal I, Rehman MI, Lightfoot E, et al. CD8 alpha beta T cells are not essential to the pathogenesis of arthritis or colitis in HLA-B27 transgenic rats. J Immunol (2003) 170(2):1099–105. doi:10.4049/jimmunol.170.2.1099

234. Turner MJ, Sowders DP, DeLay ML, Mohapatra R, Bai S, Smith JA, et al. HLA-B27 misfolding in transgenic rats is associated with activation of the unfolded protein response. J Immunol (2005) 175(4):2438–48. doi:10.4049/jimmunol.175.4.2438

235. Tran TM, Dorris ML, Satumtira N, Richardson JA, Hammer RE, Shang J, et al. Additional human beta2-microglobulin curbs HLA-B27 misfolding and promotes arthritis and spondylitis without colitis in male HLA-B27-transgenic rats. Arthritis Rheum (2006) 54(4):1317–27. doi:10.1002/art.21740

236. Turner MJ, Delay ML, Bai S, Klenk E, Colbert RA. HLA-B27 up-regulation causes accumulation of misfolded heavy chains and correlates with the magnitude of the unfolded protein response in transgenic rats: implications for the pathogenesis of spondylarthritis-like disease. Arthritis Rheum (2007) 56(1):215–23. doi:10.1002/art.22295

237. Bowness P, Ridley A, Shaw J, Chan AT, Wong-Baeza I, Fleming M, et al. Th17 cells expressing KIR3DL2+ and responsive to HLA-B27 homodimers are increased in ankylosing spondylitis. J Immunol (2011) 186(4):2672–80. doi:10.4049/jimmunol.1002653

238. Dong W, Zhang Y, Yan M, Liu H, Chen Z, Zhu P. Upregulation of 78-kDa glucose-regulated protein in macrophages in peripheral joints of active ankylosing spondylitis. Scand J Rheumatol (2008) 37(6):427–34. doi:10.1080/03009740802213310

239. Feng Y, Ding J, Fan CM, Zhu P. Interferon-gamma contributes to HLA-B27-associated unfolded protein response in spondyloarthropathies. J Rheumatol (2012) 39(3):574–82. doi:10.3899/jrheum.101257

240. Neerinckx B, Carter S, Lories RJ. No evidence for a critical role of the unfolded protein response in synovium and blood of patients with ankylosing spondylitis. Ann Rheum Dis (2014) 73(3):629–30. doi:10.1136/annrheumdis-2013-204170

241. Zeng L, Lindstrom MJ, Smith JA. Ankylosing spondylitis macrophage production of higher levels of interleukin-23 in response to lipopolysaccharide without induction of a significant unfolded protein response. Arthritis Rheum (2011) 63(12):3807–17. doi:10.1002/art.30593

242. Ciccia F, Accardo-Palumbo A, Rizzo A, Guggino G, Raimondo S, Giardina A, et al. Evidence that autophagy, but not the unfolded protein response, regulates the expression of IL-23 in the gut of patients with ankylosing spondylitis and subclinical gut inflammation. Ann Rheum Dis (2014) 73(8):1566–74. doi:10.1136/annrheumdis-2012-202925

243. Rehaume LM, Mondot S, Aguirre de Cárcer D, Velasco J, Benham H, Hasnain SZ, et al. ZAP-70 genotype disrupts the relationship between microbiota and host leading to spondyloarthritis and ileitis. Arthritis Rheumatol (2014) 66(10):2780–92. doi:10.1002/art.38773

244. Ruutu M, Thomas G, Steck R, Degli-Esposti MA, Zinkernagel MS, Alexander K, et al. beta-glucan triggers spondylarthritis and Crohn’s disease-like ileitis in SKG mice. Arthritis Rheum (2012) 64(7):2211–22. doi:10.1002/art.34423

245. Benham H, Rehaume LM, Hasnain SZ, Velasco J, Baillet AC, Ruutu M, et al. Interleukin-23 mediates the intestinal response to microbial beta-1,3-glucan and the development of spondyloarthritis pathology in SKG mice. Arthritis Rheumatol (2014) 66(7):1755–67. doi:10.1002/art.38638

246. Rayavarapu S, Coley W, Nagaraju K. An update on pathogenic mechanisms of inflammatory myopathies. Curr Opin Rheumatol (2011) 23(6):579–84. doi:10.1097/BOR.0b013e32834b41d2

247. Baechler EC, Bauer JW, Slattery CA, Ortmann WA, Espe KJ, Novitzke J, et al. An interferon signature in the peripheral blood of dermatomyositis patients is associated with disease activity. Mol Med (2007) 13(1–2):59–68. doi:10.2119/2006-00085.Baechler

248. Walsh RJ, Kong SW, Yao Y, Jallal B, Kiener PA, Pinkus JL, et al. Type I interferon-inducible gene expression in blood is present and reflects disease activity in dermatomyositis and polymyositis. Arthritis Rheum (2007) 56(11):3784–92. doi:10.1002/art.22928

249. Greenberg SA, Pinkus JL, Pinkus GS, Burleson T, Sanoudou D, Tawil R, et al. Interferon-alpha/beta-mediated innate immune mechanisms in dermatomyositis. Ann Neurol (2005) 57(5):664–78. doi:10.1002/ana.20464

250. Nagaraju K, Casciola-Rosen L, Lundberg I, Rawat R, Cutting S, Thapliyal R, et al. Activation of the endoplasmic reticulum stress response in autoimmune myositis: potential role in muscle fiber damage and dysfunction. Arthritis Rheum (2005) 52(6):1824–35. doi:10.1002/art.21103

251. Vattemi G, Engel WK, McFerrin J, Askanas V. Endoplasmic reticulum stress and unfolded protein response in inclusion body myositis muscle. Am J Pathol (2004) 164(1):1–7. doi:10.1016/S0002-9440(10)63089-1

252. Emslie-Smith AM, Arahata K, Engel AG. Major histocompatibility complex class I antigen expression, immunolocalization of interferon subtypes, and T cell-mediated cytotoxicity in myopathies. Hum Pathol (1989) 20(3):224–31. doi:10.1016/0046-8177(89)90128-7

253. Nagaraju K, Raben N, Loeffler L, Parker T, Rochon PJ, Lee E, et al. Conditional up-regulation of MHC class I in skeletal muscle leads to self-sustaining autoimmune myositis and myositis-specific autoantibodies. Proc Natl Acad Sci U S A (2000) 97(16):9209–14. doi:10.1073/pnas.97.16.9209

254. Li CK, Knopp P, Moncrieffe H, Singh B, Shah S, Nagaraju K, et al. Overexpression of MHC class I heavy chain protein in young skeletal muscle leads to severe myositis: implications for juvenile myositis. Am J Pathol (2009) 175(3):1030–40. doi:10.2353/ajpath.2009.090196

255. Psarras A, Emery P, Vital EM. Type I interferon-mediated autoimmune diseases: pathogenesis, diagnosis and targeted therapy. Rheumatology (Oxford) (2017) 56(10):1662–75. doi:10.1093/rheumatology/kew431

256. Bennett L, Palucka AK, Arce E, Cantrell V, Borvak J, Banchereau J, et al. Interferon and granulopoiesis signatures in systemic lupus erythematosus blood. J Exp Med (2003) 197(6):711–23. doi:10.1084/jem.20021553

257. Wang J, Cheng Q, Wang X, Zu B, Xu J, Xu Y, et al. Deficiency of IRE1 and PERK signal pathways in systemic lupus erythematosus. Am J Med Sci (2014) 348(6):465–73. doi:10.1097/MAJ.0000000000000328

258. Lee WS, Sung MS, Lee EG, Yoo HG, Cheon YH, Chae HJ, et al. A pathogenic role for ER stress-induced autophagy and ER chaperone GRP78/BiP in T lymphocyte systemic lupus erythematosus. J Leukoc Biol (2015) 97(2):425–33. doi:10.1189/jlb.6A0214-097R

259. Zhang H, Zhao C, Wang S, Huang Y, Wang H, Zhao J, et al. Anti-dsDNA antibodies induce inflammation via endoplasmic reticulum stress in human mesangial cells. J Transl Med (2015) 13:178. doi:10.1186/s12967-015-0536-7

260. Lenna S, Farina AG, Martyanov V, Christmann RB, Wood TA, Farber HW, et al. Increased expression of endoplasmic reticulum stress and unfolded protein response genes in peripheral blood mononuclear cells from patients with limited cutaneous systemic sclerosis and pulmonary arterial hypertension. Arthritis Rheum (2013) 65(5):1357–66. doi:10.1002/art.37891

261. Furue M, Mitoma C, Mitoma H, Tsuji G, Chiba T, Nakahara T, et al. Pathogenesis of systemic sclerosis-current concept and emerging treatments. Immunol Res (2017) 65(4):790–7. doi:10.1007/s12026-017-8926-y

262. Baek HA, Kim DS, Park HS, Jang KY, Kang MJ, Lee DG, et al. Involvement of endoplasmic reticulum stress in myofibroblastic differentiation of lung fibroblasts. Am J Respir Cell Mol Biol (2012) 46(6):731–9. doi:10.1165/rcmb.2011-0121OC

263. Heindryckx F, Binet F, Ponticos M, Rombouts K, Lau J, Kreuger J, et al. Endoplasmic reticulum stress enhances fibrosis through IRE1alpha-mediated degradation of miR-150 and XBP-1 splicing. EMBO Mol Med (2016) 8(7):729–44. doi:10.15252/emmm.201505925

264. Jacobson DL, Gange SJ, Rose NR, Graham NM. Epidemiology and estimated population burden of selected autoimmune diseases in the United States. Clin Immunol Immunopathol (1997) 84(3):223–43. doi:10.1006/clin.1997.4412

265. Harris JE. Cellular stress and innate inflammation in organ-specific autoimmunity: lessons learned from vitiligo. Immunol Rev (2016) 269(1):11–25. doi:10.1111/imr.12369

266. Schuit FC, In’t Veld PA, Pipeleers DG. Glucose stimulates proinsulin biosynthesis by a dose-dependent recruitment of pancreatic beta cells. Proc Natl Acad Sci U S A (1988) 85(11):3865–9. doi:10.1073/pnas.85.11.3865

267. Rashighi M, Agarwal P, Richmond JM, Harris TH, Dresser K, Su MW, et al. CXCL10 is critical for the progression and maintenance of depigmentation in a mouse model of vitiligo. Sci Transl Med (2014) 6(223):223ra23. doi:10.1126/scitranslmed.3007811

268. Padgett LE, Broniowska KA, Hansen PA, Corbett JA, Tse HM. The role of reactive oxygen species and proinflammatory cytokines in type 1 diabetes pathogenesis. Ann N Y Acad Sci (2013) 1281:16–35. doi:10.1111/j.1749-6632.2012.06826.x

269. Frigerio S, Junt T, Lu B, Gerard C, Zumsteg U, Holländer GA, et al. Beta cells are responsible for CXCR3-mediated T-cell infiltration in insulitis. Nat Med (2002) 8(12):1414–20. doi:10.1038/nm1202-792

270. Hassler JR, Scheuner DL, Wang S, Han J, Kodali VK, Li P, et al. The IRE1alpha/XBP1s pathway is essential for the glucose response and protection of beta cells. PLoS Biol (2015) 13(10):e1002277. doi:10.1371/journal.pbio.1002277

271. Engin F, Yermalovich A, Nguyen T, Hummasti S, Fu W, Eizirik DL, et al. Restoration of the unfolded protein response in pancreatic beta cells protects mice against type 1 diabetes. Sci Transl Med (2013) 5(211):211ra156. doi:10.1126/scitranslmed.3006534

272. Marhfour I, Lopez XM, Lefkaditis D, Salmon I, Allagnat F, Richardson SJ, et al. Expression of endoplasmic reticulum stress markers in the islets of patients with type 1 diabetes. Diabetologia (2012) 55(9):2417–20. doi:10.1007/s00125-012-2604-3

273. Allagnat F, Fukaya M, Nogueira TC, Delaroche D, Welsh N, Marselli L, et al. C/EBP homologous protein contributes to cytokine-induced pro-inflammatory responses and apoptosis in beta-cells. Cell Death Differ (2012) 19(11):1836–46. doi:10.1038/cdd.2012.67

274. Brozzi F, Nardelli TR, Lopes M, Millard I, Barthson J, Igoillo-Esteve M, et al. Cytokines induce endoplasmic reticulum stress in human, rat and mouse beta cells via different mechanisms. Diabetologia (2015) 58(10):2307–16. doi:10.1007/s00125-015-3669-6

275. Støy J, Edghill EL, Flanagan SE, Ye H, Paz VP, Pluzhnikov A, et al. Insulin gene mutations as a cause of permanent neonatal diabetes. Proc Natl Acad Sci U S A (2007) 104(38):15040–4. doi:10.1073/pnas.0707291104

276. Liu M, Hodish I, Haataja L, Lara-Lemus R, Rajpal G, Wright J, et al. Proinsulin misfolding and diabetes: mutant INS gene-induced diabetes of youth. Trends Endocrinol Metab (2010) 21(11):652–9. doi:10.1016/j.tem.2010.07.001

277. Oyadomari S, Koizumi A, Takeda K, Gotoh T, Akira S, Araki E, et al. Targeted disruption of the chop gene delays endoplasmic reticulum stress-mediated diabetes. J Clin Invest (2002) 109(4):525–32. doi:10.1172/JCI0214550

278. Zhou R, Tardivel A, Thorens B, Choi I, Tschopp J. Thioredoxin-interacting protein links oxidative stress to inflammasome activation. Nat Immunol (2010) 11(2):136–40. doi:10.1038/ni.1831

279. Wang J, Song MY, Lee JY, Kwon KS, Park BH. The NLRP3 inflammasome is dispensable for ER stress-induced pancreatic beta-cell damage in Akita mice. Biochem Biophys Res Commun (2015) 466(3):300–5. doi:10.1016/j.bbrc.2015.09.009

280. Morita S, Villalta SA, Feldman HC, Register AC, Rosenthal W, Hoffmann-Petersen IT, et al. Targeting ABL-IRE1alpha signaling spares ER-stressed pancreatic beta cells to reverse autoimmune diabetes. Cell Metab (2017) 25(5):1207. doi:10.1016/j.cmet.2017.04.026

281. Lee KP, Dey M, Neculai D, Cao C, Dever TE, Sicheri F. Structure of the dual enzyme Ire1 reveals the basis for catalysis and regulation in nonconventional RNA splicing. Cell (2008) 132(1):89–100. doi:10.1016/j.cell.2007.10.057

282. Harding HP, Zhang Y, Ron D. Protein translation and folding are coupled by an endoplasmic-reticulum-resident kinase. Nature (1999) 397(6716):271–4. doi:10.1038/16729

283. Neubert K, Meister S, Moser K, Weisel F, Maseda D, Amann K, et al. The proteasome inhibitor bortezomib depletes plasma cells and protects mice with lupus-like disease from nephritis. Nat Med (2008) 14(7):748–55. doi:10.1038/nm1763

284. Ichikawa HT, Conley T, Muchamuel T, Jiang J, Lee S, Owen T, et al. Beneficial effect of novel proteasome inhibitors in murine lupus via dual inhibition of type I interferon and autoantibody-secreting cells. Arthritis Rheum (2012) 64(2):493–503. doi:10.1002/art.33333

285. Sung C, Wei Y, Watanabe S, Lee HS, Khoo YM, Fan L, et al. Extended evaluation of virological, immunological and pharmacokinetic endpoints of CELADEN: a randomized, placebo-controlled trial of celgosivir in dengue fever patients. PLoS Negl Trop Dis (2016) 10(8):e0004851. doi:10.1371/journal.pntd.0004851

Conflict of Interest Statement: The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Smith. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

REVIEW
published: 16 May 2018
doi: 10.3389/fimmu.2018.01075

Metabolic Reprogramming in Modulating T Cell Reactive Oxygen Species Generation and Antioxidant Capacity

Josephin N. Rashida Gnanaprakasam, Ruohan Wu and Ruoning Wang*

Center for Childhood Cancer & Blood Diseases, Hematology/Oncology & BMT, The Research Institute at Nationwide Children’s Hospital, Ohio State University, Columbus, OH, United States

OPEN ACCESS

Edited by:

Heitor Affonso Paula Neto, Universidade Federal do Rio de Janeiro, Brazil

Reviewed by:

Martin Rottenberg, Karolinska Institutet (KI), Sweden
Pedro Barcellos-de-Souza, Instituto Nacional de Câncer (INCA), Brazil

*Correspondence:

Ruoning Wang
ruoning.wang@nationwidechildrens.org

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 16 December 2017
Accepted: 30 April 2018
Published: 16 May 2018

Citation:

Rashida Gnanaprakasam JN, Wu R and Wang R (2018) Metabolic Reprogramming in Modulating T Cell Reactive Oxygen Species Generation and Antioxidant Capacity. Front. Immunol. 9:1075. doi: 10.3389/fimmu.2018.01075

A robust adaptive immune response requires a phase of proliferative burst which is followed by the polarization of T cells into relevant functional subsets. Both processes are associated with dramatically increased bioenergetics demands, biosynthetic demands, and redox demands. T cells meet these demands by rewiring their central metabolic pathways that generate energy and biosynthetic precursors by catabolizing and oxidizing nutrients into carbon dioxide. Simultaneously, oxidative metabolism also produces reactive oxygen species (ROS), which are tightly controlled by antioxidants and plays important role in regulating T cell functions. In this review, we discuss how metabolic rewiring during T cell activation influence ROS production and antioxidant capacity.

Keywords: reactive oxygen species, oxidative stress, metabolism, T cell, antioxidant

INTRODUCTION

T cells are central orchestrators of antigen-specific adaptive immunity and tolerance. Upon stimulation of antigen receptors, T cells rapidly transit from naïve to an active state followed by massive clonal expansion. Depending on the nature of pathogens and the surrounding cytokine milieu, proliferating T cells can differentiate into diverse phenotypic and functional subsets to elicit a robust immune response. After the clearance of pathogens, the majority of effector T cells die through apoptosis and the remaining memory T (Tmem) cells are responsible for immunity upon re-exposure to the same pathogen. Accumulating evidence suggests that a coordinated rewiring of cellular metabolism is required for T cell activation and differentiation by fulfilling the bioenergetic, biosynthetic, and redox demands (1–9). Importantly, different phenotypic and functional T cell subsets are characterized by distinct metabolic programs (Table 1), which are largely controlled by immune modulatory signaling cascades (10–17). Naïve T (Tnai) cells, Tmem cells, and immune-suppressive regulatory T (Treg) cells predominantly rely on fatty acid oxidation (FAO) and oxidative phosphorylation (OXPHOS) to meet their relatively low-energy needs (14, 15, 18, 19). However, persistent aerobic glycolysis, the pentose phosphate pathway (PPP), and glutaminolysis are required to drive cell growth, clonal expansion, and effector functions in both CD4+ subsets and CD8+ effector T (Teff) cells (Table 1) (10, 15, 16, 18, 20–31).

TABLE 1 | The metabolic profiles of T cell subsets.

These metabolic programs actively support ATP production by providing mitochondrial OXPHOS substrates, support biomass accumulation by generating metabolic precursors for the biosynthesis of protein, lipids, and nucleic acids, and maintain redox balance through generation and elimination of reactive oxygen species (ROS).

MITOCHONDRIAL OXPHOS AND NADPH OXIDASES (NOXs) IN GENERATING ROS IN T CELLS

The mitochondria are the central metabolic hub and powerhouse of all eukaryotic cells. The oxidation of acetyl-CoA to carbon dioxide (CO_2) by the tricarboxylic acid (TCA) cycle is the central metabolic process for fueling ATP production. While glycolysis and FAO primarily provide the OXPHOX substrate, acetyl-CoA, for mitochondria in Tnai cells, Tmem cells, and Treg cells (14, 15, 18, 19), heightened mitochondrial biogenesis during T cell activation leads to higher numbers of mitochondria and likely the enhanced mitochondrial dependent metabolic flux in Teff cells compared with Tnai cells (23, 32, 33). In particular, a surplus of 3-, 4-, and 5-carbon metabolites (anaplerotic substrates) including pyruvate, malate, and α-ketoglutarate (α-KG) feed into the TCA cycle during the catabolism of glutamine and other amino acids (5, 13, 15, 34). The electron transport chain (ETC) constantly transfers electrons from NADH and FADH2 to oxygen while allowing protons (H+) to pass through the inner mitochondrial membrane to form an electrochemical proton gradient that drives ATP synthesis. However, both protons and electrons can leak from the ETC due to the uncoupling of ATP synthase from the proton gradient and a premature exit of electron before reaching cytochrome c oxidase, respectively. Electron leak largely occurs at the sites of complex I (NADH–ubiquinone oxidoreductase) and complex III (ubiquinone–cytochrome c oxidoreductase) in the ETC and results in the partial reduction of oxygen, generating superoxide (). Subsequently, mitochondrial dismutase acts to convert superoxide to hydrogen peroxide (H_2O_2), which is free to diffuse into cytosol and act as a redox signaling molecule to elicit different cellular responses (35–37). Thus, increased ROS production in T cells can occur as a result of metabolic reprogramming during T cell activation. Besides mitochondria, cytoplasmic ROS is generated by NOXs, which is also an important source of ROS in T cell. NOX family proteins are transmembrane proteins that transport the electrons from nicotinamide adenine dinucleotide (phosphate), NAD(P)H, to oxygen and generate superoxide anion as the intermediate product of oxidase and subsequently H_2O_2, as the product of dismutation of the superoxide. There are different isoforms of the NOX enzyme including NOX1, NOX2, NOX3, NOX4, NOX5, dual oxidase 1, and DUOX2, and the expression of these subunits varies among different tissues. NOX-2 is an important source of ROS in T cells (38, 39). The ROS production by NOX is regulated at various levels including the assembly of functional NOX complex, the availability of prosthetic group, flavin adenine dinucleotide, the intracellular concentration of calcium, cell surface receptor signals mediated by G protein-coupled receptors, complement, T cell receptor (TCR), and CD28 (35–37, 40, 41).

ROS SIGNALING IN REGULATING T CELL ACTIVATION AND DIFFERENTIATION

T cell activation requires ligation of TCR and the major histocompatibility complex molecules. This interaction will initiate the signaling cascade and activation of transcriptional factors such as nuclear factor of activated T cells (NFAT), activator protein 1 (AP-1), and nuclear factor of kappa light chain enhancer in B cells (42). It has been reported that TCR ligation increases the production of ROS from OXPHOS and cytoplasmic ROS from NADPH oxidases (NOXs), a family of plasma membrane-associated oxidases (36, 40, 41). ROS-mediated signaling events are required for driving T cell activation, proliferation, and differentiation (Figure 1) (36, 41). T cells with reduced production of mitochondrial ROS display impaired production of interleukin-2 and antigen-specific proliferation, indicating an essential signaling role for mitochondrial ROS in driving optimal TCR signaling. The proximal TCR signaling machinery, including zeta chain-associated protein kinase 70, linker of activated T cell, SH2 domain-containing leukocyte protein, phospholipase Cγ1, and protein kinase Cθ, is involved in driving ROS production upon T cell activation (36, 41, 43). Conversely, physiologically relevant levels of ROS facilitate the activation of oxidation-dependent transcription factors, such as NF-κB and AP-1, which are required for driving essential signaling events to support T cell-mediated immune responses (44–46). However, excessive ROS production following ablation of de novo glutathione (GSH) synthesis suppresses the activity of mammalian target of rapamycin and the expression of transcription factors NFAT and c-MYC, the latter of which control metabolic reprogramming following T cell activation (15, 47, 48). Thus, T cells fail to meet their increased energy and biosynthetic needs and display compromised proliferation (48). In addition, uncontrolled ROS production is involved in the activation-induced T-cell death by affecting expression of apoptosis related genes including Bcl-2 and FasL and mitochondrial membrane potential (43, 49–52). NOX-derived ROS modulates the function of GATA-binding protein 3, signal transducer and activator of transcription, and T-box transcription factor to collectively control T cell activation and differentiation. T cells from NOX-deficient animals showed a skewed Th17 phenotype, whereas NOX-intact cells exhibited a preferred Th1 response (39, 53–55). In CD8 T cells, NOX-derived ROS is involved in regulating the production of IFN-γ and CD39 expression through c-Jun N-terminal kinase and NFκB signaling (40, 56). Importantly, the impact of ROS on T cell activation can be extended to the later T cell differentiation stages. Fine tuning of ROS is required for polarizing T cell in part by engaging lineage-specific transcription factors and modulating cytokine profiles, and consequently directs T cell-mediated inflammatory responses (39, 40, 53–55, 57–61).

FIGURE 1 | Mitochondria and NADPH oxidases (NOX)-derived reactive oxygen species (ROS) regulates T cell activation, differentiation, and metabolism. Mitochondria and NOX are the two major sources of ROS. The stimulation of T cell receptor (TCR) initiates signaling and metabolic events that drive ROS production in cytoplasm through NOX-dependent reaction and ROS production in mitochondria via mitochondria electron transport chain (ETC). Excess ROS causes damage and cell death. However, physiologically relevant levels of ROS mediate essential redox signaling through nodulation of a wide spectrum of redox-sensitive transcription factors to drive T cell activation and function.

METABOLIC PATHWAYS IN MODULATING ANTIOXIDANT CAPACITIES

Excessive ROS production causes collateral damage to macromolecules, cellular organelles, and eventually necrosis, which can lead to uncontrolled hyper-inflammation and tissue damage. Thus, a fine-tuned balance between ROS production and antioxidant capacity ensures appropriate levels of intracellular ROS (Figure 2) (44, 55, 62). GSH, a tripeptide of glutamine, cysteine, and glycine, is the most abundant antioxidant capable of providing reducing equivalents and also serves as a versatile nucleophilic cofactor in a wide spectrum of metabolic reactions in aerobic organisms (63, 64). Thioredoxin (TXN) is a class of small redox proteins that are involved in modulating cell surface receptors and confers tolerance to oxidative stress in T cells (65–69). A reciprocal redox reaction can be coupled between these two antioxidant systems to act as a backup for each other under certain conditions (70–77). Supporting these findings, the inhibition of thioredoxin reductase (TXNRD) conferred an increased susceptibility of cancer cells to GSH depletion (78–80). Glutathione-disulfide reductase (GSR) regenerates GSH from its oxidized form, glutathione disulfide (GSSG), whereas TXNRD is responsible for the regeneration of TXN once it has been oxidized. Importantly, both GSR and TXNRD require NADPH as a reducing agent. Upon antigen stimulation, both PPP and glutaminolysis are significantly upregulated and further enhance T cell antioxidant capacities by generating NADPH through metabolic reactions that are controlled by glucose-6-phosphate dehydrogenase, phosphoglycerate dehydrogenase, malic enzyme 1, and isocitrate dehydrogenase 1. The intracellular GSH concentrations are normally in a range of three orders of magnitude higher than extracellular GSH. Even though some cells are able to recycle extracellular GSH, it may only play a minor role in maintaining intracellular GSH pool (63, 64, 81–86). By contrast, both the regeneration of GSH from GSSG (recycling pathway) and de novo synthesis of GSH, by glutamate-cysteine ligase (GCL) and glutathione synthase (GS), are required to maintain intracellular GSH levels (64, 87). The ligation of glutamate and cysteine to form dipeptide ϒ-glutamylcysteine (ϒ-GC) is the first and also the rate-limiting step of GSH de novo synthesis, which is controlled by ATP-dependent ligase GCL, a heterodimer of a catalytic subunit (GCLC) and modifier subunit (GCLM). Subsequently, GSH is formed by GS-mediated ligation of ϒ-GC and glycine (88, 89). Thus, the supply of intracellular cysteine, glycine, and glutamate must fulfill the need of de novo synthesis of GSH during T cell activation. Supporting this idea, the metabolic processes that are involved in providing three amino acids are tightly regulated upon T cell activation (13, 15, 90–92). Upon T cell activation, heightened glycolysis, PPP, and glutaminolysis intersect with the de novo synthesis of GSH through promoting cysteine uptake and providing glycine, glutamine, and NADPH (93–95). As such, the genetic abrogation of de novo synthesis of GSH, the glucose, or glutamine starvation significantly dampens T cell activation (10, 13, 15, 20, 48).

FIGURE 2 | T cell metabolic programs that link to reactive oxygen species (ROS) production and the de novo synthesis of GSH. Pyruvate that is derived from glucose via glycolysis is shuttled to the mitochondria and drives the tricarboxylic acid (TCA) cycle and fuels oxidative phosphorylation (OXPHOS). Glucose-derived glucose-6-phosphate feeds into the pentose phosphate pathway (PPP) and produces NADPH in the cytoplasm. In addition, glutamate feeds the TCA cycle through α-ketoglutarate (α-KG) to fuel OXPHOS and generate ROS. Excessive ROS production is regulated by glutathione (GSH), a tripeptide of glutamine, cysteine, and glycine, which is synthesized de novo by glutamate-cysteine ligase (GCL) and glutathione synthetase (GSS). In addition, NADPH, glutathione-disulfide reductase (GSR), and glutathione peroxidase (GPX) are involved in regenerating GSH from glutathione disulfide (GSSG), whereas thioredoxin reductase (TXNRD) is responsible for the regeneration of thioredoxin (TXN) to control oxidative stress in T cell.

GLUTAMINE CATABOLISM IN COORDINATING THE PRODUCTION OF ROS AND GSH

Glutamine has been known as a key nutrient, which supports a diverse range of cellular functions (93–102). Glutamine provides high proportions of the energy from OXPHOS, provides precursors for various biosynthetic pathways, as a key nitrogen and carbon donor, and also is catabolized to various intermediate metabolites that have signaling roles in modulating cellular processes. In specialized cells, such as the cells of the nervous system, glutamine catabolism intersects with signaling networks to support the production of central neurotransmitters including glutamate, GABA, and aspartate (103–106). To meet bioenergetic and biosynthetic demand during T cell growth and proliferation, glutaminolysis replenishes the anapleurotic substrate α-KG that fuels OXPHOS via the TCA cycle and also provides sources of nitrogen and carbon to support the biosynthesis of nonessential amino acids, lipids, nucleotides, and polyamines (13, 15, 102, 107). Similar to cancer cells, de novo synthesis of GSH in T cells, which relies on glutamine to provide precursors, plays an essential role in suppressing oxidative stress. Accordingly, glutaminolysis is a branched pathway that consists of several paths, enabling energy production through oxidation and biomolecule production, including GSH through biosynthesis (93–95). While the ATP generating capacity of glutaminolysis is considered to be redundant with glucose oxidation and/or FAO, the oxidation of glutamine is indispensable for driving T cell proliferation and differentiation (13, 15, 102). However, enhanced glutamine oxidation in the mitochondria also increases the production of its by-product, mitochondrial ROS, the main source of cellular ROS in T cells (35, 37). Therefore, glutamate represents a key branch point in glutaminolysis that can be committed toward mitochondrial oxidation to produce ATP and ROS, or toward de novo synthesis of GSH to modulate redox balance and suppress oxidative stress. In addition, the high rate of glutaminolysis ensures that the capacity to supply glutamate, the most abundant intracellular metabolite in cells, exceeds the demand for glutamate from each of the downstream metabolic branches. The branched pathways in glutaminolysis enable the production of counteracting metabolites, i.e., ROS and GSH, from a common metabolic precursor, and permit a fine-tuned coordination between the metabolic flux shunted toward GSH synthesis and the metabolic flux shunted toward OXPHOS. Consistent with this idea, the overall high consumption rate of glutamine in proliferative cells is suggested to provide a sensitive and precise regulation on intermediate metabolites that can be committed toward several metabolic branches, hence permitting rapid responses to meet the demands for energy production or antioxidant production (99, 108). In addition to increasing antioxidant capacity, T cells may adapt by shifting glucose catabolism from OXPHOS toward aerobic glycolysis, which could provide biosynthetic precursors and rapidly produce ATP by the substrate level of phosphorylation.

CONCLUSION AND PERSPECTIVE

Reactive oxygen species is not only a by-product of cellular metabolic programs but also a key signaling molecule involved in directing T cell activation and differentiation. However, uncontrolled ROS production causes collateral damage to biomolecules and cellular organelles. Under pathophysiological conditions, ROS generation from mitochondria can contribute to the initiation and progression of inflammatory and autoimmune diseases. However, oxidative stress caused by elevated ROS may also render key immune effector cells vulnerable to agents that can either modulate stress response or modulate metabolic pathways for ROS and GSH production (Figure 3). Redox signaling is essential to regulate T cell metabolism. Technological advancement in genetic models and metabolomics will allow us to understand the key metabolic processes that dictate T cell fate through modulation ROS and GSH production. Thus, further research is expected to illustrate the complex interplay between cellular metabolism and redox signaling in T cells, thereby offering novel therapies for treating inflammatory and autoimmune diseases.

FIGURE 3 | Cellular redox homeostasis is essential for mounting an effective T cell-mediated immune response. In addition to generate ATP and provide biosynthetic precursors, T cell activation-induced metabolic reprogramming actively regulates redox homeostasis. The coordination of de novo synthesis of glutathione (GSH) and the production of reactive oxygen species (ROS) ensures T cell redox balance and a fine-tuned T cell response.

AUTHOR CONTRIBUTIONS

JG and RHW wrote the manuscript. RW wrote and finalized the manuscript.

FUNDING

This work was supported by R21AI117547 and 1R01AI114581 from National Institutes of Health, V2014-001 from the V-Foundation and 128436-RSG-15-180-01-LIB from the American Cancer Society (RW).

REFERENCES

1. Finlay D, Cantrell DA. Metabolism, migration and memory in cytotoxic T cells. Nat Rev Immunol (2011) 11:109–17. doi:10.1038/nri2888

2. Wang R, Green DR. Metabolic checkpoints in activated T cells. Nat Immunol (2012) 13:907–15. doi:10.1038/ni.2386

3. Pearce EL, Pearce EJ. Metabolic pathways in immune cell activation and quiescence. Immunity (2013) 38:633–43. doi:10.1016/j.immuni.2013.04.005

4. Weinberg SE, Sena LA, Chandel NS. Mitochondria in the regulation of innate and adaptive immunity. Immunity (2015) 42:406–17. doi:10.1016/j.immuni.2015.02.002

5. O’Neill LA, Kishton RJ, Rathmell J. A guide to immunometabolism for immunologists. Nat Rev Immunol (2016) 16:553–65. doi:10.1038/nri.2016.70

6. Buck MD, Sowell RT, Kaech SM, Pearce EL. Metabolic instruction of immunity. Cell (2017) 169:570–86. doi:10.1016/j.cell.2017.04.004

7. Ma EH, Poffenberger MC, Wong AH, Jones RG. The role of AMPK in T cell metabolism and function. Curr Opin Immunol (2017) 46:45–52. doi:10.1016/j.coi.2017.04.004

8. Patel CH, Powell JD. Targeting T cell metabolism to regulate T cell activation, differentiation and function in disease. Curr Opin Immunol (2017) 46:82–8. doi:10.1016/j.coi.2017.04.006

9. Zeng H, Chi H. mTOR signaling in the differentiation and function of regulatory and effector T cells. Curr Opin Immunol (2017) 46:103–11. doi:10.1016/j.coi.2017.04.005

10. Frauwirth KA, Riley JL, Harris MH, Parry RV, Rathmell JC, Plas DR, et al. The CD28 signaling pathway regulates glucose metabolism. Immunity (2002) 16:769–77. doi:10.1016/S1074-7613(02)00323-0

11. Bensinger SJ, Bradley MN, Joseph SB, Zelcer N, Janssen EM, Hausner MA, et al. LXR signaling couples sterol metabolism to proliferation in the acquired immune response. Cell (2008) 134:97–111. doi:10.1016/j.cell.2008.04.052

12. Wofford JA, Wieman HL, Jacobs SR, Zhao Y, Rathmell JC. IL-7 promotes Glut1 trafficking and glucose uptake via STAT5-mediated activation of Akt to support T-cell survival. Blood (2008) 111:2101–11. doi:10.1182/blood-2007-06-096297

13. Carr EL, Kelman A, Wu GS, Gopaul R, Senkevitch E, Aghvanyan A, et al. Glutamine uptake and metabolism are coordinately regulated by ERK/MAPK during T lymphocyte activation. J Immunol (2011) 185:1037–44. doi:10.4049/jimmunol.0903586

14. Michalek RD, Gerriets VA, Jacobs SR, Macintyre AN, Maciver NJ, Mason EF, et al. Cutting edge: distinct glycolytic and lipid oxidative metabolic programs are essential for effector and regulatory CD4+ T cell subsets. J Immunol (2011) 186:3299–303. doi:10.4049/jimmunol.1003613

15. Wang R, Dillon CP, Shi LZ, Milasta S, Carter R, Finkelstein D, et al. The transcription factor Myc controls metabolic reprogramming upon T lymphocyte activation. Immunity (2011) 35:871–82. doi:10.1016/j.immuni.2011.09.021

16. Finlay DK, Rosenzweig E, Sinclair LV, Feijoo-Carnero C, Hukelmann JL, Rolf J, et al. PDK1 regulation of mTOR and hypoxia-inducible factor 1 integrate metabolism and migration of CD8+ T cells. J Exp Med (2012) 209:2441–53. doi:10.1084/jem.20112607

17. Kidani Y, Elsaesser H, Hock MB, Vergnes L, Williams KJ, Argus JP, et al. Sterol regulatory element-binding proteins are essential for the metabolic programming of effector T cells and adaptive immunity. Nat Immunol (2013) 14:489–99. doi:10.1038/ni.2570

18. Pearce EL, Walsh MC, Cejas PJ, Harms GM, Shen H, Wang LS, et al. Enhancing CD8 T-cell memory by modulating fatty acid metabolism. Nature (2009) 460:103–7. doi:10.1038/nature08097

19. Shi LZ, Wang R, Huang G, Vogel P, Neale G, Green DR, et al. HIF1alpha-dependent glycolytic pathway orchestrates a metabolic checkpoint for the differentiation of TH17 and Treg cells. J Exp Med (2011) 208:1367–76. doi:10.1084/jem.20110278

20. Rathmell JC, Farkash EA, Gao W, Thompson CB. IL-7 enhances the survival and maintains the size of naive T cells. J Immunol (2001) 167:6869–76. doi:10.4049/jimmunol.167.12.6869

21. Jacobs SR, Herman CE, Maciver NJ, Wofford JA, Wieman HL, Hammen JJ, et al. Glucose uptake is limiting in T cell activation and requires CD28-mediated Akt-dependent and independent pathways. J Immunol (2008) 180:4476–86. doi:10.4049/jimmunol.180.7.4476

22. Gerriets VA, Rathmell JC. Metabolic pathways in T cell fate and function. Trends Immunol (2012) 33:168–73. doi:10.1016/j.it.2012.01.010

23. van der Windt GJ, Everts B, Chang CH, Curtis JD, Freitas TC, Amiel E, et al. Mitochondrial respiratory capacity is a critical regulator of CD8(+) T cell memory development. Immunity (2012) 36(1):68–78. doi:10.1016/j.immuni.2011.12.007

24. Kolev M, Dimeloe S, Le Friec G, Navarini A, Arbore G, Povoleri GA, et al. Complement regulates nutrient influx and metabolic reprogramming during Th1 cell responses. Immunity (2015) 42:1033–47. doi:10.1016/j.immuni.2015.05.024

25. Patsoukis N, Bardhan K, Chatterjee P, Sari D, Liu B, Bell LN, et al. PD-1 alters T-cell metabolic reprogramming by inhibiting glycolysis and promoting lipolysis and fatty acid oxidation. Nat Commun (2015) 6:6692. doi:10.1038/ncomms7692

26. Ray JP, Staron MM, Shyer JA, Ho P-C, Marshall HD, Gray SM, et al. The interleukin-2-mTORc1 kinase axis defines the signaling, differentiation, and metabolism of T helper 1 and follicular B helper T cells. Immunity (2015) 43:690–702. doi:10.1016/j.immuni.2015.08.017

27. Angela M, Endo Y, Asou HK, Yamamoto T, Tumes DJ, Tokuyama H, et al. Fatty acid metabolic reprogramming via mTOR-mediated inductions of PPARgamma directs early activation of T cells. Nat Commun (2016) 7:13683. doi:10.1038/ncomms13683

28. Buck MD, O’Sullivan D, Klein Geltink RI, Curtis JD, Chang CH, Sanin DE, et al. Mitochondrial dynamics controls T cell fate through metabolic programming. Cell (2016) 166:63–76. doi:10.1016/j.cell.2016.05.035

29. Wang Y, Bi Y, Chen X, Li C, Li Y, Zhang Z, et al. Histone deacetylase SIRT1 negatively regulates the differentiation of interleukin-9-producing CD4+ T cells. Immunity (2016) 44:1337–49. doi:10.1016/j.immuni.2016.05.009

30. Beckermann KE, Dudzinski SO, Rathmell JC. Dysfunctional T cell metabolism in the tumor microenvironment. Cytokine Growth Factor Rev (2017) 35:7–14. doi:10.1016/j.cytogfr.2017.04.003

31. Binger KJ, Côrte-Real BF, Kleinewietfeld M. Immunometabolic regulation of interleukin-17-producing T helper cells: uncoupling new targets for autoimmunity. Front Immunol (2017) 8:311. doi:10.3389/fimmu.2017.00311

32. D’Souza AD, Parikh N, Kaech SM, Shadel GS. Convergence of multiple signaling pathways is required to coordinately up-regulate mtDNA and mitochondrial biogenesis during T cell activation. Mitochondrion (2007) 7:374–85. doi:10.1016/j.mito.2007.08.001

33. Ron-Harel N, Santos D, Ghergurovich JM, Sage PT, Reddy A, Lovitch SB, et al. Mitochondrial biogenesis and proteome remodeling promote one-carbon metabolism for T cell activation. Cell Metab (2016) 24:104–17. doi:10.1016/j.cmet.2016.06.007

34. Geiger R, Rieckmann JC, Wolf T, Basso C, Feng Y, Fuhrer T, et al. L-arginine modulates T cell metabolism and enhances survival and anti-tumor activity. Cell (2016) 167:829–842.e13. doi:10.1016/j.cell.2016.09.031

35. Murphy MP. How mitochondria produce reactive oxygen species. Biochem J (2009) 417:1–13. doi:10.1042/BJ20081386

36. Kaminski MM, Sauer SW, Kaminski M, Opp S, Ruppert T, Grigaravicius P, et al. T cell activation is driven by an ADP-dependent glucokinase linking enhanced glycolysis with mitochondrial reactive oxygen species generation. Cell Rep (2012) 2:1300–15. doi:10.1016/j.celrep.2012.10.009

37. Schieber M, Chandel NS. ROS function in redox signaling and oxidative stress. Curr Biol (2014) 24:R453–62. doi:10.1016/j.cub.2014.03.034

38. Bedard K, Krause K-H. The NOX family of ROS-generating NADPH oxidases: physiology and pathophysiology. Physiol Rev (2007) 87:245–313. doi:10.1152/physrev.00044.2005

39. Hubert MT, Thayer TC, Steele C, Cuda CM, Morel L, Piganelli JD, et al. NADPH oxidase deficiency regulates Th lineage commitment and modulates autoimmunity. J Immunol (2010) 185:5247–58. doi:10.4049/jimmunol.1001472

40. Jackson SH, Devadas S, Kwon J, Pinto LA, Williams MS. T cells express a phagocyte-type NADPH oxidase that is activated after T cell receptor stimulation. Nat Immunol (2004) 5:818–27. doi:10.1038/ni1096

41. Sena LA, Li S, Jairaman A, Prakriya M, Ezponda T, Hildeman DA, et al. Mitochondria are required for antigen-specific T cell activation through reactive oxygen species signaling. Immunity (2013) 38:225–36. doi:10.1016/j.immuni.2012.10.020

42. Smith-Garvin JE, Koretzky GA, Jordan MS. T cell activation. Annu Rev Immunol (2009) 27:591–619. doi:10.1146/annurev.immunol.021908.132706

43. Kamiński M, Kießling M, Süss D, Krammer PH, Gülow K. Novel role for mitochondria: protein kinase Cθ-dependent oxidative signaling organelles in activation-induced T-cell death. Mol Cell Biol (2007) 27:3625–39. doi:10.1128/MCB.02295-06

44. Devadas S, Zaritskaya L, Rhee SG, Oberley L, Williams MS. Discrete generation of superoxide and hydrogen peroxide by T cell receptor stimulation. J Exp Med (2002) 195:59–70. doi:10.1084/jem.20010659

45. Kaminski MM, Roth D, Krammer PH, Gulow K. Mitochondria as oxidative signaling organelles in T-cell activation: physiological role and pathological implications. Arch Immunol Ther Exp (Warsz) (2013) 61:367–84. doi:10.1007/s00005-013-0235-0

46. Murphy MP, Siegel RM. Mitochondrial ROS fire up T cell activation. Immunity (2013) 38:201–2. doi:10.1016/j.immuni.2013.02.005

47. Klein Geltink RI, O’Sullivan D, Pearce EL. Caught in the cROSsfire: GSH controls T cell metabolic reprogramming. Immunity (2017) 46:525–7. doi:10.1016/j.immuni.2017.03.022

48. Mak TW, Grusdat M, Duncan GS, Dostert C, Nonnenmacher Y, Cox M, et al. Glutathione primes T cell metabolism for inflammation. Immunity (2017) 46:675–89. doi:10.1016/j.immuni.2017.03.019

49. Hildeman DA, Mitchell T, Teague TK, Henson P, Day BJ, Kappler J, et al. Reactive oxygen species regulate activation-induced T cell apoptosis. Immunity (1999) 10:735–44. doi:10.1016/S1074-7613(00)80072-2

50. Li-Weber M, Weigand MA, Giaisi M, Süss D, Treiber MK, Baumann S, et al. Vitamin E inhibits CD95 ligand expression and protects T cells from activation-induced cell death. J Clin Invest (2002) 110:681. doi:10.1172/JCI0215073

51. Hildeman DA. Regulation of T-cell apoptosis by reactive oxygen species. Free Radic Biol Med (2004) 36:1496–504. doi:10.1016/j.freeradbiomed.2004.03.023

52. Takahashi A, Hanson MG, Norell HR, Havelka AM, Kono K, Malmberg K-J, et al. Preferential cell death of CD8+ effector memory (CCR7− CD45RA−) T cells by hydrogen peroxide-induced oxidative stress. J Immunol (2005) 174:6080–7. doi:10.4049/jimmunol.174.10.6080

53. Purushothaman D, Sarin A. Cytokine-dependent regulation of NADPH oxidase activity and the consequences for activated T cell homeostasis. J Exp Med (2009) 206:1515–23. doi:10.1084/jem.20082851

54. Shatynski KE, Chen H, Kwon J, Williams MS. Decreased STAT5 phosphorylation and GATA-3 expression in NOX2-deficient T cells: role in T helper development. Eur J Immunol (2012) 42:3202–11. doi:10.1002/eji.201242659

55. Belikov AV, Schraven B, Simeoni L. T cells and reactive oxygen species. J Biomed Sci (2015) 22:85. doi:10.1186/s12929-015-0194-3

56. Bai A, Moss A, Rothweiler S, Longhi MS, Wu Y, Junger WG, et al. NADH oxidase-dependent CD39 expression by CD8+ T cells modulates interferon gamma responses via generation of adenosine. Nat Commun (2015) 6:8819. doi:10.1038/ncomms9819

57. Zhi L, Ustyugova IV, Chen X, Zhang Q, Wu MX. Enhanced Th17 differentiation and aggravated arthritis in IEX-1-deficient mice by mitochondrial reactive oxygen species-mediated signaling. J Immunol (2012) 189:1639–47. doi:10.4049/jimmunol.1200528

58. Gerriets VA, Kishton RJ, Nichols AG, Macintyre AN, Inoue M, Ilkayeva O, et al. Metabolic programming and PDHK1 control CD4+ T cell subsets and inflammation. J Clin Invest (2015) 125:194–207. doi:10.1172/JCI76012

59. Abimannan T, Peroumal D, Parida JR, Barik PK, Padhan P, Devadas S. Oxidative stress modulates the cytokine response of differentiated Th17 and Th1 cells. Free Radic Biol Med (2016) 99:352–63. doi:10.1016/j.freeradbiomed.2016.08.026

60. Padgett LE, Tse HM. NADPH oxidase-derived superoxide provides a third signal for CD4 T cell effector responses. J Immunol (2016) 197:1733–42. doi:10.4049/jimmunol.1502581

61. Xu T, Stewart KM, Wang X, Liu K, Xie M, Kyu Ryu J, et al. Metabolic control of TH17 and induced Treg cell balance by an epigenetic mechanism. Nature (2017) 548:228–33. doi:10.1038/nature23475

62. Sena LA, Chandel NS. Physiological roles of mitochondrial reactive oxygen species. Mol Cell (2012) 48:158–67. doi:10.1016/j.molcel.2012.09.025

63. Kosower NS, Kosower EM. The glutathione status of cells. Int Rev Cytol (1978) 54:109–60. doi:10.1016/S0074-7696(08)60166-7

64. Meister A. Metabolism and function of glutathione: an overview. Biochem Soc Trans (1982) 10:78–9. doi:10.1042/bst0100078

65. Tagaya Y, Wakasugi H, Masutani H, Nakamura H, Iwata S, Mitsui A, et al. Role of ATL-derived factor (ADF) in the normal and abnormal cellular activation: involvement of dithiol related reduction. Mol Immunol (1990) 27:1279–89. doi:10.1016/0161-5890(90)90032-U

66. Wakasugi N, Tagaya Y, Wakasugi H, Mitsui A, Maeda M, Yodoi J, et al. Adult T-cell leukemia-derived factor/thioredoxin, produced by both human T-lymphotropic virus type I- and Epstein-Barr virus-transformed lymphocytes, acts as an autocrine growth factor and synergizes with interleukin 1 and interleukin 2. Proc Natl Acad Sci U S A (1990) 87:8282–6. doi:10.1073/pnas.87.21.8282

67. Rosen A, Lundman P, Carlsson M, Bhavani K, Srinivasa BR, Kjellstrom G, et al. A CD4+ T cell line-secreted factor, growth promoting for normal and leukemic B cells, identified as thioredoxin. Int Immunol (1995) 7:625–33. doi:10.1093/intimm/7.4.625

68. Matthias LJ, Yam PT, Jiang XM, Vandegraaff N, Li P, Poumbourios P, et al. Disulfide exchange in domain 2 of CD4 is required for entry of HIV-1. Nat Immunol (2002) 3:727–32. doi:10.1038/ni815

69. Mougiakakos D, Johansson CC, Jitschin R, Bottcher M, Kiessling R. Increased thioredoxin-1 production in human naturally occurring regulatory T cells confers enhanced tolerance to oxidative stress. Blood (2011) 117:857–61. doi:10.1182/blood-2010-09-307041

70. Mustacich D, Powis G. Thioredoxin reductase. Biochem J (2000) 346(Pt 1):1–8. doi:10.1042/0264-6021:3460001

71. Johansson C, Lillig CH, Holmgren A. Human mitochondrial glutaredoxin reduces S-glutathionylated proteins with high affinity accepting electrons from either glutathione or thioredoxin reductase. J Biol Chem (2004) 279:7537–43. doi:10.1074/jbc.M312719200

72. Hanschmann EM, Lonn ME, Schutte LD, Funke M, Godoy JR, Eitner S, et al. Both thioredoxin 2 and glutaredoxin 2 contribute to the reduction of the mitochondrial 2-Cys peroxiredoxin Prx3. J Biol Chem (2010) 285:40699–705. doi:10.1074/jbc.M110.185827

73. Tan SX, Greetham D, Raeth S, Grant CM, Dawes IW, Perrone GG. The thioredoxin-thioredoxin reductase system can function in vivo as an alternative system to reduce oxidized glutathione in Saccharomyces cerevisiae. J Biol Chem (2010) 285:6118–26. doi:10.1074/jbc.M109.062844

74. Du Y, Zhang H, Lu J, Holmgren A. Glutathione and glutaredoxin act as a backup of human thioredoxin reductase 1 to reduce thioredoxin 1 preventing cell death by aurothioglucose. J Biol Chem (2012) 287:38210–9. doi:10.1074/jbc.M112.392225

75. Lu J, Holmgren A. The thioredoxin antioxidant system. Free Radic Biol Med (2014) 66:75–87. doi:10.1016/j.freeradbiomed.2013.07.036

76. Couto N, Wood J, Barber J. The role of glutathione reductase and related enzymes on cellular redox homoeostasis network. Free Radic Biol Med (2016) 95:27–42. doi:10.1016/j.freeradbiomed.2016.02.028

77. Lei XG, Zhu JH, Cheng WH, Bao Y, Ho YS, Reddi AR, et al. Paradoxical roles of antioxidant enzymes: basic mechanisms and health implications. Physiol Rev (2016) 96:307–64. doi:10.1152/physrev.00010.2014

78. Lu J, Chew EH, Holmgren A. Targeting thioredoxin reductase is a basis for cancer therapy by arsenic trioxide. Proc Natl Acad Sci U S A (2007) 104:12288–93. doi:10.1073/pnas.0701549104

79. Mandal PK, Schneider M, Kolle P, Kuhlencordt P, Forster H, Beck H, et al. Loss of thioredoxin reductase 1 renders tumors highly susceptible to pharmacologic glutathione deprivation. Cancer Res (2010) 70:9505–14. doi:10.1158/0008-5472.CAN-10-1509

80. Wang Y, Lu H, Wang D, Li S, Sun K, Wan X, et al. Inhibition of glutathione synthesis eliminates the adaptive response of ascitic hepatoma 22 cells to nedaplatin that targets thioredoxin reductase. Toxicol Appl Pharmacol (2012) 265:342–50. doi:10.1016/j.taap.2012.09.001

81. Hwang C, Sinskey AJ, Lodish HF. Oxidized redox state of glutathione in the endoplasmic reticulum. Science (1992) 257:1496–502. doi:10.1126/science.1523409

82. Ganguly D, Srikanth CV, Kumar C, Vats P, Bachhawat AK. Why is glutathione (a tripeptide) synthesized by specific enzymes while TSH releasing hormone (TRH or thyroliberin), also a tripeptide, is produced as part of a prohormone protein? IUBMB Life (2003) 55:553–4. doi:10.1080/15216540310001623064

83. Perrone GG, Grant CM, Dawes IW. Genetic and environmental factors influencing glutathione homeostasis in Saccharomyces cerevisiae. Mol Biol Cell (2005) 16:218–30. doi:10.1091/mbc.E04-07-0560

84. Bennett BD, Kimball EH, Gao M, Osterhout R, Van Dien SJ, Rabinowitz JD. Absolute metabolite concentrations and implied enzyme active site occupancy in Escherichia coli. Nat Chem Biol (2009) 5:593–9. doi:10.1038/nchembio.186

85. Morgan B, Sobotta MC, Dick TP. Measuring E(GSH) and H2O2 with roGFP2-based redox probes. Free Radic Biol Med (2011) 51:1943–51. doi:10.1016/j.freeradbiomed.2011.08.035

86. Park JO, Rubin SA, Xu YF, Amador-Noguez D, Fan J, Shlomi T, et al. Metabolite concentrations, fluxes and free energies imply efficient enzyme usage. Nat Chem Biol (2016) 12:482–9. doi:10.1038/nchembio.2077

87. Lu SC. Regulation of glutathione synthesis. Mol Aspects Med (2009) 30:42–59. doi:10.1016/j.mam.2008.05.005

88. Chen Y, Shertzer HG, Schneider SN, Nebert DW, Dalton TP. Glutamate cysteine ligase catalysis: dependence on ATP and modifier subunit for regulation of tissue glutathione levels. J Biol Chem (2005) 280:33766–74. doi:10.1074/jbc.M504604200

89. Franklin CC, Backos DS, Mohar I, White CC, Forman HJ, Kavanagh TJ. Structure, function, and post-translational regulation of the catalytic and modifier subunits of glutamate cysteine ligase. Mol Aspects Med (2009) 30:86–98. doi:10.1016/j.mam.2008.08.009

90. Angelini G, Gardella S, Ardy M, Ciriolo MR, Filomeni G, Di Trapani G, et al. Antigen-presenting dendritic cells provide the reducing extracellular microenvironment required for T lymphocyte activation. Proc Natl Acad Sci U S A (2002) 99:1491–6. doi:10.1073/pnas.022630299

91. Levring TB, Hansen AK, Nielsen BL, Kongsbak M, Von Essen MR, Woetmann A, et al. Activated human CD4+ T cells express transporters for both cysteine and cystine. Sci Rep (2012) 2:266. doi:10.1038/srep00266

92. Ma EH, Bantug G, Griss T, Condotta S, Johnson RM, Samborska B, et al. Serine is an essential metabolite for effector T cell expansion. Cell Metab (2017) 25:345–57. doi:10.1016/j.cmet.2017.01.014

93. Gorrini C, Harris IS, Mak TW. Modulation of oxidative stress as an anticancer strategy. Nat Rev Drug Discov (2013) 12:931–47. doi:10.1038/nrd4002

94. Hensley CT, Wasti AT, Deberardinis RJ. Glutamine and cancer: cell biology, physiology, and clinical opportunities. J Clin Invest (2013) 123:3678–84. doi:10.1172/JCI69600

95. Altman BJ, Stine ZE, Dang CV. From Krebs to clinic: glutamine metabolism to cancer therapy. Nat Rev Cancer (2016) 16:619–34. doi:10.1038/nrc.2016.71

96. Kvamme E, Svenneby G. Effect of anaerobiosis and addition of keto acids on glutamine utilization by Ehrlich ascites-tumor cells. Biochim Biophys Acta (1960) 42:187–8. doi:10.1016/0006-3002(60)90779-4

97. Lund P. Glutamine metabolism in the rat. FEBS Lett (1980) 117:K86–92. doi:10.1016/0014-5793(80)80573-4

98. Kovacevic Z, McGivan JD. Mitochondrial metabolism of glutamine and glutamate and its physiological significance. Physiol Rev (1983) 63:547–605. doi:10.1152/physrev.1983.63.2.547

99. Newsholme EA, Crabtree B, Ardawi MS. Glutamine metabolism in lymphocytes: its biochemical, physiological and clinical importance. Q J Exp Physiol (1985) 70:473–89. doi:10.1113/expphysiol.1985.sp002935

100. DeBerardinis RJ, Mancuso A, Daikhin E, Nissim I, Yudkoff M, Wehrli S, et al. Beyond aerobic glycolysis: transformed cells can engage in glutamine metabolism that exceeds the requirement for protein and nucleotide synthesis. Proc Natl Acad Sci U S A (2007) 104:19345–50. doi:10.1073/pnas.0709747104

101. Tannahill GM, Curtis AM, Adamik J, Palsson-Mcdermott EM, Mcgettrick AF, Goel G, et al. Succinate is an inflammatory signal that induces IL-1beta through HIF-1alpha. Nature (2013) 496:238–42. doi:10.1038/nature11986

102. Klysz D, Tai X, Robert PA, Craveiro M, Cretenet G, Oburoglu L, et al. Glutamine-dependent alpha-ketoglutarate production regulates the balance between T helper 1 cell and regulatory T cell generation. Sci Signal (2015) 8:ra97. doi:10.1126/scisignal.aab2610

103. Daikhin Y, Yudkoff M. Compartmentation of brain glutamate metabolism in neurons and glia. J Nutr (2000) 130:1026S–31S. doi:10.1093/jn/130.4.1026S

104. Bak LK, Schousboe A, Waagepetersen HS. The glutamate/GABA-glutamine cycle: aspects of transport, neurotransmitter homeostasis and ammonia transfer. J Neurochem (2006) 98:641–53. doi:10.1111/j.1471-4159.2006.03913.x

105. Belanger M, Allaman I, Magistretti PJ. Brain energy metabolism: focus on astrocyte-neuron metabolic cooperation. Cell Metab (2011) 14:724–38. doi:10.1016/j.cmet.2011.08.016

106. Hertz L. The glutamate-glutamine (GABA) cycle: importance of late postnatal development and potential reciprocal interactions between biosynthesis and degradation. Front Endocrinol (2013) 4:59. doi:10.3389/fendo.2013.00059

107. Wang R, Green DR. Metabolic reprogramming and metabolic dependency in T cells. Immunol Rev (2012) 249:14–26. doi:10.1111/j.1600-065X.2012.01155.x

108. Newsholme EA, Crabtree B, Ardawi MS. The role of high rates of glycolysis and glutamine utilization in rapidly dividing cells. Biosci Rep (1985) 5:393–400. doi:10.1007/BF01116556

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Rashida Gnanaprakasam, Wu and Wang. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

REVIEW
published: 23 May 2018
doi: 10.3389/fimmu.2018.00935

Autophagy and Its Interaction With Intracellular Bacterial Pathogens

Mariana da Silva Siqueira, Renato de Moraes Ribeiro and Leonardo H. Travassos*

Laboratory of Immunoreceptors and Signaling, Immunobiology Program, Institute of Biophysics Carlos Chagas Filho, Federal University of Rio de Janeiro, Rio de Janeiro, Brazil

OPEN ACCESS

Edited by:

Thomas A. Kufer, University of Hohenheim, Germany

Reviewed by:

Klaus Ruckdeschel, Universitätsklinikum Hamburg-Eppendorf, Germany
Serge Mostowy, Imperial College London, United Kingdom

*Correspondence:

Leonardo H. Travassos
leo.travassos@biof.ufrj.br

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 20 January 2018
Accepted: 16 April 2018
Published: 23 May 2018

Citation:

Siqueira MS, Ribeiro RM and Travassos LH (2018) Autophagy and Its Interaction With Intracellular Bacterial Pathogens. Front. Immunol. 9:935. doi: 10.3389/fimmu.2018.00935

Cellular responses to stress can be defined by the overwhelming number of changes that cells go through upon contact with and stressful conditions such as infection and modifications in nutritional status. One of the main cellular responses to stress is autophagy. Much progress has been made in the understanding of the mechanisms involved in the induction of autophagy during infection by intracellular bacteria. This review aims to discuss recent findings on the role of autophagy as a cellular response to intracellular bacterial pathogens such as, Streptococcus pyogenes, Mycobacterium tuberculosis, Shigella flexneri, Salmonella typhimurium, Listeria monocytogenes, and Legionella pneumophila, how the autophagic machinery senses these bacteria directly or indirectly (through the detection of bacteria-induced nutritional stress), and how some of these bacterial pathogens manage to escape from autophagy.

Keywords: autophagy, infection, Mycobacterium tuberculosis, Shigella flexneri, Listeria monocytogenes, Salmonella typhimurium, Streptococcus pyogenes, Legionella pneumophila

INTRODUCTION

Autophagy is a homeostatic and highly conserved survival mechanism in which portions of the cytoplasm such as long-lived proteins and damaged organelles are sequestered in double-membrane vesicles (called autophagosomes). Then, autophagosomes fuse with lysosomes, leading to the degradation of the sequestered content and recycling of functional blocks for anabolic processes, especially during nutrient shortages (1). Indeed, for many years, autophagy was mainly considered as a breakdown process to degrade macromolecules to generate energy during nutrient deprivation. To date, three types of autophagy have been described, chaperone-mediated autophagy, microautophagy, and macroautophagy (1). Here, we discuss the interaction of the best-characterized type of autophagy (macroautophagy, hereafter autophagy), with intracellular bacterial pathogens, a process designated xenophagy.

The first report demonstrating induction of autophagy by bacteria was published in 1984. In this study, Rikihisa described the presence of vesicles containing glycogen granules and rickettsiae in Guinea pig polymorphonuclear (PMNs) cells infected with the bacteria (2). Despite this initial study, it was only after the studies from Nakagawa et al. and Gutierrez et al. that autophagy was regarded as an important cell autonomous arm of the innate immune system against intracellular bacteria. In their seminal and independent studies, Nakagawa et al. and Gutierrez et al. demonstrated a crucial role for autophagy in the sequestration and degradation of group A Streptococcus (GAS) and Mycobacterium bovis BCG, respectively (3, 4). Since then, an amazing number of elegant studies have demonstrated a key role of autophagy in the control of infection by different bacterial pathogens and also how some of these most well-succeeded pathogens circumvent or even use autophagy to establish replicative niches inside different cell types (5–7).

The Autophagosome Formation Core Machinery

Possibly one of the most exciting areas in the field of autophagy, the mechanisms involved in the formation of autophagosomes, the hallmark of this process, have been the focus of many research groups. Morphologically, autophagy begins with the formation of a cup-shaped double-membrane structure that surrounds the cargo. Upon its complete closure, the phagophore is now called an autophagosome, a transient organelle that delivers its content for degradation in lysosomes (8). After extensive work from several groups, the proteins that participate in autophagosome biogenesis can be categorized into complexes that take place in different steps of the autophagosome formation (1). Below, we will summarize the different steps of the autophagic process and the major protein groups that take part in each step of the whole process and discuss critical findings linking these proteins with bacterial-induced autophagy. For extensive literature on autophagosome formation machinery, please refer to Suzuki et al. (9) and Yin et al. (10).

Signal Induction

The ULK Complex and Autophagy Induction

The uncoordinated-51-like kinase (ULK1) complex comprising ULK1, ATG13, FAK family kinase-interacting protein of 200 kDa (FIP200), and ATG101 is responsible for sensing changes in nutrient status within the cell. Its activation is instrumental in the initiation of autophagy. This complex works downstream mammalian target of rapamycin complex 1 (mTORC1) and under, nutrient-rich conditions is phosphorylated by mTOR, which inhibits ULK1 recruitment to the phagophore assembly site (PAS).

Under nutrient starvation, however, mTORC1 is inactivated, and ULK1 is released, allowing FIP200 phosphorylation and translocation of the complex to PAS for the recruitment of ATG proteins, required for autophagosome formation (11). Interestingly, components of the ULK complex have also been shown to target bacterial vacuoles during infection with intracellular bacteria (12). This is the case of FIP200 during infection with Salmonella typhimurium. Experiments performed by Kageyama et al. suggest that this protein is recruited to the vicinity of vacuoles containing S. typhimurium. See below for more detailed information regarding autophagy induced by this pathogen.

Nucleation

Class III Phosphatidylinositol 3-Kinase (PtdIns3K) Complex and Trafficking of Atg9 for Autophagosome Nucleation

The class III PtdIns3K complex consisting of Beclin 1, ATG14L, phosphoinositide 3-kinase regulatory subunit 4 (PIK3R4) are recruited to PAS to initiate phagophore membrane nucleation through the activation of PtdIns3-kinase class III (PtdIns3KC3). As a result, PtdIns3P is generated at this site, and the PtdIns3P-binding protein WD-repeat domain phosphoinositide-interacting 1 (WIPI1) and 2 (WIPI2) are recruited to the PAS, allowing ATG proteins to be recruited later on (13). Mammalian Atg9 (mAtg9) is another protein required for the assembly of phagophore, although its role is still not completely understood. It has been demonstrated that mAtg9 is not necessary for LC3 recruitment to phagophore, but essential for its generation following infection with Salmonella typhimurium (12).

Expansion

Ubiquitin-Like Conjugation Systems and Autophagosome Expansion

Pivotal for the formation of autophagosomes are two ubiquitin-like conjugation systems: Atg8/LC3 and Atg12. The Atg8/LC3 system modifies the core autophagy protein microtubule-associated 1 light chain 3 (LC3). LC3 has a diffuse cytosolic distribution pattern and is cleaved at its C-terminus by the cysteine protease Atg4 to form LC3-I, which has a C-terminal glycine residue. Upon autophagy induction, LC3-I is sequentially modified by the E1-like enzyme Atg7 and the E2-like enzyme Atg3 to form LC3-II after the conjugation of LC3-I to phosphatidylethanolamine (PE). This lipidated form of LC3 is attached to both outer and inner phagophore membrane being eventually removed from the autophagosomal membrane by Atg4 before the fusion with late endosomes/lysosomes (1, 14). In the Atg12 conjugation system, Atg5 and Atg12 proteins form a complex through the covalent binding of Atg12 to the C-terminus of Atg5 in a reaction involving Atg7 and Atg10. Then, the scaffold protein Atg16L1 is conjugated to Atg5 via its N-terminus, forming the 800 kDa Atg12–Atg5–Atg16L1 complex. It has been proposed that the Atg16L1 complex works as an E3-like enzyme to target LC3-I to its membrane site of lipid conjugation (15). Data from the literature suggest that these two systems work coordinately as in Atg3-deficient cells, where no LC3-II is found, Atg12–Atg5 conjugation is dramatically reduced (16).

Alternative (non-canonical) forms of autophagy have been identified and reported to target invading bacteria (17–19). In this review, however, we will focus on xenophagy and its implication in intracellular bacterial infections.

Cargo Selection During Infection With Bacterial Pathogens

Invasion of host cytosol by bacteria imposes a significant challenge to homeostasis and triggers several cellular and immune responses such as proinflammatory cascades and cell-autonomous in an attempt to control of bacterial replication, such as xenophagy.

In addition to the steps discussed above, autophagy has an additional and essential step that is cargo selection. One of the central questions regarding xenophagy relates to its specificity and how autophagy machinery specifically recognizes bacteria. This is of major importance as xenophagy, which eventually aims to reduce not only bacterial load but also prevent cellular stress resulting, for instance, from bacteria-induced amino acid starvation (see later in this review). To explain the central mechanisms involved in the selection of intracellular bacteria by the autophagy machinery, we will focus on bacterial models that helped us shape the field.

Mycobacterium tuberculosis

Mycobacterium tuberculosis is the causative agent of tuberculosis (TB), possibly one of the oldest human pathogens and still among the top 10 causes of death worldwide (20). M. tuberculosis is a non-motile and facultative intracellular pathogen of macrophages. In this regard, the infection of alveolar macrophages is a crucial requisite toward the establishment of a successful replicative niche. Experiments using mice depleted for resident alveolar macrophages have shown that these animals become protected from M. tuberculosis (21). One of the main features of TB pathogenesis is the ability of M. tuberculosis to survive within alveolar macrophages through the interference with phagolysosome biogenesis (3, 22).

In the last decade, autophagy emerged as an essential protective strategy employed by the host to restrict the spread of M. tuberculosis. The first piece of evidence on the role of autophagy in the control of Mycobacterium was provided by the cornerstone study of Gutierrez et al. (3). The authors demonstrated that upon the induction of autophagy by starvation or rapamycin M. tuberculosis variant bovis BCG colocalized to LC3+ compartments in RAW 264.7 macrophages. Moreover, BCG phagosomes were shown to be positive for markers of acidification such as cathepsin D and Lamp-1, suggesting that xenophagy induction was able to override the blockade in phagosome maturation by BCG, with a clear impact on bacterial killing (Figure 1). Interferon-γ (IFN-γ) is essential for resistance to infection, by interfering with the transcription of more than 2,000 genes (23). In a more physiological context, Gutierrez et al. demonstrated that IFN-γ, a potent activator of macrophages, was able to mimic the effects of rapamycin or starvation on the induction of autophagy, through the immunity-related p47 guanosine triphosphatases (IRG) Irgm1 (LRG-47) (3) (Figure 1). These results put autophagy on the center stage of the immune mechanisms involved in the protection against M. tuberculosis infection. After their initial discoveries, in a subsequent study, the same group demonstrated that both Irgm1 and its human ortholog IRGM are necessary for the induction of autophagy, generating large autolysosomes that contributed with M. tuberculosis intracellular growth restriction upon macrophage activation by IFN-γ (24). The mechanism behind IRGM restriction of M. tuberculosis seems to rely on its interaction with cardiolipin in mitochondria to generate ROS and mitochondrial fission, both necessary for M. tuberculosis killing (25). The role of IFN-γ in autophagy also involves the participation of interferon-induced guanylate-binding (GBP), which are also upregulated in the presence of the cytokine. It has been demonstrated that GBPs promote oxidative killing and the delivery of antimicrobial peptides to autophagolysosomes, contributing to M. tuberculosis intracellular replication control (26) (Figure 1). Altogether, these studies demonstrated an essential in vitro role for xenophagy and its induction by IFN-γ in the control of M. tuberculosis intracellular replication.

FIGURE 1 | Autophagy targets Mycobacterium tuberculosis (Mtb) by different mechanisms. Stimulation with IFN-γ or Toll-like receptors (TLRs) ligands leads to an increase in the localization of (Mtb) into autophagosomes (left). 6-kDa early secretory antigenic target (ESAT-6) secretion system 1 (ESX-1)-induced phagosomal damage induces the exposure of Mtb to cytosolic autophagy adaptors such as Optneurin, p62, NDP52 and NBR1 which bind to ubiquitin associated with Mtb as a consequence of the E3-ligases SMURF-1 and Parkin, culminating with targeting of the bacteria for autophagic degradation (center). Extracellular bacterial DNA from Mtb is detected by STING to activate TBK1 and lead to Mtb ubiquitination and recruitment of p62 and NDP52 (middle-left). IRGM-induced increase in ROS provokes autophagic targeting of Mtb (right).

Although the link between deficiencies in ATG proteins and defective xenophagy has been widely reported upon infection with M. tuberculosis, ATG5 have also been described to play a critical autophagy-independent role in an in vivo TB mouse model. Kimmey et al. have demonstrated that the deletion of Atg14L, Atg12, Atg16L1, Atg7, and Atg3 in the myeloid compartment did not affect the outcome of M. tuberculosis infection, suggesting that the loss of autophagy is not implicated with the progression of the disease. In sharp contrast, the authors reported that the loss of ATG5 in PMN but not in alveolar macrophages led to exacerbated imunopathology, sensitizing mice to M. tuberculosis. Together, these findings suggest that ATG5 has unique autophagy-independent features that are not shared with other ATG proteins, pointing for a reinterpretation of the role of ATG5 in the control of M. tuberculosis infection in vivo (27).

One primary open question that remained to be answered was how eukaryotic cells sense M. tuberculosis infection to induce autophagy. Toll-like receptors (TLRs) detect a myriad of extracellular and endolysosome located microbial products. It has been reported that Poly (I:C), LPS, and ssRNA, ligands for TLR3, TLR4, and TLR7, respectively, induce autophagosome formation through MyD88-dependent pathways. Interestingly, activation of TLR7 by its ligand increased the ability of macrophages to kill BCG (28) (Figure 1). However, it was not clear how TLRs would be able to detect BCG to induce autophagy in the absence of exogenous stimulation with their cognate ligands. The first molecular evidence of the detection of M. tuberculosis-derived microbial-associated molecular pattern (MAMP) triggering autophagy demonstrated that stimulation of interferon genes (STING), an important adaptor of TANK-binding kinase (TBK1) in the interferon stimulatory DNA pathway, senses the presence of cytosolic DNA to trigger the ubiquitination of M. tuberculosis after phagosome damage. Upon sensing of extracellular DNA from M. tuberculosis by STING, M. tuberculosis is ubiquitinated, leading to the recruitment by the autophagic adaptors p62/SQSTM1 (hereafter p62), a multi-domain protein that functions as an autophagic adaptor. p62 possesses an LC3-interacting protein region (LIR) and a C-terminal ubiquitin-associated (UBA) domain that binds ubiquitinated substrates and an LIR. Together with p62 and nuclear dot protein 52 kDa (NDP52) work to link ubiquitinated substrates to LC3 recruitment, ensuring the efficient delivery of M. tuberculosis to autophagosomes (29) (Figure 1). Although the sequestration of Mycobacteria by xenophagy has been demonstrated to be mostly dependent on ATG proteins, one report has been shown that sequestration of ubiquitinated mycobacteria can occur in ATG5-independent manner. The authors found that following 6-kDa early secretory antigenic target (ESAT-6) secretion system 1 (ESX-1)-mediated phagosome escape, ubiquitinated bacteria were resequestered by structures that resembled autophagosomes and localized to Lamp-1+ compartments. Notably, ubiquitinated M. marinum were never decorated with LC3 and ATG5 deficiency and did not affect bacterial counts. It remains to be elucidated if the finding that M. marinum did not localize to LC3+ compartments represents a potential specific mechanism of escape from autophagy (30).

Upon phagosome damage mediated by ESX-1, M. tuberculosis is ubiquitinated, in an essential step required for the recruitment of the autophagic adaptors p62 and NDP52 and LC3. Although it has not been determined, which bacterial or host proteins (or both) are ubiquitinated during xenophagy, much progress has been made in the identification of host proteins that mediate ubiquitination involved in xenophagy. Several ubiquitin-ligases have been described as participants of bacterial ubiquitination. Parkin has a well-established role in mitophagy where it promotes the ubiquitination of mitochondrial surface proteins prior to the recruitment of p62 in order to direct malfunctioning mitochondria for autophagic degradation. In 2013, Parkin was also reported to be crucial in the conjugation of K63-ubiquitin chains to M. tuberculosis inside macrophages. In line with this finding, Park2−/− displayed increased M. tuberculosis replication in an in vivo TB model (31) (Figure 1). Of note, Parkin has also been demonstrated to participate in ubiquitination of other mycobacterial species such as M. leprae (32). Similarly, SMAD-specific E3 ubiquitin-ligase protein 1 (Smurf1) has been demonstrated to mediate K48- but not K63-ubiquitination and the recruitment of the autophagy adaptor NBR1 during M. tuberculosis infection to control its replication in human macrophages and to associate with bacteria present in the lung of patients with pulmonary TB (33). In their study, Franco et al. reported that Smurf1- but not Parkin-dependent ubiquitination is necessary for the recruitment of proteasome and NBR1 for the vicinity of M. tuberculosis. In contrast, K63 ubiquitination by Parkin but not Smurf1 is required for the recruitment of p62 to the bacterial surface (Figure 1). It remains to be elucidated why host cells employ different ubiquitin-ligases with apparent redundant roles for targeting M. tuberculosis for xenophagy. One possibility is that the apparent redundancy of Smurf-1 and Parkin is a countermeasure resulting from the ability of M. tuberculosis to escape from autophagy. Also, the different ubiquitin moieties added to M. tuberculosis surface could help in the recruitment of various adaptors. Indeed, Smurf1-mediated ubiquitination recruits the adaptor NBR1, which is not recruited by Parkin-mediated activity.

Several recent studies have reported that M. tuberculosis uses sophisticated mechanisms to escape xenophagy and replicate inside host cells. In addition to the induction of miR33 and miR33* expression to manipulate cellular metabolism and energy levels (34) and miRNA125a to inhibit UVRAG expression (35) (discussed later in this review), M. tuberculosis also induces the expression of other microRNAs (miRNAs) to circumvent xenophagy by interfering with different aspects of cellular physiology. This is the case of miR30A that has its expression increased during infection with M. tuberculosis to decrease Beclin 1 expression levels, leading to inhibition autophagosome elongation to promote intracellular survival of M. tuberculosis (36). Similarly, miR144* inhibits antimicrobial responses against M. tuberculosis in monocytes by targeting the expression of DNA damage-regulated autophagy modulator 2, allowing M. tuberculosis replication (37). In contrast, miR155 has been demonstrated to play a pro-autophagic role during M. tuberculosis infection. Wang et al. reported that miR155 targets Ras homolog enriched in brain (Rheb), a negative regulator of autophagy to accelerate the process of xenophagy. Inhibition of autophagy by M. tuberculosis seems to aim not only xenophagy but other essential steps of the immune response as well. It has been recently demonstrated that the bacterial factor PE_PGRS47 inhibits autophagy through an unknown mechanism to block MHC II antigen presentation and dampen adaptative immune responses against M. tuberculosis (38). Altogether, these studies provide compelling evidence that despite the crucial role of xenophagy as an antimycobacterial mechanism, M. tuberculosis has developed means to escape autophagy and replicate within macrophages.

Streptococcus pyogenes

Streptococcus pyogenes is the causative agent of a variety of infections, ranging from such as pharyngitis and skin infections to life-threatening necrotizin fasciitis and bacteremia (39). In 2004, Nakagawa et al. provide one of the first definitive evidence of the role of autophagy as a cell-autonomous antimicrobial mechanism. In this study, HeLa cells were shown to specifically target cytosolic GAS. This process was dependent on the toxin streptolysin O (SLO), a cholesterol-dependent pore-forming cytolysin (40). Nakagawa et al. demonstrated that the majority of the cytosolic population of GAS colocalized to LC3+ compartments, in contrast to SLO-deficient mutants in which no colocalization with LC3 was found (4) (Figure 2). As demonstrated for other intracellular bacteria, the adaptors p62, NDP52, and NBR1 are essential for recognition of ubiquitin decorated GAS and recruitment of LC3 before autophagic degradation (41, 42). Evasion of xenophagy by GAS has been reported, and GAS has been shown to evade ubiquitin recognition by the abovementioned autophagic adaptors. Barnett et al. have found that the globally disseminated serotype M1T1 (strain 5448) clone of GAS can avoid xenophagy to replicate in the cytosol. This is achieved by the expression of SpeB, a cysteine protease that degrades p62, NDP52, and NBR1. M1T1 ΔspeB mutants fail to evade recognition by these proteins and are efficiently degraded through xenophagy (41) (Figure 2). These findings reveal a new mechanism by which GAS evades elimination by xenophagy. Notably, data from the literature demonstrate that xenophagy efficiently eliminates other GAS serotypes such as M6, M49, and M89. GAS is a successful human bacterial pathogen that causes a vast array of diseases and the work of Barnett et al. uncovers autophagy evasion as a determinant feature for the dissemination of GAS. The mechanisms employed by autophagy to target intracellular GAS also include the engagement of the CD46 pathogen receptor (43). CD46 is a glycoprotein expressed by all nucleated human cells that physically binds several pathogens such as adenoviruses B and D, human herpesvirus 6, Neisseria, and GAS (44) (Figure 2). Although several innate immune receptors such as TLRs have been described to trigger xenophagy upon infection or ligand stimulation, how these receptors are connected to the selective targeting of intracellular bacteria to lysosomes is still unclear. The findings from Joubert et al. provide an important piece of data to this open question. One possibility that needs to be experimentally tested is that CD46 might be concomitantly activated together with TLRs to promote xenophagy. Another known host factor that has been reported to participate in GAS targeting for xenophagy is 8-nitroguanosine 3’,5’-cyclic monophosphate (8-nitro-cGMP), a downstream mediator of nitric oxide that has been shown to promote protein S-guanylation on bacterial surface, which are then K63 ubiquitinated prior to the recruitment of LC3 (45). Although these findings shed light into a new xenophagy targeting mechanism during infection with GAS, some open questions remain, such as (i) is this mechanism specific for GAS? and (ii) which autophagy adaptors and ubiquitin-ligases participate in this process. One interesting question regarding the induction of xenophagy by GAS is the role of endothelial cells in this process. Despite different reports showing that xenophagy plays an important role in the clearance of intracellular GAS, in endothelial cells, the results are contrasting. While, Cutting et al. demonstrated the ability of endothelial cells to upregulate xenophagy in order to control GAS infection, a recent study from Lu et al. reports that endothelial cells fail to target GAS for degradation due to an intrinsic defect in the ubiquitination of intracellular bacteria (46, 47). Even though much progress has been done in the understanding of the mechanisms of GAS-induced autophagy, further studies are required in order to clarify whether endothelial cells are in fact defective in xenophagy, if this defect is specific for infection with GAS or if GAS can halt xenophagy in these cells and not in epithelial cells.

FIGURE 2 | GAS is targeted by xenophagy by different mechanisms. Following activation of CD46, GAS is directed to autophagosomes. Streptolysin O promotes escape from phagosomes and ubiquitination and recognition by autophagic adaptors p62, NDP52 and NBR1. SpeB producing strains are able to degrade such adaptors to escape from xenophagy. GAS can also undergo modifications by ROS/NO-induced 8-nitro-cGMP via S-guanylation of its surface proteins followed by ubiquitination and targeting to autophagosomes.

Shigella flexneri

Shigella spp. are Gram-negative and highly invasive enteropathogens and a significant cause of disease, especially in children under the age of 5 years, causing approximately one million deaths worldwide (48). A few minutes after its invasion of epithelial cells and macrophages, S. flexneri is able to lyse the phagocytic vacuole and access the cytosolic compartment where it replicates (49). As countermeasures, host cells trigger autophagy to restrict S. flexneri intracellular growth and cell-to-cell spreading. The first evidence of an interaction between Shigella and autophagy was provided by a study dating from 2005. In this study, it was demonstrated that wild-type S. flexneri can escape from autophagic targeting by employing IcsB, one of the effectors of its type 3 secretion system (T3SS). Ogawa et al. observed that deletion mutants for IcsB, which is secreted by cytosolic bacteria and localizes to the bacterial surface were more efficiently targeted by autophagosomes. These results suggest that S. flexneri is able to escape from xenophagy. According to this study, the escape mechanism employed by S. flexneri involves IcsA/VirG, a 52 kDa protein that requires the bacterial chaperone IpgA for its stability, activates the complex-related proteins (Arp) 2/3 complex through the recruitment and activation of N-WASP, to induce actin polymerization and bacterial motility within the cell (50–54). Mechanistically, the study of Ogawa et al. demonstrated that, in ΔicsB mutants, IcsA/VirG triggers autophagy by binding to ATG5. According to the authors, IcsB inhibits IcsA/VirG affinity for ATG5. Thus, in wild-type S. flexneri, IcsB reduces IcsA/VirG affinity for ATG5 to initiate xenophagy (52) (Figure 3). More recently, a study added more complexity to the role of IcsB as a factor contributing to S. flexneri escape from autophagy. Baxt and Goldberg reported that IcsB also contributes to S. flexneri escape from xenophagy by recruiting transducer of CDC42-dependent actin assembly 1 (Toca-1) to prevent the recruitment of the adaptor NDP52 and LC3 (55).

FIGURE 3 | Shigella flexneri employs different mechanisms to escape from autophagy. During bacterial entry into host cells, Nod1 and Nod2 recruit ATG16L1 to initiate autophagosome formation in order to restrict S. flexneri replication (left). Vacuole damage leads to β-glycan exposure and recognition by Galectins 3 and 8 and recruitment of NDP52, followed by bacterial ubiquitination and binding to p62 and NDP52, culminating to S. flexneri targeting for autophagic degradation (center). IcsB plays a central role in disguising autophagic machinery. This protein competes with IcsA/VirG for binding to ATG5, preventing p62, and NDP52 binding, septin caging and autophagosome formation (right). Septin caging and further recruitment of autophagic adaptors are blocked by IcsB expression.

The ubiquitination of S. flexneri has been reported to be essential for the recruitment of the adaptors p62 and NDP52 (56). However, in contrast to S. typhimurium and M. tuberculosis for which several ubiquitin-ligases that ubiquitinate bacterial surface have been described, the mechanism used to host cells to tag S. flexneri with ubiquitin is not clear. LUBAC is an ubiquitin-ligase that mediates the formation of M1-linked ubiquitin chains that culminate with xenophagy and bacterial degradation of S. typhimurium (see below). In contrast, LUBAC was reported to play no major role in the trafficking of S. flexneri to autophagosomes (57). According to this study, S. flexneri escapes from LUBAC-dependent ubiquitination by secreting the effector E3-ligase H1.4 to antagonize the activity of LUBAC (57). Despite the lack of substantial data on the how S. flexneri is ubiquitinated, different adaptors that bind ubiquitin are independently recruited to the bacterial surface. For instance, p62 and NDP52 have been demonstrated to be recruited to S. flexneri surface and to regulate xenophagy mediated by each other. In agreement with the notion of its anti-autophagic role, IcsB also contributed to S. flexneri escape from autophagy by hiding IcsA/VirG from ubiquitin coating (56) (Figure 3). The reason why S. flexneri recruits different adaptors S. flexneri is not clear. However, it is possible that p62 and NDP52 may recognize different ubiquitin linkages as a result of the activity of different ubiquitin-ligases. Another hypothesis is that their LIR domains could be able to recruit different LC3 homologs and different adaptors that could contribute to membrane recruitment from various sources for the formation of autophagosome around bacteria. These hypotheses still lack experimental confirmation.

Shortly after S. flexneri entry in epithelial cells, the phagocytic vacuole is ruptured, membrane remnants expose host sugars in the cytosol, and galectin 3 promotes ubiquitination and recruitment of p62 to support xenophagy (58). In contrast to other reports from the literature, the authors did not observe increased recruitment of p62 in ΔicsB.

Interestingly, members of the NF-κB pathway such as TRAF6 and NEMO and the peptidoglycan receptor Nod1 were reported to localize to these membrane remnants (58). Similarly, NLRP3, NLRC4, ASC, and Caspase-1 were also found associated with Shigella vacuolar membrane remnants. The physiological meaning of these findings is still to be defined. One possibility is that these membrane portions might be used for the activation of inflammatory cascades and that this process is likely to be regulated by autophagy. Another possibility is that by recruiting these proteins to its vicinity, S. flexneri modulates NF-κB activation and inflammation to favor its replication and spread.

Septins are conserved GTP-binding proteins that play critical roles in cell division, cytoskeletal dynamics, and membrane remodeling (59). These proteins have been shown to form cages around S. flexneri actively polymerizing actin. Interestingly, colocalization of septins, p62, and LC3 on S. flexneri bacterial surface has been demonstrated and depletion of septins markedly reduced xenophagy of S. flexneri, suggesting an intimate relationship between these two processes (60). More recently, the precise mechanisms involved in S. flexneri-cage assembly were revealed. Sirianni et al. have found that mitochondrial proteins associate with S. flexneri-septin cages and that mitochondria promote the formation of septin cage assembly around S. flexneri for antibacterial xenophagy (61). S. flexneri has been demonstrated to induce mitochondrial damage and in the study by Siriani et al., this aspect was linked to dampening of septin cages and escape (61, 62). Of note, IcsB contributes to masking S. flexneri from septin caging (60). These results demonstrate that IcsB dampens xenophagy by at least three different mechanisms: competing with IcsA/VirG for binding to ATG5, by avoiding septin caging and targeting to autophagosomes, and by recruiting Toca-1 to inhibit the recruitment of NDP52 and LC3.

In addition to direct interaction of its virulence factors and autophagy proteins, pattern-recognition receptors also seem to participate in the interplay between S. flexneri and autophagic pathways. It has been demonstrated that the infection of macrophages by S. flexneri induces a robust activation of Caspase-1 that leads to inflammasome activation and cell death by pyroptosis in an NLRC4-dependent but ASC-independent mechanism (63). Interestingly, both Caspase-1 and NLRC4 were shown to negatively regulate autophagosome formation in macrophages infected with S. flexneri as demonstrated by studies in which bone marrow-derived macrophages (BMDMs) from knockout mice for the genes encoding these proteins were shown to induce the formation of GFP-LC3 positive membranes around bacteria in contrast to wild-type BMDMs (63). In contrast to previous studies, IcsA/VirG was not implicated in autophagy induction (52, 63), which can be explained by the different cell types used in these studies. In contrast to negative regulation of autophagy by NLRC4, NLRC1 (Nod1), and NLRC2 (Nod2), the founding members of the NLR family have been linked to autophagy induction. Nod1 and Nod2 are sensors of intracellular peptidoglycan that upon engagement lead to the activation of NF-κB activation through the recruitment of the adaptor protein RIP2 (64). Both Nod1 and Nod2 have been shown to recruit ATG16L1 at early stages of infection by S. flexneri to initiate autophagosome formation. As a result, Nod1- and Nod2-deficient MEFs display decreased numbers of GFP-LC3 positive bacteria, and interestingly, these findings did not rely on recruitment of RIP2 or NF-κB activation. Notably, in this study, the most common Nod2 mutation associated with Crohn disease (CD) resulted in impaired recruitment of ATG16L1 to the bacterial entry site and much less xenophagy, underscoring the notion that dysregulation of bacterial autophagy is likely to play an important role in the pathogenesis of CD (49) (Figure 3). It remains to be clarified if and in which conditions Nod1/2-dependent pro-autophagic signals would prevail over NLRC4-dependent anti-autophagy ones and vice versa.

Salmonella typhimurium

Salmonella typhimurium is a pathogenic Gram-negative bacterium found in the intestinal lumen and a major cause of gastroenteritis in humans and other mammals (65). This pathogen uses two T3SS, encoded by Salmonella pathogenicity island 1 and 2 (SPI2) to enter non-phagocytic cells and establish a replicative niche within vacuoles termed Salmonella-containing vacuole (SCV). In 2006, it was first reported that a fraction of the bacterial population within the SCV previously demonstrated to form pores in this compartment was able to reach the cytosol being immediately targeted by LC3 and ATG proteins. In this study, the authors showed that Atg5-deficient MEFs infected with S. typhimurium had decreased fusion of LC3+ bacteria colocalized with Lamp1, suggesting diminished bacterial degradation in lysosomes. Indeed, these cells harbored increased bacterial numbers, confirming the role of autophagy in the control of S. typhimurium infection (66) (Figure 4). Importantly, xenophagy has been reported to be essential in the control of S. typhimurium in other models such as Caenorhabditis elegans and Dictyostelium discoideum, suggesting that the role of xenophagy as an anti-S. typhimurium mechanism has been conserved throughout evolution (67).

FIGURE 4 | Mechanisms of autophagy induction by Salmonella Typhimurium. Upon entry in epithelial cells, S. Typhimurium resides in a specialized compartment, the Salmonella-containing vacuole (SCV). A fraction of the bacterial population damages and/or escapes from SCV and initiates to replicate in the cytosol. Either still within the SCV or free in the cytosolic compartment, S. Typhimurium triggers autophagy by several means. β-glycan present in vacuole remnants is recognized by Galectin-8 (Gal-8) and targets bacteria to autophagosomes. S. Typhimurium can also be ubiquitinated by the E3-ligases LRSAM or LUBAC, allowing its recognition by autophagic adaptors Optneurin, p62 or NDP52. RNF166, another E3-ligase, ubiquitinates p62 to increase the ability of this protein to bind bacteria-associated ubiquitin. Diacylglycerol (DAG) recognition and autophagy induction upon S. Typhimurium infection are not depicted here.

Although S. typhimurium targeting by autophagy was demonstrated, the means by which autophagosome formation machinery recognizes cytosolic Salmonella is not completely clear and has been the subject of many studies. Cytosolic, but not SCV residing bacteria, have been shown to be decorated with ubiquitin early during infection (66, 68). Interestingly, Ub+ bacteria colocalize with LC3, suggesting that the autophagic machinery can detect ubiquitinated substrates. Indeed, p62 has been shown to play a crucial role in the recognition, targeting to lysosomes and restriction of cytosolic ubiquitinated S. typhimurium (69). Other ubiquitin-binding proteins have also been reported to participate in Salmonella-induced autophagy. Similarly to p62, Optineurin harbors LIR and UBA domains and was shown to be necessary for the control of S. typhimurium. Interestingly, this mechanism requires Optineurin to be phosphorylated by TBK1 on serine-177 in order to enhance ubiquitin- and LC3-binding affinity to promote bacterial clearance (70). One aspect of S. typhimurium recognition by autophagy machinery that remained elusive was which bacterial substrates are ubiquitinated prior to detection by the adaptors p62, NDP52, and Optineurin. A recent study from Fiskin et al. in which ubiquitination site profiling was performed during infection with S. typhimurium revealed that outer membrane proteins are targets for ubiquitination (71).

Several ubiquitin-ligases have been reported as necessary for ubiquitination of S. typhimurium. Leucine-rich repeat and sterile α-motif-containing 1 (LRSAM1) was shown to play an essential role in the autophagic degradation of S. typhimurium. This E3-ligase was found to localize to cytosolic Salmonella upon infection of epithelial cells to ensure proper ubiquitination and autophagic control of bacterial replication. In line with these findings, a cohort study reported that lymphoblasts from patients with Charcot-Marie-Tooth disease, which harbor a frameshift mutation that truncates the RING domain of LRSAM1, present limited antibacterial activity as compared to cells from control individuals (72, 73). Another E3-ligase demonstrated to be involved in autophagic targeting of S. typhimurium is RNF166. This gene was identified in a screening for human E3-ligases as necessary for the recruitment of p62, NDP52, and LC3 for the bacterial surface in order to limit S. typhimurium replication. A unique feature of RNF166 is that, rather than tagging bacteria, it drives K29- and K33-linked ubiquitination of p62 at K91 and K189. According to the authors, this step is essential for p62-dependent bacterial targeting for autophagosomes (74) (Figure 4). More recently, the role of LUBAC, another E3-ligase, has been described. LUBAC generates linear (M1-linked) polyubiquitin patches on the surface of S. typhimurium to recruit the adaptors Optineurin, NDP52 and p62 and direct bacteria for autophagic degradation. Indeed, MEFs from cpdm−/− mice, which harbor a spontaneous mutation in LUBAC or MEFs silenced for the protein, display an increased time-dependent replication of S. typhimurium in comparison to wild-type or control-silenced cells, respectively (57). In addition to the recruitment of autophagy adaptors, LUBAC was reported to be crucial in triggering pro-inflammatory roles during infection with S. typhimurium (Figure 4). LUBAC-dependent generation of M1-linked polyubiquitin chains on the surface of the bacteria also recruits NEMO to this site (57, 75). These findings are of particular interest as it suggests that bacterial surface can provide mechanical support for the assembly of signaling platforms such as NF-κB activation, a major transcription factor that controls the production of inflammatory mediators such as cytokines and chemokines. Given that LRSAM1 was found to be only partially responsible for S. typhimurium ubiquitination, which RNF166 ubiquitinates p62 rather than bacteria and that LUBAC required an upstream E3-ligase, Polajnar et al. hypothesized that other ubiquitin ligases were involved in the ubiquitination of S. typhimurium and identified Ring-between-Ring E3 ligase ARIH1 (also known as HHARI) as an important protein ubiquitin-ligase for targeting this pathogen to autophagosomes (76). Notably, this study demonstrated that depletion of LRSAM1 and ARIH1 led to an enhancement in LUBAC-dependent ubiquitination and NF-κB activation, culminating with increased bacterial replication, in contrast to previous findings, reporting that NF-κB activation led to bacterial growth restriction (57, 75, 76). Together, these data indicate that recruitment of different ubiquitin-ligases (with different ubiquitin linkage abilities) to the bacterial surface may endow cells with several layers of protection against the replication of cytosolic S. typhimurium.

In addition to bacterial ubiquitination, lipid second messengers have also been reported to be required for efficient targeting of S. typhimurium. Shahnazari et al. demonstrated that diacylglycerol (DAG) is produced during infection with S. typhimurium in a phospholipase D- and phosphatidic acid phosphatase-dependent manner. DAG localization in bacteria-containing phagosomes seemed to be a requisite for autophagy and may occur in parallel to independent p62 and NDP52 recruitment, once again suggesting several layers of proteins involved in bacterial targeting (77).

The detection of damage in the SCV has been demonstrated to be an important step in the targeting of S. typhimurium for autophagic degradation (66). Galectin-8 is a β-galactoside-binding lectin that has been reported to monitor endosomal and lysosomal integrity and detects bacterial invasion by binding host glycans exposed on damaged SCVs. Recently, it has been demonstrated that among galectins 1–4, 7–10, and 12–14, only Galectin-8 colocalized to S. typhimurium during infection of HeLa cells. Interestingly, NDP52 was recruited to cytosolic exposed S. typhimurium, directly binding to Galectin-8 to restrict bacterial replication. These and previous findings lead to a model in which, upon SCV damage, host sugar molecules such as β-galactoside, usually confined to the lumen of endosomes are exposed in the cytosol and sensed by Galectin-8 that in turn recruits NDP52 and LC3 to SCV to initiate lysosomal degradation of S. typhimurium (42, 78). Despite their role in mediating S. typhimurium-induced autophagic degradation, p62 and NDP52 show independent targeting activity. In a study in which HeLa cells were silenced for p62 or NDP52, there was no interference in the number of NDP52+ or p62+ bacteria, respectively (Figure 4). Interestingly, it was demonstrated that these adaptors recognize ubiquitin deposited in distinct microdomains at the bacterial surface that could result from the activity from different ubiquitin-ligases (79). Future studies must provide explanations if and why cells preferably decide toward the employment of one or the other ubiquitin-ligase and autophagy adaptors.

Listeria monocytogenes

Listeria monocytogenes is a Gram-positive bacterial pathogen that causes listeriosis, a self-limiting disease in healthy individuals that become severe in immunocompromised or elderly individuals and pregnant women (80). One of the main features of L. monocytogenes is its ability to replicate within several cell types during infection, including macrophages, a cell type usually able to kill the majority of intracellular bacteria (81).

Before its replication in the cytosol, L. monocytogenes must escape from the phagosome. This is achieved through the expression of several virulence factors rapidly upon entry. Possibly, the main bacterial factor associated with phagosome escape, listeriolysin O (LLO), is a cholesterol-dependent, pore-forming cytolysin that form pores in the phagosomal membrane immediately after bacteria uptake (82–85). In experiments with fluorescently labeled molecules of increasing sizes, it has been demonstrated that the pores grow in size until large enough to allow bacterial escape (86). In addition to LLO pore-forming activity, two C-type phospholipases, phosphatidylinositol-specific (PI-PLC, plcB), and a broad-range phosphatidylcholine (PC-PLC, plcA) also contribute to L. monocytogenes escape from phagosome, likely digesting its membrane (86).

In order to successfully replicate in the cytosol, L. monocytogenes needs to circumvent several layers of host defense. Autophagy has been reported to contribute to the control of infection, although several studies show that the bacteria are able to escape from autophagic degradation (87, 88). Infection of RAW 264.7 macrophages of wild-type L. monocytogenes showed that ~40% of the intracellular bacterial population was targeted by LC3 by 1 h postinfection (p.i) in an LLO-dependent manner (Figure 5). However, at 8 h p.i, only 10% of the bacterial population was LC3-positive. These results suggest that L. monocytogenes was able to escape from autophagic degradation. Indeed, after initial targeting by LC3, replication rates robustly increased, in line with the drop in bacterial colocalization with LC3 observed at later stages of infection (89).

FIGURE 5 | ActA and phospholipases (plc) A and B are major Listeria monocytogenes virulence factors involved in the escape from autophagy. Upon Listeriolysin O-dependent escape from phagosome wild-type L. monocytogenes escapes from autophagic targeting due to the expression of ActA, that blocks the recruitment of p62 and NDP52 to the bacterial surface. The expression of plcA/B stalls the formation of autophagosomal membranes. Galectin recognition of damaged vacuoles and autophagy induction are not shown here.

ActA, a key virulence factor of L. monocytogenes involved in intracellular motility, has also been implicated in autophagy evasion. In vitro studies demonstrate contradictory results with ΔactA mutants in different genetic backgrounds and cell types. While EGDe ΔactA mutants infecting Hela cells show time-dependent increase in the colocalization with LC3, 10403S ΔactA mutants in the 10403S background infecting macrophages loses its staining for LC3 at later time points during infection (88, 90). It remains to be elucidated whether the differences observed for both genetic backgrounds are related or not to the different cell types used. Despite this controversy, it is important to note that ΔactA mutants in both backgrounds display comparable replication in vitro (88–90). Importantly, ActA-dependent escape of autophagy does not rely on its ability to mediate bacterial motility. Using a series of ActA truncated mutants, Yoshikawa et al. demonstrated that as long as the capacity of ActA to recruit actin-related proteins (Arp) 2/3 complex, vasodilator-stimulated phosphoprotein or actin, is retained, bacteria are able to disguise autophagic recognition (88) (Figure 5).

The ubiquitination of L. monocytogenes and the involvement of autophagy adaptors such as p62 and NDP52 have been reported in the targeting L. monocytogenes to autophagosomes. In HeLa cells, p62 and NDP52 were shown to be recruited independently during the infection with the ΔactA EGDe (56). Recently, these strains were compared in regards to LC3, p62, and Ub during infection of macrophages. Although ΔactA mutants in EGDe and 10403S genetic backgrounds were reported to be sharply different regarding colocalization with LC3, p62, and Ub recruitment and replication were identical for both strains suggesting that ΔactA can block xenophagy downstream of ubiquitination and LC3 recruitment (90). The ubiquitin-ligases Parkin and SMURF1 were demonstrated to play a role in the ubiquitination of L. monocytogenes. Park2−/− mice infected with L. monocytogenes showed up to 20-fold higher bacterial load relative to wild-type animals (31). Smurf1−/− macrophages infected with ΔactA L. monocytogenes do not show recruitment of K48-ubiquitin to the bacterial surface while K63-linked ubiquitination was not affected. In line with this finding, Smurf1−/− mice infected with L. monocytogenes were shown to harbor significantly higher bacterial burdens in comparison to wild-type (33). As ubiquitination of cytosolic bacteria has been known to be essential for autophagic degradation, it is assumed that the higher bacterial burden in Smurf1−/− mice is a consequence of dampened antibacterial autophagy. More recently, NEDD4 (neuronal precursor cell expressed, developmentally downregulated 4), another ubiquitin-ligase has been implicated autophagic degradation of L. monocytogenes (Figure 5). However, in contrast to Parkin and Smurf1, NEDD4 does not recruit ubiquitin to the bacterial surface but enhances the mediated K6- and K27-linkage ubiquitination of BECN1, leading to higher stability of BECN1 and increased autophagy (91).

Listeria monocytogenes has been reported to induce amino acid starvation and activation of the general control nonderepressible 2 (GCN2)-eIF2α pathway upstream mTOR. GCN2 is one of four “stress kinases” that block translation by phosphorylating eIF2α. GCN2 is thought to bind uncharged tRNAs to “sense” amino acids availability (92). Upon detection of a decrease in the amino acid pool, mTOR activity is reduced leading to autophagy activation to normalize this condition. Unlike what is observed during the infection of epithelial cells with S. flexneri, in L. monocytogenes-infected cells, autophagy is kept repressed, suggesting that L. monocytogenes possesses other virulence weapons to block autophagy (93–95). In addition to ΔactA-mediated escape from autophagy, L. monocytogenes employs its two C-type phospholipases to disrupt the autophagosome elongation step in order to inhibit autophagy-dependent degradation. In an in vitro study, it was observed that L. monocytogenes deleted for plcA and plcB were more strongly targeted to autophagosomes than wild-type bacteria at later time points of infection. In parallel, wild-type bacteria induced the accumulation of granules positive for LC3, ATG16L1, and as well as WIPI-2, a phosphatidylinositol 3-phosphate-binding protein that is present on maturing phagophores, suggesting blockade of pre-autophagosome structures. Interestingly, the authors demonstrate that in plcA/plcB L. monocytogenes mutants, the accumulation of such structures was not observed (93, 95). These results, together with the previous findings of Mitchell et al. point toward the combined effects of ActA and L. monocytogenes phospholipases in the escape from autophagy (87) (Figure 4).

The detection of MAMPs has also been described as an autophagy trigger during the infection of L. monocytogenes. In 2008, a study using Drosophila melanogaster as a model for L. monocytogenes infection reported that peptidoglycan-recognition protein (PGRP-LE) mediated autophagy-dependent control of bacterial replication in vitro and in vivo (96). Interestingly, the intracellular peptidoglycan receptor Nod1 has also been linked to xenophagy of L. monocytogenes in vitro. MEFs from Nod1-deficient mice were demonstrated to be defective in targeting L. monocytogenes to autophagosomes, indicating an important role for peptidoglycan recognition in the induction of autophagy during infection with this bacterium in mammals as well (49).

Legionella pneumophila

The Gram-negative bacterium L. pneumophila was first identified as the causative agent of an epidemic of pneumonia at an American Legion convention in Philadelphia, PA, USA in 1976 (97). This disease is characterized by the inhalation of aerosols containing high numbers of L. pneumophila (98). Although usually found in freshwater protozoa and amebae, L. pneumophila can accidentally replicate in alveolar macrophages in human lung, especially in immune-compromised patients (99, 100). In order to replicate within its eukaryotic host, L. pneumophila employs strategies that involve blocking the fusion of phagosomes with lysosomes after phagocytic ingestion of the bacteria and the generation of endoplasmic reticulum (ER)-like compartment that affords its replication (7, 101). The L. pneumophila-containing vacuoles (LCVs) present features that are shared by autophagosomes, including its close association with ER membrane (100, 102). This led to the speculation that the formation of biogenesis could involve the autophagy machinery (103, 104). Initial studies that focused on the characterization of the LCV reported that this compartment did not fuse with acidic vesicles since proteins that localize to endolysosomes, such as Lamp-1 and Rab7 were absent in LCV membrane and that the ability of L. pneumophila to evade phagosomal maturation was dependent on its viability (98, 105). Further studies using avirulent strains of L. pneumophila identified the intracellular multiplication (icm) and defect in organelle trafficking (dot) loci as the genetic loci determinants required for intracellular multiplication and evasion phagosome–lysosome fusion (106, 107). The emergence of autophagy as an antimicrobial effector led to the examination of the role of this process in the pathogenesis of L. pneumophila infection. Since LC3 is a major marker for autophagosome membranes, several cell biology approaches aimed to analyze the recruitment of LC3+ compartments to LCVs (100). Interestingly, following infection of macrophages with L. pneumophila, the formation of autophagosomes was blunted. In line with the role of Dot/Icm in the virulence of this bacterium, infection of macrophages with an isogenic Dot/Icm-deficient dotA mutant was unable to induce defects in autophagy induction (108). To identify the bacterial factors involved in autophagy inhibition, Choy et al. conducted a genetic screen that mapped the defect in autophagy to a chromosomal region encoding for 10 effectors. Analysis of the effects of the individual effectors revealed the protein RavZ as necessary and sufficient for blocking autophagy (108). In vitro analysis demonstrated that RavZ, which displays cysteine-protease activity, acts to deconjugate LC3 from autophagosomes and block its reconjugation (108). Furthermore, recent reports demonstrate that RavZ might participate not only in the deconjugation of LC3 but also in other steps that interfere with xenophagy. Kubori et al. have found in co-infection experiments with L. pneumophila and S. typhimurium that the recruitment of ubiquitin, p62, and NDP52 to the surface of S. typhimurium was dampened, suggesting a deubiquitinase-like enzymatic activity for RavZ (109). The resolution of the crystal structure of RavZ yielded new clues to its mechanisms. According to this study, by targeting autophagosomes through PIP3- and curvature-sensing motifs, RavZ limits its activity only to LC3 that is bound to autophagosomes (110) (Figure 6). Other RavZ-independent mechanisms for L. pneumophila evasion of autophagy have been described as well. Phylogenetic analyses suggested a high degree of similarity between one L. pneumophila and the eukaryotic sphingosine-1 phosphate lyase (SPL) (111). The L. pneumophila SPL homolog (LpSlp) has similar enzymatic activities to the eukaryotic SPL, which finely regulates intracellular levels of sphingosine-1-phosphate (S1P) (112), which have been shown to stimulate autophagy (111, 113). Infection of macrophages with wild-type L. pneumophila but not the LpSpl-deficient mutant leads to a depletion in S1P levels and inhibition of autophagy, indicating that L. pneumophila uses molecular mimicry to block autophagy and replicate within macrophages (111).

FIGURE 6 | Legionella pneumophila disrupts autophagy to create a replicative niche. L. pneumophila secretes RavZ through its Dot/Icm apparatus in order to deconjugate LC3 from ER-derived vesicles and block autophagy in order to escape from autophagic degradation.

The Role AMP-Dependent Protein Kinase (AMPK) Activation and Bacterial-Induced Amino Acid Starvation in Bacterial Xenophagy

Living organisms obtain energy from the catabolism of nutrients whose molecular blocks are then converted into ATP and NADPH. The fact that cells are continually synthesizing ATP keeps its level close to maximal, with only small variations (114). However, under nutrient stress, when ATP levels drop, adenylate kinase shifts to an ATP synthesis mode to restore its levels. In turn, AMP levels increase significantly and, physiologically, changes in AMP concentrations are much higher than those observed to ATP (115), which makes the AMP/ATP ratio the most reliable marker of the cellular energetic status (114). Under such conditions, AMPK detects tiny changes in AMP levels and represents the principal cellular metabolism regulator (114). One of the main direct consequences of AMPK engagement is the activation of ULK1, suppressing mTORC1 inhibitory activity to allow the formation of autophagosomes (116).

In addition to its crucial role as a metabolic sensor, AMPK has also been widely reported to be involved in the activation of autophagy by bacteria. In a bacterial peritonitis-induced sepsis model, the use of the AMPK activator aminoimidazole carboxamide ribonucleotide (AICAR) increased bacterial killing, suggesting the implication of AMPK in the enhancement of the activity of phagocytic cells. Indeed, the use of these activators led to increased chemotaxis, phagocytosis, and bacterial killing of neutrophils infected with Escherichia coli (117). Evidence from the literature demonstrates that activation of AMPK by treatment with AICAR can also increase targeting of M. tuberculosis to LC3-positive compartments. Of note, when key autophagic proteins such as ATG7 were silenced, this effect was not observed, suggesting AICAR promotes the targeting of M. tuberculosis to autophagosomes. Moreover, AICAR-induced xenophagy was shown to contribute to bacterial killing in vitro, in a mechanism involving mTOR inhibition and increased mitochondrial biogenesis and ATP generation, likely as a result of energy drop during M. tuberculosis infection. Since it has been previously demonstrated that peroxisome proliferator-activated receptor-gamma, coactivator 1α (PPARGC1A) is important for the regulation of mitochondrial gene expression and glucose metabolism, it was speculated that PPARGC1A was involved in AICAR-induced xenophagy of M. tuberculosis. When PPARGC1A expression was silenced in macrophages infected with M. tuberculosis, the robust increase in mitochondrial biogenesis, ATP generation, and decreased M. tuberculosis replication induced by AICAR treatment were not observed (118, 119). The findings of this study support those from Gutierrez et al. (3) demonstrating that induction of autophagy through rapamycin enhances antimicrobial defenses against M. tuberculosis. Although AMPK activation was found to be involved in the efficient xenophagy-dependent control of M. tuberculosis, this pathogen developed sophisticated mechanisms to manipulate AMPK activity in order to favor its replication. In another recent evidence, miRNAs emerged as important “fine-tuners” of gene expression in response to pathophysiological stimuli. These RNAs bind to the 3’-untranslated region of specific mRNAs to reduce protein expression by blocking mRNA translation or inducing its degradation (120). Accumulating evidence shows that many miRNAs regulate the complex interplay between mycobacterial survival strategies and host innate immune and metabolic pathways (121). One of these miRNAs, miR33 has been shown to the regulation of fatty acid metabolism and insulin signaling (122). M. tuberculosis seems to use the expression of miRNAs to subvert autophagy to create a favorable replicative niche. M. tuberculosis infection of macrophages induces the expression of miR-33 and its passenger strand miR-33* to dampen mitochondrial fatty acid oxidation and lipophagy (autophagy of lipid droplets) to increase cellular lipid content, which is essential for the bacilli as a nutrient source during infection (34). According to this study, autophagy inhibition was achieved by inhibition of AMPK, which controls transcription factor EB and Forkhead box transcription factor class O (FOXO3), transcriptional regulators of autophagy and lysosomal biogenesis gene programs, respectively (123, 124). Altogether, these studies demonstrate that AMPK is activated during infection with intracellular bacteria.

The mechanisms by which intracellular bacteria initiate xenophagy are not completely elucidated, but compelling evidence from the literature suggests that these pathogens trigger energy imbalance and cellular nutritional stress that result in the activation of cellular responses culminating in the upregulation of autophagic activity (93). It has been reported that the infection of epithelial cells with S. flexneri infection induces a general and persistent loss of amino acids, leading to amino acid starvation-induced stress (94). In contrast, S. typhimurium induces only a rapid and transient depletion of amino acid pools. Of note, during S. flexneri- and S. typhimurium-induced amino acid depletion, robust relocalization of mTORC1 is observed. While in S. flexneri-infected cells, S6K1 and 4EBP1, two major targets of mTOR, are downregulated, and mTORC1 dispersed in the cytosol throughout infection, during the infection with S. typhimurium, this is observed only in early time-points, suggesting that this bacterial pathogen developed means to manipulate mTOR signaling to favor its intracellular survival (94) (Figure 7). Indeed, in a recent study, Ganesan et al. demonstrated that despite sustained low levels of ATP in macrophages infected by S. typhimurium, AMPK was only transiently activated at early time-points and then returned to basal levels (125). AMPK activation is known to be regulated by a cytosolic complex consisting of liver kinase B1 (LKB1) and Sirtuin-1 (Sirt1), where Sirt1 is necessary for deacetylation and activation of LKB1 (126). Interestingly, the study by Ganesan et al. reports that S. typhimurium induces the lysosomal degradation of AMPK, LKB1, and Sirt1 to reactivate mTORC1 activity in order to inhibit autophagosome formation and escape from xenophagy. Notably, this reactivation was shown to be dependent on SsrB, a regulator of pathogenicity island 2 (SPI2) encoded virulence factors (127), and SsaV a component of the SPI2 type III secretion apparatus (128). S. typhimurium mutants lacking SsrB and SsaV failed to induce lysosomal degradation of the AMPK/LKB1/Sirt1 circuit and are efficiently targeted to autophagosomes (125). Together, the studies from Tattoli et al. and Ganesan et al. demonstrate that nutritional cellular stress induced by bacterial infection triggers xenophagy to control bacterial replication (in the case of S. flexneri infection) and that S. typhimurium developed mechanisms to escape from autophagy by reactivating mTORC1 activity. Together, the studies from Tattoli et al. and Ganesan et al. demonstrate that nutritional cellular stress induced by bacterial infection triggers xenophagy to control bacterial replication (in the case of S. flexneri infection) and that S. typhimurium developed mechanisms to escape from autophagy by reactivating mTORC1 activity.

FIGURE 7 | Bacteria-induced amino acid starvation stress triggers xenophagy. Shigella flexneri induces a persistent amino acid starvation that leads to GCN2 activation and inhibition of mTORC1 to allow the formation of bacteria targeted autophagosomes (left). In contrast, Salmonella Typhimurium (center) and Listeria monocytogenes (right) trigger only a transient amino acid starvation and inhibition of mTORC1, allowing its reactivation at later time-points to block the formation of autophagosomes.

Listeria monocytogenes has also been reported to induce amino acid starvation-induced cellular stress and activation of the GCN2-eIF2α pathway upstream mTOR. Upon de detection of a decrease in the amino acid pool, mTOR activity is reduced leading to autophagy activation in order to normalize this condition. Unlike what is observed during the infection of epithelial cells with S. flexneri, in L. monocytogenes cells, autophagy is kept repressed, suggesting that L. monocytogenes possesses other virulence weapons to block autophagy (93–95) (Figure 7). Finally, AMPK has also been implicated in the enhancement of xenophagy during the infection with E. coli. According to this study, E. coli infection leads to an increase in intracellular calcium levels, which activates Ca(2+)/calmodulin-dependent protein kinase kinase β (CaMKKβ) to promote AMPK activation. AMPK was undoubtfully implicated in CaMKKβ-mediated xenophagy when macrophages were silenced for AMPK and control of E. coli replication was dampened (129).

CONCLUDING REMARKS

Xenophagy has been widely reported to target bacteria for autophagic degradation, with clear impact on intracellular bacterial handling. Even with major advances in our understanding of the mechanisms involved in cargo selection, many questions remain unanswered. For example, why so many different mechanisms to target bacterial pathogens exposed to the cytosol? Still, why different autophagic adaptors and ubiquitin-ligases with apparent redundant functions? Although no evidence in this direction has been reported, we cannot exclude that different types of autophagosomes exist. Thus, p62, NDP52, NBR1, and Optineurin would function as sorters for different autophagosomes. It is possible that the different ubiquitin-ligases work in this direction as well by adding different ubiquitin linkages to the bacterial surface. Regarding bacteria-induced nutritional stress and autophagy induction, it is still to be elucidated whether amino acid starvation is induced upon infection with bacterial pathogens other than Shigella, Salmonella, and Listeria. Also, why bacteria induce amino acid starvation that leads to autophagy to subsequently inhibit it?

In in vitro studies, it is clear that only a fraction of the intracellular bacterial population is targeted to autophagosomes, with modest impact in bacterial replication control following autophagy ablation. This is in sharp contrast to in vivo studies, which demonstrate much more pronounced differences in bacterial replication in the absence of autophagy. How would these differences be explained?

Future goals in the field must address these open questions to provide a full understanding of the role of autophagy in bacterial infections.

AUTHOR CONTRIBUTIONS

LT, MS, and RM wrote the paper. LT also revised the final document.

FUNDING

Research in Leonardo Travassos’ Lab was funded by Fundação Carlos Chagas Filho de Amparo à Pesquisa do Estado do Rio de Janeiro (Faperj) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq). LT is research fellow from Faperj (Cientista do Nosso Estado) and CNPq (1D). MS was supported by a fellowship from Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES). RM was supported by a fellowship from CNPQ.

REFERENCES

1. Hussey S, Travassos LH, Jones NL. Autophagy as an emerging dimension to adaptive and innate immunity. Semin Immunol (2009) 21:233–41. doi:10.1016/j.smim.2009.05.004

2. Rikihisa Y. Glycogen autophagosomes in polymorphonuclear leukocytes induced by rickettsiae. Anat Rec (1984) 208:319–27. doi:10.1002/ar.1092080302

3. Gutierrez MG, Master SS, Singh SB, Taylor GA, Colombo MI, Deretic V. Autophagy is a defense mechanism inhibiting BCG and Mycobacterium tuberculosis survival in infected macrophages. Cell (2004) 119:753–66. doi:10.1016/j.cell.2004.11.038

4. Nakagawa I, Amano A, Mizushima N, Yamamoto A, Yamaguchi H, Kamimoto T, et al. Autophagy defends cells against invading group A Streptococcus. Science (2004) 306:1037–40. doi:10.1126/science.1103966

5. Amano A, Nakagawa I, Yoshimori T. Autophagy in innate immunity against intracellular bacteria. J Biochem (2006) 140:161–6. doi:10.1093/jb/mvj162

6. Huang J, Brumell JH. Bacteria-autophagy interplay: a battle for survival. Nat Rev Microbiol (2014) 12:101–14. doi:10.1038/nrmicro3160

7. Kagan JC, Stein MP, Pypaert M, Roy CR. Legionella subvert the functions of Rab1 and Sec22b to create a replicative organelle. J Exp Med (2004) 199:1201–11. doi:10.1084/jem.20031706

8. Yang Z, Klionsky DJ. Mammalian autophagy: core molecular machinery and signaling regulation. Curr Opin Cell Biol (2010) 22:124–31. doi:10.1016/j.ceb.2009.11.014

9. Suzuki H, Osawa T, Fujioka Y, Noda NN. Structural biology of the core autophagy machinery. Curr Opin Struct Biol (2017) 43:10–7. doi:10.1016/j.sbi.2016.09.010

10. Yin Z, Pascual C, Klionsky DJ. Autophagy: machinery and regulation. Microb Cell (2016) 3:457–65. doi:10.15698/mic2016.12.546

11. Jung CH, Jun CB, Ro SH, Kim YM, Otto NM, Cao J, et al. ULK-Atg13-FIP200 complexes mediate mTOR signaling to the autophagy machinery. Mol Biol Cell (2009) 20:1992–2003. doi:10.1091/mbc.E08-12-1249

12. Kageyama S, Omori H, Saitoh T, Sone T, Guan J-L, Akira S, et al. The LC3 recruitment mechanism is separate from Atg9L1-dependent membrane formation in the autophagic response against Salmonella. Mol Biol Cell (2011) 22:2290–300. doi:10.1091/mbc.E10-11-0893

13. Nair U, Cao Y, Xie Z, Klionsky DJ. Roles of the lipid-binding motifs of Atg18 and Atg21 in the cytoplasm to vacuole targeting pathway and autophagy. J Biol Chem (2010) 285:11476–88. doi:10.1074/jbc.M109.080374

14. Carneiro LAM, Travassos LH. The interplay between NLRs and autophagy in immunity and inflammation. Front Immunol (2013) 4:361. doi:10.3389/fimmu.2013.00361

15. Fujita N, Itoh T, Omori H, Fukuda M, Noda T, Yoshimori T. The Atg16L complex specifies the site of LC3 lipidation for membrane biogenesis in autophagy. Mol Biol Cell (2008) 19:2092–100. doi:10.1091/mbc.E07-12-1257

16. Sou YS, Waguri S, Iwata J, Ueno T, Fujimura T, Hara T, et al. The Atg8 conjugation system is indispensable for proper development of autophagic isolation membranes in mice. Mol Biol Cell (2008) 19:4762–75. doi:10.1091/mbc.E08-03-0309

17. Heckmann BL, Boada-Romero E, Cunha LD, Magne J, Green DR. LC3-associated phagocytosis and inflammation. J Mol Biol (2017) 429:3561–76. doi:10.1016/j.jmb.2017.08.012

18. Martinez J, Malireddi RK, Lu Q, Cunha LD, Pelletier S, Gingras S, et al. Molecular characterization of LC3-associated phagocytosis reveals distinct roles for Rubicon, NOX2 and autophagy proteins. Nat Cell Biol (2015) 17:893–906. doi:10.1038/ncb3192

19. Sanjuan MA, Dillon CP, Tait SWG, Moshiach S, Dorsey F, Connell S, et al. Toll-like receptor signalling in macrophages links the autophagy pathway to phagocytosis. Nature (2007) 450:1253–7. doi:10.1038/nature06421

20. Espert L, Beaumelle B, Vergne I. Autophagy in Mycobacterium tuberculosis and HIV infections. Front Cell Infect Microbiol (2015) 5:49. doi:10.3389/fcimb.2015.00049

21. Lam A, Prabhu R, Gross CM, Riesenberg LA, Singh V, Aggarwal S. Role of apoptosis and autophagy in tuberculosis. Am J Physiol Lung Cell Mol Physiol (2017) 313:L218–L229. doi:10.1152/ajplung.00162.2017

22. Vergne I, Chua J, Singh SB, Deretic V. Cell biology of Mycobacterium tuberculosis phagosome. Annu Rev Cell Dev Biol (2004) 20:367–94. doi:10.1146/annurev.cellbio.20.010403.114015

23. Zhang SY, Boisson-Dupuis S, Chapgier A, Yang K, Bustamante J, Puel A, et al. Inborn errors of interferon (IFN)-mediated immunity in humans: insights into the respective roles of IFN-α/β, IFN-γ, and IFN-λ in host defense. Immunol Rev (2008) 226:29–40. doi:10.1111/j.1600-065X.2008.00698.x

24. Singh SB, Davis AS, Taylor GA, Deretic V. Human IRGM induces autophagy to eliminate intracellular mycobacteria. Science (2006) 313:1438–41. doi:10.1126/science.1129577

25. Singh SB, Ornatowski W, Vergne I, Naylor J, Delgado M, Roberts E, et al. Human IRGM regulates autophagy and cell-autonomous immunity functions through mitochondria. Nat Cell Biol (2010) 12:1154–65. doi:10.1038/ncb2119

26. Kim B-H, Shenoy AR, Kumar P, Das R, Tiwari S, Macmicking JD. A family of IFN-γ-inducible 65-kD GTPases protects against bacterial infection. Science (2011) 332:717–21. doi:10.1126/science.1201711

27. Kimmey JM, Huynh JP, Weiss LA, Park S, Kambal A, Debnath J, et al. Unique role for ATG5 in neutrophil-mediated immunopathology during M. tuberculosis infection. Nature (2015) 528:565–9. doi:10.1038/nature16451

28. Delgado MA, Elmaoued RA, Davis AS, Kyei G, Deretic V. Toll-like receptors control autophagy. EMBO J (2008) 27:1110–21. doi:10.1038/emboj.2008.31

29. Watson RO, Manzanillo PS, Cox JS. Extracellular M. tuberculosis DNA targets bacteria for autophagy by activating the host DNA-sensing pathway. Cell (2012) 150:803–15. doi:10.1016/j.cell.2012.06.040

30. Collins CA, De Maziere A, Van Dijk S, Carlsson F, Klumperman J, Brown EJ. Atg5-independent sequestration of ubiquitinated mycobacteria. PLoS Pathog (2009) 5:e1000430. doi:10.1371/journal.ppat.1000430

31. Manzanillo PS, Ayres JS, Watson RO, Collins AC, Souza G, Rae CS, et al. The ubiquitin ligase parkin mediates resistance to intracellular pathogens. Nature (2013) 501:512–6. doi:10.1038/nature12566

32. De Leseleuc L, Orlova M, Cobat A, Girard M, Huong NT, Ba NN, et al. PARK2 mediates interleukin 6 and monocyte chemoattractant protein 1 production by human macrophages. PLoS Negl Trop Dis (2013) 7:e2015. doi:10.1371/journal.pntd.0002015

33. Franco LH, Nair VR, Scharn CR, Xavier RJ, Torrealba JR, Shiloh MU, et al. The ubiquitin ligase smurf1 functions in selective autophagy of Mycobacterium tuberculosis and anti-tuberculous host defense. Cell Host Microbe (2017) 21:59–72. doi:10.1016/j.chom.2016.11.002

34. Ouimet M, Koster S, Sakowski E, Ramkhelawon B, Van Solingen C, Oldebeken S, et al. Mycobacterium tuberculosis induces the miR-33 locus to reprogram autophagy and host lipid metabolism. Nat Immunol (2016) 17:677–86. doi:10.1038/ni.3434

35. Kim JK, Yuk JM, Kim SY, Kim TS, Jin HS, Yang CS, et al. MicroRNA-125a inhibits autophagy activation and antimicrobial responses during mycobacterial infection. J Immunol (2015) 194:5355–65. doi:10.4049/jimmunol.1402557

36. Chen Z, Wang T, Liu Z, Zhang G, Wang J, Feng S, et al. Inhibition of autophagy by MiR-30A induced by mycobacteria tuberculosis as a possible mechanism of immune escape in human macrophages. Jpn J Infect Dis (2015) 68:420–4. doi:10.7883/yoken.JJID.2014.466

37. Kim JK, Lee HM, Park KS, Shin DM, Kim TS, Kim YS, et al. MIR144* inhibits antimicrobial responses against Mycobacterium tuberculosis in human monocytes and macrophages by targeting the autophagy protein DRAM2. Autophagy (2017) 13:423–41. doi:10.1080/15548627.2016.1241922

38. Saini NK, Baena A, Ng TW, Venkataswamy MM, Kennedy SC, Kunnath-Velayudhan S, et al. Suppression of autophagy and antigen presentation by Mycobacterium tuberculosis PE_PGRS47. Nat Microbiol (2016) 1:16133. doi:10.1038/nmicrobiol.2016.133

39. Carapetis JR, Steer AC, Mulholland EK, Weber M. The global burden of group A streptococcal diseases. Lancet Infect Dis (2005) 5:685–94. doi:10.1016/S1473-3099(05)70267-X

40. Sakurai A, Maruyama F, Funao J, Nozawa T, Aikawa C, Okahashi N, et al. Specific behavior of intracellular Streptococcus pyogenes that has undergone autophagic degradation is associated with bacterial streptolysin O and host small G proteins Rab5 and Rab7. J Biol Chem (2010) 285:22666–75. doi:10.1074/jbc.M109.100131

41. Barnett TC, Liebl D, Seymour LM, Gillen CM, Lim JY, Larock CN, et al. The globally disseminated M1T1 clone of group A Streptococcus evades autophagy for intracellular replication. Cell Host Microbe (2013) 14:675–82. doi:10.1016/j.chom.2013.11.003

42. Thurston TL, Ryzhakov G, Bloor S, Von Muhlinen N, Randow F. The TBK1 adaptor and autophagy receptor NDP52 restricts the proliferation of ubiquitin-coated bacteria. Nat Immunol (2009) 10:1215–21. doi:10.1038/ni.1800

43. Joubert PE, Meiffren G, Gregoire IP, Pontini G, Richetta C, Flacher M, et al. Autophagy induction by the pathogen receptor CD46. Cell Host Microbe (2009) 6:354–66. doi:10.1016/j.chom.2009.09.006

44. Cattaneo R. Four viruses, two bacteria, and one receptor: membrane cofactor protein (CD46) as pathogens’ magnet. J Virol (2004) 78:4385–8. doi:10.1128/JVI.78.9.4385-4388.2004

45. Ito C, Saito Y, Nozawa T, Fujii S, Sawa T, Inoue H, et al. Endogenous nitrated nucleotide is a key mediator of autophagy and innate defense against bacteria. Mol Cell (2013) 52:794–804. doi:10.1016/j.molcel.2013.10.024

46. Cutting AS, Rosario Y, Mu R, Rodriguez A, Till A, Subramani S, et al. The role of autophagy during group B Streptococcus infection of blood-brain barrier endothelium. J Biol Chem (2014) 289:35711–23. doi:10.1074/jbc.M114.588657

47. Lu SL, Kawabata T, Cheng YL, Omori H, Hamasaki M, Kusaba T, et al. Endothelial cells are intrinsically defective in xenophagy of Streptococcus pyogenes. PLoS Pathog (2017) 13:e1006444. doi:10.1371/journal.ppat.1006444

48. Kotloff KL, Riddle MS, Platts-Mills JA, Pavlinac P, Zaidi AKM. Shigellosis. Lancet (2017) 391(10122):801–12. doi:10.1016/S0140-6736(17)33296-8

49. Travassos LH, Carneiro LAM, Ramjeet M, Hussey S, Kim Y-G, Magalhães JG, et al. Nod1 and Nod2 direct autophagy by recruiting ATG16L1 to the plasma membrane at the site of bacterial entry. Nat Immunol (2009) 11:55–62. doi:10.1038/ni.1823

50. Egile C, Loisel TP, Laurent V, Li R, Pantaloni D, Sansonetti PJ, et al. Activation of the CDC42 effector N-WASP by the Shigella flexneri IcsA protein promotes actin nucleation by Arp2/3 complex and bacterial actin-based motility. J Cell Biol (1999) 146:1319–32. doi:10.1083/jcb.146.6.1319

51. Ogawa M, Suzuki T, Tatsuno I, Abe H, Sasakawa C. IcsB, secreted via the type III secretion system, is chaperoned by IpgA and required at the post-invasion stage of Shigella pathogenicity. Mol Microbiol (2003) 48:913–31. doi:10.1046/j.1365-2958.2003.03489.x

52. Ogawa M, Yoshimori T, Suzuki T, Sagara H, Mizushima N, Sasakawa C. Escape of intracellular Shigella from autophagy. Science (2005) 307:727–31. doi:10.1126/science.1106036

53. Suzuki T, Miki H, Takenawa T, Sasakawa C. Neural Wiskott-Aldrich syndrome protein is implicated in the actin-based motility of Shigella flexneri. EMBO J (1998) 17:2767–76. doi:10.1093/emboj/17.10.2767

54. Suzuki T, Mimuro H, Miki H, Takenawa T, Sasaki T, Nakanishi H, et al. Rho family GTPase Cdc42 is essential for the actin-based motility of Shigella in mammalian cells. J Exp Med (2000) 191:1905–20. doi:10.1084/jem.191.11.1905

55. Baxt LA, Goldberg MB. Host and bacterial proteins that repress recruitment of LC3 to Shigella early during infection. PLoS One (2014) 9:e94653. doi:10.1371/journal.pone.0094653

56. Mostowy S, Sancho-Shimizu V, Hamon M, Simeone R, Brosch R, Johansen T, et al. p62 and NDP52 Proteins target intracytosolic Shigella and Listeria to different autophagy pathways. J Biol Chem (2011) 286:26987–95. doi:10.1074/jbc.M111.223610

57. Noad J, Von Der Malsburg A, Pathe C, Michel MA, Komander D, Randow F. LUBAC-synthesized linear ubiquitin chains restrict cytosol-invading bacteria by activating autophagy and NF-κB. Nat Microbiol (2017) 2. doi:10.1038/nmicrobiol.2017.63

58. Dupont N, Lacas-Gervais S, Bertout J, Paz I, Freche B, Van Nhieu GT, et al. Shigella phagocytic vacuolar membrane remnants participate in the cellular response to pathogen invasion and are regulated by autophagy. Cell Host Microbe (2009) 6:137–49. doi:10.1016/j.chom.2009.07.005

59. Srougi MC, Burridge K. Undressing a cellular corset: septins exposed. Nat Cell Biol (2009) 11:9–10. doi:10.1038/ncb0109-9

60. Mostowy S, Bonazzi M, Hamon MAA, Tham TN, Mallet A, Lelek M, et al. Entrapment of intracytosolic bacteria by septin cage-like structures. Cell Host Microbe (2010) 8:433–44. doi:10.1016/j.chom.2010.10.009

61. Sirianni A, Krokowski S, Lobato-Marquez D, Buranyi S, Pfanzelter J, Galea D, et al. Mitochondria mediate septin cage assembly to promote autophagy of Shigella. EMBO Rep (2016) 17:1029–43. doi:10.15252/embr.201541832

62. Carneiro LAM, Travassos LH, Soares F, Tattoli I, Magalhaes JG, Bozza MT, et al. Shigella induces mitochondrial dysfunction and cell death in nonmyleoid cells. Cell Host Microbe (2009) 5:123–36. doi:10.1016/j.chom.2008.12.011

63. Suzuki T, Franchi L, Toma C, Ashida H, Ogawa M, Yoshikawa Y, et al. Differential regulation of caspase-1 activation, pyroptosis, and autophagy via Ipaf and ASC in Shigella-infected macrophages. PLoS Pathog (2007) 3:e111. doi:10.1371/journal.ppat.0030111

64. Inohara N, Koseki T, Lin J, Del Peso L, Lucas PC, Chen FF, et al. An induced proximity model for NF-kappa B activation in the Nod1/RICK and RIP signaling pathways. J Biol Chem (2000) 275:27823–31. doi:10.1074/jbc.M003415200

65. Haselbeck AH, Panzner U, Im J, Baker S, Meyer CG, Marks F. Current perspectives on invasive nontyphoidal Salmonella disease. Curr Opin Infect Dis (2017) 30:498–503. doi:10.1097/QCO.0000000000000398

66. Birmingham CL, Smith AC, Bakowski MA, Yoshimori T, Brumell JH. Autophagy controls Salmonella infection in response to damage to the Salmonella-containing vacuole. J Biol Chem (2006) 281:11374–83. doi:10.1074/jbc.M509157200

67. Jia K, Thomas C, Akbar M, Sun Q, Adams-Huet B, Gilpin C, et al. Autophagy genes protect against Salmonella typhimurium infection and mediate insulin signaling-regulated pathogen resistance. Proc Natl Acad Sci U S A (2009) 106:14564–9. doi:10.1073/pnas.0813319106

68. Perrin AJ, Jiang X, Birmingham CL, So N, Brumell JH. Recognition of bacteria in the cytosol of mammalian cells by the ubiquitin system. Curr Biol (2004) 14:806–11. doi:10.1016/j.cub.2004.04.033

69. Zheng YT, Shahnazari S, Brech A, Lamark T, Johansen T, Brumell JH. The adaptor protein p62/SQSTM1 targets invading bacteria to the autophagy pathway. J Immunol (2009) 183:5909–16. doi:10.4049/jimmunol.0900441

70. Wild P, Farhan H, Mcewan DG, Wagner S, Rogov VV, Brady NR, et al. Phosphorylation of the autophagy receptor optineurin restricts Salmonella growth. Science (2011) 333:228–33. doi:10.1126/science.1205405

71. Fiskin E, Bionda T, Dikic I, Behrends C. Global analysis of host and bacterial ubiquitinome in response to Salmonella typhimurium infection. Mol Cell (2016) 62:967–81. doi:10.1016/j.molcel.2016.04.015

72. Guernsey DL, Jiang H, Bedard K, Evans SC, Ferguson M, Matsuoka M, et al. Mutation in the gene encoding ubiquitin ligase LRSAM1 in patients with Charcot-Marie-Tooth disease. PLoS Genet (2010) 6:e1001081. doi:10.1371/journal.pgen.1001081

73. Huett A, Heath RJ, Begun J, Sassi SO, Baxt LA, Vyas JM, et al. The LRR and RING domain protein LRSAM1 is an E3 ligase crucial for ubiquitin-dependent autophagy of intracellular Salmonella typhimurium. Cell Host Microbe (2012) 12:778–90. doi:10.1016/j.chom.2012.10.019

74. Heath RJ, Goel G, Baxt LA, Rush JS, Mohanan V, Paulus GLCLC, et al. RNF166 determines recruitment of adaptor proteins during antibacterial autophagy. Cell Rep (2016) 17:2183–94. doi:10.1016/j.celrep.2016.11.005

75. Van Wijk SJL, Fricke F, Herhaus L, Gupta J, Hotte K, Pampaloni F, et al. Linear ubiquitination of cytosolic Salmonella typhimurium activates NF-kappaB and restricts bacterial proliferation. Nat Microbiol (2017) 2:17066. doi:10.1038/nmicrobiol.2017.66

76. Polajnar M, Dietz MS, Heilemann M, Behrends C. Expanding the host cell ubiquitylation machinery targeting cytosolic Salmonella. EMBO Rep (2017) 18:1572–85. doi:10.15252/embr.201643851

77. Shahnazari S, Yen W-LL, Birmingham CL, Shiu J, Namolovan A, Zheng YT, et al. A diacylglycerol-dependent signaling pathway contributes to regulation of antibacterial autophagy. Cell Host Microbe (2010) 8:137–46. doi:10.1016/j.chom.2010.07.002

78. Thurston TLM, Wandel MP, Muhlinen N, Foeglein Á, Randow F. Galectin 8 targets damaged vesicles for autophagy to defend cells against bacterial invasion. Nature (2012) 482:414–8. doi:10.1038/nature10744

79. Cemma M, Kim PK, Brumell JH. The ubiquitin-binding adaptor proteins p62/SQSTM1 and NDP52 are recruited independently to bacteria-associated microdomains to target Salmonella to the autophagy pathway. Autophagy (2011) 7:341–5. doi:10.4161/auto.7.3.14046

80. Lam GY, Czuczman MA, Higgins DE, Brumell JH. Interactions of Listeria monocytogenes with the autophagy system of host cells. Adv Immunol (2012) 113:7–18. doi:10.1016/B978-0-12-394590-7.00008-7

81. Radoshevich L, Cossart P. Listeria monocytogenes: towards a complete picture of its physiology and pathogenesis. Nat Rev Microbiol (2018) 16:32–46. doi:10.1038/nrmicro.2017.126

82. Beauregard KE, Lee KD, Collier RJ, Swanson JA. pH-dependent perforation of macrophage phagosomes by listeriolysin O from Listeria monocytogenes. J Exp Med (1997) 186:1159–63. doi:10.1084/jem.186.7.1159

83. Cossart P, Vicente MF, Mengaud J, Baquero F, Perez-Diaz JC, Berche P. Listeriolysin O is essential for virulence of Listeria monocytogenes: direct evidence obtained by gene complementation. Infect Immun (1989) 57:3629–36.

84. Henry R, Shaughnessy L, Loessner MJ, Alberti-Segui C, Higgins DE, Swanson JA. Cytolysin-dependent delay of vacuole maturation in macrophages infected with Listeria monocytogenes. Cell Microbiol (2006) 8:107–19. doi:10.1111/j.1462-5822.2005.00604.x

85. Portnoy DA, Jacks PS, Hinrichs DJ. Role of hemolysin for the intracellular growth of Listeria monocytogenes. J Exp Med (1988) 167:1459–71. doi:10.1084/jem.167.4.1459

86. Shaughnessy LM, Hoppe AD, Christensen KA, Swanson JA. Membrane perforations inhibit lysosome fusion by altering pH and calcium in Listeria monocytogenes vacuoles. Cell Microbiol (2006) 8:781–92. doi:10.1111/j.1462-5822.2005.00665.x

87. Mitchell G, Ge L, Huang Q, Chen C, Kianian S, Roberts MF, et al. Avoidance of autophagy mediated by PlcA or ActA is required for Listeria monocytogenes growth in macrophages. Infect Immun (2015) 83:2175–84. doi:10.1128/IAI.00110-15

88. Yoshikawa Y, Ogawa M, Hain T, Yoshida M, Fukumatsu M, Kim M, et al. Listeria monocytogenes ActA-mediated escape from autophagic recognition. Nat Cell Biol (2009) 11:1233–40. doi:10.1038/ncb1967

89. Birmingham CL, Canadien V, Gouin E, Troy EB, Yoshimori T, Cossart P, et al. Listeria monocytogenes evades killing by autophagy during colonization of host cells. Autophagy (2007) 3:442–51. doi:10.4161/auto.4450

90. Cemma M, Lam GY, Stöckli M, Higgins DE, Brumell JH. Strain-specific interactions of Listeria monocytogenes with the autophagy system in host cells. PLoS One (2015) 10:e0125856. doi:10.1371/journal.pone.0125856

91. Pei G, Buijze H, Liu H, Moura-Alves P, Goosmann C, Brinkmann V, et al. The E3 ubiquitin ligase NEDD4 enhances killing of membrane-perturbing intracellular bacteria by promoting autophagy. Autophagy (2017) 13(12):2041–55. doi:10.1080/15548627.2017.1376160

92. Van De Velde LA, Guo XJ, Barbaric L, Smith AM, Oguin TH III, Thomas PG, et al. Stress kinase GCN2 controls the proliferative fitness and trafficking of cytotoxic T cells independent of environmental amino acid sensing. Cell Rep (2016) 17:2247–58. doi:10.1016/j.celrep.2016.10.079

93. Tattoli I, Sorbara MT, Philpott DJ, Girardin SE. Stalling autophagy: a new function for Listeria phospholipases. Microb Cell (2014) 1:48–50. doi:10.15698/mic2014.01.124

94. Tattoli I, Sorbara MT, Vuckovic D, Ling A, Soares F, Carneiro LA, et al. Amino acid starvation induced by invasive bacterial pathogens triggers an innate host defense program. Cell Host Microbe (2012) 11:563–75. doi:10.1016/j.chom.2012.04.012

95. Tattoli I, Sorbara MT, Yang C, Tooze SA, Philpott DJ, Girardin SE. Listeria phospholipases subvert host autophagic defenses by stalling pre-autophagosomal structures. EMBO J (2013) 32:3066–78. doi:10.1038/emboj.2013.234

96. Yano T, Mita S, Ohmori H, Oshima Y, Fujimoto Y, Ueda R, et al. Autophagic control of Listeria through intracellular innate immune recognition in Drosophila. Nat Immunol (2008) 9:908–16. doi:10.1038/ni.1634

97. Fraser DW, Tsai TR, Orenstein W, Parkin WE, Beecham HJ, Sharrar RG, et al. Legionnaires’ disease: description of an epidemic of pneumonia. N Engl J Med (1977) 297:1189–97. doi:10.1056/NEJM197712012972201

98. Horwitz MA. Formation of a novel phagosome by the Legionnaires’ disease bacterium (Legionella pneumophila) in human monocytes. J Exp Med (1983) 158:1319–31. doi:10.1084/jem.158.4.1319

99. Jo E-K, Yuk J-M, Shin D-M, Sasakawa C. Roles of autophagy in elimination of intracellular bacterial pathogens. Front Immunol (2013) 4. doi:10.3389/fimmu.2013.00097

100. Sherwood RK, Roy CR. Autophagy evasion and endoplasmic reticulum subversion: the Yin and Yang of Legionella intracellular infection. Annu Rev Microbiol (2016) 70:413–33. doi:10.1146/annurev-micro-102215-095557

101. Kagan JC, Roy CR. Legionella phagosomes intercept vesicular traffic from endoplasmic reticulum exit sites. Nat Cell Biol (2002) 4:945–54. doi:10.1038/ncb883

102. Eskelinen E-L. Maturation of autophagic vacuoles in mammalian cells. Autophagy (2005) 1:1–10. doi:10.4161/auto.1.1.1270

103. Amer AO, Swanson MS. Autophagy is an immediate macrophage response to Legionella pneumophila. Cell Microbiol (2005) 7:765–78. doi:10.1111/j.1462-5822.2005.00509.x

104. Dubuisson JF, Swanson MS. Mouse infection by Legionella, a model to analyze autophagy. Autophagy (2006) 2:179–82. doi:10.4161/auto.2831

105. Joshi AD, Sturgill-Koszycki S, Swanson MS. Evidence that Dot-dependent and -independent factors isolate the Legionella pneumophila phagosome from the endocytic network in mouse macrophages. Cell Microbiol (2001) 3:99–114. doi:10.1046/j.1462-5822.2001.00093.x

106. Berger KH, Isberg RR. Two distinct defects in intracellular growth complemented by a single genetic locus in Legionella pneumophila. Mol Microbiol (1993) 7:7–19. doi:10.1111/j.1365-2958.1993.tb01092.x

107. Marra A, Blander SJ, Horwitz MA, Shuman HA. Identification of a Legionella pneumophila locus required for intracellular multiplication in human macrophages. Proc Natl Acad Sci U S A (1992) 89:9607–11. doi:10.1073/pnas.89.20.9607

108. Choy A, Dancourt J, Mugo B, O’connor TJ, Isberg RR, Melia TJ, et al. The Legionella effector RavZ inhibits host autophagy through irreversible Atg8 deconjugation. Science (2012) 338:1072–6. doi:10.1126/science.1227026

109. Kubori T, Bui XT, Hubber A, Nagai H. Legionella RavZ plays a role in preventing ubiquitin recruitment to bacteria-containing vacuoles. Front Cell Infect Microbiol (2017) 7:384. doi:10.3389/fcimb.2017.00384

110. Horenkamp FA, Kauffman KJ, Kohler LJ, Sherwood RK, Krueger KP, Shteyn V, et al. The Legionella anti-autophagy effector RavZ targets the autophagosome via PI3P- and curvature-sensing motifs. Dev Cell (2015) 34:569–76. doi:10.1016/j.devcel.2015.08.010

111. Rolando M, Escoll P, Nora T, Botti J, Boitez V, Bedia C, et al. Legionella pneumophila S1P-lyase targets host sphingolipid metabolism and restrains autophagy. Proc Natl Acad Sci U S A (2016) 113:1901–6. doi:10.1073/pnas.1522067113

112. Takabe K, Paugh SW, Milstien S, Spiegel S. “Inside-out” signaling of sphingosine-1-phosphate: therapeutic targets. Pharmacol Rev (2008) 60:181–95. doi:10.1124/pr.107.07113

113. Bedia C, Levade T, Codogno P. Regulation of autophagy by sphingolipids. Anticancer Agents Med Chem (2011) 11:844–53. doi:10.2174/187152011797655131

114. Hardie DG. AMP-activated protein kinase: an energy sensor that regulates all aspects of cell function. Genes Dev (2011) 25:1895–908. doi:10.1101/gad.17420111

115. Ataullakhanov FI, Vitvitsky VM. What determines the intracellular ATP concentration. Biosci Rep (2002) 22:501–11. doi:10.1023/A:1022069718709

116. Kim J, Kundu M, Viollet B, Guan K-L. AMPK and mTOR regulate autophagy through direct phosphorylation of Ulk1. Nat Cell Biol (2011) 13:132–41. doi:10.1038/ncb2152

117. Park DW, Jiang S, Tadie JM, Stigler WS, Gao Y, Deshane J, et al. Activation of AMPK enhances neutrophil chemotaxis and bacterial killing. Mol Med (2013) 19:387–98. doi:10.2119/molmed.2013.00065

118. Leick L, Fentz J, Bienso RS, Knudsen JG, Jeppesen J, Kiens B, et al. PGC-1{alpha} is required for AICAR-induced expression of GLUT4 and mitochondrial proteins in mouse skeletal muscle. Am J Physiol Endocrinol Metab (2010) 299:E456–65. doi:10.1152/ajpendo.00648.2009

119. Yang CS, Kim JJ, Lee HM, Jin HS, Lee SH, Park JH, et al. The AMPK-PPARGC1A pathway is required for antimicrobial host defense through activation of autophagy. Autophagy (2014) 10:785–802. doi:10.4161/auto.28072

120. Rottiers V, Najafi-Shoushtari SH, Kristo F, Gurumurthy S, Zhong L, Li Y, et al. MicroRNAs in metabolism and metabolic diseases. Cold Spring Harb Symp Quant Biol (2011) 76:225–33. doi:10.1101/sqb.2011.76.011049

121. Kim JK, Kim TS, Basu J, Jo E-KK. MicroRNA in innate immunity and autophagy during mycobacterial infection. Cell Microbiol (2017) 19. doi:10.1111/cmi.12687

122. Davalos A, Goedeke L, Smibert P, Ramirez CM, Warrier NP, Andreo U, et al. miR-33a/b contribute to the regulation of fatty acid metabolism and insulin signaling. Proc Natl Acad Sci U S A (2011) 108:9232–7. doi:10.1073/pnas.1102281108

123. He C, Klionsky DJ. Regulation mechanisms and signaling pathways of autophagy. Annu Rev Genet (2008) 43:67–93. doi:10.1146/annurev-genet-102808-114910

124. Settembre C, Di Malta C, Polito VA, Garcia Arencibia M, Vetrini F, Erdin S, et al. TFEB links autophagy to lysosomal biogenesis. Science (2011) 332:1429–33. doi:10.1126/science.1204592

125. Ganesan R, Hos NJ, Gutierrez S, Fischer J, Stepek JM, Daglidu E, et al. Salmonella typhimurium disrupts Sirt1/AMPK checkpoint control of mTOR to impair autophagy. PLoS Pathog (2017) 13:e1006227. doi:10.1371/journal.ppat.1006227

126. Lan F, Cacicedo JM, Ruderman N, Ido Y. SIRT1 modulation of the acetylation status, cytosolic localization, and activity of LKB1. Possible role in AMP-activated protein kinase activation. J Biol Chem (2008) 283:27628–35. doi:10.1074/jbc.M805711200

127. Walthers D, Carroll RK, Navarre WW, Libby SJ, Fang FC, Kenney LJ. The response regulator SsrB activates expression of diverse Salmonella pathogenicity island 2 promoters and counters silencing by the nucleoid-associated protein H-NS. Mol Microbiol (2007) 65:477–93. doi:10.1111/j.1365-2958.2007.05800.x

128. Grant AJ, Morgan FJ, Mckinley TJ, Foster GL, Maskell DJ, Mastroeni P. Attenuated Salmonella typhimurium lacking the pathogenicity island-2 type 3 secretion system grow to high bacterial numbers inside phagocytes in mice. PLoS Pathog (2012) 8:e1003070. doi:10.1371/journal.ppat.1003070

129. Liu X, Wang N, Zhu Y, Yang Y, Chen X, Chen Q, et al. Extracellular calcium influx promotes antibacterial autophagy in Escherichia coli infected murine macrophages via CaMKKbeta dependent activation of ERK1/2, AMPK and FoxO1. Biochem Biophys Res Commun (2016) 469:639–45. doi:10.1016/j.bbrc.2015.12.052

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Siqueira, Ribeiro and Travassos. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

REVIEW
published: 07 June 2018
doi: 10.3389/fimmu.2018.01306

Integrated Stress Responses to Bacterial Pathogenesis Patterns

Larissa O. C. P. Rodrigues, Rodrigo S. F. Graça and Leticia A. M. Carneiro*

Laboratório de Inflamação e Imunidade, Instituto de Microbiologia Paulo de Góes, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil

OPEN ACCESS

Edited by:

Francesca Granucci, Università degli studi di Milano Bicocca, Italy

Reviewed by:

Michaela Semeraro, INSERM U1151 Institut Necker Enfants Malades Centre de médecine moléculaire (INEM), France
Barbara M. Bröker, University of Greifswald, Germany

*Correspondence:

Leticia A. M. Carneiro
leticiac@micro.ufrj.br

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 20 February 2018
Accepted: 25 May 2018
Published: 07 June 2018

Citation:

Rodrigues LOCP, Graça RSF and Carneiro LAM (2018) Integrated Stress Responses to Bacterial Pathogenesis Patterns. Front. Immunol. 9:1306. doi: 10.3389/fimmu.2018.01306

Activation of an appropriate innate immune response to bacterial infection is critical to limit microbial spread and generate cytokines and chemokines to instruct appropriate adaptive immune responses. Recognition of bacteria or bacterial products by pattern recognition molecules is crucial to initiate this response. However, it is increasingly clear that the context in which this recognition occurs can dictate the quality of the response and determine the outcome of an infection. The cross talk established between host and pathogen results in profound alterations on cellular homeostasis triggering specific cellular stress responses. In particular, the highly conserved integrated stress response (ISR) has been shown to shape the host response to bacterial pathogens by sensing cellular insults resulting from infection and modulating transcription of key genes, translation of new proteins and cell autonomous antimicrobial mechanisms such as autophagy. Here, we review the growing body of evidence demonstrating a role for the ISR as an integral part of the innate immune response to bacterial pathogens.

Keywords: eukaryotic translation initiation factor 2 alpha, cellular stress, bacterial pathogens, heme-regulated eIF2α kinase, general control non-derepressible 2, PKR-like ER kinase, PKR

INTRODUCTION

Microbial sensing by pattern recognition molecules (PRMs) triggers a robust innate immune response with the production of cytokines, chemokines, and antimicrobial factors (1–4). In the last decade, the concept that, in addition to microbial-associated molecular patterns recognition by PRMs, the host response can be tuned by the recognition of alterations in homeostasis induced by pathogens during progression of disease has been established (5–7). Such alterations on cell homeostasis allow the host to differentiate pathogenic organisms from those that do not represent a threat and, thus, adequate the immune responses to deal with the attack being mounted accordingly. It is interesting that despite the multitude of virulence mechanisms among bacterial species, most of them converge to few common “patterns of pathogenesis” that include membrane damage, access to the cytosol, disruption of cytoskeleton, and protein aggregation among others (5–10). In a sense, these patterns of pathogenesis would align with the concept of danger-associated molecular patterns (DAMPs), which are host molecules whose presence indicate that there has been tissue damage such as, for example, extracellular ATP or the chromatin-associated protein high-mobility group box 1 (11–13). But they are not exactly the same as DAMPs as these are molecules that are released as a result of cellular death and that are recognized by receptors on other cells whereas patterns of pathogenesis induce alterations of cell homeostasis during infection and provides the infected cell with information to mount a more refined response and to adapt and, in many cases recover from the insult.

In this context, the cellular mechanisms to sense and respond to stress can be regarded as an integral part of the innate immune response. The integrated stress response (ISR), a common adaptive pathway that eukaryotic cells activate in response to diverse stress stimuli is one such mechanism. The core event in this pathway is the phosphorylation of eukaryotic translation initiation factor 2 alpha (eIF2α) by one or more of four members of the eIF2α kinase family (6). The phosphorylation of eIF2α results in a marked decrease in global protein synthesis accompanied by the induction of selected genes, including the transcription factor ATF4, both of which are important to promote cellular recovery (6, 7, 14). This type of response to stress mediated by the eIF2α kinases, parallels those mediated by the mTOR pathway or by autophagy in the sense that are highly conserved signaling modules that regulate essential metabolic circuits, both in homeostatic and stress conditions, from yeast to mammals (6, 8, 15). In the context of an infection, the power of this type of “sensing system” relies on the fact that it does not recognizes pathogens per se but rather utilizes an ancient system that detects cellular stress/damage to sense insults that are caused by pathogenic bacteria regardless of its specific virulence factors.

In the present review, we focus on the emerging role of the ISR on host response to bacterial pathogens, which only recently began to be appreciated, in contrast to its well-established role in response to viruses. As obligate intracellular pathogens that highjack the host cell machinery to produce its own proteins, the link between viruses and the ISR is more obvious and more generally accepted. The impact of the ISR on viral infections has been extensively reviewed elsewhere (16–21). Here, we discuss recent data that implicate the ISR as an important component of cell autonomous anti-bacterial responses. As an emerging topic, there are still many gaps in our understanding of the mechanisms underlying this process but we believe that our current knowledge already provides a conceptual framework to work with. As much as we tried to bring together evidence of a role for ISR in different bacterial infections, this is by no means an exhaustive review.

eIF2α AND eIF2α-KINASES

Regulation of translation may be useful to coordinate several innate immune functions such as microbial sensing, microbial replication control, and induction of inflammatory cytokines. Translation shut down can help cells to cope with stress conditions and prevent further damage until the insult is gone. However, this happens in a context where cells still need to communicate that they are under attack in order to prevent infection spread and initiate adequate immune responses. Among metabolic sensors, eIF2α kinases have major roles in adjusting the protein synthesis machinery to enhance translation of mRNAs that are relevant to deal with the source of stress, including those induced by PRM activation, while shutting down the translation of unrelated proteins (6, 7, 21). This ability to screen and modulate host protein synthesis can affect the quality of the innate immune responses both at the transcriptional and translational levels. In addition, the gene expression program induced during ISR adjusts the stress response according to cellular context, nature, and intensity of stress stimuli (6, 7). Finally, although ISR is primarily a homeostatic-preserving program by which cells adapt to survive, severe and/or long-lasting stress can tip the balance toward cell death signaling by regulating the cell autonomous processes of autophagy and apoptosis (6, 7, 15).

The eIF2α kinases act as early responders to alterations in cellular homeostasis which is mainly due to the fact that these proteins are at the same time the sensors of stress and the kinases that phosphorylate eIF2α (6, 10, 20, 22). Each kinase dimerizes and autophosphorilates for full activation in response to distinct environmental and physiological types of stress. Double-stranded RNA (dsRNA)-dependent protein kinase (PKR) is activated mainly by dsRNA during viral infection but also by oxidative and ER stress, growth factor deprivation, cytokines, bacterial infections, and ribotoxic stress (23–27). Interestingly, caspase activity in the early stages of apoptosis was also shown to activate PKR, indicating a role for protein synthesis inhibition in apoptosis (28). PKR-like ER kinase (PERK) is activated by accumulation of unfolded proteins in the ER or perturbations in calcium homeostasis, cellular energy, or redox status (29–31). It has also been reported to respond to ATP depletion and subsequent sarcoplasmic/ER Ca2+-ATPase pump inhibition in the context of glucose deprivation in neuronal cells and in pancreatic β cells (32, 33). Heme-regulated eIF2α kinase (HRI) is a sensor for low levels of intracellular heme as well as arsenite-induced oxidative stress, heat shock, nitric oxide, 26S proteasome inhibition, and osmotic stress (34–37). This array of types of stress activate HRI independently of heme but require the presence of heat shock proteins HSP90 and HSP70 (37). General control non-derepressible 2 (GCN2) is highly conserved from yeasts to humans and is activated in response to amino acid deprivation when it binds to deacylated transfer RNAs (tRNAs) via histidyl-tRNA synthetase-related domain (38, 39). As one can appreciate, some types of stress can potentially activate more than one of these four kinases. Most likely, the eIF2α kinases act cooperatively to specifically tune cellular responses stress. Of note, all of these kinases have been reported to have roles independent of eIF2α phosphorylation but here we will focus on the ISR, which signals through eIF2α phosphorylation.

The common signaling hub for all the stress stimuli that activate ISR is phosphorylation of the subunit α of eIF2 on serine 51 (6, 10, 20, 22). eIF2 is constituted by three subunits (α, β, and γ). When bound to GTP and Met-tRNAiMet (initiator methionyl-tRNA), eIF2 form a ternary complex that delivers the initiator tRNA to the 40S ribosomal subunit. eIF2 is released from the ribosome bound to a GDP and to be ready for another round of translation initiation, the eIF2 complex must be recycled back to its active GTP-bound form. The guanine nucleotide exchange factor eIF2B exchanges GDP for GTP on the γ subunit and maintains the levels of the ternary complex available for new rounds of translation. Under a variety of stress conditions, however, phosphorylation of the α subunit of eIF2 at Ser51 blocks general translation initiation, as it converts eIF2 to a competitive inhibitor of eIF2B by blocking the GDP–GTP exchange reaction and reducing the dissociation rate of eIF2 from eIF2B (6, 40, 41). Phosphorylation of eIF2α leads to a global arrest in translation but it does not affect all mRNA transcripts alike. A subset of mRNAs that contain upstream open reading frames and often encode proteins that are important for stress recovery and re-establishment of homeostasis have selective increased translation (6).

One of the genes that are upregulated following eIF2α phosphorylation is the transcription factor ATF4. Studies using ATF4-deficient mice have shown it has critical roles in the regulation of normal metabolic as well as redox processes such as regulation of obesity, glucose homeostasis, energy expenditure, and neural plasticity (42–44). Under stress conditions, increased ATF4 expression represents a signature of the ISR and is mainly due to translational control, as Atf4 is one of those mRNAs that have its translation augmented upon eIF2α phosphorylation in contrast with the general translational arrest observed for most transcripts (6, 45). As a transcription factor, ATF4 can activate several transcriptional programs that will ultimately determine the cell fate—from cell death to re-establishment of homeostasis. The ability of ATF4 to interact with multiple other transcription factors allows it to generate distinct tailored responses to different types of cellular stress. Thus, despite ATF4 being a master common regulator of ISR, its target genes will be highly dependent on stress intensity and cellular context (45–47). For example, when acting in combination with ATF3, ATF4 is a part of a program that aims to re-establish cellular homeostasis and promote survival (48). Conversely, when interacting with CHOP, ATF4 promotes cell death following ER stress (49). In addition to the interacting partners that cooperate with ATF4 to promote transcription of target genes, another set of interacting partners prevent ATF4 transcriptional activity as is the case for PHD3 during hypoxia and TRIB3 during amino acid starvation and ER (50–52).

ISR AND BACTERIAL INFECTIONS

Eukaryotes have evolved in a context of constant interactions with prokaryotes and it is clear that the latter have contributed to shape those organisms throughout evolution. A human being harbors more than 1,000 bacterial species as part of their microbiota and interacts with another incalculable number of bacterial species during its lifetime (53). The vast majority of these interactions does not result in disease and, in many cases, they are actually beneficial to the hosts. However, despite representing less than 1% of the total number of estimated bacterial species in our planet, pathogenic bacteria still cause millions of deaths every year.

In general, those bacteria that are considered as pathogenic are the ones endowed with certain attributes that allow them to (1) colonize the host; (2) find a nutritionally compatible niche in the host body; (3) avoid, subvert, or circumvent the host innate and adaptive immune responses; (4) replicate, using host resources; and (5) exit and spread to a new host (54). However, even though some bacteria display very well-defined virulence attributes, the pathogenic potential of a given bacterium can only really be observed upon interaction with its host. The final outcome of an infection is never the result of bacterial virulence alone but rather a cross talk between the host and the pathogen. This complicates the definition of “true pathogen” as the same bacterial pathogen can have different impact in different individuals. Thus, for the host, it is critical to be able to assess the potential threat that a given pathogen represents in order to establish an appropriate response.

During a bacterial infection, a multitude of signals exchanged by the two organisms establishes a cross talk that will ultimately determine the outcome of the infectious process. Many known bacterial virulence factors are only synthesized when bacteria go through major changes in metabolism in order to adapt to the dynamic conditions of the host environment (55). While doing that, bacterial pathogens may have profound effects on host cell homeostasis that, in turn, trigger cellular stress responses. Bellow, we will discuss how the ISR can be triggered by cellular alterations caused by bacterial infections and the impact of this response on host–pathogen interactions. The data discussed in the next sections are summarized in Figure 1.

FIGURE 1 | Integrated stress response (ISR) activation by bacterial pathogenesis patterns. This figure summarizes how cellular damage induced by different bacterial species is sensed by one or more eukaryotic translation initiation factor 2 alpha (eIF2α) kinases to activate defense mechanisms and homeostatic programs. We intentionally included a simplified representation of pattern recognition molecules (PRMs) recognition of microbes, PAMPs, and danger-associated molecular patterns (DAMPs) in all cartoons to strengthen the notion that these system act together to refine the cell response to the infection. (A) Pathogenesis pattern: bacterial growth; (B) pathogenesis pattern: membrane damage; (C) pathogenesis pattern: access to cytosol; (D) pathogenesis pattern: cytoskeleton disruption and protein aggregation.

BACTERIAL GROWTH

The ability to survive and grow inside the host upon infection is one of the most common pathogenesis patterns as it represents the ability of a given pathogen to scape the host response and establish a replicative niche. For the host, being able to differentiate growing and dying bacteria, especially in the context of an acute infection, is key to mount a proper response. Molecules whose presence could indicate bacterial growth include peptidoglycan fragments released during bacterial cell division, quorum-sensing inducers that are produced once the bacterial population reaches a certain density and bacterial pyrophosphates such as HMB-PP (5, 56–58). Alternatively, instead of direct detection of a molecule, bacterial growth sensing could be achieved by sensing of altered local levels of cellular nutrients such amino acids or oxygen (59–61).

Recently, the definition of PAMPs has been updated to allow the classification of those produced specifically by living microorganisms, the so-called Vita-PAMPs, and those that represent the degradation products of dead microorganisms, named PAPMs-postmortem (PAMPs-PM), as two different categories that have different biological activities (10, 62). Moretti et al. (10) has recently identified cyclic-di-adenosine monophosphate (c-di-AMP), a second messenger that is produced by live Gram-positive bacteria, as a Vita-PAMP. The authors show that phagocytes are able to discriminate live and dead Listeria innocua by sensing this Vita-PAMP through the innate immune sensor stimulator of interferon genes resulting in ER stress, PERK and eIF2α phosphorylation. Subsequently, an autophagic response ensued to sequester stressed ER membranes and prevent stress-induced cell death while also inducing an IFN-dependent response. Importantly, this response was blunted in phagocytes lacking PERK. Finally, following L. monocytogenes infection, mice engineered to have PERK-deficient macrophages presented lower systemic levels of IFN-I and higher bacterial burden on both liver and spleen when compared with WT controls. In this model, at a cellular level, there was no difference between the response induced by live L. innocua, a non-pathogenic bacteria, and live L. monocytogenes, but there was differences when these were compared to dead bacteria. The response to any infection is multilayered and dependent on the interaction of multiple sensing systems—each one of these systems provides the cells with different information that when combined determine the cells response and, ultimately, its fate. In this case, the ISR provided the cells with the ability to distinguish live from dead bacteria, which is crucial to mount appropriate response even though it was not able to differentiate between a pathogenic from a non-pathogenic species. Of note, when the authors tested their hypothesis in vivo, they used only L monocytogenes, most likely because L. innocua would have been readily cleared given its lack of virulence and would have not generated any of the responses observed against L. monocytogenes. This, once again, illustrates how important the context is: in the natural course of a real infection, L. innocua would probably have never caused the systemic infection that L. monocytogenes does and would have not reached circulating phagocytic cells or the liver or the spleen. However, when given directly to these cells in vitro, it induced the same response that the bacterial species that would have encountered these cells during infection. It would also be interesting to investigate if this response is restricted to phagocytic cells or can occur in other cell types.

A contrasting study showed that PERK activation and IFN-I production by myeloid cells during infection with L. monocytogens or treatment with the pore-forming toxin LLO is actually detrimental to the host. In this model, the PERK pathway is amplified by IFN-I resulting in the activation of another eIF2α-kinase, PKR. This, in turn, served as an amplification loop for PERK-signaling leading to excessive ER stress and cell death. Consistent with this, mice deficient on CHOP, a pro-apoptotic factor that is downstream of PERK, are more resistant to L. monocytogenes infection than WT controls (63). This model could provide a partial explanation for why mice lacking IFN-I receptor have been consistently reported to be more resistant to L. monocytogenes than WT mice (64–66).

The opportunistic Gram-negative Pseudomonas aeruginosa causes both acute and chronic infections, especially in the respiratory tract (31, 67). Its ability to scape or subvert the host immune response constitutes its main virulence attribute. P. aeruginosa is able to form biofilms, a complex biological system that protects the bacteria from host immune defense mechanisms and promotes persistent infection. This bacterium coordinates the production of biofilms and other virulence factors using quorum sensing, a cell-to-cell communication system that allow bacteria to perceive their population density by producing and sensing diffusible signal molecules. One of the quorum-sensing auto inducers produced by P. aeruginosa to regulate gene expression and communicate is N-(30oxododecanoyl)-homoserin lactone (HSL-C12) (31, 67). HSL-C12 is a lipid-like diffusible molecule that has multiple effects on mammalian cells including apoptosis and release of Ca2+ from the ER stores. By perturbing ER homeostasis, HSL-C12 induces the activation of PERK and eIF2α phosphorylation resulting in protein synthesis inhibition (31). If in the one hand this inhibition results in increased NF-κB activation and transcription of pro-inflammatory genes because IκB re-synthesis is blocked, on the other hand it prevents the translation into proteins of the transcribed genes resulting in an overall downregulation of the host response and, thus, can be considered a pathogen scape mechanism. This would be one instance where the pathogen evolved to manipulate and take advantage of a cell host sensing system.

As mentioned above, nutrient availability is a critical limitation for invading microorganisms. Iron is a nutrient indispensable for growth of almost living organisms and is unlikely to be readily available for invading microorganisms resulting in fierce competition between host and pathogens (59). Like many other bacteria, P. aeruginosa has developed several mechanisms to acquire iron during infection. In a recent study, it was demonstrated that the iron-chelating siderophore pyoverdine produced by P. aeruginosa limits the concentration of iron in the cell medium resulting in the activation of HRI, eIF2α phosphorylation, and induction of Gadd34 transcription in human bronchial epithelial cells. This response had cytoprotective effect and was turned off when the medium was supplemented with iron (61).

These few examples demonstrate that host cells can detect growth of bacteria by sensing molecules that accumulate as the number of bacteria increase including those that bacteria use to communicate with each other, such as quorum-sensing auto inducers and second messengers as well as molecules that bacteria use to acquire nutrients.

MEMBRANE INTEGRITY

The detection of this type of stress is highly conserved. Damage of the plasma membrane is an archaic threat that needs to be faced with efficient cell autonomous defense mechanisms (5). Recently, a pivotal role for GCN2 in the response to membrane damage has been uncovered in different models. For example, it has been demonstrated that membrane permeabilization by the detergent digitonin induces a robust response characterized by GCN2 phosphorilation and ATF3 expression (68). In Drosophila, the damage caused by Pseudomonas entomophila in gut cells induces a starvation-like state, resulting in GCN2 and eIF2α phosphorylation and concomitant inhibition of the mTOR pathway by the AMP-activated kinase (AMPK). In this model, these two stress response pathways together shut down translation of new proteins and trigger innate immune responses (69).

In mammalian cells, disturbance of membrane integrity caused by bacterial pathogens can also trigger stress responses (8, 15, 68). Pore-forming toxins represent an important class of bacterial exoproducts that can induce membrane damage leading to stress responses (70). In human epithelial cells, the α-toxin produced by Staphylococcus aureus induces the formation of pores on cellular membranes resulting in potassium efflux, failure of nutrient transport and loss of ATP which, in turn, activates both GCN2 and the energy sensor AMPK, with subsequent eIF2α phosphorylation and mTORC1 deactivation (similar to what was reported in Drosophila) (15, 71). Low intracellular concentrations of potassium is known to trigger several responses in infected or stressed cells including the activation of inflammasomes and caspases (15, 71), as well as activation of multiple kinases such as p38 and CREB, in addition to the aforementioned AMPK and GCN2 (71–73). Activation of GCN2 induced by potassium efflux caused by membrane perforation indicates that cells may exploit the dependence of nutrient transport across the plasma membrane on physiological ion gradients to indirectly sense perturbations on ion concentration. Both removal of the pore from the plasma membrane by dynamin-dependent endocytosis and the metabolic reprogramming activated by the ISR are essential for cellular recovery as cells that are not able to activate this program are more susceptible to α-toxin (74, 75).

Invasive bacteria such as Salmonella Tiphymurium, Shigella flexneri, and Listeria monocytogenes also cause membrane damage during their internalization process. Similar to what was described above, all three bacteria trigger an acute intracellular amino acid starvation program that induces stress responses dependent on GCN2 and eIF2α phosphorylation at the same time as it disarms mTOR signaling unleashing an autophagic response (8, 68). However, the response that ensues is different for these three bacteria. (i) During infection with S. flexneri, a Gram-negative bacterium that escapes to and replicates in the host cell cytoplasm, amino acid starvation persists up to 4 h after infection allowing not only the induction of autophagy but also GCN2- and eIF2α-dependent formation of stress granules in the cytosol as well as reprogramming of the transcriptional response orchestrated by ATF3 (8, 76). (ii) L. monocytogenes, a Gram-positive bacterium that similar to S. flexneri escapes to and replicates in the cytosol, also triggers a state of amino acid starvation characterized by activation of GCN2, eIF2α phosphorylation, and transcriptional upregulation of ATF3. In this case, however, this response is very transient peaking at 1 h and is completely normalized after 4 h post-infection. The kinetics of this response parallels the kinetics of the pore-forming toxin LLO-dependent scape from the internalization vacuole and coincides with the maximal targeting of L. monocytogenes to autophagosomes (68). (iii) Salmonella, in contrast to the bacteria described above, remains in vesicles known as Salmonella-containing vacuoles (SCV) after its internalization. The damages to the SCV membranes trigger the same GCN2-dependent early amino acid starvation program described above. However, following Salmonella infection membrane integrity and cytosolic amino acid concentration are readily normalized allowing mTOR to be reactivated at the surface of the SCV and promoting bacterial scape from autophagy (8). Thus, these three model invasive bacteria all induce GCN2-dependent ISR during their entry processes but each one of them deal with it in different ways once again highlighting that bacteria have also evolved to counteract ISR-mediated responses.

Adherent-invasive Escherichia coli (AIEC), which is abnormally abundant in the intestinal mucosa of Crohn’s disease patients, also induces phosphorylation of GCN2 with subsequent eIF2α phosphorylation and increased ATF4 levels. Upon activation of this pathway, ATF4 binds to promoters of multiple autophagy-related genes including MAP1LC3B, Becn1, SQSTM1, ATG3, and ATG7. This is necessary to initiate autophagy and restrict bacterial growth as depleting cells from GCN2 resulted in impaired autophagy, increased bacterial replication, and elevated pro-inflammatory cytokine production both in vitro and in vivo. The authors go on to show that the GCN2–eIF2α–ATF4 pathway is activated in ileal biopsies from patients with noninflamed Crohn’s disease but not on those with inflamed Crohn’s disease, indicating that failure to activate this stress response could be one of the mechanisms contributing to active disease (77).

Thus, it appears that a nutrient sensor, GCN2, may also function as a sentinel of membrane integrity and that the responses it triggers are essential to prevent abyssal ATP loss and irreversible damage. In addition, in the case of invasive pathogens, this response might affect their ability to replicate within the host cell due to increase in autophagic activity as a consequence of amino acid starvation as well as production of inflammatory factors induced by the stress transcription factors ATF3 and ATF4.

ACCESS TO CYTOSOL

Many pathogens are able to deliver molecules directly into the cytosol of host cell. This may be achieved by AB-toxins when the B subunit binds to specific receptors on the surface of the cells and translocates the active subunit A into the cell (78), by pore-forming toxins such as listeriolysin O (mentioned above) and streptolysin O (70), or secretion systems such as the type III secretion systems of Yersinia and Salmonella (79), the type IV secretion system of Legionella, Coxiella, and Brucella (80), and the type VI secretion system of Pseudomonas and Vibrio (81, 82).

Shiga-toxigenic E. coli produces Shiga toxin (Stx) 1 and 2 that cause hemorrhagic colitis and hemolytic uremic syndrome. A newly described toxin, namely subtilase cytotoxin (SubAB), was shown to bind to and be internalized by target cells through clathrin-, lipid rafts-, and actin-dependent pathways. Once it reaches the ER, SubAB cleaves the chaperone Bip/Grp78 initiating an ER-stress induced ISR resulting in cytotoxicity. This response also included the formation of stress granules induced not only by PERK but also as a result of PKR activation (14).

Yang et al. (83) show that P. aeruginosa infection induces a strong activation of the GCN2–eIF2α–ATF4 pathway that is largely dependent on production of pyocianin during initial infection and that ultimately results in bacterial clearance through autophagy. Pyocianin is a cell permeable toxin considered to be a major virulence factor for P aeruginosa. In vivo, in rats, infection with a mutant bacterial strain that does not produce pyocianin and, thus, does not activate of the GCN2–eIF2α–ATF4 pathway results in higher number of colony-forming units in the lungs, more extensive alveolar wall thickening and higher mortality when compared to infection with the WT strain. Although indirect, these data suggest a role for the ISR in preventing prolonged infection and immunopathology. Interestingly, reduction of pyocianin production by P. aeruginosa in chronic airways infections has been associated with better host adaptation and worse outcomes in cystic fibrosis patients (84).

ACTIN CYTOSKELETON DISRUPTION

Another common feature employed by various highly divergent pathogenic bacterial species is the disruption of the host cell cytoskeleton. Invasive bacteria such as S. flexneri, L. monocytogenes, Mycobacterium marinum, and Rickettsial species exploit the actin-based motility to move inside the cell and from one cell to the other without never being exposed to immune defenses outside the cells (5, 85). Other bacterial pathogens, such as E. coli and Citrobacter freundii, produce hallmark attaching and effacing lesions that are characterized by localized destruction of the brush border villi of enterocytes, intimate attachment of bacteria to the residual apical membrane and formation of a dense plaque of actin cytoskeletal filaments beneath adherent bacteria that is essential for their pathogenesis (86, 87). Finally, some pathogens manipulate host actin cytoskeleton to either induce their own uptake or to avoid phagocytosis (88–90).

Polysomes, mRNAs, elongations factors, and aminoacyl-tRNA synthetases are found associated with actin filaments indicating that the cytoskeleton might actually act as a platform to facilitate the assembly of components involved translation (91–93). GCN2 has been recently implicated as a sensor of F-actin depolymerization. Disruption of the actin cytoskeleton by drugs such as latruculin-B and cytochalasin-D induces GCN2 activation followed by eIF2α phosphorylation, attenuation of global translation, and augmented ATF4 and CHOP expression (94). In nutrient-replete cells, GCN2 is kept in a latent state by the interaction with other proteins such as the eukaryotic elongation factor 1A (eEF1A) that delivers aminoacyl-tRNAs to ribosomes during the elongation step of protein synthesis (95, 96). During starvation periods, however, uncharged tRNA displaces eEF1A from GCN2 allowing its autophosphorilation and eIF2α phosphorylation (97, 98). Another binding partner of eEF1A is F-actin. In yeasts, the same mutations that affect binding of eEF1A to aminoacyl-tRNAs also result in actin binding and buding defects that lead to GCN2-dependent eIF2α phosphorylation (99, 100). Thus, it has been proposed that upon F-actin disruption eEF1A is displaced from GCN2 and bound to F-actin leaving GCN2 free to initiate the ISR (94). In addition, F-actin disruption also leads to deacylated tRNA accumulation, which in turn might also contribute to the activation of GCN2 resulting in global protein synthesis arrest and reduction of amino acylated tRNA levels (94).

As mentioned above, two invasive pathogens—L. monocytogenes and S. flexneri—that exploit the actin cytoskeleton of the cell to move around the cell and infect neighboring cells were shown to induce a GCN2-dependent starvation program as a consequence of membrane damage. It is possible that disruption of F-actin could impede the proper function of amino acid transporters on the plasma membrane triggering this response. In summary, infection with L. monocytogenes and S. flexneri could potentially activate GCN2 in multiple ways: when bacteria escape from the vacuole into the cytosol causing membrane damage (as it has been experimentally demonstrated) or by disrupting the actin cytoskeleton.

PROTEIN AGGREGATION

Heme-regulated eIF2α kinase is able to sense and respond to a variety of types of cellular stress including heme deprivation, oxidative stress, heat shock, and proteasome inhibition, all of which are known to result in accumulation of misfolded protein aggregates in the cytosol (35, 36, 101, 102). As it has been previously shown that infection with bacterial pathogens trigger the formation of large PRM oligomeric complexes in the cytosol, one may speculate that this is the common feature among all these types of stresses that is actually sensed by HRI (103–106). This could serve as a sensing system to monitor misfolding of large protein complexes and formation of toxic aggregates in the cytosol and trigger damage control mechanisms such as ISR and autophagy.

CONCLUDING REMARKS

Even at a single cell level, the response to an infection is multilayered and involves sensing, effector, and homeostatic mechanisms. Each one of these elements has, in itself, multiple layers of complexity and, together, they generate a full-blown response. Sensing of microbes or their products by PRMs is pivotal and activates robust inflammatory responses. Since the discovery of PRMs, there has been much debate on how the cells can tailor the response to specific pathogens using a limited number of receptors that recognize structures that are present in many different microorganisms, including non-pathogenic. This can be achieved by different means including the combinatorial effect of several PRMs (107), the compartmentalization of PRMs that only allows recognition of certain PAMPs when presented in specific compartments of the cell (108) and the sensing of vita-PAMPs versus PAMPs-PM (62). The recognition of pathogenesis patterns by the ISR represents another layer in the host response. Sensing alterations on homeostasis and cell damaged caused by infection can instruct the host to generate a more refined and specific response while triggering protective gene expression programs that enable cells to recover from the initial stress and re-establish homeostasis. Given its origins early on evolution, stress responses may actually represent an ancient innate defense mechanism against invading pathogens.

In this review, we discussed evidence showing that the ISR can have an important role in shaping the autonomous cell response to bacteria with varying levels of virulence. In this context, the ISR acts in concert with other sensing systems to adequate the response to the threat. Thus, the ISR during bacterial infection cannot be analyzed isolated from the context. This generates a complexity that represents a challenge for dissecting the precise role and the relevance of each component in the final response. While there are still many gaps to be filled before we have a more comprehensive overview, the picture that emerges is that the ISR can influence the quality of the response initiated by innate immune recognition.

For the most part, the studies discussed in this review show that several bacteria are able to activate or manipulate the ISR during infection, through different eIF2α kinases and signaling pathways, resulting in specific transcriptional programs. However, in many cases, it is yet to be defined how this affects the outcome of the infection. In some cases, it is clear that it can affect the ensuing immune response. For example, ISR activation in phagocytic cells infected by Listeria was shown to be critical for IFN-I production and bacterial clearance. On the other hand, eIF2α phosphorylation induced by the HSL-C12 from P. aeruginosa results in downregulation of translation of pro-inflammatory cytokines such as IL-6 and KC. We believe that understanding how the ISR can affect qualitatively the response to a given pathogen is a major avenue for future work. In this sense, as we progress, it would be important to determine how the ISR impact on how cells communicate infection to the neighboring cells as well as to immune cells and how this can qualitatively affect the immune response, including in subsequent exposures to the same pathogen. Finally, it would be interesting to see if the homeostatic adaptations during infection can lead to persistent alterations in the infected cell rendering it more resistant to following infections.

The interplay between ISR and autophagy is also a common theme in most of the studies mentioned here. Interestingly, two studies discussed above showed that in the absence of the GCN2–eIF2α–ATF4-autophagy pathway, opportunistic bacteria such as AIEC and P. aeruginosa establish persistent infection that perpetuate inflammation contributing to worsen the pathology in Crohn’s disease and cystic fibrosis, respectively. It will interesting to investigate how ISR could affect the development of chronic complex diseases that are known to have a microbial component to it such as the two mentioned above.

As we mentioned above, co-evolution of pathogens and their hosts have shaped (and continue to do so) their interactions. In this constant arms race, both sides try to adapt in order to survive. Thus, it should come as no surprise that some bacteria might be able to scape or even take advantage of the ISR to manipulate the host cell response. Indeed, being able to subvert host responses is part of the very definition of what a pathogen is. In this case, failure to activate the ISR properly could lead the host to underestimate the infectious threat. As it is well documented for several viruses, we expect that as our knowledge increases, we will uncover many bacterial strategies to tamper with the ISR.

Because of the significant overlap between the eIF2α kinases in addition to the complexity of many host–pathogen interactions, at this point, it is difficult to clear define the role of each of the ISR sensors in response to bacterial pathogens and most likely a combination of them are responsible for an appropriate response. Future work will help us understand how these pathways are activated and manipulated by bacterial pathogens and how can we use this knowledge to develop new treatments to prevent or cure infection. For example, if we are able to safely increase the signals generated by cells of the innate immunity by manipulating the ISR, we might be able to improve the adaptive immunity generated by vaccines.

AUTHOR CONTRIBUTIONS

LR, RG, and LC wrote the manuscript. LC edited and revised the final version of the manuscript.

FUNDING

Work in the lab of LAMC is supported by Fundação Carlos Chagas de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq). LR and RG were supported by studentships from PIBIC/UFRJ and CNPq, respectively.

REFERENCES

1. Medzhitov R. Recognition of microorganisms and activation of the immune response. Nature (2007) 449:819–26. doi:10.1038/nature06246

2. Carneiro LA, Magalhaes JG, Tattoli I, Philpott DJ, Travassos LH. Nod-like proteins in inflammation and disease. J Pathol (2008) 214:136–48. doi:10.1002/path.2271

3. Iwasaki A, Medzhitov R. Control of adaptive immunity by the innate immune system. Nat Immunol (2015) 16:343–53. doi:10.1038/ni.3123

4. Kumar S, Ingle H, Prasad DV, Kumar H. Recognition of bacterial infection by innate immune sensors. Crit Rev Microbiol (2013) 39:229–46. doi:10.3109/1040841X.2012.706249

5. Vance RE, Isberg RR, Portnoy DA. Patterns of pathogenesis: discrimination of pathogenic and nonpathogenic microbes by the innate immune system. Cell Host Microbe (2009) 6:10–21. doi:10.1016/j.chom.2009.06.007

6. Pakos-Zebruca K, Koryga I, Mnich K, Ljujic M, Samali A, Gorman AM. The integrated stress response. EMBO Rep (2016) 17:1374–95. doi:10.15252/embr.201642195

7. Argüello RJ, Rodrigues CR, Gatti E, Pierre P. Protein synthesis regulation, a pillar of strength for innate immunity? Curr Opin Immunol (2015) 32:28–25. doi:10.1016/j.coi.2014.12.001

8. Tattoli I, Sorbara MT, Vuckovic D, Ling A, Soares F, Carneiro LA, et al. Amino acid starvation induced by invasive bacterial pathogens triggers an innate host defense program. Cell Host Microbe (2012) 11:563–75. doi:10.1016/j.chom.2012.04.012

9. Keestra-Gounder AM, Byndloss MX, Seyffert N, Young BM, Chávez-Arroyo A, Tsai AY, et al. NOD1 and NOD2 signaling links ER stress with inflammation. Nature (2016) 532:394–7. doi:10.1038/nature17631

10. Moretti J, Roy S, Bozec D, Martinez J, Chapman JR, Ueberheide B, et al. STING senses microbial viability to orchestrate stress-mediated autophagy of the endoplasmic reticulum. Cell (2017) 171:809–823.e13. doi:10.1016/j.cell.2017.09.034

11. Andersson U, Yang H, Harris H. High-mobility group box 1 protein (HMGB1) operates as an alarmin outside as well as inside cells. Semin Immunol (2018) 5323:30076–83. doi:10.1016/j.smim.2018.02.011

12. Fleshner M, Crane CR. Exosomes, DAMPs and miRNA: features of stress physiology and immune homeostasis. Trends Immunol (2017) 38:768–76. doi:10.1016/j.it.2017.08.002

13. Adinolfi E, Giuliani AL, De Marchi E, Pegoraro A, Orioli E, Di Virgilio F. The P2X7 receptor: a main player in inflammation. Biochem Pharmacol (2018) 151:234–44. doi:10.1016/j.bcp.2017.12.021

14. Tsutsuki H, Yahiro K, Ogura K, Ichimura K, Iyoda S, Ohnishi M, et al. Subtilase cytotoxin produced by locus of enterocyte effacement-negative Shiga-toxigenic Escherichia coli induces stress granule formation. Cell Microbiol (2016) 18:1024–40. doi:10.1111/cmi12565

15. von Hoven G, Kloft N, Neukirch C, Ebinger S, Bobkiewicz W, Weis S, et al. Modulation of translation and induction of autophagy by bacterial exoproducts. Med Microbiol Immunol (2012) 201:401–18. doi:10.1007/s00430-012-0271-0

16. Thomas MG, Loschi M, Desbats MA, Boccaccio GL. RNA granules: the good, the bad and the ugly. Cell Signal (2011) 23:324–34. doi:10.1016/j.cellsig.2010.08.011

17. Jheng JR, Ho JY, Horng JT. ER stress, autophagy, and RNA viruses. Front Microbiol (2014) 5:388–95. doi:10.3389/fmicb.2014.00388

18. Montero H, Trujillo-Alonso V. Stress granules in the viral replication cycle. Viruses (2011) 3:2328–3238. doi:10.3390/v3112328

19. Carrasco L, Sanz MA, González-Almela E. The regulation of translation in alphavirus-infected cells. Viruses (2018) 10:70. doi:10.3390/v10020070

20. McCormick C, Khaperskyy DA. Translation inhibition and stress granules in the antiviral immune response. Nat Rev Immunol (2017) 17:647–60. doi:10.1038/nri.2017.63

21. Hoang HD, Graber TE, Alain T. Battling for ribosomes: translational control at the forefront of the antiviral response. J Mol Biol (2018) 2836:30362–70. doi:10.1016/j.jmb.2018.04.040

22. Donnelly N, Gorman AM, Gupta S, Samali A. The eIF2α kinases: their structures and functions. Cell Mol Life Sci (2013) 70:3493–511. doi:10.1007/s00018-012-1252-6

23. Clemens MJ, Elia A. The double-stranded RNA-dependent protein kinase PKR: structure and function. J Interferon Cytokine Res (1997) 17:503–24. doi:10.1089/jir.1997.17.503

24. Lemaire PA, Anderson E, Lary J, Cole JL. Mechanism of PKR activation by dsRNA. J Mol Biol (2008) 381:351–60. doi:10.1016/j.jmb.2008.05.056

25. Shimazawa M, Hara H. Inhibitor of double stranded RNA-dependent protein kinase protects against cell damage induced by ER stress. Neurosci Lett (2006) 409:192–5. doi:10.1016/j.neulet.2006.09.074

26. Garcia MA, Gil J, Ventoso I, Guerra S, Domingo E, Rivas C, et al. Impact of protein kinase PKR in cell biology: from antiviral to antiproliferative action. Microbiol Mol Biol Rev (2006) 70:1032–60. doi:10.1128/MMBR.00027-06

27. Zhou HR, He K, Landgraf J, Pan X, Pestka JJ. Direct activation of ribosome-associated double-stranded RNA-dependent protein kinase (PKR) by deoxynivalenol, anisomycin and ricin: a new model for ribotoxic stress response induction. Toxins (2014) 6:3406–25. doi:10.3390/toxins6123406

28. Saelens X, Kalai M, Vandenabeele P. Translation inhibition in apoptosis: caspase-dependent PKR activation and eIF2-alpha phosphorylation. J Biol Chem (2001) 276:41620–8. doi:10.1074/jbc.M103674200

29. Korennykh A, Walter P. Structural basis of the unfolded protein response. Annu Rev Cell Dev Biol (2012) 28:251–77. doi:10.1146/annurev-cellbio-101011-155826

30. Wang M, Kaufman RJ. Protein misfolding in the endoplasmic reticulum as a conduit to human disease. Nature (2016) 529:326–35. doi:10.1038/nature17041

31. Grabiner MA, Fu Z, Wu T, Barry KC, Scharzer C, Machen TE. Pseudomonas aeruginosa quorum-sensing molecule homoserine lactone modulates inflammatory signaling through PERK and eI-F2. J Immunol (2014) 193:1459–67. doi:10.4049/jimmunol.1303437

32. de la Cadena SG, Hernandez-Fonseca K, Camacho-Arroyo I, Massieu L. Glucose deprivation induces reticulum stress by the PERK pathway and caspase-7- and calpain-mediated caspase-12 activation. Apoptosis (2014) 19:414–27. doi:10.1007/s10495-013-0930-7

33. Moore CE, Omikorede O, Gomez E, Willars GB, Herbert TP. PERK activation at low glucose concentration is mediated by SERCA pump inhibition and confers preemptive cytoprotection to pancreatic beta-cells. Mol Endocrinol (2011) 25:315–26. doi:10.1210/me.2010-0309

34. Ill-Raga G, Tajes M, Busquets-Garcia A, Ramos-Fernandez E, Vargas LM, Bosch-Morato M, et al. Physiological control of nitric oxide in neuronal BACE1 translation by heme-regulated eIF2alpha kinase HRI induces synaptogenesis. Antioxid Redox Signal (2015) 22:1295–307. doi:10.1089/ars.2014.6080

35. McEwen E, Kedersha N, Song B, Scheuner D, Gilks N, Han A, et al. Heme-regulated inhibitor kinase-mediated phosphorylation of eukaryotic translation initiation factor 2 inhibits translation, induces stress granule formation, and mediates survival upon arsenite exposure. J Biol Chem (2005) 280:16925–33. doi:10.1074/jbc.M412882200

36. Yerlikaya A, Kimball SR, Stanley BA. Phosphorylation of eIF2alpha in response to 26S proteasome inhibition is mediated by the haem-regulated inhibitor (HRI) kinase. Biochem J (2008) 412:579–88. doi:10.1042/BJ20080324

37. Lu L, Han AP, Chen JJ. Translation initiation control by heme-regulated eukaryotic initiation factor 2alpha kinase in erythroid cells under cytoplasmic stresses. Mol Cell Biol (2001) 21:7971–80. doi:10.1128/MCB.21.23.7971-7980.2001

38. Castilho BA, Shanmugam R, Silva RC, Ramesh R, Himme BM, Sattlegger E. Keeping the eIF2 alpha kinase Gcn2 in check. Biochim Biophys Acta (2014) 1843:1948–68. doi:10.1016/j.bbamcr.2014.04.006

39. Vazquez de Aldana CR, Wek RC, Segundo PS, Truesdell AG, Hinnebusch AG. Multicopy tRNA genes functionally suppress mutations in yeast eIF-2 alpha kinase GCN2: evidence for separate pathways coupling GCN4 expression to unchanged tRNA. Mol Cell Biol (1994) 14:7920–7793. doi:10.1128/MCB.14.12.7920

40. Kepp O, Semeraro M, Bravo-San Pedro JM, Bloy N, Buqué A, Huang X, et al. eIF2α phosphorylation as a biomarker of immunogenic cell death. Semin Cancer Biol (2015) 33:86–92. doi:10.1016/j.semcancer.2015.02.004

41. Kashiwagi K, Ito T, Yokoyama S. Crystal structure of eIF2B and insights into eIF2-eIF2B interactions. FEBS J (2017) 284:868–74. doi:10.1111/febs.13896

42. Wortel IMN, van der Meer LT, Kilberg MS, van Leeuwen FN. Surviving stress: modulation of ATF4-mediated stress responses in normal and malignant cells. Trends Endocrinol Metab (2017) 28:794–806. doi:10.1016/j.tem.2017.07.003

43. Wei N, Zhu LQ, Liu D. ATF4: a novel potential therapeutic target for Alzheimer’s disease. Mol Neurobiol (2015) 52:1765–70. doi:10.1007/s12035-014-8970-8

44. Wang C, Guo F. Effects of activating transcription factor 4 deficiency on carbohydrate and lipid metabolism in mammals. IUBMB Life (2012) 64:226–30. doi:10.1002/iub.605

45. B’Chir W, Maurin AC, Carraro V, Averous J, Jousse C, Muranishi Y, et al. The eIF2alpha/ATF4 pathway is essential for stress-induced autophagy gene expression. Nucleic Acids Res (2013) 41:7683–99. doi:10.1093/nar/gkt563

46. Harding HP, Zhang Y, Zeng H, Novoa I, Lu PD, Calfon M, et al. An integrated stress response regulates amino acid metabolism and resistance to oxidative stress. Mol Cell (2003) 11:619–33. doi:10.1016/S1097-2765(03)00105-9

47. Karpinski BA, Morle GD, Huggenvik J, Uhler MD, Leiden JM. Molecular cloning of human CREB-2: an ATF/CREB transcription factor that can negatively regulate transcription from the cAMP response element. Proc Natl Acad Sci U S A (1992) 89:4820–4. doi:10.1073/pnas.89.11.4820

48. Ohoka N, Yoshii S, Hattori T, Onozaki K, Hayashi H. TRB3, a novel ER stress-inducible gene, is induced via ATF4-CHOP pathway and is involved in cell death. EMBO J (2005) 24:1243–55. doi:10.1038/sj.emboj.7600596

49. Wang Q, Mora-Jensen H, Weniger MA, Perez-Galan P, Wolford C, Hai T, et al. ERAD inhibitors integrate ER stress with an epigenetic mechanism to activate BH3-only protein NOXA in cancer cells. Proc Natl Acad Sci U S A (2009) 106:2200–5. doi:10.1073/pnas.0807611106

50. Hiwatashi Y, Kanno K, Takasaki C, Goryo K, Sato T, Torii S, et al. PHD1 interacts with ATF4 and negatively regulates its transcriptional activity without prolyl hydroxylation. Exp Cell Res (2011) 317:2789–99. doi:10.1016/j.yexcr.2011.09.005

51. Koditz J, Nesper J, Wottawa M, Stiehl DP, Camenisch G, Franke C, et al. Oxygen-dependent ATF-4 stability is mediated by the PHD3 oxygen sensor. Blood (2007) 110:3610–7. doi:10.1182/blood-2007-06-094441

52. Jousse C, Deval C, Maurin AC, Parry L, Cherasse Y, Chaveroux C, et al. TRB3 inhibits the transcriptional activation of stress-regulated genes by a negative feedback on the ATF4 pathway. J Biol Chem (2007) 282:15851–61. doi:10.1074/jbc.M611723200

53. Nava GM, Stappenbeck TS. Diversity of the autochthonous colonic microbiota. Gut Microbes (2011) 2:99–104. doi:10.4161/gmic.2.2.15416

54. Finlay BB, Falkow S. Common themes in microbial pathogenicity. Microbiol Rev (1989) 53:210–30.

55. Orji FA, Ugbogu OC, Ugbogu EA, Barbabosa-Pliego A, Monroy JC, Elghandour MMMY, et al. Pathogenic flora composition and overview of the trends used for bacterial pathogenicity identifications. Microb Pathog (2018) 5:139–46. doi:10.1016/j.micpath.2018.05.006

56. Carneiro LA, Travassos LH, Soares F, Tattoli I, Magalhaes JG, Bozza MT, et al. Shigella induces mitochondrial dysfunction and cell death in nonmyleoid cells. Cell Host Microbe (2009) 5:123–36. doi:10.1016/j.chom.2008.12.011

57. Zimmermann S, Wagner C, Müller W, Brenner-Weiss G, Hug F, Prior B, et al. Induction of neutrophil chemotaxis by the quorum-sensing molecule N-(3-oxododecanoyl)-L-homoserine lactone. Infect Immun (2006) 74:5687–92. doi:10.1128/IAI.01940-05

58. Hintz M, Reichenberg A, Altincicek B, Bahr U, Gschwind R, Kollas AK, et al. Identification of (E)-4-hydroxy-3-methyl-but-2-enyl pyrophosphate as a major activator for human γδ T cells in Escherichia coli. FEBS Letters (2001) 509:317–22. doi:10.1016/S0014-5793(01)03191-X

59. Ganz T, Nemeth E. Iron homeostasis in host defence and inflammation. Nat Rev Immunol (2015) 15:500–10. doi:10.1038/nri3863

60. Vasil ML, Ochsner UA. The response of Pseudomonas aeruginosa to iron: genetics, biochemistry and virulence. Mol Microbiol (1999) 34:399–341. doi:10.1046/j.1365-2958.1999.01586.x

61. van ‘t Wout EF, van Schadewijk A, van Boxtel R, Dalton LE, Clarke HJ, Tommassen J, et al. Virulence factors of Pseudomonas aeruginosa induce both the unfolded protein and integrated stress responses in airway epithelial cells. PLoS Pathog (2015) 11(6):e1004946. doi:10.1371/journal.ppat.1004946

62. Blander JM, Sander LE. Beyond pattern recognition: five immune checkpoints for scaling the microbial threat. Nat Rev Immunol (2012) 12:215–25. doi:10.1038/nri3167

63. Valderrama C, Clark A, Urano F, Unanue ER, Carrero JA. Listeria monocytogenes induces an interferon-enhanced activation of the integrated stress response that is detrimental for resolution of infection in mice. Eur J Immunol (2017) 47:830–40. doi:10.1002/eji.201646856

64. Carrero J, Calderon B, Unanue E. Type I interferon sensitizes lymphocytes to apoptosis and reduces resistance to Listeria infection. J Exp Med (2004) 200:535–40. doi:10.1084/jem.20040769

65. O’Connell RM, Saha SK, Vaidya SA, Bruhn KW, Miranda GA, Zarnegar B, et al. Type I interferon production enhances susceptibility to Listeria monocytogenes infection. J Exp Med (2004) 200:437–45. doi:10.1084/jem.20040712

66. Auerbuch V, Brockstedt DG, Meyer-Morse N, O’Riordan M, Portnoy DA. Mice lacking the type I interferon receptor are resistant to Listeria monocytogenes. J Exp Med (2004) 200:527–33. doi:10.1084/jem.20040976

67. Rasamiravaka T, El Jaziri M. Quorum-sensing mechanisms and bacterial response to antibiotics in P. aeruginosa. Curr Microbiol (2016) 73:747–53. doi:10.1007/s00284-016-1101-1

68. Tattoli I, Sorbara MT, Yang C, Tooze SA, Philpott DJ, Girardin SE. Listeria phospholipases subvert host autophagic defenses by stalling pre-autophagosomal structures. EMBO J (2013) 32:3066–78. doi:10.1038/emboj.2013.234

69. Chakrabarti S, Liehl P, Buchon N, Lemaitre B. Infection-induced host translational blockage inhibits immune responses and epithelial renewal in the Drosophila gut. Cell Host Microbe (2012) 12:60–70. doi:10.1016/j.chom.2012.06.001

70. Dal Peraro M, van der Goot FG. Pore-forming toxins: ancient, but never really out of fashion. Nat Rev Microbiol (2016) 14:77–92. doi:10.1038/nrmicro.2015.3

71. Kloft N, Neukirch C, Bobkiewicz W, Veerachato G, Busch T, von Hoven G, et al. Pro-autophagic signal induction by bacterial pore-forming toxins. Med Microbiol Immunol (2010) 199:299–309. doi:10.1007/s00430-010-0163-0

72. Kloft N, Busch T, Neukirch C, Weis S, Boukhallouk F, Bobkiewicz W, et al. Pore-forming toxins activate MAPK p38 by causing loss of cellular potassium. Biochem Biophys Res Commun (2009) 385:503–6. doi:10.1016/j.bbrc.2009.05.121

73. Gonzalez MR, Bischofberger M, Freche B, Ho S, Parton RG, van der Goot FG. Pore-forming toxins induce multiple cellular responses promoting survival. Cell Microbiol (2011) 13:1026–43. doi:10.1111/j.1462-5822.2011.01600.x

74. Kloft N, Neukirch C, Von Hoven G, Bobkiewicz W, Weis S, Boller K, et al. A subunit of eukaryotic translation initiation factor 2alpha-phosphatase (CreP/PPP1R15B) regulates membrane traffic. J Biol Chem (2012) 287:35299–317. doi:10.1074/jbc.M112.379883

75. von Hoven G, Neukirch C, Meyenburg M, Füser S, Petricna MB, Rivas AJ, et al. eIF2α confers cellular tolerance to S. aureus α0-toxin. Front Immunol (2015) 6:383.103389/fimmu.2015.00383

76. Travassos LH, Carneiro LA, Ramjeet M, Hussey S, Kim YG, Magalhães JG, et al. Nod1 and Nod2 direct autophagy by recruiting ATG16L1 to the plasma membrane at the site of bacterial entry. Nat Immunol (2010) 11:55–62. doi:10.1038/ni.1823

77. Bretin A, Carriere J, Dalmasso G, Bergougnoux A, B’chir W, Maurin AC, et al. Activation of the EIF2AK4-EIF2A/eIF2a-ATF4 pathway triggers autophagy response to Crohn disease-associated adherent-invasive Escherichia coli infection. Autophagy (2016) 12:770–83. doi:10.1080/15548627.2016.1156823

78. do Vale A, Cabanes D, Sousa S. Bacterial toxins as pathogen weapons against phagocytes. Front Microbiol (2016) 7:42–9. doi:10.3389/fmicb.2016.00042

79. Deng W, Marshall NC, Rowland JL, McCoy JM, Worrall LJ, Santos AS, et al. Assembly, structure, function and regulation of type III secretion systems. Nat Rev Microbiol (2017) 15:323–37. doi:10.1038/nrmicro.2017.20

80. Grohmann E, Christie PJ, Waksman G, Backert S. Type IV secretion in Gram-negative and Gram-positive bacteria. Mol Microbiol (2018) 107:455–71. doi:10.1111/mmi.13896

81. Sana TG, Berni B, Bleves S. The T6SSs of Pseudomonas aeruginosa strain PAO1 and their effectors: beyond bacterial-cell targeting. Front Cell Infect Microbiol (2016) 6:61–6. doi:10.3389/fcimb.2016.00061

82. Joshi A, Kostiuk B, Rogers A, Teschler J, Pukatzki S, Yildiz FH. Rules of engagement: the type VI secretion system in Vibrio cholerae. Trends Microbiol (2017) 25:267–79. doi:10.1016/j.tim.2016.12.003

83. Yang ZS, Ma LQ, Zhu K, Yan JY, Bian L, Zhang KQ, et al. Pseudomonas toxin pyocyanin triggers autophagy: implications for pathoadaptive mutations. Autophagy (2016) 12:1015–28. doi:10.1080/15548627.2016.1170256

84. Rada B, Leto TL. Pyocyanin effects on respiratory epithelium: relevance in Pseudomonas aeruginosa airway infections. Trends Microbiol (2013) 21:73–81. doi:10.1016/j.tim.2012.10.004

85. Gouin E, Welch MD, Cossart P. Actin-based motility of intracellular pathogens. Curr Opin Microbiol (2005) 8:35–45. doi:10.1016/j.mib.2004.12.013

86. Gaytán MO, Martínez-Santos VI, Soto E, González-Pedrajo B. Type three secretion system in attaching and effacing pathogens. Front Cell Infect Microbiol (2016) 6:129–35. doi:10.3389/fcimb.2016.00129

87. Schmidt MA. LEEways: tales of EPEC, ATEC and EHEC. Cell Microbiol (2010) 12:1544–52. doi:10.1111/j.1462-5822.2010.01518.x

88. Galan JE, Wolf-Watz H. Protein delivery into eukaryotic cells by type III secretion machines. Nature (2006) 444:567–73. doi:10.1038/nature05272

89. Viboud GI, Bliska JB. Yersinia outer proteins: role in modulation of host cell signaling responses and pathogenesis. Annu Rev Microbiol (2005) 59:69–89. doi:10.1146/annurev.micro.59.030804.121320

90. Heasman SJ, Ridley AJ. Mammalian Rho GTPases: new insights into their functions from in vivo studies. Nat Rev Mol Cell Biol (2008) 9:690–701. doi:10.1038/nrm2476

91. Kim S, Coulombe PA. Emerging role for the cytoskeleton as an organizer and regulator of translation. Nat Rev Mol Cell Biol (2010) 11:75–81. doi:10.1038/nrm2818

92. Sattlegger E, Chernova TA, Gogoi NM, Pillai IV, Chernoff YO, Munn AL. Yeast studies reveal moonlighting functions of the ancient actin cytoskeleton. IUBMB Life (2014) 66:538–45. doi:10.1002/iub.1294

93. Sotelo-Silveira J, Crispino M, Puppo A, Sotelo JR, Koenig E. Myelinated axons contain beta-actin mRNA and ZBP-1 in periaxoplasmic ribosomal plaques and depend on cyclic AMP and F-actin integrity for in vitro translation. J Neurochem (2008) 104:545–57. doi:10.1111/j.1471-4159.2007.04999.x

94. Silva RC, Sattlegger E, Castilho BA. Perturbations in actin dynamics reconfigure protein complexes that modulate GCN2 activity and promote an eIF2 response. J Cell Sci (2016) 129:4521–33. doi:10.1242/jcs.194738

95. Garriz A, Qiu H, Dey M, Seo E-J, Dever TE, Hinnebusch AG. A network of hydrophobic residues impeding helix alphaC rotation maintains latency of kinase Gcn2, which phosphorylates the alpha subunit of translation initiation factor 2. Mol Cell Biol (2009) 29:1592–607. doi:10.1128/MCB.01446-08

96. Lageix S, Zhang J, Rothenburg S, Hinnebusch AG. Interaction between the tRNA-binding and C-terminal domains of Yeast Gcn2 regulates kinase activity in vivo. PLoS Genet (2015) 11:e1004991. doi:10.1371/journal.pgen.1004991

97. Marton MJ, Crouch D, Hinnebusch AG. GCN1, a translational activator of GCN4 in Saccharomyces cerevisiae, is required for phosphorylation of eukaryotic translation initiation factor 2 by protein kinase GCN2. Mol Cell Biol (1993) 13:3541–56. doi:10.1128/MCB.13.6.3541

98. Sattlegger E, Hinnebusch AG. Separate domains in GCN1 for binding protein kinase GCN2 and ribosomes are required for GCN2 activation in amino acid-starved cells. EMBO J (2000) 19:6622–33. doi:10.1093/emboj/19.23.6622

99. Gross SR, Kinzy TG. Improper organization of the actin cytoskeleton affects protein synthesis at initiation. Mol Cell Biol (2007) 27:1974–89. doi:10.1128/MCB.00832-06

100. Perez WB, Kinzy TG. Translation elongation factor 1A mutants with altered actin bundling activity show reduced aminoacyl-tRNA binding and alter initiation via eIF2alpha phosphorylation. J Biol Chem (2014) 289:20928–38. doi:10.1074/jbc.M114.570077

101. Han AP, Yu C, Lu L, Fujiwara Y, Browne C, Chin G, et al. Heme-regulated eIF2alpha kinase (HRI) is required for translational regulation and survival of erythroid precursors in iron deficiency. EMBO J (2001) 20:6909–18. doi:10.1093/emboj/20.23.6909

102. Ranu RS. Regulation of protein synthesis in rabbit reticulocyte lysates: the hemeregulated protein kinase (HRI) and double stranded RNA induced protein kinase (dRI) phosphorylate the same site(s) on initiation factor eIF-2. Biochem Biophys Res Commun (1979) 91:1437–44. doi:10.1016/0006-291X(79)91227-0

103. Caruso R, Warner N, Inohara N, Nunez G. NOD1 and NOD2: signaling, host defense, and inflammatory disease. Immunity (2014) 41:898–908. doi:10.1016/j.immuni.2014.12.010

104. Gaudet RG, Sintsova A, Buckwalter CM, Leung N, Cochrane A, Li J, et al. Cytosolic detection of the bacterial metabolite HBP activates TIFA-dependent innate immunity. Science (2015) 348:1251–5. doi:10.1126/science.aaa4921

105. Hou F, Sun L, Zheng H, Skaug B, Jiang QX, Chen ZJ. MAVS forms functional prion-like aggregates to activate and propagate antiviral innate immune response. Cell (2011) 146:448–61. doi:10.1016/j.cell.2011.06.041

106. Maekawa S, Ohto U, Shibata T, Miyake K, Shimizu T. Crystal structure of NOD2 and its implications in human disease. Nat Commun (2016) 7:11813–8. doi:10.1038/ncomms11813

107. Creagh EM, O’Neill LA. TLRs, NLRs and RLRs: a trinity of pathogen sensors that co-operate in innate immunity. Trends Immunol (2006) 27:352–7. doi:10.1016/j.it.2006.06.003

108. Brubaker SW, Bonham KS, Zanoni I, Kagan JC. Innate immune pattern recognition: a cell biological perspective. Annu Rev Immunol (2015) 33:257–90. doi:10.1146/annurev-immunol-032414-112240

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Rodrigues, Graça and Carneiro. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

REVIEW
published: 16 October 2018
doi: 10.3389/fimmu.2018.02379

Pattern Recognition Receptors and the Host Cell Death Molecular Machinery

Gustavo P. Amarante-Mendes1,2[*], Sandy Adjemian3,4, Laura Migliari Branco5,6, Larissa C. Zanetti[7], Ricardo Weinlich[7][*] and Karina R. Bortoluci5,6[*]

1Departamento de Imunologia, Instituto de Ciências Biomédicas, Universidade de São Paulo, São Paulo, Brazil

2Instituto de Investigação em Imunologia, Instituto Nacional de Ciência e Tecnologia (INCT), São Paulo, Brazil

3Molecular Signaling and Cell Death Unit, Inflammation Research Center, VIB, Ghent, Belgium

4Department of Biomedical Molecular Biology, Ghent University, Ghent, Belgium

5Departamento de Ciências Biológicas, Universidade Federal de São Paulo, Diadema, Brazil

6Centro de Terapia Celular e Molecular (CTC-Mol), Universidade Federal de São Paulo, São Paulo, Brazil

7Instituto Israelita de Ensino e Pesquisa, Hospital Israelita Albert Einstein, São Paulo, Brazil

Edited by:
Leticia A. Carneiro, Universidade Federal do Rio de Janeiro, Brazil

Reviewed by:
Fabianno Ferreira Dutra, Universidade Federal do Estado do Rio de Janeiro, Brazil
Marco Di Gioia, Boston Children's Hospital, Harvard University, United States

*Correspondence: Gustavo P. Amarante-Mendes, gpam@usp.br
Ricardo Weinlich, ricardo.weinlich@einstein.br
Karina R. Bortoluci, kbortoluci@gmail.com

Specialty section: This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 02 August 2018
Accepted: 25 September 2018
Published: 16 October 2018

Citation: Amarante-Mendes GP, Adjemian S, Branco LM, Zanetti LC, Weinlich R and Bortoluci KR (2018) Pattern Recognition Receptors and the Host Cell Death Molecular Machinery. Front. Immunol. 9:2379. doi: 10.3389/fimmu.2018.02379

Pattern Recognition Receptors (PRRs) are proteins capable of recognizing molecules frequently found in pathogens (the so-called Pathogen-Associated Molecular Patterns—PAMPs), or molecules released by damaged cells (the Damage-Associated Molecular Patterns—DAMPs). They emerged phylogenetically prior to the appearance of the adaptive immunity and, therefore, are considered part of the innate immune system. Signals derived from the engagement of PRRs on the immune cells activate microbicidal and pro-inflammatory responses required to eliminate or, at least, to contain infectious agents. Molecularly controlled forms of cell death are also part of a very ancestral mechanism involved in key aspects of the physiology of multicellular organism, including the elimination of unwanted, damaged or infected cells. Interestingly, each form of cell death has its particular effect on inflammation and on the development of innate and adaptive immune responses. In this review article, we discuss some aspects of the molecular interplay between the cell death machinery and signals initiated by the activation of PRRs by PAMPs and DAMPs.

Keywords: PRR, pathogen recognition receptor, apoptosis, necroptosis, pyroptosis, inflammation

INTRODUCTION

In 1989, Charles Janeway Jr. proposed the existence of a collection of receptors expressed by innate immune cells responsible for detecting conserved products of microbial origin (1). After 25 years of intense research, fierce debates, and a Nobel Prize granted on this subject, it is unquestionable that Janeway's ingenious idea has revolutionized our understanding of the immune system. Indeed, his seminal article is considered as one of the pillars of immunology (2).

The so-called Pattern Recognition Receptors (PRRs) are proteins capable of recognizing molecules frequently associated with pathogens (aka Pathogen-Associated Molecular Patterns—PAMPs). A more comprehensive description of PRRs and their signaling transduction pathways can be found elsewhere (3). Briefly, PRRs can be found associated to subcellular compartments, such as the cellular and endosomal membranes, the cytosol, as well as extracellularly, in secreted forms present in the bloodstream and interstitial fluids (3). There are four major sub-families of PRRs—the Toll-like receptors (TLRs), the nucleotide-binding oligomerization domain (NOD)- Leucin Rich Repeats (LRR)-containing receptors (NLR), the retinoic acid-inducible gene 1 (RIG-1) -like receptors (RLR; aka RIG-1-like helicases—RLH), and the C-type lectin receptors (CLRs) (4). As predicted by Janeway, the engagement of PRRs on the innate immune cells induces co-stimulatory signals for the adaptive immune cells (particularly T lymphocytes) (5). In addition, they activate microbicidal and pro-inflammatory responses required to eliminate (or at least to contain) infectious agents, including the induction of infected cell death (6), as discussed below.

Another ingenious idea came from Polly Matzinger (7), who proposed that the immune system is less concerned with the origin of the antigens (self vs. non-self) than with the context of their encounter with our body (tissue damage vs. tissue homeostasis). In her “Danger Theory,” Matzinger suggested that during tissue stress or damage, endogenous molecules are released or activated and initiate or propagate the inflammatory response, which, among other things, empower antigen-presenting cells to activate the adaptive immune response. Today, these molecules are collectively known as DAMPs (Damage-Associated Molecular Patterns). Importantly, soon enough it became clear that similarly to PAMPs, DAMPs could also engage PRRs.

These two theories together put forward the idea that our body is equipped to distinguish “healthy,” homeostatic tissue turnover or encounters with foreign “friendly” microorganisms, from potential “danger” that may come from pathogens and/or tissue damage.

CELL DEATH PROGRAMS

Molecularly controlled forms of cell death are part of a very ancestral mechanism involved in key aspects of the physiology of multicellular organism, including the elimination of unwanted, damaged or infected cells. Importantly to our discussion, cell death can have a direct or an indirect impact upon the course of infection, as the elimination of infected cells may eradicate or at least restrain the growth of a given pathogen. Moreover, the recognition of dying cells or their by-products modulates both inflammatory and immune responses. In the following sections, we will briefly describe the mechanisms that govern the three major types of molecularly controlled forms of cell death, namely apoptosis, necroptosis and pyroptosis, that participate in host defense through elimination of infected cells, and how they are regulated by signals derived from PRRs. For information regarding other cell death modes please refer to the work published by the Nomenclature on Cell Death Committee 2018 (8)

Apoptosis

Apoptosis was the first type of programmed cell death to be described, initially based on morphological features that distinguished it from necrosis, an uncontrolled, accidental form of cell death observed upon extreme physicochemical insults (9). In this regard, apoptosis is characterized by chromatin condensation, nuclear fragmentation, cell shrinkage with formation of cellular membrane blebs, and, finally, cellular disintegration into fragments known as apoptotic bodies (10). Importantly, during apoptosis, the plasma membrane integrity is preserved, avoiding the release of intracellular contents to the extracellular milieu. This feature contributes to the concept that apoptosis is an (relatively) inflammatory-silent form of cell death. Indeed, recognition and elimination of apoptotic cells during physiological circumstances, such as tissue/organ sculpture during development and tissue homeostasis, occurs without the cardinal signs of inflammation. In addition, it is well established that recognition of apoptotic cells by macrophages, in particular, results in the production of anti-inflammatory molecules, such as TGF-β and PGE_2 (11). On the other hand, it is also known that apoptotic cells release a series of so-called “find-me” signals, such as extracellular ATP and lysophosphatidylcholine (LPC), capable of recruiting phagocytes to the site of apoptotic corpses, characterizing, therefore, at least one aspect of an inflammatory reaction (12, 13). Besides, more recently, it was shown that apoptosis initiated via the FAS/CD95 death receptor is associated with the release of chemokines and other immunologically active proteins that coordinates the migration of phagocytes and proper removal of apoptotic cells (14). Taken together, it is reasonable to say that although not completely “silent,” apoptosis is a form of cell death that does not trigger an overt inflammatory response.

From the molecular point of view, much of our knowledge about the regulation of apoptosis came from works with the nematode Caenorhabditis elegans. In a series of elegant studies, Bob Horvitz and colleagues identified four crucial genes (Ced-3, Ced-4, Ced-9, and Egl-1) responsible for the control of developmental cell death in C. elegans (15), which granted him the Nobel Prize in Physiology or Medicine in 2002, together with John Sulston and Sidney Brenner, “for their discoveries concerning genetic regulation of organ development and programmed cell death.” Soon after Horvitz discoveries, it became clear that cell death in C. elegans and apoptosis in mammals shared a very similar, phylogenetically conserved mechanism. Apoptosis is executed by certain members of a family of cysteine aspartate-specific proteases called caspases (16–18). Importantly, not all caspases induces apoptosis. Caspases-1, -4, -5, -11, -12, -13, and-14 are inflammatory caspases not related to the initiation or execution of the apoptotic program. Caspases are produced as an inactive pro-form (zymogen) that can be activated either through proteolytic processing by upstream caspases (in the case of caspases-3, -6, and-7) or via dimerization in the context of multimolecular platforms, such as the apoptosome (caspase-9), the DISC (death-inducing signaling complex) (caspases-8 and-10), the PIDDosome (caspase-2), and the inflammasome (caspase-1 and-11) (16). Executioner or effector caspases, such as caspase-3, -6, and-7 (and CED-3 in C. elegans), are responsible for the induction of the morphological as well as the biochemical features associated with apoptosis, including oligonucleosomal DNA fragmentation and externalization of phosphatidylserine (PS) residues from the inner to the outer leaflet of the plasma membrane (19). Interestingly, in mammals, although the inhibition of effector caspases prevents apoptosis, it does not preclude cell death, which proceeds with different morphological and biochemical characteristics (20). Because of this, it has been proposed that apoptosis in mammals may not be actually a cell death mechanism, but perhaps a termination step of a cell-death program aimed to properly dispose damaged or unwanted cells without initiating inflammatory responses (18).

There are two signaling pathways of apoptosis (Figure 1). The intrinsic pathway deals with signals derived from intracellular stress, such as DNA damage, oxidative stress, dysregulation of Ca2+ homeostasis, interference with the cytoskeleton structure, endoplasmic reticulum stress, etc. Its first layer of regulation comprises the differential expression/activation of BCL-2 family members, responsible for controlling the mitochondria outer membrane permeabilization (MOMP) (21). When the pro-apoptotic stress is too strong for a given cell, MOMP allows the selective release of certain mitochondrial proteins, such as SMAC (second mitochondria-derived activator of caspases)/Diablo (direct IAP binding protein with low pI), HtrA2 (high temperature requirement protein A2)/Omi, and cytochrome c to the cytosol. Cytochrome c associates with APAF-1 (apoptosis-activating factor-1), the mammalian CED-4 homolog, and pro-caspase-9, thereby assembling the apoptosome and enabling caspase-9 to activate the downstream effector caspases. SMAC/Diablo and HtrA2/Omi facilitate apoptosis by preventing the inhibitory action of the inhibitors of apoptosis proteins (IAPs) on the effector caspases. The extrinsic pathway, in comparison, is initiated by the interaction of trimeric, extracellular ligands (TNF-α, CD95L, and TRAIL) to their cognate receptors (TNFR1, CD95 and TRAILRI, or TRAILRII, respectively) present on the plasma membrane (10, 22, 23). The stimulation of these so-called death receptors (DRs) leads to the recruitment of adaptor molecules, such as TRADD (Tumor necrosis factor receptor type 1-associated death domain protein) and/or FADD (Fas-associated protein with death domain), and the pro-caspase-8, giving rise to the conventional DISC. Next, caspase-8 directly activates the effector caspases or amplifies the cell death signal by engaging BID (BH3 interacting-domain death agonist), a pro-apoptotic member of the BCL-2 (B-cell lymphoma 2) family, leading to MOMP, cytochrome c release and assembly of the apoptosome (Figure 1). It is important to mention that the activation of caspase-8 in the context of DISC can be regulated by c-FLIP (cellular FLICE-like inhibitory protein), a catalytically-dead caspase-8 homolog (24).

FIGURE 1. Apoptosis pathways. The Intrinsic Pathway of apoptosis is activated when intracellular “stresses,” such as DNA or cytoskeleton damage or absence of growth/survival factors, are “perceived” by BH3-only members of the Bcl-2 family. These molecules become activated and migrate to the mitochondria where they facilitate or actively induce the release of apoptogenic factors, such as cytochrome c and SMAC/Diablo, to the cytosol. Cytochrome c associates with APAF-1 and pro-caspase-9 to form the apoptosome, resulting in the activation of caspase-9, which activates the effector caspases-3, -6, and -7, responsible for the biochemical and morphological modifications associated to apoptosis. SMAC/Diablo participates by preventing inhibition of caspases by IAPs. The extrinsic pathway of apoptosis initiates by the engagement of Death Receptor by their cognate Death Receptor Ligands causing the formation of the Death-inducing signaling complex (DISC). DISC is formed by the intracellular portion of the Death Receptors, the adaptor proteins TRADD and/or FADD and the pro-caspase-8 (or pro-caspase-10). Activated caspase-8 may directly activate the effector caspases or process the BH3-only protein Bid. Truncated Bid migrates to mitochondria and activates the extrinsic pathway of apoptosis.

In some instances, apoptosis can also be triggered by TLR stimulation, as a defense mechanism against infection. TLR2 was the first PRR to be associated with induction of apoptosis, by virtue of its ability to recruit FADD via MyD88 (Myeloid differentiation primary response 88), and the consequent activation of caspase-8 (25). Likewise, bacterial lipoproteins were reported to trigger apoptosis through this TLR2 pathway (26, 27) and Mycobacterium tuberculosis was also shown to induce TLR-2/caspase-8-dependent apoptosis in macrophages (28). Interestingly, TLR3-induced apoptosis is mediated via TRIF (TIR-domain-containing adapter-inducing interferon-β), which interacts with RIPK1 (Receptor Interacting Serine/Threonine Kinase 1) through its RHIM (RIP homotypic interaction motif) domain (please refer to necroptosis section for further information on these protein-protein interactions). FADD is then recruited, and activates caspase-8 leading to apoptosis (25, 29). In human keratinocytes, poly I:C-induced apoptosis required the stimulation of TLR3 and its adaptor TRIF, thus inducing caspase-8 activation (30); the same molecules were shown to induce apoptosis in human breast cancer cells (31). Not surprisingly, TLR4 can induce apoptosis either via MyD88 or TRIF, and depending on the cell type or conditions engage the extrinsic or intrinsic pathways. For instance, Yersinia was shown to induce TLR4-mediated apoptosis of macrophages through TRIF (32, 33). TRIF-mediated apoptosis seems to be executed through the extrinsic pathway, with no evidence of the involvement of the mitochondrial pathway (34). Interestingly, UV irradiation was shown to induce apoptosis in murine macrophages through TLR4 and MyD88 (35). Despite these observations and a number of other examples that we have not presented here, it is important to emphasize that PRR-induced apoptosis is a relatively minor event compared to all other triggers of apoptosis and that PRR activation leads preferentially to other forms of regulated cell death, as we will discuss below.

Necroptosis

Evidence of a molecularly controlled necrotic cell death was first provided by studies showing that Tumor Necrosis Factor Receptor 1 (TNFR1) and CD95 ligation were capable of inducing necrosis, particularly when caspase activity was inhibited (36, 37). This idea was further supported by a study that demonstrated that the cowpox virus could induce necrosis in porcine kidney cells when it harbored the caspase inhibitor CrmA (cytokine response modifier A) (38). This cell death mode was named “Necroptosis,” as it reflects the existence of a molecular pathway (like apoptosis) but with a necrotic phenotype.

The first molecule to be identified in the necroptotic pathway was RIPK1 as its kinase activity inhibitor, necrostatin-1 (Nec-1), was shown to suppress cell death triggered by caspase inhibition during TNFR1/Fas stimulation (39). RIPK1 has been previously involved in apoptotic and survival pathways, functioning as a scaffold protein to the assembly of the respective signaling platforms (40). Contrastingly, the RIPK1 kinase activity is indispensible for death receptor-triggered necroptosis, as its auto-phosphorylation induces a conformational change that allows RIPK1 to recruit, via their respective RHIM domains, the next member of this pathway, namely RIPK3 (41–43). Once recruited, RIPK3 gets activated by auto-phosphorylation and forms an amyloid-like structure, which promotes the recruitment and activation of Mixed Lineage Kinase Domain-Like (MLKL) (42, 44–47). RIPK3-phosphorylated MLKL oligomerizes and translocates to the plasma membrane, where it interacts with phosphatidylinositides and induces plasma membrane disruption [(48–51); Figure 2]. Distinct effector mechanisms were raised to account for the MLKL-driven permeabilization of the plasma membrane, either directly by pore or cation channel formation, or indirectly, by activation of TRPM or other ion channels (48–52). It is still unclear, however, which of these mechanisms are physiologically relevant. Nonetheless, in all cases, MLKL induces a loss of osmolality control, which causes cell swelling and membrane rupture. Recently, ESCRT-III machinery was suggested to counter these effects by shedding out the MLKL-damaged plasma membrane regions (53).

FIGURE 2. Necroptosis signaling. Death Receptor (DR)-induced necroptosis requires RIPK1 kinase activity to recruit RIPK3 that, in turn, recruits and activates MLKL via phosphorylation of its pseudokinase domain. Once phosphorylated, MLKL oligomerizes and migrates to the plasma membrane, where it interacts with phosphatidylinositol phosphates and induces membrane destabilization and rupture. Necroptosis signaling mediated by TRIF, IFNR, and DAI can directly activate RIPK3 and, in this case, RIPK1 acts as a negative regulator, mostly by recruiting to the signaling platform the suppressive complex containing Caspase-8, FADD and c-FLIP.

Necroptosis can be initiated by a variety of signals. The first to be described and most thoroughly studied was TNFR1 ligation (36). Upon its ligation, TNFR1 typically assembles a multimolecular complex (Complex I) composed by TRADD, RIPK1, TRAF2, TRAF5, cIAP1, cIAP2, and LUBAC (linear ubiquitin chain assembly complex), which is involved in NF-κB activation, pro-inflammatory cytokines synthesis and cell survival (54). Sustained TNFRI ligation leads to CYLD-mediated deubiquitination of this complex, which disassembles, allowing the formation of a secondary complex (Complex II) in the cytosol, constituted by TRADD, FADD, RIPK1, caspase-8, and occasionally c-FLIP (54). As pointed out above, when c-FLIP levels are low, caspase-8 forms active homodimers and triggers downstream events that culminate in apoptosis. However, in the absence of FADD, c-FLIP or a functional caspase-8, TNFRI signaling results in the recruitment of TRADD and RIPK1, forming a platform called complex IIb or necrosome, wherein RIPK3 and MLKL are activated to execute necroptosis (55). Although slightly differing on how RIPK1 is brought to the complex, this molecule has also a central role in Fas and TRAILR-induced necroptosis, as RIPK1 is, in all these cases, mandatory to recruit RIPK3 via their RHIM homotypic domain interactions (54).

Necroptosis can also be triggered by PRRs, such as TLR3 and TLR4, intracellular sensing proteins, such as DAI, RIG-I and MDA-5 as well as interferon signaling [(56–59); Figure 2]. Intriguingly, however, RIPK1 is dispensable for or even inhibitory of the necrosome formation during TLR3-, TLR4-, DAI-, and interferon-mediated necroptosis (57, 60). In these cases, RIPK3 is directly recruited to the signaling platforms, and the presence of RIPK1 slows down or halts the RIPK3-mediated activation of MLKL (60, 61). The ability of RIPK1 to recruit FADD, and consequently, caspase-8 and FLIP accounts, at least in part, for its inhibitory property. Therefore, from the molecular point of view, necroptosis ought to be defined as a RIPK3-dependent form of cell death.

Many other stimuli have been described as capable to induce necroptosis, ranging from UV irradiation, chemotherapeutic drugs (such as cisplatin, etoposide, and staurosporine), natural compounds (such as shikonin and its analogs), to DNA damage, hypoxia, ischemia/reperfusion and oxidative stress (62). The signaling pathways that lead to necroptosis in each of these cases are still to be fully elucidated. Further studies are required to evaluate whether they are dependent on RIPK1 and also whether they directly signal to a RIPK3-activating platform or indirectly, via up regulation of a classic necroptotic inducer, such as TNF or FasL. For example, UV irradiation was reported to induce necroptosis via TNF upregulation but also via spontaneous aggregation of RIPK1 and RIPK3, independently of any death receptor ligation (29, 63). Particularly puzzling is the fact that shikonin, a naphthoquinone compound obtained from a plant extract, can induce necroptosis even in the absence of FADD/caspase-8/FLIP inhibition, which is thought to be mandatory for this type of cell death (64). Thus, either this compound can itself somehow block their activity, or it shall be instrumental to decipher alternative ways in which MLKL is activated and necroptosis is executed.

Nonetheless, despite the different mechanisms that initiate necroptosis, in all cases cells undergo rapid MLKL-mediated plasma membrane permeabilization with consequent release of intracellular contents, including many DAMPs, such as lysosomal proteases, DNA, mtDNA, ATP, and HMGB1 [(55); Table 1). Therefore, similarly to pyroptosis (see below), necroptosis is considered a pro-inflammatory form of cell death. Even so, it is still to be determined whether the pro-inflammatory properties of necroptotic cells are the result of the intracellular content leakage or, rather, they can actively produce and/or modify specific DAMPs. Evidence for the latter comes from ESCRT-III-deficient cells that undergo necroptosis much faster, which limits the amount of inflammatory cytokines and chemokines produced and hinders antigen cross-presentation (53). Moreover, both RIPK3 and MLKL have been associated with inflammasome and NF-κB activation, supporting the notion that the pro-inflammatory potential of necroptotic cells goes beyond the passive release of their intracellular content (110–112).

Table 1. DAMPs released by cell death and its role in the immune system.

Necroptotic cells not only induce a potent inflammatory response but they are also highly immunogenic, which may be instrumental against infection and during anti-tumoral responses. For example, mice injected with necroptotic cells present a higher CD8+ T cell cross-priming and increased tumor immunity when compared with animals injected with apoptotic cells (113, 114). Likewise, RIPK3 deficiency in mice inhibits immune cell infiltration and attenuates organ injury during sepsis (115). Therefore, given that necroptosis is highly immunogenic, disruption in the necroptotic pathway would be expected in some pathophysiological conditions. Indeed, it was reported that most of the in vitro transformed cells as well as human tumor samples have low or no expression of RIPK3 (116), and a cohort of chronic lymphocytic leukemia patients present down regulation of CYLD (117). Furthermore, patients with lower expression of RIPK3 or MLKL have worse prognosis for breast cancer or ovarian cancer, respectively (116, 118), suggesting that resistance to necroptosis is positively selected during tumor growth and/or development. This may be associated with an increased ability to evade immune attack, either by prolonging the lifespan of the transformed cells, by decreasing the availability of DAMPs, or by avoiding the activation of antigen-presenting cells during the immune responses. Therefore, induction of necroptosis in tumors may change its immunogenicity and promote a better immune response against it. This is particularly exciting, as we are currently witnessing novel and promising approaches in tumor treatment that are based on stimulation of the immune system. On the other hand, it is possible that the inflammation generated by necroptosis may promote tumor development by stimulating angiogenesis and metastasis (119). Therefore, thorough investigation of the benefits and pitfalls of inducing inflammatory cell death for each cancer type will be required in order to determine whether inducing necroptosis is indeed a good option in the specific cancer treatment.

Besides its impact on tumorigenesis and tumor progression, deficient necroptotic signaling can be detrimental during viral infection. Mice lacking RIPK3 are highly sensitive to vaccinia virus due to widespread infection (120). Likewise, RIPK3-deficient mice are more susceptible to Influenza A virus (IAV) than the wild-type animals (121). Remarkably, seasonal IAV, but not the 1918 and 2009 pandemic IAV strains, induces RIPK3-mediated immunogenic death of dendritic cells (122). The pandemic strains' ability to suppress necroptosis was mapped to the hemagglutinin (HA) genomic segment (122), indicating that either the pandemic strains' HA do not induce necroptosis or it may directly interfere with the necroptotic signaling pathway.

Keeping with the notion that suppressing necroptosis is advantageous to the infectious agent, there is accumulating evidence that viruses can encode molecules that are able to directly interfere with the necroptotic signaling. vIRA, a molecule expressed by MCMV that contains a RHIM-like domain blocks RIPK3 recruitment to RIPK1 and to DAI (57). MCMV expressing vIRA mutated in its RHIM domain produces an attenuated viremia in wild-type mice, which is reverted in RIPK3-deficient animals (57). Likewise, HSV-1 and HSV-2 express ICP-6 and ICP-10, respectively, which are able to suppress necroptosis in human cells through a similar RHIM-dependent mechanism (123, 124). Curiously, in mice, ICP-6 was shown to promote necroptosis through direct aggregation with RIPK3, restricting virus propagation (124, 125). A different mode of action was reported for the IE1-regulated gene product expressed by HCMV, which suppresses necroptosis downstream of RIPK3 activation and MLKL recruitment (126).

Bacteria can also induce necroptosis, at least in vitro. It is less clear, though, whether necroptosis plays a central role in bacterial infections in vivo. Loss of RIPK3 in combination with deletion or inhibition of caspase-8 or FADD renders mice susceptible to a number of pathogens, including Yersinia and Citrobacter (127, 128). However, the relative contribution of necroptosis and caspase-8-mediated apoptosis in these models were not yet tested, as caspase-8- or FADD–deficient animals are not viable (129–131).

Necroptosis, though, may not always be protective against infection. Macrophage death by necroptosis correlates with increased susceptibility to Salmonella infection (132). Also, HIV-specific CD8+ T cell response, which is a key indicator of infection control, is impaired due to increased necroptosis levels in this cell population (133). Taken together, necroptosis seems to be detrimental when it eliminates the population that is central for the immune control of the infection. In the other cases, necroptosis limits infection, mostly likely by destroying the pathogen's replicative niche through a cell death mode that generates a pro-inflammatory and immunogenic environment. However, it is important to note that, as mentioned above, RIPK3 and MLKL were shown to participate of additional signaling platforms, including inflammasome activation, NF-κB signaling and even apoptosis induction (134). Therefore, in the light of these novel RIPK3 and MLKL roles, it is essential to reevaluate the relative contribution of necroptosis to the phenotypes observed. A good illustration comes from the fact that while RIPK3-deficient mice are more susceptible to IAV, MLKL-deficient animals are not, indicating that necroptosis is not the sole RIPK3-mediated mechanism important in IAV control (121). In fact, it was shown that IAV also triggers RIPK3-mediated apoptosis, via recruitment of RIPK1, FADD and caspase-8. This was further supported by the fact that MLKL-caspase-8 double deficient mice present similar levels of susceptibility to IAV infection observed with the RIPK3-deficient animals (121). Another example is that RIPK3-deficient mice are less susceptible to Staphylococcus aureus lung damage and present reduced bacterial loads and inflammation, while MLKL-deficient animals present an opposite outcome, suggesting that these molecules have independent, non-necroptotic roles (135).

Pyroptosis

Pyroptosis is a necrotic form of regulated cell death distinct from necroptosis, mainly due to the requirement of inflammatory caspase-1 and/or caspase-11 (murine caspase-11 corresponds to caspases-4 and -5 in humans) [(136); Figure 3). It is the result of pore formation in the plasma membrane that increases osmotic pressure ensuing in osmotic lysis and, consequently, the release of the intracellular content, including pro-inflammatory cytokines and DAMPs (137). Although distinct from the typical oligonucleosomal fragmentation observed during apoptosis, DNA fragmentation is also a hallmark of pyroptosis, which seems to occur independently of the caspase-activated DNase (CAD) (138).

FIGURE 3. Molecular basis of pyroptosis. (A) Canonical inflammasome assembly upon sensing of PAMPs, DAMPs or other cytosolic disturbs leads to the recruitment and activation of caspase-1 directly or via the recruitment of the adaptor protein ASC. Caspase-1 induces the maturation of pro-IL-1β and pro-IL-18 into their active forms as well as cleavage of Gasdermin D (GSDMD). The GSDMD pore form domain (PFD) interacts with the plasma membrane to form the GSDMD pore, leading to the release of the intracellular content, including IL-1β and IL-18. (B) Non-canonical inflammasome activation is initiated by the detection of cytosolic LPS from gram-negative bacteria by the pro-caspase-11 itself. Activated caspase-11 (caspase-4 or caspase-5 in humans), in turn, induces GSDMD cleavage and consequent pyroptosis.

Pyroptosis is a form of cell death initiated in response to the engagement of certain members of the PRRs, which are capable of assembling complex structures called inflammasomes. These platforms are composed by a sensor protein, either from the NLR or the pyrin and HIN domain-containing protein (PYHIN) families of cytosolic PRRs, in addition to the adaptor molecule apoptosis-associated speck-like protein containing a caspase activating and recruitment domain (ASC) and pro-caspase-1. There are five major types of the so-called canonical inflammasomes—NLR pyrin domain-containing 3 (NLRP3), NLRP1, neuronal apoptosis inhibitory protein (NAIP)/NLR CARD-containing 4 (NLRC4), absent in melanoma 2 (AIM2), and PYRIN inflammasomes (139). Activation of one of these cytosolic sensors in response to PAMPs, DAMPs or cytosolic disturbances such as ionic imbalance leads to the recruitment and activation of caspase-1 either directly or through the ASC adaptor molecule [(140); Figure 3]. Besides the induction of pyroptosis, caspase-1 also leads to the processing and release of the pro-inflammatory cytokines IL-1β and IL-18 (139).

In contrast to the canonical inflammasomes, which require multicomplex structures, the non-canonical inflammasome seems to be composed solely by pro-caspase-11, which plays the role of the sensor as well as the executor (141, 142). During intracellular gram-negative bacteria infections, Lipid A, a component of LPS, can directly bind to the CARD domain of pro-caspase-11 (143), which gets activated and induces pyroptosis. Interestingly, the non-canonical caspase-11 inflammasome acts independently of LPS recognition by TLR4 and does not directly induce IL1-β and IL-18 maturation [(141, 142); Figure 3]. In monocytes, however, non-canonical inflammasome stimulation may result in minor production of IL-1β and IL-18 through the bystander induction of NLRP3 activation (144). Interestingly, LPS-induced lethal shock is driven by the activation of the non-canonical inflammasome. Since IL-1β and IL-18 release are not a major outcome of caspase-11 activation, the exacerbated inflammatory response observed in sepsis seems to be mainly driven by pyroptosis, probably due to the efflux of DAMPs, such as High Mobility Group Box 1 (HMGB1) and IL-1α [Table 1; (145)].

Such non-canonical inflammasome-mediated responses have drawn the attention of different research groups that became interested in unraveling the relevant as well as the pathogenic caspase-11 downstream targets. In 2015, two concurrent studies reported that Gasdermin D (GSDMD), a member of the GSDM family, was the effector component of the non-canonical inflammasome pathway (146, 147), which was later confirmed by a third study (148). Kayagaki and colleagues performed N-ethyl-N-nitrosourea mutagenesis screening for mutations that compromised LPS-induced IL-1β release and pyroptosis (146) while Shi and colleagues employed the clustered regularly interspaced palindromic repeat (CRISPR)/Cas9 genome-editing screens in TLR4 deficient mouse bone marrow-derived macrophages for guide RNAs that protected from LPS-induced cell death (147). Both studies hit GSDMD as a substrate for caspase-11 and the effector of pyroptosis. GSDMD is composed by a C-terminal and a N-terminal domain linked by a long loop. Caspase-11 cleaves an aspartate residue within the linking loop, releasing the N-terminal fragment from the inhibitory C-terminus (146, 147). The N-terminal domain, also called Pore-Forming Domain (PFD) (149) oligomerizes and associates with lipids in the inner plasma membrane to form 10–33 nm pores leading to cell swelling and eventually to cell lysis (150–153). Importantly, it was also demonstrated that caspase-1 cleaves GSDMD at the same site as caspase-11, establishing that GSDMD is also required for the canonical inflammasome-driven pyroptosis (147).

Until the discovery of GSDMD as the pyroptosis executioner, the physiological function of GSDM proteins was largely unknown. However, recent studies described that the PFD is highly conserved among several members the GSMD family. Indeed, expression of PFD from GSDMA, GSDMA3, GSDMB, GSDMC, GSDME, or GSDMA3 in HEK293 was able to induce pore formation and a cell death phenotype similar to pyroptosis (147, 151). Moreover, GSDMA3 cleavage by caspase-3 in HEK293 and macrophages results in a secondary necrotic cell death after apoptosis (154). This necrotic cell death might contribute to hearing loss in GSDMA3 spontaneous mutations that are associated with deafness (155). Thus, given the cytotoxic activity of different GSDM PFD, some authors have proposed a redefinition of pyroptosis as a GSDM-mediated cell death (146). However, it is controversial how other GSDM members are activated and whether these proteins participate in cell death pathways. Also, GSDMD seems not to be required for pyroptosis during prolonged inflammasome activation in response to the classical agonists, ATP, and flagellin (146). Moreover, in the absence of caspase-1 protease activity, caspase-8 accounts for GSDMD-independent cell death in response to inflammasome agonists (156–158). Since some of these processes share features of pyroptosis, it is hard to define pyroptosis solely as being a process of cell death regulated by inflammatory caspases or mediated by GSDM proteins, since we can find exceptions to the rules that govern both concepts.

From the biological point-of-view, cell death by pyroptosis results in a fast removal of infected cell leading to the elimination of the replication niche. Conversely to the previous idea of liberation of bacteria to the extracellular milieu by pyroptotic cells (159), the current knowledge predicts that, instead, the damaged bacteria remain trapped within the pyroptotic corpses. This structure is called pore-induced trap (PIT) and it prevents bacterial dissemination (160, 161). Despite that PIT does not directly kill intracellular bacteria, pyroptosis renders them more susceptible to H_20_2, to the antimicrobial peptide polymyxin B and to the antibiotic ciprofloxacin (157). As a consequence, the recovered bacteria from PIT are less capable to infect neighbor cells.

The inflammatory milieu created by the release of the intracellular content from pyroptotic cells recruits circulating phagocytes to the infectious site. Subsequently, neutrophils efferocyte the PIT and kill the pathogen by a mechanism dependent on reactive oxygen species (ROS) (161). Extracellular bacteria can also be controlled by the action of antimicrobial peptides (160, 161) and potentially by the GSDMD N-terminal domain released during cell lysis due to its affinity to cardiolipin and phosphatidylserine expressed in some bacterial cell membranes, such as Escherichia coli and Listeria monocytogenes (152, 162). Interestingly, canonical and non-canonical inflammasomes are required for intestinal epithelial cells (IECs) responses to infections (163, 164). The activation of NLRC4 inflammasomes in IECs results in a lytic cell death prior to a non-conventional process of cell expulsion that contributes to control bacterial replication. Although caspase-1 and Gasdermin-D were required for IEC pyroptosis, both molecules were dispensable for cell expulsion, demonstrating that coordinated inflammasome responses in IECs are important to prevent bacterial translocation to deeper tissues (163, 164).

Interestingly, neutrophils seems to be more resistant to pyroptosis than macrophages in response to Salmonella and are able to maintain a sustained IL-1β production and secretion, which could be important to control the infection (165). However, Kambara et al. (166) recently described that a specific neutrophil elastase (ELANE) is able to cleave GSDMD independently of caspases activity, promoting a lytic cell death in these cells. Interestingly, these authors demonstrated that GSDMD-dependent neutrophil death impairs the control of extracellular bacteria E. coli, thus suggesting that GSDMD could exert an anti-inflammatory role depending on the infection context.

In addition to its role in the elimination of replicative niche, the pyroptosis machinery is involved in IL-1β and IL-18 release. As these cytokines lack the signal peptide, their release is considered to occur by non-conventional pathways (167). Among the different pathways that have been proposed to explain their secretion, mechanisms involving cell death are particularly subject to intense debate in the literature. Growing evidences suggest that IL-1β can be released by viable monocytes (168), dendritic cells (DCs) (169), and macrophages (170). GSDMD pore is large enough to allow IL-1β release concomitant with the influx of cationic ions (148). Notably, in viable cells, GSDMD seems to be required for IL-1β translocation to the extracellular space in response to stimuli that hyperactivate phagocytes, such as oxidized phospholipids (oxPAPC) in DCs or LPS in human monocytes (170–172). Nonetheless, it is difficult to establish whether the cells were actually viable, since cell death can precede cell lysis, thus suggesting that pyroptosis and cell lysis can be uncoupled events (173). Moreover, the assessment of cell death by the detection of lactate dehydrogenase release (LDH), used in several studies as the only viability assay, might be insufficient to discriminate viable cells from dying cells since both viable and unviable cells can release LDH to the cell culture (170, 173, 174).

Although many studies have demonstrated the requirement of canonical and non-canonical inflammasomes to host defense against pathogens, the precise contribution of pyroptosis and other inflammasome-related mechanisms are poorly understood and arose mainly from in vitro assays or bacterial infection models in mice deficient for molecules that compose these platforms (159, 175). In L. monocytogenes, S. typhimurium, and B. thailandensis in vivo infections, the lack of caspase-1/11 was more deleterious to the host than IL-1β/IL-18 deficiency (176–179), while in mice infected with F. novicida, the treatment with recombinant IL-1β/IL-18 only partially recovered the resistance to infection, suggesting that cytokine secretion was not sufficient to protect the host (180). The susceptibility of GSDMD-deficient mice seems to correlate with that demonstrated by caspase-1/11-deficient mice, although the deletion of GSDMD also culminated in reduced IL-1β/IL-18 release (146–148). Even though (a) IL-1β/IL-18 secretion might occur independently of pyroptosis and (b) caspase-1 or caspase-11 deletion is more severe than IL-1β/IL-18 ablation in some bacterial infections, usually cytokine release and cell death are overlapping events necessary for optimal host defense (159). In any case, the susceptibility of GSDMD and other GSDM deficient mouse strains to infectious agents and its comparison to mice lacking caspase-1, caspase-11, IL-1β, and/or IL-18 remains to be established, especially in non-bacterial infection contexts. For example, despite clear evidences of the involvement of inflammatory caspases in the host control of some fungal infections such as Candida albicans, Aspergillus fumigatus, and Paracoccidioides brasiliensis (172), the requirement of GSDMD to cell death and the consequences to the host resistance against these infections is still to be elucidated.

Notwithstanding, the highly pro inflammatory outcome of pyroptosis as well as the cell loss can be prejudicial to the host during the response to pathogens. In HIV patients, the quiescent CD4 T cells depletion seems to be mainly mediated by pyroptosis (181, 182). During HIV abortive infection, the engagement of the interferon-gamma-inducible-protein 16 (IFI16) in response to cytosolic viral DNA leads to inflammasome assembly and caspase-1 mediated CD4 T cells pyroptosis in lymphoid tissues (181, 182). Interestingly, co-cultivation of lymphoid-derived cells sensitizes blood-derived CD4 T cells to HIV-induced pyroptosis (183). Moreover, pyroptotic peripheral blood CD14+CD16− monocytes from HIV-infected patients release ASC specks, a hallmark of inflammasome activation. Therefore, besides the depletion of CD4 T cells, pyroptosis of CD4 T cells and monocytes contributes to the chronic inflammation that characterizes the disease (184).

The identification of the non-canonical inflammasome and the discovery of GSDMD as the executioner of pyroptosis have expanded our understanding of the mechanisms driving this type of cell death. However, further studies are necessary to elucidate the precise role of inflammatory and non-inflammatory caspases and the participation of members from GSDM family and/or other effector proteases in the molecular regulation of pyroptosis. In addition, the understanding of its role during infection or inflammatory processes in vivo will contribute to better understand the biological relevance of this regulated cell death induced in response to the PRRs activation.

PRR SENSING OF CELL DEATH AND CELL DEATH PRODUCTS

The notion that cells undergoing cell death release or expose several intracellular molecules regardless of the accidental nature or the different regulated death programs (apoptosis, necrosis or pyroptosis) is widely recognized. Although mainly non-inflammatory in the intracellular space, molecules released/exposed from damaged cells can participate in the activation of inflammation and immune responses. Indeed, a broad range of receptors, including PRR, sense these DAMPs and alert the immune system by inducing immune cell migration, increasing phagocytosis by macrophages and DCs, stimulating the production of pro-inflammatory cytokines or even contributing to the maturation of DCs, among other key functions (72, 80, 185). A number of studies have been dedicated to the characterization of putative DAMPs, and it became apparent that the type of cell death, as well as the nature of cell death stimuli, influence the quality and quantity of DAMPs release (Table 1).

Importantly, the stress or damage before the cellular demise itself is determinant to set in motion a sequence of events leading to an immunogenic cell death (ICD). The sensing of this stress regulates the cell death process thus initiating signaling pathways that will actively—or not—generate danger signals (186). Other DAMPs will be passively released as a result of membrane rupture during necroptosis or pyroptosis. These DAMPs define in part the immunogenicity of cell death, but are not sufficient to elicit a specific anti-tumor immune response, for instance. Indeed, they are released or exposed by the dying cells and act as adjuvant providing that antigens are exhibited conjointly (187). In contrast, a non-immunogenic cell death does not provide the required levels of DAMPs and antigens to evoke an adaptive immune response (187).

Together, these concepts redefined the widely accepted paradigm stating that apoptosis is always a silent cell death modality as opposed to necrosis, which is inflammatory and immunogenic. Therefore, a non-immunogenic apoptosis is characterized by the absence of plasma membrane leakage and the rapid phagocytosis of apoptotic bodies prevents the release of DAMPs and the consequent inflammatory reaction. Indeed the apoptotic process reduce cell immunogenicity by diverse ways including, (1) preservation of intracellular structures and plasma membrane, thereby blocking the release of DAMPS; (2) reduction of cellular volume, by condensation of the nucleus and shedding of small vesicles, which favors its rapid elimination by the surrounding tissue; (3) expression of “find-me” and “eat-me” signals, which increases the speed of cell clearance (12, 188); (4) inhibition of the production of interferons and pro-inflammatory cytokines, as the DNA is being chopped and condensed (189); (5) induction of an antigenic tolerance in the engulfing APCs (190); and (6) post-translational modifications on DAMPS and alarmins that decrease their pro-inflammatory potential (191, 192).

Interestingly, depending on the trigger, apoptosis can be immunogenic. Indeed, some chemotherapeutic agents, such as anthracyclines, as well as radiation and hypericin-based photodynamic therapy, were found to strongly prime immune responses through the induction of ICD (65, 185). Among these, immunogenic chemotherapies are well characterized and involve the emission of a number of danger signals. The pre-apoptotic release or exposure on the plasma membrane of ER-chaperones, such as calreticulin and Heat Shock Proteins (HSPs), constitutes an early event of ICD, which relies on the induction of an ER-stress. Calreticulin promotes the uptake of dying cells by DCs (72) and the inhibition of its exposure during anthracycline-induced apoptosis of murine tumor cell lines abolished their immunogenic potential (72). The early apoptotic secretion of ATP, which binds to P2X7 or P2Y2 purinergic receptor on DCs, stimulates the formation of the NLRP3 inflammasome, thereby inducing the release of IL-1β (66). Moreover, ATP released by dying cells undergoing ICD is responsible for the recruitment and differentiation of myeloid precursors into inflammatory DCs, mediating a specific antitumor immune response (193). Passive release of the nuclear protein HMGB1 occurs during secondary necrosis (i.e., late-stage apoptotic cells), which interacts with TLR4 on DCs, and through Myd88 signaling, enables efficient tumor antigen processing and cross-presentation (80). Additionally, anthracyclines have been shown to induce the release of RNA, thereby stimulating TLR3 as a mimic of viral infection. Activation of TLR3 is then responsible for type I IFNs production that acts in an autocrine and paracrine manner to promote the secretion of CXCL10 (194). Release of Annexin A1 has also been described after anthracyclines treatment, stimulating the Formyl Peptide Receptor 1 (FPR1), thus directing the final trajectory of DCs to dying tumor cells (195).

Besides chemotherapeutic agents, bacterial and viral infection can also trigger an immunogenic apoptosis. In this case, PAMPs, such as LPS or double-stranded RNA, expressed by the pathogen can stimulate TLR signaling and induce an immune response. Indeed, phagocytosis of infected apoptotic cells could trigger TLR activation and lead to IL-6 and TGF-β production that induce the development of infection-specific as well as self-reactive TH17 cells, linking infection to autoimmunity (196, 197). Finally, defects in mechanisms of apoptotic cell clearance are linked to autoimmunity disorders, including lupus and rheumatoid arthritis, likely due to the increased risk of loss of cell integrity with the consequent release of DAMPs and increased availability of circulating self-antigens (198, 199).

Accidental or programmed forms of necrosis are responsible for the release of an usually larger panel of DAMPs compared to apoptotic cells, mainly due to plasma membrane permeabilization. Also, RIPK1-mediated activation of the NF-κB pathway, through upregulation of pro-inflammatory cytokines and increased antigen presentation, was reported to play a role in the immunogenicity of necroptotic cell death mode (186). Recently, it was show that necroptosis is accompanied by the release of the classical and potent DAMPs—HSPs, ATP, and HMGB1 (200, 201). While HSPs were shown to stimulate TLR2 and 4, inducing DCs maturation, HMGB1 was reported to interact with TLR3, 4 and 9, as well as RAGE, to activate DCs and macrophages (202–204). Necrotic cells can potentially release intact mitochondria, a major source of ATP, which may activate the NLRP3 inflammasome resulting in IL-1β secretion and neutrophil recruitment (67, 96, 205). Moreover, mitochondrial DAMPs, such as formyl peptides and mitochondrial DNA, can potentially act on FPR1 and TLR9, respectively, inducing neutrophils recruitment and degranulation (97, 115). Additionally, Mincle, the C-type lectin receptor 4E was reported to interact with the necrotic DAMP SAP130 (spliceosome-associated protein 130), normally involved in spliceosomes assembly. The stimulation of this PRR was also reported to induce recruitment of neutrophils (106). Uric acid has been described as a product of accidental necrosis (108). Once released, this DAMP can precipitate and form monosodium urate (MSU) crystals able to induce the expression of costimulatory molecules by DCs, as well as activating the NLRP3 inflammasome to trigger the production of IL-1β and IL-18 (206, 207). RNA and double-stranded DNA (dsDNA) are also released during necrotic cell death, and while RNA stimulates TLR3, dsDNA stimulates TLR9 and two members of the RLR family, RIG-I and MDA5, responsible for the release of IFN-β and CXCL10 through IRF3 and NF-κb pathways. The AIM2 inflammasome can sense dsDNA released by necrotic cells, thereby inducing IL-1β secretion (204). Finally, it is important to mention that some proteins considered DAMPs also stimulate receptors that are not PRRs. For instance, IL-1α acts on IL-1R1 to trigger an inflammatory response mediated by the Myd88 pathway (208) and full-length IL-33 is another DAMP released during necrosis, with immunological property due to the absence of caspase processing (191).

CONCLUSIONS

The protective response of our body against pathogens and tumor cells depends on proper activation of both innate and adaptive immunity. Particularly, macrophages and DCs reside on the center of these two arms of immunity. They are powerful antigen-presenting cells that may elicit effector T cell responses (protection) or induce T cells to become regulatory (tolerance), depending on their activation status. They express PRRs, which are very ancient proteins that help us identify and react to pathogens and danger signals. Upon engagement, through the interaction with conserved molecular patterns frequently associated with pathogens (PAMPs), PRRs trigger a series of biochemical signaling cascades that activates pro-inflammatory programs on DCs that enable the differentiation of antigen-specific T cells into protective effector TH1, TH2, and TH17 cells. PRR engagement also triggers programs of cell death, particularly necroptosis and pyroptosis, the necrotic forms of cell death associated with a pro-inflammatory outcome (Table 2). These forms of cell death release larger amounts of DAMPs, which in turn, stimulate surrounding cells via PRRs, thus constituting a positive feedback loop capable of amplifying host defense mechanisms (Figure 4). Apoptosis, on the other hand, is a cell death program mostly related to non-inflammatory outcomes and likely to take major role in the maintenance of homeostasis by “silently” eliminating unwanted or damaged cells. However, apoptosis may also participate in elimination of infectious agents or tumor cells. Therefore, recognition of pathogen- and damage/danger-associated molecules by the same set of immune receptors (PRRs) is a powerful strategy that bridges intrinsic cell death programs and complex immune cell interactions to preserve homeostasis and at the same time protects the organism against infection and cellular transformation (Figure 4).

Table 2. PRR agonists and consequent cell death program.

FIGURE 4. Interplay between PRRs and cell death mechanisms. The engagement of PRRs in response to PAMPs induces the activation of different cell death machineries in order to promote tissue homeostasis and host-defense against pathogens. Importantly, cell death products known collectively as DAMPs forms a feedback loop that stimulate PRRs to induce inflammatory/immune responses.

AUTHOR CONTRIBUTIONS

LM designed the figures. LZ and SA prepared Table. GA-M, RW, and KB designed the review organization. All authors contributed to the writing, reviewed, and approved the manuscript.

ACKNOWLEDGMENTS

LM and LZ are recipients of Ph.D. fellowships from FAPESP (Fundação de Amparo à Pesquisa do Estado de São Paulo, Brazil). SA is recipient of a postdoctoral fellowship from FWO (Fonds voor Wetenschappelijk Onderzoek—Vlaanderen, Belgium). GA-M, KB, and RW are supported by grants from FAPESP and from the Brazilian Research Council (CNPq).

REFERENCES

1. Janeway CA Jr. Approaching the asymptote? Evolution and revolution in immunology. Cold Spring Harb Symp Quant Biol. (1989) 54(Pt 1):1–13. doi: 10.1101/SQB.1989.054.01.003

2. Janeway CA Jr. Pillars article: approaching the asymptote? Evolution and revolution in immunology. Cold spring harb symp quant biol. 1989. 54: 1–13. J Immunol. (2013) 191:4475–4487.

3. Medzhitov R, Janeway CA Jr. Innate immunity: impact on the adaptive immune response. Curr Opin Immunol. (1997) 9:4–9.

4. Walsh D, McCarthy J, O'Driscoll C, Melgar S. Pattern recognition receptors–molecular orchestrators of inflammation in inflammatory bowel disease. Cytokine Growth Factor Rev. (2013) 24:91–104. doi: 10.1016/j.cytogfr.2012.09.003

5. Medzhitov R, Preston-Hurlburt P, Janeway CA Jr. A human homologue of the Drosophila Toll protein signals activation of adaptive immunity. Nature (1997) 388:394–7. doi: 10.1038/41131

6. Bortoluci KR, Medzhitov R. Control of infection by pyroptosis and autophagy: role of TLR and NLR. Cell Mol Life Sci. (2010) 67:1643–51. doi: 10.1007/s00018-010-0335-5

7. Matzinger P. Tolerance, danger, and the extended family. Annu Rev Immunol. (1994) 12:991–1045. doi: 10.1146/annurev.iy.12.040194.005015

8. Galluzzi L, Vitale I, Aaronson SA, Abrams JM, Adam D, Agostinis P, et al. Molecular mechanisms of cell death: recommendations of the Nomenclature Committee on Cell Death 2018. Cell Death Differ. (2018) 25:486–541. doi: 10.1038/s41418-017-0012-4

9. Kerr JF, Wyllie AH, Currie AR. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer (1972) 26:239–57. doi: 10.1038/bjc.1972.33

10. Amarante-Mendes GP, Green DR. The regulation of apoptotic cell death. Braz J Med Biol Res. (1999) 32:1053–61.

11. Fadok VA, Bratton DL, Konowal A, Freed PW, Westcott JY, Henson PM. Macrophages that have ingested apoptotic cells in vitro inhibit proinflammatory cytokine production through autocrine/paracrine mechanisms involving TGF-beta, PGE2, and PAF. J Clin Invest. (1998) 101:890–8. doi: 10.1172/JCI1112

12. Medina CB, Ravichandran KS. Do not let death do us part: ‘find-me' signals in communication between dying cells and the phagocytes. Cell Death Differ. (2016) 23:979–89. doi: 10.1038/cdd.2016.13

13. Hochreiter-Hufford A, Ravichandran KS. Clearing the dead: apoptotic cell sensing, recognition, engulfment, and digestion. Cold Spring Harb Perspect Biol. (2013) 5:a008748. doi: 10.1101/cshperspect.a008748

14. Cullen SP, Henry CM, Kearney CJ, Logue SE, Feoktistova M, Tynan GA, et al. Fas/CD95-induced chemokines can serve as “find-me” signals for apoptotic cells. Mol Cell (2013) 49:1034–48. doi: 10.1016/j.molcel.2013.01.025

15. Metzstein MM, Stanfield GM, Horvitz HR. Genetics of programmed cell death in C. elegans: past, present and future. Trends Genet. (1998) 14:410–6. doi: 10.1016/S0168-9525(98)01573-X

16. Pop C, Salvesen GS. Human caspases: activation, specificity, and regulation. J Biol Chem. (2009) 284:21777–81. doi: 10.1074/jbc.R800084200

17. McIlwain DR, Berger T, Mak TW. Caspase functions in cell death and disease. Cold Spring Harb Perspect Biol. (2013) 5:a008656. doi: 10.1101/cshperspect.a008656

18. Pereira WO, Amarante-Mendes GP. Apoptosis: a programme of cell death or cell disposal? Scand J Immunol. (2011) 73:401–7. doi: 10.1111/j.1365-3083.2011.02513.x

19. Martin SJ, Finucane DM, Amarante-Mendes GP, O'Brien GA, Green DR. Phosphatidylserine externalization during CD95-induced apoptosis of cells and cytoplasts requires ICE/CED-3 protease activity. J Biol Chem. (1996) 271:28753–6. doi: 10.1074/jbc.271.46.28753

20. Amarante-Mendes GP, Finucane DM, Martin SJ, Cotter TG, Salvesen GS, Green DR. Anti-apoptotic oncogenes prevent caspase-dependent and independent commitment for cell death. Cell Death Differ. (1998) 5:298–306. doi: 10.1038/sj.cdd.4400354

21. Tait SW, Green DR. Mitochondrial regulation of cell death. Cold Spring Harb Perspect Biol. (2013) 5:a008706. doi: 10.1101/cshperspect.a008706

22. Wertz IE, Dixit VM. Regulation of death receptor signaling by the ubiquitin system. Cell Death Differ. (2010) 17:14–24. doi: 10.1038/cdd.2009.168

23. Guicciardi ME, Gores GJ. Life and death by death receptors. FASEB (2009) 23:1625–37. doi: 10.1096/fj.08-111005

24. Yu JW, Shi Y. FLIP and the death effector domain family. Oncogene (2008) 27:6216–27. doi: 10.1038/onc.2008.299

25. Salaun B, Romero P, Lebecque S. Toll-like receptors' two-edged sword: when immunity meets apoptosis. Eur J Immunol. (2007) 37:3311–8. doi: 10.1002/eji.200737744

26. Aliprantis AO, Yang RB, Mark MR, Suggett S, Devaux B, Radolf JD, et al. Cell activation and apoptosis by bacterial lipoproteins through toll-like receptor-2. Science (1999) 285:736–9. doi: 10.1126/science.285.5428.736

27. Aliprantis AO, Yang RB, Weiss DS, Godowski P, Zychlinsky A. The apoptotic signaling pathway activated by Toll-like receptor-2. EMBO J. (2000) 19:3325–36. doi: 10.1093/emboj/19.13.3325

28. Lopez M, Sly LM, Luu Y, Young D, Cooper H, Reiner NE. The 19-kDa Mycobacterium tuberculosis protein induces macrophage apoptosis through Toll-like receptor-2. J Immunol. (2003) 170:2409–16. doi: 10.4049/jimmunol.170.5.2409

29. Feoktistova M, Geserick P, Kellert B, Dimitrova DP, Langlais C, Hupe M, et al. cIAPs block Ripoptosome formation, a RIP1/caspase-8 containing intracellular cell death complex differentially regulated by cFLIP isoforms. Mol Cell (2011) 43:449–63. doi: 10.1016/j.molcel.2011.06.011

30. Weber A, Kirejczyk Z, Besch R, Potthoff S, Leverkus M, Hacker G. Proapoptotic signalling through Toll-like receptor-3 involves TRIF-dependent activation of caspase-8 and is under the control of inhibitor of apoptosis proteins in melanoma cells. Cell Death Differ. (2010) 17:942–51. doi: 10.1038/cdd.2009.190

31. Salaun B, Coste I, Rissoan MC, Lebecque SJ, Renno T. TLR3 can directly trigger apoptosis in human cancer cells. J Immunol. (2006) 176:4894–901. doi: 10.4049/jimmunol.176.8.4894

32. Ruckdeschel K, Pfaffinger G, Haase R, Sing A, Weighardt H, Hacker G, et al. Signaling of apoptosis through TLRs critically involves toll/IL-1 receptor domain-containing adapter inducing IFN-beta, but not MyD88, in bacteria-infected murine macrophages. J Immunol. (2004) 173:3320–8. doi: 10.4049/jimmunol.173.5.3320

33. Haase R, Kirschning CJ, Sing A, Schrottner P, Fukase K, Kusumoto S, et al. A dominant role of Toll-like receptor 4 in the signaling of apoptosis in bacteria-faced macrophages. J Immunol. (2003) 171:4294–303. doi: 10.4049/jimmunol.171.8.4294

34. Kaiser WJ, Offermann MK. Apoptosis induced by the toll-like receptor adaptor TRIF is dependent on its receptor interacting protein homotypic interaction motif. J Immunol. (2005) 174:4942–52. doi: 10.4049/jimmunol.174.8.4942

35. Harberts E, Fishelevich R, Liu J, Atamas SP, Gaspari AA. MyD88 mediates the decision to die by apoptosis or necroptosis after UV irradiation. Innate Immun. (2014) 20:529–39. doi: 10.1177/1753425913501706

36. Laster SM, Wood JG, Gooding LR. Tumor necrosis factor can induce both apoptic and necrotic forms of cell lysis. J Immunol. (1988) 141:2629–34.

37. Vercammen D, Brouckaert G, Denecker G, Van de Craen M, Declercq W, Fiers W, et al. (1998). Dual signaling of the Fas receptor: initiation of both apoptotic and necrotic cell death pathways. J Exp Med. (1998) 188:919–30. doi: 10.1084/jem.188.5.919

38. Ray CA, Pickup DJ. The mode of death of pig kidney cells infected with cowpox virus is governed by the expression of the crmA gene. Virology (1996) 217:384–91. doi: 10.1006/viro.1996.0128

39. Degterev A, Huang Z, Boyce M, Li Y, Jagtap P, Mizushima N, et al. Chemical inhibitor of nonapoptotic cell death with therapeutic potential for ischemic brain injury. Nat Chem Biol. (2005) 1:112–9. doi: 10.1038/nchembio711

40. Hsu H, Huang J, Shu HB, Baichwal V, Goeddel DV. TNF-dependent recruitment of the protein kinase RIP to the TNF receptor-1 signaling complex. Immunity (1996) 4:387–96. doi: 10.1016/S1074-7613(00)80252-6

41. Zhang DW, Shao J, Lin J, Zhang N, Lu BJ, Lin SC, et al. RIP3, an energy metabolism regulator that switches TNF-induced cell death from apoptosis to necrosis. Science (2009) 325:332–6. doi: 10.1126/science.1172308

42. Li J, McQuade T, Siemer AB, Napetschnig J, Moriwaki K, Hsiao YS, et al. The RIP1/RIP3 necrosome forms a functional amyloid signaling complex required for programmed necrosis. Cell (2012) 150:339–50. doi: 10.1016/j.cell.2012.06.019

43. Sun X, Yin J, Starovasnik MA, Fairbrother WJ, Dixit VM. Identification of a novel homotypic interaction motif required for the phosphorylation of receptor-interacting protein (RIP) by RIP3. J Biol Chem. (2002) 277:9505–11. doi: 10.1074/jbc.M109488200

44. Murphy JM, Czabotar PE, Hildebrand JM, Lucet IS, Zhang JG, Alvarez-Diaz S, et al. The pseudokinase MLKL mediates necroptosis via a molecular switch mechanism. Immunity (2013) 39:443–53. doi: 10.1016/j.immuni.2013.06.018

45. Sun L, Wang H, Wang Z, He S, Chen S, Liao D, et al. Mixed lineage kinase domain-like protein mediates necrosis signaling downstream of RIP3 kinase. Cell (2012) 148:213–27. doi: 10.1016/j.cell.2011.11.031

46. Zhao J, Jitkaew S, Cai Z, Choksi S, Li Q, Luo J, et al. Mixed lineage kinase domain-like is a key receptor interacting protein 3 downstream component of TNF-induced necrosis. Proc Natl Acad Sci USA. (2012) 109:5322–7. doi: 10.1073/pnas.1200012109

47. Wu J, Huang Z, Ren J, Zhang Z, He P, Li Y, et al. Mlkl knockout mice demonstrate the indispensable role of Mlkl in necroptosis. Cell Res. (2013) 23:994–1006. doi: 10.1038/cr.2013.91

48. Cai Z, Jitkaew S, Zhao J, Chiang HC, Choksi S, Liu J, et al. Plasma membrane translocation of trimerized MLKL protein is required for TNF-induced necroptosis. Nat Cell Biol. (2014) 16:55–65. doi: 10.1038/ncb2883

49. Chen X, Li W, Ren J, Huang D, He WT, Song Y, et al. Translocation of mixed lineage kinase domain-like protein to plasma membrane leads to necrotic cell death. Cell Res. (2014) 24:105–21. doi: 10.1038/cr.2013.171

50. Dondelinger Y, Declercq W, Montessuit S, Roelandt R, Goncalves A, Bruggeman I, et al. MLKL compromises plasma membrane integrity by binding to phosphatidylinositol phosphates. Cell Rep. (2014) 7:971–81. doi: 10.1016/j.celrep.2014.04.026

51. Wang H, Sun L, Su L, Rizo J, Liu L, Wang LF, et al. Mixed lineage kinase domain-like protein MLKL causes necrotic membrane disruption upon phosphorylation by RIP3. Mol Cell (2014) 54:133–46. doi: 10.1016/j.molcel.2014.03.003

52. Xia B, Fang S, Chen X, Hu H, Chen P, Wang H, et al. MLKL forms cation channels. Cell Res. (2016) 26:517–28. doi: 10.1038/cr.2016.26

53. Gong YN, Guy C, Olauson H, Becker JU, Yang M, Fitzgerald P, et al. ESCRT-III acts downstream of MLKL to regulate necroptotic cell death and its consequences. Cell (2017). 169:286.e16–300. doi: 10.1016/j.cell.2017.03.020

54. Weinlich R, Green DR. The two faces of receptor interacting protein kinase-1. Mol Cell (2014) 56:469–80. doi: 10.1016/j.molcel.2014.11.001

55. Grootjans S, Vanden Berghe T, Vandenabeele P. Initiation and execution mechanisms of necroptosis: an overview. Cell Death Differ. (2017) 24:1184–95. doi: 10.1038/cdd.2017.65

56. De Carvalho DD, Binato R, Pereira WO, Leroy JM, Colassanti MD, Proto-Siqueira R, et al. BCR-ABL-mediated upregulation of PRAME is responsible for knocking down TRAIL in CML patients. Oncogene (2011) 30:223–33. doi: 10.1038/onc.2010.409

57. Upton JW, Kaiser WJ, Mocarski ES. DAI/ZBP1/DLM-1 complexes with RIP3 to mediate virus-induced programmed necrosis that is targeted by murine cytomegalovirus vIRA. Cell Host Microbe (2012) 11:290–7. doi: 10.1016/j.chom.2012.01.016

58. Schock SN, Chandra NV, Sun Y, Irie T, Kitagawa Y, Gotoh B, et al. Induction of necroptotic cell death by viral activation of the RIG-I or STING pathway. Cell Death Differ. (2017) 24:615–25. doi: 10.1038/cdd.2016.153

59. Thapa RJ, Nogusa S, Chen P, Maki JL, Lerro A, Andrake M, et al. Interferon-induced RIP1/RIP3-mediated necrosis requires PKR and is licensed by FADD and caspases. Proc Natl Acad Sci USA. (2013) 110:E3109–18. doi: 10.1073/pnas.1301218110

60. Dillon CP, Weinlich R, Rodriguez DA, Cripps JG, Quarato G, Gurung P, et al. RIPK1 blocks early postnatal lethality mediated by caspase-8 and RIPK3. Cell (2014) 157:1189–202. doi: 10.1016/j.cell.2014.04.018

61. Orozco S, Yatim N, Werner MR, Tran H, Gunja SY, Tait SW, et al. RIPK1 both positively and negatively regulates RIPK3 oligomerization and necroptosis. Cell Death Differ. (2014) 21:1511–21. doi: 10.1038/cdd.2014.76

62. Vanden Berghe T, Hassannia B, Vandenabeele P. An outline of necrosome triggers. Cell Mol Life Sci. (2016) 73:2137–52. doi: 10.1007/s00018-016-2189-y

63. Tenev T, Bianchi K, Darding M, Broemer M, Langlais C, Wallberg F, et al. The Ripoptosome, a signaling platform that assembles in response to genotoxic stress and loss of IAPs. Mol Cell (2011) 43:432–48. doi: 10.1016/j.molcel.2011.06.006

64. Han W, Li L, Qiu S, Lu Q, Pan Q, Gu Y, et al. Shikonin circumvents cancer drug resistance by induction of a necroptotic death. Mol Cancer Ther. (2007) 6:1641–9. doi: 10.1158/1535-7163.MCT-06-0511

65. Garg AD, Krysko DV, Verfaillie T, Kaczmarek A, Ferreira GB, Marysael T, et al. A novel pathway combining calreticulin exposure and ATP secretion in immunogenic cancer cell death. EMBO J. (2012) 31:1062–79. doi: 10.1038/emboj.2011.497

66. Ghiringhelli F, Apetoh L, Tesniere A, Aymeric L, Ma Y, Ortiz C, et al. Activation of the NLRP3 inflammasome in dendritic cells induces IL-1beta-dependent adaptive immunity against tumors. Nat Med. (2009) 15:1170–8. doi: 10.1038/nm.2028

67. Iyer SS, Pulskens WP, Sadler JJ, Butter LM, Teske GJ, Ulland TK, et al. Necrotic cells trigger a sterile inflammatory response through the Nlrp3 inflammasome. Proc Natl Acad Sci USA. (2009) 106:20388–93. doi: 10.1073/pnas.0908698106

68. Adamczak SE, de Rivero Vaccari JP, Dale G, Brand FJ III, Nonner D, Bullock MR, et al. Pyroptotic neuronal cell death mediated by the AIM2 inflammasome. J Cereb Blood Flow Metab. (2014) 34:621–9. doi: 10.1038/jcbfm.2013.236

69. Yang H, Ma Y, Chen G, Zhou H, Yamazaki T, Klein C, et al. Contribution of RIP3 and MLKL to immunogenic cell death signaling in cancer chemotherapy. Oncoimmunology (2016) 5:e1149673. doi: 10.1080/2162402X.2016.1149673

70. Weyd H, Abeler-Dorner L, Linke B, Mahr A, Jahndel V, Pfrang S, et al. Annexin A1 on the surface of early apoptotic cells suppresses CD8+ T cell immunity. PLoS ONE (2013) 8:e62449. doi: 10.1371/journal.pone.0062449

71. Franklin BS, Bossaller L, De Nardo D, Ratter JM, Stutz A, Engels G, et al. The adaptor ASC has extracellular and ‘prionoid' activities that propagate inflammation. Nat Immunol. (2014) 15:727–37. doi: 10.1038/ni.2913

72. Obeid M, Tesniere A, Ghiringhelli F, Fimia GM, Apetoh L, Perfettini JL, et al. Calreticulin exposure dictates the immunogenicity of cancer cell death. Nat Med. (2007) 13:54–61. doi: 10.1038/nm1523

73. Panaretakis T, Kepp O, Brockmeier U, Tesniere A, Bjorklund AC, Chapman DC, et al. Mechanisms of pre-apoptotic calreticulin exposure in immunogenic cell death. EMBO J. (2009) 28:578–90. doi: 10.1038/emboj.2009.1

74. Kim H, Kim WJ, Jeon ST, Koh EM, Cha HS, Ahn KS, et al. Cyclophilin A may contribute to the inflammatory processes in rheumatoid arthritis through induction of matrix degrading enzymes and inflammatory cytokines from macrophages. Clin Immunol. (2005) 116:217–24. doi: 10.1016/j.clim.2005.05.004

75. Dear JW, Simpson KJ, Nicolai MP, Catterson JH, Street J, Huizinga T, et al. Cyclophilin A is a damage-associated molecular pattern molecule that mediates acetaminophen-induced liver injury. J Immunol. (2011) 187:3347–52. doi: 10.4049/jimmunol.1100165

76. Miles K, Clarke DJ, Lu W, Sibinska Z, Beaumont PE, Davidson DJ, et al. Dying and necrotic neutrophils are anti-inflammatory secondary to the release of alpha-defensins. J Immunol. (2009) 183:2122–32. doi: 10.4049/jimmunol.0804187

77. Calderwood SK, Gong J, Murshid A. Extracellular HSPs: the complicated roles of extracellular HSPs in Immunity. Front Immunol. (2016) 7:159. doi: 10.3389/fimmu.2016.00159

78. Basu S, Binder RJ, Suto R, Anderson KM, Srivastava PK. Necrotic but not apoptotic cell death releases heat shock proteins, which deliver a partial maturation signal to dendritic cells and activate the NF-kappa B pathway. Int Immunol. (2000) 12:1539–46. doi: 10.1093/intimm/12.11.1539

79. Pouwels SD, Zijlstra GJ, van der Toorn M, Hesse L, Gras R, Ten Hacken NH, et al. Cigarette smoke-induced necroptosis and DAMP release trigger neutrophilic airway inflammation in mice. Am J Physiol Lung Cell Mol Physiol. (2016) 310:L377–86. doi: 10.1152/ajplung.00174.2015

80. Apetoh L, Ghiringhelli F, Tesniere A, Obeid M, Ortiz C, Criollo A, et al. Toll-like receptor 4-dependent contribution of the immune system to anticancer chemotherapy and radiotherapy. Nat Med. (2007) 13:1050–9. doi: 10.1038/nm1622

81. Bell CW, Jiang W, Reich CF III, Pisetsky DS. The extracellular release of HMGB1 during apoptotic cell death. Am J Physiol Cell physiol. (2006) 291:C1318–25. doi: 10.1152/ajpcell.00616.2005

82. Hou L, Yang Z, Wang Z, Zhang X, Zhao Y, Yang H, et al. NLRP3/ASC-mediated alveolar macrophage pyroptosis enhances HMGB1 secretion in acute lung injury induced by cardiopulmonary bypass. Lab Invest. (2018) 98:1052–64. doi: 10.1038/s41374-018-0073-0

83. Martinotti S, Patrone M, Ranzato E. Emerging roles for HMGB1 protein in immunity, inflammation, and cancer. Immunotargets Ther. (2015) 4:101–9. doi: 10.2147/ITT.S58064

84. Li H, Bo H, Wang J, Shao H, Huang S. Separation of supercoiled from open circular forms of plasmid DNA, and biological activity detection. Cytotechnology (2011) 63:7–12. doi: 10.1007/s10616-010-9322-9

85. Yang D, Postnikov YV, Li Y, Tewary P, de la Rosa G, Wei F, et al. High-mobility group nucleosome-binding protein 1 acts as an alarmin and is critical for lipopolysaccharide-induced immune responses. J Exp Med. (2012) 209:157–71. doi: 10.1084/jem.20101354

86. Murakami T, Hu Z, Tamura H, Nagaoka I. Release mechanism of high mobility group nucleosome binding domain 1 from lipopolysaccharide-stimulated macrophages. Mol Med Rep. (2016) 13:3115–20. doi: 10.3892/mmr.2016.4893

87. Kaczmarek A, Vandenabeele P, Krysko DV. Necroptosis: the release of damage-associated molecular patterns and its physiological relevance. Immunity (2013) 38:209–23. doi: 10.1016/j.immuni.2013.02.003

88. Kim B, Lee Y, Kim E, Kwak A, Ryoo S, Bae SH, et al. The interleukin-1alpha precursor is biologically active and is likely a key alarmin in the IL-1 family of cytokines. Front Immunol. (2013) 4:391. doi: 10.3389/fimmu.2013.00391

89. England H, Summersgill HR, Edye ME, Rothwell NJ, Brough D. Release of interleukin-1alpha or interleukin-1beta depends on mechanism of cell death. J Biol Chem. (2014) 289:15942–50. doi: 10.1074/jbc.M114.557561

90. Gross O, Yazdi AS, Thomas CJ, Masin M, Heinz LX, Guarda G, et al. Inflammasome activators induce interleukin-1alpha secretion via distinct pathways with differential requirement for the protease function of caspase-1. Immunity (2012) 36:388–400. doi: 10.1016/j.immuni.2012.01.018

91. Schmitz J, Owyang A, Oldham E, Song Y, Murphy E, McClanahan TK, et al. IL-33, an interleukin-1-like cytokine that signals via the IL-1 receptor-related protein ST2 and induces T helper type 2-associated cytokines. Immunity (2005) 23:479–90. doi: 10.1016/j.immuni.2005.09.015

92. Bonilla WV, Frohlich A, Senn K, Kallert S, Fernandez M, Johnson S, et al. The alarmin interleukin-33 drives protective antiviral CD8+ T cell responses. Science (2012) 335:984–9. doi: 10.1126/science.1215418

93. Vanden Berghe T, Kalai M, Denecker G, Meeus A, Saelens X, Vandenabeele P. Necrosis is associated with IL-6 production but apoptosis is not. Cell Signal. (2006) 18:328–35. doi: 10.1016/j.cellsig.2005.05.003

94. Coutant F, Perrin-Cocon L, Agaugue S, Delair T, Andre P, Lotteau V. Mature dendritic cell generation promoted by lysophosphatidylcholine. J Immunol. (2002) 169:1688–95. doi: 10.4049/jimmunol.169.4.1688

95. Rickard JA, O'Donnell JA, Evans JM, Lalaoui N, Poh AR, Rogers T, et al. RIPK1 regulates RIPK3-MLKL-driven systemic inflammation and emergency hematopoiesis. Cell (2014) 157:1175–88. doi: 10.1016/j.cell.2014.04.019

96. Maeda A, Fadeel B. Mitochondria released by cells undergoing TNF-alpha-induced necroptosis act as danger signals. Cell Death Dis. (2014) 5:e1312. doi: 10.1038/cddis.2014.277

97. Zhang Q, Raoof M, Chen Y, Sumi Y, Sursal T, Junger W, et al. Circulating mitochondrial DAMPs cause inflammatory responses to injury. Nature (2010) 464:104–7. doi: 10.1038/nature08780

98. Carlos D, Costa FR, Pereira CA, Rocha FA, Yaochite JN, Oliveira GG, et al. Mitochondrial DNA activates the NLRP3 inflammasome and predisposes to Type 1 diabetes in murine model. Front Immunol. (2017) 8:164. doi: 10.3389/fimmu.2017.00164

99. Yu J, Nagasu H, Murakami T, Hoang H, Broderick L, Hoffman HM, et al. Inflammasome activation leads to Caspase-1-dependent mitochondrial damage and block of mitophagy. Proc Natl Acad Sci USA. (2014) 111:15514–9. doi: 10.1073/pnas.1414859111

100. Crouser ED, Shao G, Julian MW, Macre JE, Shadel GS, Tridandapani S, et al. Monocyte activation by necrotic cells is promoted by mitochondrial proteins and formyl peptide receptors. Crit Care Med. (2009) 37:2000–9. doi: 10.1097/CCM.0b013e3181a001ae

101. Noll F, Behnke J, Leiting S, Troidl K, Alves GT, Muller-Redetzky H, et al. Self-extracellular RNA acts in synergy with exogenous danger signals to promote inflammation. PLoS ONE (2017) 12:e0190002. doi: 10.1371/journal.pone.0190002

102. Choi JJ, Reich CF III, Pisetsky DS. Release of DNA from dead and dying lymphocyte and monocyte cell lines in vitro. Scand J Immunol. (2004) 60:159–66. doi: 10.1111/j.0300-9475.2004.01470.x

103. Matsumura K, Iwai H, Kato-Miyazawa M, Kirikae F, Zhao J, Yanagawa T, et al. Peroxiredoxin 1 contributes to host defenses against Mycobacterium tuberculosis. J Immunol. (2016) 197:3233–44. doi: 10.4049/jimmunol.1601010

104. Chen B, Miller AL, Rebelatto M, Brewah Y, Rowe DC, Clarke L, et al. S100A9 induced inflammatory responses are mediated by distinct damage associated molecular patterns (DAMP) receptors in vitro and in vivo. PLoS ONE (2015) 10:e0115828. doi: 10.1371/journal.pone.0115828

105. Lotfi R. Necrosis-Associated Factors (DAMPs) including S100A4 used to pulse Dendritic Cells (DCs) induce regulatory T cells. J Cell Sci Ther. (2013) 3:3–8. doi: 10.4172/2157-7013.1000134

106. Yamasaki S, Ishikawa E, Sakuma M, Hara H, Ogata K, Saito T. Mincle is an ITAM-coupled activating receptor that senses damaged cells. Nat Immunol. (2008) 9:1179–88. doi: 10.1038/ni.1651

107. Seifert L, Werba G, Tiwari S, Giao Ly NN, Alothman S, Alqunaibit D, et al. The necrosome promotes pancreatic oncogenesis via CXCL1 and Mincle-induced immune suppression. Nature (2016) 532:245–9. doi: 10.1038/nature17403

108. Kono H, Chen CJ, Ontiveros F, Rock KL. Uric acid promotes an acute inflammatory response to sterile cell death in mice. J Clin Invest. (2010) 120:1939–49. doi: 10.1172/JCI40124

109. Braga TT, Forni MF, Correa-Costa M, Ramos RN, Barbuto JA, Branco P, et al. Soluble uric acid activates the NLRP3 inflammasome. Sci Rep. (2017) 7:39884. doi: 10.1038/srep39884

110. Moriwaki K, Balaji S, McQuade T, Malhotra N, Kang J, Chan FK. The necroptosis adaptor RIPK3 promotes injury-induced cytokine expression and tissue repair. Immunity (2014) 41:567–78. doi: 10.1016/j.immuni.2014.09.016

111. Vince JE, Wong WW, Gentle I, Lawlor KE, Allam R, O'Reilly L, et al. Inhibitor of apoptosis proteins limit RIP3 kinase-dependent interleukin-1 activation. Immunity (2012) 36:215–27. doi: 10.1016/j.immuni.2012.01.012

112. Lawlor KE, Khan N, Mildenhall A, Gerlic M, Croker BA, D'Cruz AA, et al. RIPK3 promotes cell death and NLRP3 inflammasome activation in the absence of MLKL. Nat Commun. (2015) 6:6282. doi: 10.1038/ncomms7282

113. Yatim N, Jusforgues-Saklani H, Orozco S, Schulz O, Barreira R, da Silva C, et al. RIPK1 and NF-kappaB signaling in dying cells determines cross-priming of CD8+ T cells. Science (2015) 350:328–34. doi: 10.1126/science.aad0395

114. Aaes TL, Kaczmarek A, Delvaeye T, De Craene B, De Koker S, Heyndrickx L, et al. Vaccination with necroptotic cancer cells induces efficient anti-tumor immunity. Cell Rep. (2016) 15:274–87. doi: 10.1016/j.celrep.2016.03.037

115. Duprez L, Takahashi N, Van Hauwermeiren F, Vandendriessche B, Goossens V, Vanden Berghe T, et al. RIP kinase-dependent necrosis drives lethal systemic inflammatory response syndrome. Immunity (2011) 35:908–18. doi: 10.1016/j.immuni.2011.09.020

116. Koo GB, Morgan MJ, Lee DG, Kim WJ, Yoon JH, Koo JS, et al. Methylation-dependent loss of RIP3 expression in cancer represses programmed necrosis in response to chemotherapeutics. Cell Res. (2015) 25:707–25. doi: 10.1038/cr.2015.56

117. Liu P, Xu B, Shen W, Zhu H, Wu W, Fu Y, et al. Dysregulation of TNFalpha-induced necroptotic signaling in chronic lymphocytic leukemia: suppression of CYLD gene by LEF1. Leukemia (2012) 26:1293–300. doi: 10.1038/leu.2011.357

118. He L, Peng K, Liu Y, Xiong J, Zhu FF. Low expression of mixed lineage kinase domain-like protein is associated with poor prognosis in ovarian cancer patients. Onco Targets Ther. (2013) 6:1539–43. doi: 10.2147/OTT.S52805

119. Su Z, Yang Z, Xie L, DeWitt JP, Chen Y. Cancer therapy in the necroptosis era. Cell Death Differ. (2016) 23:748–56. doi: 10.1038/cdd.2016.8

120. Cho YS, Challa S, Moquin D, Genga R, Ray TD, Guildford M, et al. Phosphorylation-driven assembly of the RIP1-RIP3 complex regulates programmed necrosis and virus-induced inflammation. Cell (2009) 137:1112–23. doi: 10.1016/j.cell.2009.05.037

121. Nogusa S, Thapa RJ, Dillon CP, Liedmann S, Oguin TH III, Ingram JP, et al. RIPK3 activates parallel pathways of MLKL-driven necroptosis and FADD-mediated apoptosis to protect against influenza A virus. Cell Host Microbe (2016) 20:13–24. doi: 10.1016/j.chom.2016.05.011

122. Hartmann BM, Albrecht RA, Zaslavsky E, Nudelman G, Pincas H, Marjanovic N, et al. Pandemic H1N1 influenza A viruses suppress immunogenic RIPK3-driven dendritic cell death. Nat Commun. (2017) 8:1931. doi: 10.1038/s41467-017-02035-9

123. Guo H, Omoto S, Harris PA, Finger JN, Bertin J, Gough PJ, et al. Herpes simplex virus suppresses necroptosis in human cells. Cell Host Microbe (2015) 17:243–51. doi: 10.1016/j.chom.2015.01.003

124. Huang Z, Wu SQ, Liang Y, Zhou X, Chen W, Li L, et al. RIP1/RIP3 binding to HSV-1 ICP6 initiates necroptosis to restrict virus propagation in mice. Cell Host Microbe (2015) 17:229–42. doi: 10.1016/j.chom.2015.01.002

125. Wang X, Li Y, Liu S, Yu X, Li L, Shi C, et al. Direct activation of RIP3/MLKL-dependent necrosis by herpes simplex virus 1 (HSV-1) protein ICP6 triggers host antiviral defense. Proc Natl Acad Sci USA. (2014) 111:15438–43. doi: 10.1073/pnas.1412767111

126. Omoto S, Guo H, Talekar GR, Roback L, Kaiser WJ, Mocarski ES. Suppression of RIP3-dependent necroptosis by human cytomegalovirus. J Biol Chem. (2015) 290:11635–48. doi: 10.1074/jbc.M115.646042

127. Weng D, Marty-Roix R, Ganesan S, Proulx MK, Vladimer GI, Kaiser WJ, et al. Caspase-8 and RIP kinases regulate bacteria-induced innate immune responses and cell death. Proc Natl Acad Sci USA. (2014) 111:7391–6. doi: 10.1073/pnas.1403477111

128. Pearson JS, Giogha C, Muhlen S, Nachbur U, Pham CL, Zhang Y, et al. EspL is a bacterial cysteine protease effector that cleaves RHIM proteins to block necroptosis and inflammation. Nat Microbiol. (2017) 2:16258. doi: 10.1038/nmicrobiol.2016.258

129. Yeh WC, de la Pompa JL, McCurrach ME, Shu HB, Elia AJ, Shahinian A, et al. FADD: essential for embryo development and signaling from some, but not all, inducers of apoptosis. Science (1998) 279:1954–8. doi: 10.1126/science.279.5358.1954

130. Walsh CM, Wen BG, Chinnaiyan AM, O'Rourke K, Dixit VM, Hedrick SM. A role for FADD in T cell activation and development. Immunity (1998) 8:439–49. doi: 10.1016/S1074-7613(00)80549-X

131. Varfolomeev EE, Schuchmann M, Luria V, Chiannilkulchai N, Beckmann JS, Mett IL, et al. Targeted disruption of the mouse Caspase 8 gene ablates cell death induction by the TNF receptors, Fas/Apo1, and DR3 and is lethal prenatally. Immunity (1998) 9:267–76. doi: 10.1016/S1074-7613(00)80609-3

132. Hu ZQ, Zhao WH. Type 1 interferon-associated necroptosis: a novel mechanism for Salmonella enterica Typhimurium to induce macrophage death. Cell Mol Immunol. (2013) 10:10–2. doi: 10.1038/cmi.2012.54

133. Gaiha GD, McKim KJ, Woods M, Pertel T, Rohrbach J, Barteneva N, et al. Dysfunctional HIV-specific CD8+ T cell proliferation is associated with increased caspase-8 activity and mediated by necroptosis. Immunity (2014) 41:1001–12. doi: 10.1016/j.immuni.2014.12.011

134. Weinlich R, Oberst A, Beere HM, Green DR. Necroptosis in development, inflammation and disease. Nat Rev Mol Cell Biol. (2017) 18:127–36. doi: 10.1038/nrm.2016.149

135. Kitur K, Parker D, Nieto P, Ahn DS, Cohen TS, Chung S, et al. Toxin-induced necroptosis is a major mechanism of Staphylococcus aureus lung damage. PLoS Pathog. (2015) 11:e1004820. doi: 10.1371/journal.ppat.1004820

136. Man SM, Karki R, Kanneganti TD. Molecular mechanisms and functions of pyroptosis, inflammatory caspases and inflammasomes in infectious diseases. Immunol Rev. (2017) 277:61–75. doi: 10.1111/imr.12534

137. Fink SL, Cookson BT. Caspase-1-dependent pore formation during pyroptosis leads to osmotic lysis of infected host macrophages. Cell Microbiol. (2006) 8:1812–25. doi: 10.1111/j.1462-5822.2006.00751.x

138. Bergsbaken T, Fink SL, Cookson BT. Pyroptosis: host cell death and inflammation. Nat Rev Microbiol. (2009) 7:99–109. doi: 10.1038/nrmicro2070

139. Broz P, Dixit VM. Inflammasomes: mechanism of assembly, regulation and signalling. Nat Rev Immunol. (2016) 16:407–20. doi: 10.1038/nri.2016.58

140. Broz P, von Moltke J, Jones JW, Vance RE, Monack DM. Differential requirement for Caspase-1 autoproteolysis in pathogen-induced cell death and cytokine processing. Cell Host Microbe (2010) 8:471–83. doi: 10.1016/j.chom.2010.11.007

141. Hagar JA, Powell DA, Aachoui Y, Ernst RK, Miao EA. Cytoplasmic LPS activates caspase-11: implications in TLR4-independent endotoxic shock. Science (2013) 341:1250–3. doi: 10.1126/science.1240988

142. Kayagaki N, Wong MT, Stowe IB, Ramani SR, Gonzalez LC, Akashi-Takamura S, et al. Noncanonical inflammasome activation by intracellular LPS independent of TLR4. Science (2013) 341:1246–9. doi: 10.1126/science.1240248

143. Shi J, Zhao Y, Wang Y, Gao W, Ding J, Li P, et al. Inflammatory caspases are innate immune receptors for intracellular LPS. Nature (2014) 514:187–92. doi: 10.1038/nature13683

144. Yang D, He Y, Munoz-Planillo R, Liu Q, Nunez G. Caspase-11 requires the Pannexin-1 channel and the purinergic P2X7 pore to mediate pyroptosis and endotoxic shock. Immunity (2015) 43:923–32. doi: 10.1016/j.immuni.2015.10.009

145. Lamkanfi M, Dixit VM. Inflammasomes and their roles in health and disease. Annu Rev Cell Dev Biol. (2012) 28:137–61. doi: 10.1146/annurev-cellbio-101011-155745

146. Kayagaki N, Stowe IB, Lee BL, O'Rourke K, Anderson K, Warming S, et al. Caspase-11 cleaves gasdermin D for non-canonical inflammasome signalling. Nature (2015) 526:666–71. doi: 10.1038/nature15541

147. Shi J, Zhao Y, Wang K, Shi X, Wang Y, Huang H, et al. Cleavage of GSDMD by inflammatory caspases determines pyroptotic cell death. Nature (2015) 526:660–5. doi: 10.1038/nature15514

148. He WT, Wan H, Hu L, Chen P, Wang X, Huang Z, et al. Gasdermin D is an executor of pyroptosis and required for interleukin-1beta secretion. Cell Res. (2015) 25:1285–98. doi: 10.1038/cr.2015.139

149. Kovacs SB, Miao EA. Gasdermins: effectors of pyroptosis. Trends Cell Biol. (2017) 27:673–84. doi: 10.1016/j.tcb.2017.05.005

150. Aglietti RA, Estevez A, Gupta A, Ramirez MG, Liu PS, Kayagaki N, et al. GsdmD p30 elicited by caspase-11 during pyroptosis forms pores in membranes. Proc Natl Acad Sci USA. (2016) 113:7858–63. doi: 10.1073/pnas.1607769113

151. Ding J, Wang K, Liu W, She Y, Sun Q, Shi J, et al. Pore-forming activity and structural autoinhibition of the gasdermin family. Nature (2016) 535:111–6. doi: 10.1038/nature18590

152. Liu X, Zhang Z, Ruan J, Pan Y, Magupalli VG, Wu H, et al. Inflammasome-activated gasdermin D causes pyroptosis by forming membrane pores. Nature (2016) 535:153–8. doi: 10.1038/nature18629

153. Sborgi L, Ruhl S, Mulvihill E, Pipercevic J, Heilig R, Stahlberg H, et al. GSDMD membrane pore formation constitutes the mechanism of pyroptotic cell death. EMBO J. (2016) 35:1766–78. doi: 10.15252/embj.201694696

154. Rogers C, Fernandes-Alnemri T, Mayes L, Alnemri D, Cingolani G, Alnemri ES. Cleavage of DFNA5 by caspase-3 during apoptosis mediates progression to secondary necrotic/pyroptotic cell death. Nat Commun. (2017) 8:14128. doi: 10.1038/ncomms14128

155. Wang Y, Gao W, Shi X, Ding J, Liu W, He H, et al. Chemotherapy drugs induce pyroptosis through caspase-3 cleavage of a gasdermin. Nature (2017) 547:99–103. doi: 10.1038/nature22393

156. Van Opdenbosch N, Van Gorp H, Verdonckt M, Saavedra PHV, de Vasconcelos NM, Gonçalves A, et al. Caspase-1 engagement and TLR-induced c-FLIP expression suppress ASC/Caspase-8-dependent apoptosis by inflammasome sensors NLRP1b and NLRC4. Cell Rep. (2017) 21:3427–44. doi: 10.1016/j.celrep.2017.11.088

157. Schneider KS, Gross CJ, Dreier RF, Saller BS, Mishra R, Gorka O, et al. The inflammasome drives GSDMD-independent secondary pyroptosis and IL-1 release in the absence of Caspase-1 protease activity. Cell Rep. (2017) 21:3846–59. doi: 10.1016/j.celrep.2017.12.018

158. Mascarenhas DPA, Cerqueira DM. Pereira SF, Castanheira FVS, Fernandes TD, Manin GZ, et al. Inhibition of caspase-1 or gasdermin-D enable caspase-8 activation in the Naip5/NLRC4/ASC inflammasome. PLoS Pathog. (2017) 13:e1006502. doi: 10.1371/journal.ppat.1006502

159. Jorgensen I, Rayamajhi M, Miao EA. Programmed cell death as a defence against infection. Nat Rev Immunol. (2017) 17:151–64. doi: 10.1038/nri.2016.147

160. Jorgensen I, Lopez JP, Laufer SA, Miao EA. IL-1β, IL-18, and eicosanoids promote neutrophil recruitment to pore-induced intracellular traps following pyroptosis. Eur J Immunol. (2016) 46:2761–6. doi: 10.1002/eji.201646647

161. Jorgensen I, Zhang Y, Krantz BA, Miao EA. Pyroptosis triggers pore-induced intracellular traps (PITs) that capture bacteria and lead to their clearance by efferocytosis. J Exp Med. (2016) 213:2113–28. doi: 10.1084/jem.20151613

162. Wright JA, Bryant CE. The killer protein Gasdermin D. Cell Death Differ. (2016) 23:1897–8. doi: 10.1038/cdd.2016.100

163. Rauch I, Deets KA, Ji DX, von Moltke J, Tenthorey JL, Lee AY, et al. NAIP-NLRC4 Inflammasomes Coordinate Intestinal Epithelial Cell Expulsion with Eicosanoid and IL-18 Release via Activation of Caspase-1 and−8. Immunity (2017) 46:649–59. doi: 10.1016/j.immuni.2017.03.016

164. Knodler LA, Crowley SM, Sham HP, Yang H, Wrande M, Ma C, et al. Noncanonical inflammasome activation of caspase-4/caspase-11 mediates epithelial defenses against enteric bacterial pathogens. Cell Host Microbe (2014) 16:249–56. doi: 10.1016/j.chom.2014.07.002

165. Chen KW, Gross CJ, Sotomayor FV, Stacey KJ, Tschopp J, Sweet MJ, et al. The neutrophil NLRC4 inflammasome selectively promotes IL-1beta maturation without pyroptosis during acute Salmonella challenge. Cell Rep. (2014) 8:570–82. doi: 10.1016/j.celrep.2014.06.028

166. Kambara H, Liu F, Zhang X, Liu P, Bajrami B, Teng Y, et al. Gasdermin D exerts anti-inflammatory effects by promoting neutrophil death. Cell Rep. (2018) 22:2924–36. doi: 10.1016/j.celrep.2018.02.067

167. Monteleone M, Stow JL, Schroder K. Mechanisms of unconventional secretion of IL-1 family cytokines. Cytokine (2015) 74:213–8. doi: 10.1016/j.cyto.2015.03.022

168. Elliott EI, Sutterwala FS. Monocytes take their own path to IL-1beta. Immunity (2016) 44:713–5. doi: 10.1016/j.immuni.2016.03.015

169. Zanoni I, Tan Y, Di Gioia M, Broggi A, Ruan J, Shi J, et al. An endogenous caspase-11 ligand elicits interleukin-1 release from living dendritic cells. Science (2016) 352:1232–6. doi: 10.1126/science.aaf3036

170. Evavold CL, Ruan J, Tan Y, Xia S, Wu H, Kagan JC. The pore-forming protein gasdermin D regulates interleukin-1 secretion from living macrophages. Immunity (2018) 48:35.e6–44. doi: 10.1016/j.immuni.2017.11.013

171. Gaidt MM, Ebert TS, Chauhan D, Schmidt T, Schmid-Burgk JL, Rapino F, et al. Human monocytes engage an alternative inflammasome pathway. Immunity (2016) 44:833–46. doi: 10.1016/j.immuni.2016.01.012

172. Russo HM, Rathkey J, Boyd-Tressler A, Katsnelson MA, Abbott DW, Dubyak GR. Active Caspase-1 induces plasma membrane pores that precede pyroptotic lysis and are blocked by Lanthanides. J Immunol. (2016) 197:1353–67. doi: 10.4049/jimmunol.1600699

173. DiPeso L, Ji DX, Vance RE, Price JV. Cell death and cell lysis are separable events during pyroptosis. Cell Death Discov. (2017) 3:17070. doi: 10.1038/cddiscovery.2017.70

174. Lage SL, Amarante-Mendes GP, Bortoluci KR. Evaluation of pyroptosis in macrophages using cytosolic delivery of purified flagellin. Methods (2013) 61:110–6. doi: 10.1016/j.ymeth.2013.02.010

175. Jorgensen I, Miao EA. Pyroptotic cell death defends against intracellular pathogens. Immunol Rev. (2015) 265:130–42. doi: 10.1111/imr.12287

176. Aachoui Y, Leaf IA, Hagar JA, Fontana MF, Campos CG, Zak DE, et al. Caspase-11 protects against bacteria that escape the vacuole. Science (2013) 339:975–8. doi: 10.1126/science.1230751

177. Sauer JD, Pereyre S, Archer KA, Burke TP, Hanson B, Lauer P, et al. Listeria monocytogenes engineered to activate the Nlrc4 inflammasome are severely attenuated and are poor inducers of protective immunity. Proc Natl Acad Sci USA. (2011) 108:12419–24. doi: 10.1073/pnas.1019041108

178. Warren SE, Duong H, Mao DP, Armstrong A, Rajan J, Miao EA, et al. Generation of a Listeria vaccine strain by enhanced caspase-1 activation. Eur J Immunol. (2011) 41:1934–40. doi: 10.1002/eji.201041214

179. Miao EA, Leaf IA, Treuting PM, Mao DP, Dors M, Sarkar A, et al. Caspase-1-induced pyroptosis is an innate immune effector mechanism against intracellular bacteria. Nat Immunol. (2010) 11:1136–42. doi: 10.1038/ni.1960

180. Mariathasan S, Weiss DS, Dixit VM, Monack DM. Innate immunity against Francisella tularensis is dependent on the ASC/caspase-1 axis. J Exp Med. (2005) 202:1043–9. doi: 10.1084/jem.20050977

181. Monroe KM, Yang Z, Johnson JR, Geng X, Doitsh G, Krogan NJ, et al. IFI16 DNA sensor is required for death of lymphoid CD4 T cells abortively infected with HIV. Science (2014) 343:428–32. doi: 10.1126/science.1243640

182. Doitsh G, Galloway NL, Geng X, Yang Z, Monroe KM, Zepeda O, et al. Cell death by pyroptosis drives CD4 T-cell depletion in HIV-1 infection. Nature (2014) 505:509–14. doi: 10.1038/nature12940

183. Munoz-Arias I, Doitsh G, Yang Z, Sowinski S, Ruelas D, Greene WC. Blood-derived CD4 T cells naturally resist pyroptosis during abortive HIV-1 infection. Cell Host Microbe (2015) 18:463–70. doi: 10.1016/j.chom.2015.09.010

184. Ahmad F, Mishra N, Ahrenstorf G, Franklin BS, Latz E, Schmidt RE, et al. Evidence of inflammasome activation and formation of monocyte-derived ASC specks in HIV-1 positive patients. AIDS (2018) 32:299–307. doi: 10.1097/QAD.0000000000001693

185. Casares N, Pequignot MO, Tesniere A, Ghiringhelli F, Roux S, Chaput N, et al. Caspase-dependent immunogenicity of doxorubicin-induced tumor cell death. J Exp Med. (2005) 202:1691–701. doi: 10.1084/jem.20050915

186. Yatim N, Cullen S, Albert ML. Dying cells actively regulate adaptive immune responses. Nat Rev Immunol. (2017) 17:262–75. doi: 10.1038/nri.2017.9

187. Galluzzi L, Buque A, Kepp O, Zitvogel L, Kroemer G. Immunogenic cell death in cancer and infectious disease. Nat Rev Immunol. (2017) 17:97–111. doi: 10.1038/nri.2016.107

188. Ravichandran KS. Find-me and eat-me signals in apoptotic cell clearance: progress and conundrums. J Exp Med. (2010) 207:1807–17. doi: 10.1084/jem.20101157

189. Nagata S. Apoptotic DNA fragmentation. Exp Cell Res. (2000) 256:12–8. doi: 10.1006/excr.2000.4834

190. Trahtemberg U, Mevorach D. Apoptotic cells induced signaling for immune homeostasis in macrophages and dendritic cells. Front Immunol. (2017) 8:1356. doi: 10.3389/fimmu.2017.01356

191. Luthi AU, Cullen SP, McNeela EA, Duriez PJ, Afonina IS, Sheridan C, et al. Suppression of interleukin-33 bioactivity through proteolysis by apoptotic caspases. Immunity (2009) 31:84–98. doi: 10.1016/j.immuni.2009.05.007

192. Kazama H, Ricci JE, Herndon JM, Hoppe G, Green DR, Ferguson TA. Induction of immunological tolerance by apoptotic cells requires caspase-dependent oxidation of high-mobility group box-1 protein. Immunity (2008) 29:21–32. doi: 10.1016/j.immuni.2008.05.013

193. Ma Y, Adjemian S, Mattarollo SR, Yamazaki T, Aymeric L, Yang H, et al. Anticancer chemotherapy-induced intratumoral recruitment and differentiation of antigen-presenting cells. Immunity (2013) 38:729–41. doi: 10.1016/j.immuni.2013.03.003

194. Sistigu A, Yamazaki T, Vacchelli E, Chaba K, Enot DP, Adam J, et al. Cancer cell-autonomous contribution of type I interferon signaling to the efficacy of chemotherapy. Nat Med. (2014) 20:1301–9. doi: 10.1038/nm.3708

195. Vacchelli E, Ma Y, Baracco EE, Sistigu A, Enot DP, Pietrocola F, et al. Chemotherapy-induced antitumor immunity requires formyl peptide receptor 1. Science (2015) 350:972–8. doi: 10.1126/science.aad0779

196. Torchinsky MB, Garaude J, Martin AP, Blander JM. Innate immune recognition of infected apoptotic cells directs T(H)17 cell differentiation. Nature (2009) 458:78–82. doi: 10.1038/nature07781

197. Campisi L, Barbet G, Ding Y, Esplugues E, Flavell RA, Blander JM. Apoptosis in response to microbial infection induces autoreactive TH17 cells. Nat Immunol. (2016) 17:1084–92. doi: 10.1038/ni.3512

198. Fond AM, Ravichandran KS. Clearance of dying cells by phagocytes: mechanisms and implications for disease pathogenesis. Adv Exp Med Biol. (2016) 930:25–49. doi: 10.1007/978-3-319-39406-0_2

199. Poon IK, Lucas CD, Rossi AG, Ravichandran KS. Apoptotic cell clearance: basic biology and therapeutic potential. Nat Rev Immunol. (2014) 14:166–80. doi: 10.1038/nri3607

200. El Mezayen R, El Gazzar M, Seeds MC, McCall CE, Dreskin SC, Nicolls MR. Endogenous signals released from necrotic cells augment inflammatory responses to bacterial endotoxin. Immunol Lett. (2007) 111:36–44. doi: 10.1016/j.imlet.2007.04.011

201. Nystrom S, Antoine DJ, Lundback P, Lock JG, Nita AF, Hogstrand K, et al. TLR activation regulates damage-associated molecular pattern isoforms released during pyroptosis. EMBO J. (2013) 32:86–99. doi: 10.1038/emboj.2012.328

202. Bonnet MC, Preukschat D, Welz PS, van Loo G, Ermolaeva MA, Bloch W, et al. The adaptor protein FADD protects epidermal keratinocytes from necroptosis in vivo and prevents skin inflammation. Immunity (2011) 35:572–82. doi: 10.1016/j.immuni.2011.08.014

203. Vabulas RM, Ahmad-Nejad P, da Costa C, Miethke T, Kirschning CJ, Hacker H, et al. Endocytosed HSP60s use toll-like receptor 2 (TLR2) and TLR4 to activate the toll/interleukin-1 receptor signaling pathway in innate immune cells. J Biol Chem. (2001) 276:31332–9. doi: 10.1074/jbc.M103217200

204. Zitvogel L, Kepp O, Kroemer G. Decoding cell death signals in inflammation and immunity. Cell (2010) 140:798–804. doi: 10.1016/j.cell.2010.02.015

205. Wang Q, Imamura R, Motani K, Kushiyama H, Nagata S, Suda T. Pyroptotic cells externalize eat-me and release find-me signals and are efficiently engulfed by macrophages. Int Immunol. (2013) 25:363–72. doi: 10.1093/intimm/dxs161

206. Shi Y, Evans JE, Rock KL. Molecular identification of a danger signal that alerts the immune system to dying cells. Nature (2003) 425:516–21. doi: 10.1038/nature01991

207. Martinon F, Petrilli V, Mayor A, Tardivel A, Tschopp J. Gout-associated uric acid crystals activate the NALP3 inflammasome. Nature (2006) 440:237–41. doi: 10.1038/nature04516

208. Chen CJ, Kono H, Golenbock D, Reed G, Akira S, Rock KL. Identification of a key pathway required for the sterile inflammatory response triggered by dying cells. Nat Med. (2007) 13:851–6. doi: 10.1038/nm1603

209. Abdi J, Mutis T, Garssen J, Redegeld FA. Toll-like receptor (TLR)-1/2 triggering of multiple myeloma cells modulates their adhesion to bone marrow stromal cells and enhances bortezomib-induced apoptosis. PLoS ONE (2014) 9:e96608. doi: 10.1371/journal.pone.0096608

210. Eriksson M, Pena-Martinez P, Ramakrishnan R, Chapellier M, Hogberg C, Glowacki G, et al. Agonistic targeting of TLR1/TLR2 induces p38 MAPK-dependent apoptosis and NFkappaB-dependent differentiation of AML cells. Blood Adv. (2017) 1:2046–57. doi: 10.1182/bloodadvances.2017006148

211. Kaiser WJ, Sridharan H, Huang C, Mandal P, Upton JW, Gough PJ, et al. Toll-like receptor 3-mediated necrosis via TRIF, RIP3, and MLKL. J Biol Chem. (2013) 288:31268–79. doi: 10.1074/jbc.M113.462341

212. Zhang Y, Bliska JB. Role of Toll-like receptor signaling in the apoptotic response of macrophages to Yersinia infection. Infect Immun. (2003) 71:1513–9. doi: 10.1128/IAI.71.3.1513-1519.2003

213. He S, Liang Y, Shao F, Wang X. Toll-like receptors activate programmed necrosis in macrophages through a receptor-interacting kinase-3-mediated pathway. Proc Natl Acad Sci USA. (2011) 108:20054–9. doi: 10.1073/pnas.1116302108

214. Sun R, Zhang Y, Lv Q, Liu B, Jin M, Zhang W, et al. Toll-like receptor 3 (TLR3) induces apoptosis via death receptors and mitochondria by up-regulating the transactivating p63 isoform alpha (TAP63alpha). J Biol Chem. (2011) 286:15918–28. doi: 10.1074/jbc.M110.178798

215. Huang Z, Zhou T, Sun X, Zheng Y, Cheng B, Li M, et al. Necroptosis in microglia contributes to neuroinflammation and retinal degeneration through TLR4 activation. Cell Death Differ. (2018) 25:180–9. doi: 10.1038/cdd.2017.141

216. Ding HS, Yang J, Chen P, Yang J, Bo SQ, Ding JW, et al. The HMGB1-TLR4 axis contributes to myocardial ischemia/reperfusion injury via regulation of cardiomyocyte apoptosis. Gene (2013) 527:389–93. doi: 10.1016/j.gene.2013.05.041

217. Hornung V, Bauernfeind F, Halle A, Samstad EO, Kono H, Rock KL, et al. Silica crystals and aluminum salts activate the NALP3 inflammasome through phagosomal destabilization. Nat Immunol. (2008) 9:847–56. doi: 10.1038/ni.1631

218. Dostert C, Petrilli V, Van Bruggen R, Steele C, Mossman BT, Tschopp J. Innate immune activation through Nalp3 inflammasome sensing of asbestos and silica. Science (2008) 320:674–7. doi: 10.1126/science.1156995

219. Shio MT, Eisenbarth SC, Savaria M, Vinet AF, Bellemare MJ, Harder KW, et al. Malarial hemozoin activates the NLRP3 inflammasome through Lyn and Syk kinases. PLoS Pathog. (2009) 5:e1000559. doi: 10.1371/annotation/abca067d-b82b-4de6-93c5-0fcc38e3df05

220. Duewell P, Kono H, Rayner KJ, Sirois CM, Vladimer G, Bauernfeind FG, et al. NLRP3 inflammasomes are required for atherogenesis and activated by cholesterol crystals. Nature (2010) 464:1357–61. doi: 10.1038/nature08938

221. Mariathasan S, Weiss DS, Newton K, McBride J, O'Rourke K, Roose-Girma M, et al. Cryopyrin activates the inflammasome in response to toxins and ATP. Nature (2006) 440:228–32. doi: 10.1038/nature04515

222. Craven RR, Gao X, Allen IC, Gris D, Bubeck Wardenburg J, McElvania-Tekippe E, et al. Staphylococcus aureus alpha-hemolysin activates the NLRP3-inflammasome in human and mouse monocytic cells. PLoS ONE (2009) 4:e7446. doi: 10.1371/journal.pone.0007446

223. Gurcel L, Abrami L, Girardin S, Tschopp J, van der Goot FG. Caspase-1 activation of lipid metabolic pathways in response to bacterial pore-forming toxins promotes cell survival. Cell (2006) 126:1135–45. doi: 10.1016/j.cell.2006.07.033

224. Harder J, Franchi L, Munoz-Planillo R, Park JH, Reimer T, Nunez G. Activation of the Nlrp3 inflammasome by Streptococcus pyogenes requires streptolysin O and NF-kappa B activation but proceeds independently of TLR signaling and P2X7 receptor. J Immunol. (2009) 183:5823–9. doi: 10.4049/jimmunol.0900444

225. Kanneganti TD, Ozoren N, Body-Malapel M, Amer A, Park JH, Franchi L, et al. Bacterial RNA and small antiviral compounds activate caspase-1 through cryopyrin/Nalp3. Nature (2006) 440:233–6. doi: 10.1038/nature04517

226. Franchi L, Eigenbrod T, Munoz-Planillo R, Ozkurede U, Kim YG, Arindam C, et al. Cytosolic double-stranded RNA activates the NLRP3 inflammasome via MAVS-induced membrane permeabilization and K+ efflux. J Immunol. (2014) 193:4214–22. doi: 10.4049/jimmunol.1400582

227. Wen H, Gris D, Lei Y, Jha S, Zhang L, Huang MT, et al. Fatty acid-induced NLRP3-ASC inflammasome activation interferes with insulin signaling. Nat Immunol. (2011) 12:408–15. doi: 10.1038/ni.2022

228. Kofoed EM, Vance RE. Innate immune recognition of bacterial ligands by NAIPs determines inflammasome specificity. Nature (2011) 477:592–5. doi: 10.1038/nature10394

229. Zhao Y, Yang J, Shi J, Gong YN, Lu Q, Xu H, et al. The NLRC4 inflammasome receptors for bacterial flagellin and type III secretion apparatus. Nature (2011) 477:596–600. doi: 10.1038/nature10510

230. Franchi L, Amer A, Body-Malapel M, Kanneganti TD, Ozoren N, Jagirdar R, et al. Cytosolic flagellin requires Ipaf for activation of caspase-1 and interleukin 1beta in salmonella-infected macrophages. Nat Immunol. (2006) 7:576–82. doi: 10.1038/ni1346

231. Miao EA, Alpuche-Aranda CM, Dors M, Clark AE, Bader MW, Miller SI, et al. Cytoplasmic flagellin activates caspase-1 and secretion of interleukin 1beta via Ipaf. Nat Immunol. (2006) 7:569–75. doi: 10.1038/ni1344

232. Rauch I, Tenthorey JL, Nichols RD, Al Moussawi K, Kang JJ, Kang C, et al. NAIP proteins are required for cytosolic detection of specific bacterial ligands in vivo. J Exp Med. (2016) 213:657–65. doi: 10.1084/jem.20151809

233. Zhao Y, Shi J, Shi X, Wang Y, Wang F, Shao F. Genetic functions of the NAIP family of inflammasome receptors for bacterial ligands in mice. J Exp Med. (2016) 213:647–56. doi: 10.1084/jem.20160006

234. Hornung V, Ablasser A, Charrel-Dennis M, Bauernfeind F, Horvath G, Caffrey DR, et al. AIM2 recognizes cytosolic dsDNA and forms a caspase-1-activating inflammasome with ASC. Nature (2009) 458:514–8. doi: 10.1038/nature07725

235. Boyden ED, Dietrich WF. Nalp1b controls mouse macrophage susceptibility to anthrax lethal toxin. Nat Genet. (2006) 38:240–4. doi: 10.1038/ng1724

236. Faustin B, Lartigue L, Bruey JM, Luciano F, Sergienko E, Bailly-Maitre B, et al. Reconstituted NALP1 inflammasome reveals two-step mechanism of caspase-1 activation. Mol Cell (2007) 25:713–24. doi: 10.1016/j.molcel.2007.01.032

237. Xu H, Yang J, Gao W, Li L, Li P, Zhang L, et al. Innate immune sensing of bacterial modifications of Rho GTPases by the Pyrin inflammasome. Nature (2014) 513:237–41. doi: 10.1038/nature13449

238. Salvestrini V, Orecchioni S, Talarico G, Reggiani F, Mazzetti C, Bertolini F, et al. Extracellular ATP induces apoptosis through P2X7R activation in acute myeloid leukemia cells but not in normal hematopoietic stem cells. Oncotarget (2017) 8:5895–908. doi: 10.18632/oncotarget.13927

239. Placido R, Auricchio G, Falzoni S, Battistini L, Colizzi V, Brunetti E, et al. P2X(7) purinergic receptors and extracellular ATP mediate apoptosis of human monocytes/macrophages infected with Mycobacterium tuberculosis reducing the intracellular bacterial viability. Cell Immunol. (2006) 244:10–8. doi: 10.1016/j.cellimm.2007.02.001

240. Ishibashi O, Ali MM, Luo SS, Ohba T, Katabuchi H, Takeshita T, et al. Short RNA duplexes elicit RIG-I-mediated apoptosis in a cell type- and length-dependent manner. Sci Signal. (2011) 4:ra74. doi: 10.1126/scisignal.2001614

241. Thapa RJ, Ingram JP, Ragan KB, Nogusa S, Boyd DF, Benitez AA, et al. DAI senses influenza A virus genomic RNA and activates RIPK3-dependent cell death. Cell Host Microbe (2016) 20:674–81. doi: 10.1016/j.chom.2016.09.014

Conflict of Interest Statement: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

The reviewer FD and handling Editor declared their shared affiliation at the time of review.

Copyright © 2018 Amarante-Mendes, Adjemian, Branco, Zanetti, Weinlich and Bortoluci. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

ORIGINAL RESEARCH
published: 15 September 2017
doi: 10.3389/fimmu.2017.01127

Systems Approach Reveals Nuclear Factor Erythroid 2-Related Factor 2/Protein Kinase R Crosstalk in Human Cutaneous Leishmaniasis

Áislan de Carvalho Vivarini[1], Teresa Cristina Calegari-Silva[1], Alessandra Mattos Saliba[2], Viviane Sampaio Boaventura3,4, Jaqueline França-Costa3,4, Ricardo Khouri3,4, Tim Dierckx[5], Karina Luiza Dias-Teixeira[1], Nicolas Fasel[6], Aldina Maria Prado Barral3,4, Valéria Matos Borges3,4, Johan Van Weyenbergh[5]* and Ulisses Gazos Lopes[1]*

1 Laboratory of Molecular Parasitology, Carlos Chagas Filho Biophysics Institute, Center of Health Science, Federal University of Rio de Janeiro, Rio de Janeiro, Rio de Janeiro, Brazil

2 Department of Microbiology, Immunology and Parasitology – FCM/UERJ, State University of Rio de Janeiro, Rio de Janeiro, Rio de Janeiro, Brazil

3 Gonçalo Moniz Institute, Oswaldo Cruz Foundation, Salvador, Bahia, Brazil

4 Federal University of Bahia, Salvador, Bahia, Brazil

5 Department of Microbiology and Immunology, Rega Institute for Medical Research, KU Leuven, Leuven, Belgium

6 Faculty of Biology and Medicine, Department of Biochemistry, University of Lausanne, Lausanne, Switzerland

OPEN ACCESS

Edited by:

Leticia A. Carneiro, Federal University of Rio de Janeiro, Brazil

Reviewed by:

Min Zhang, Indiana University School of Medicine, United States
Peter Epeh Kima, University of Florida, United States

*Correspondence:

Johan Van Weyenbergh
j.vw@live.be;
Ulisses Gazos Lopes
lopesu@biof.ufrj.br

Specialty section:

This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology

Received: 29 May 2017
Accepted: 28 August 2017
Published: 15 September 2017

Citation:

Vivarini ÁC, Calegari-Silva TC, Saliba AM, Boaventura VS, França-Costa J, Khouri R, Dierckx T, Dias-Teixeira KL, Fasel N, Barral AMP, Borges VM, Van Weyenbergh J and Lopes UG (2017) Systems Approach Reveals Nuclear Factor Erythroid 2-Related Factor 2/Protein Kinase R Crosstalk in Human Cutaneous Leishmaniasis. Front. Immunol. 8:1127. doi: 10.3389/fimmu.2017.01127

Leishmania parasites infect macrophages, causing a wide spectrum of human diseases, from cutaneous to visceral forms. In search of novel therapeutic targets, we performed comprehensive in vitro and ex vivo mapping of the signaling pathways upstream and downstream of antioxidant transcription factor [nuclear factor erythroid 2-related factor 2 (Nrf2)] in cutaneous leishmaniasis (CL), by combining functional assays in human and murine macrophages with a systems biology analysis of in situ (skin biopsies) CL patient samples. First, we show the PKR pathway controls the expression and activation of Nrf2 in Leishmania amazonensis infection in vitro. Nrf2 activation also required PI3K/Akt signaling and autophagy mechanisms. Nrf2- or PKR/Akt-deficient macrophages exhibited increased levels of ROS/RNS and reduced expression of Sod1 Nrf2-dependent gene and reduced parasite load. L. amazonensis counteracted the Nrf2 inhibitor Keap1 through the upregulation of p62 via PKR. This Nrf2/Keap1 observation was confirmed in situ in skin biopsies from Leishmania-infected patients. Next, we explored the ex vivo transcriptome in CL patients, as compared to healthy controls. We found the antioxidant response element/Nrf2 signaling pathway was significantly upregulated in CL, including downstream target p62. In silico enrichment analysis confirmed upstream signaling by interferon and PI3K/Akt, and validated our in vitro findings. Our integrated in vitro, ex vivo, and in silico approach establish Nrf2 as a central player in human cutaneous leishmaniasis and reveal Nrf2/PKR crosstalk and PI3K/Akt pathways as potential therapeutic targets.

Keywords: Leishmania, macrophage, nuclear factor erythroid 2-related factor 2, PKR, Sod1

INTRODUCTION

Human cutaneous leishmaniasis (CL) is spread worldwide, and the incidence is estimated to be from 0.7 to 1.2 million cases each year (1). Different clinical manifestations occur in humans due to the immune response and the infection by distinct Leishmania species (2). Leishmania parasites exhibit a plethora of adaptive mechanisms that interfere with several macrophage functions through the manipulation of host signaling pathways (3).

The imbalance between oxidative stress and cytoprotective systems of detoxification dictates the outcome of intracellular parasitic infections (4). The transcription factor [nuclear factor erythroid 2-related factor 2 (Nrf2)] is a master regulator of phase II defense gene expression that may protect cells from oxidative stress. The DNA promoter sequences of phase II defense genes share the canonical antioxidant response element (ARE), which is recognized by Nrf2 (5). Nrf2-dependent expression leads to profound effects on the suppression of the inflammatory response and immune activation through Toll-like receptors (6, 7).

The control of Nrf2 activation is dictated by different post-translational modifications. Multiple sites on the Nrf2 protein are phosphorylated by kinases, such as PERK, members of the MAPK family, PKCζ, and GSK3β, increasing the nuclear translocation and binding of the protein to ARE elements on the promoters of target genes (8–10).

The PI3K/Akt pathway modulates Nrf2 signaling (11) and, importantly, recent reports have demonstrated the modulation of the PI3K/Akt pathway upon Leishmania amazonensis infection (12). The enzyme GSK3, a target of Akt1, phosphorylates the Nh6 domain of Nrf2 and facilitates the action of ubiquitin ligase, leading to proteasomal degradation. The inhibition of GSK3 by phosphorylation allows the nuclear translocation of Nrf2 (13).

Kelch-like ECH-associated protein 1 (Keap1) is a major inhibitor of Nrf2 that constitutively induces the ubiquitination of the Nh2 domain, directing Nrf2 to proteasomal degradation. Keap1 is uncoupled from Nrf2 because of post-translational modifications due to oxidative stress, releasing Nrf2 for nuclear translocation (14).

Autophagy may modulate Nrf2 activation via Keap1 degradation (15). The components of the autophagy pathway are sensors of oxidative stress (16), and the increase in the expression of p62 (Sqstm1), an Nrf2 target autophagy gene (17), favors the cellular capacity to process proteins destined for the autophagosome, decreasing oxidative stress (18).

The phosphorylation of p62 allows its binding to several cargo proteins, including Keap1, leading to autophagy as well as the Nrf2 stability and activation (19). The activation of Nrf2 through the p62/autophagy non-canonical pathway has also been demonstrated in macrophages treated with LPS, PolyI:C and peptidoglycan (PGN) upon the engagement of TLR4, TLR3, and TLR2, respectively (20).

Double-stranded RNA-dependent protein kinase [protein kinase R (PKR)] has in the Nh2-terminal domain two double-stranded RNA-binding motifs, and its kinase catalytic domain is located in the carboxyl-terminal (21, 22). On binding dsRNA, PKR dimerizes and undergoes autophosphorylation at multiple sites (23). Expression of catalytically defective mutant PKR (K296R) in cells inhibited the autophosphorylation and subsequent the activation of its major substrate eIF2-α (24, 25). PKR-mediated signaling may promote autophagy through eIF2-α phosphorylation (26). In several viral infections, PKR plays an essential role in the autophagy trigger (27). In STAT3−/− cells, PKR is able to induce autophagy through LC3-I to LC3-II conversion and the formation of vacuole compartments (28). In cells treated with type I interferon (IFN-I), both LC3 activation and p62 expression are increased (29).

In this work, we unveil the mechanisms that regulate Nrf2 gene expression in a PKR-dependent fashion. We describe for the first time the signaling pathway that coordinates Nrf2 activation during Leishmania infection. Finally, the induction of cytoprotective genes through the novel PKR/Nrf2 pathway may represent a prominent therapeutic mechanism for treatment and guide the development of novel targets in both infectious and inflammatory diseases.

MATERIALS AND METHODS

Reagents

Chloroquine diphosphate salt, DL-sulforaphane (SFN), phorbol-12 myristate-13 acetate (PMA), N-acetyl-L-cysteine (NAC), Wortmannin, and LY294002 hydrochloride were purchased from Sigma-Aldrich (St. Louis, MO, USA). AKTi (AKT inhibitor VIII, Akt1/2) was purchased from Santa Cruz Biotechnology (Dallas, TX, USA). Poly (cytidylic-inosic) acid potassium salt (PolyI:C) and the PKR inhibitor CAS 608512-97-6 were purchase from Calbiochem-Millipore (Darmstadt, Germany). Human recombinant interferon-alpha 2b was obtained from Blausiegel (Cotia, SP, Brazil).

Cell Lines and Culture

The mouse macrophage leukemia cell line RAW 264.7 (TIB-71; American Type Culture Collection (ATCC), Manassas, VA, USA), the human monocytic leukemia cell line THP-1 (ATCC:TIB202TM) and the human embryonic kidney cell line HEK-293T (ATCC:CRL-11268) were maintained in DMEM medium with high glucose (Vitrocell Embriolife, Campinas, SP, Brazil) supplemented with 10% heat-inactivated fetal bovine serum (Sigma-Aldrich, St. Louis, MO, USA). THP-1 cells were differentiated to macrophages with 40 ng/mL of PMA for 3 days. Afterward, the cells were washed three times with PBS and incubated with fresh medium for an additional 3 days. RAW 264.7 cells expressing either empty vector (RAW-Bla cells) or a dominant-negative PKR K296R (RAW-DN-PKR cells) were donated by Dr. Aristóbolo Silva, Federal University of Minas Gerais, Brazil.

Peritoneal Macrophages

Ten-week-old male 129/SvEv PKR−/− (PKR-ko) and their respective wild-type littermates (WT) were used for experiments. Briefly, 4 days before peritoneal lavage, 2 mL of 3% thioglycolate were intra-peritoneally injected in each mouse. Thioglycolate-elicited peritoneal macrophages from wild-type or PKR-knockout 129Sv/Ev were obtained by injecting 8 mL of serum-free DMEM into the peritoneal cavity. After 1 h, the cells were washed once in PBS and then plated in in DMEM medium supplemented with 10% FBS on glass coverslips at 2 × 10^5/well in 6-well or 24-well polystyrene plates for subsequent Leishmania infection assays.

Cell Treatment

To induce the activation of Nrf2, 10 mM SFN were used as positive controls. For the inhibition of PKR activity, we pretreated the cells for 1 h with 300 nM of the PKR inhibitor (PKRi). To induce PKR activation, poly(inosinic-cytidylic-) acid potassium salt (PolyI:C) at a final concentration of 25 µg/mL or recombinant IFNα-2b at 1,000 U/mL were used. PI3K/Akt inhibition was accomplished by cell treatment with 10 µM LY294002, 10 mM Wortmannin or 5 mM AKTI (AKT inhibitor VIII Akt1/2). To inhibit autophagy, we used 40 µM chloroquine. N-acetylcysteine (NAC) was used at a concentration of 10 mM.

Parasites, Culture Conditions, and Infection

Leishmania (Leishmania) amazonensis (WHOM/BR/75/Josefa) and Leishmania (Viannia) braziliensis (BA788) were used in this study. The L. (L.) amazonensis strains obtained from biopsies of patients with diffuse cutaneous leishmaniasis (DCL) (Ba276, Ba336, and Ba760) or localized cutaneous leishmaniasis (Ba69, Ba73, and Ba125) were also used in vitro assays. The promastigote forms were grown at 26°C in Schneider’s Insect Medium (Sigma-Aldrich) with 10% fetal bovine serum, and metacyclic promastigotes were collected from stationary cultures and used for cell infections. Macrophages were infected with Leishmania promastigotes at a parasite:cell ratio of 10:1 at 37°C. Infected macrophages were counted in a Neubauer Chamber by light microscopy to assess the infection index, which was calculated by multiplying the percentage of infected macrophages by the average number of parasites per macrophage in Giemsa-stained slides.

Immunoblotting

THP-1 cells (1 × 10^6 cells) were washed twice with ice-cold PBS and then lysed in 100 µL of lysis buffer (50 mM Tris-HCl, pH 7.5, 5 mM EDTA, 10 mM EGTA, 50 mM NaF, 20 mM β-glycerophosphate, 250 mM NaCl, 0.1% Triton X-100, 1 µg/mL BSA, and a 1:100 dilution of protease inhibitor cocktail, Sigma-Aldrich, St. Louis, MO, USA) for total protein extraction. For nuclear protein extraction, after infection and/or treatment, the cells were washed twice with 1x PBS and then lysed with 100 µL of buffer A (HEPES 10 mM pH 7.9. 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, NP-40 0,25% (v/v); cocktail of protease inhibitors) for 10 min on ice. The lysed cells were centrifuged at 14,000 g for 1 min at 4°C, and the pellet was resuspended in 60 µL of buffer C (20 mM HEPES pH 7.9, 0.4 M NaCl, 1 mM EDTA, 1 mM EGTA, 20% glycerol, protease inhibitor cocktail) and incubated on ice for 20 min. The lysate was centrifuged at 14,000 g for 5 min, and the supernatant containing nuclear proteins was collected in a new tube. The protein extracts were subjected to electrophoresis on 10% SDS-polyacrylamide gels and transferred to nitrocellulose membranes (Amersham Biosciences, Piscataway, NJ, USA). After blocking with 5% non-fat dry milk in TBS with 0.1% Tween-20 (TBS-T), the blots were incubated over-night with antibodies against PKR (12297), Nrf2 (12721), GSK3 (9369), Sqstm1/p62 (5114), LC3B (2775), phospho-GSK3β-Ser9 (9336), phospho-Akt-Ser473 (9271), phospho-eIF2α-Ser51 (9721), α-Tubulin (2144), β-Tubulin (2146), and Lamin A/C (2032) from Cell Signaling Technology; phospho-PKR Th451 (07-886) from Millipore; keap1 (150654) from Abcam; and β-actin (47778), Sod1 (8637), followed by anti-rabbit (2004) or anti-mouse (2005) horseradish peroxidase-conjugated IgG (1:4,000) from Santa Cruz Biotechnology. The membranes were then submitted to three washes with 0.1% TBS-T after each incubation, and the proteins were detected using the ECL chemiluminescent detection system (Amersham Biosciences).

Immunohistochemistry

To validate the differential expression of Nrf2 (C20—Santa Cruz Biotechnology) and keap1 (150654—Abcam) in DCL and LCL samples, immunohistochemistry was performed on formalin-fixed, paraffin-embedded (FFPE) sections. Briefly, after deparaffinization, rehydration and target retrieval (DAKO Corporation, Hamburg, Germany), slides from five DCL and five LCL cases were incubated with serum-free protein block reagent and then incubated overnight with Nrf2 or Keap1 (4and 10 mg/mL, respectively, both from Abcam, Cambridge, United Kingdom) or anti-rabbit isotype control antibodies. After the sequential application of a peroxidase-blocking reagent, DAKO EnVision + System-HRP (DAKO Corporation, Hamburg, Germany), digital images of the tissue sections were captured using a Nikon E600 light microscope and a Q-color 1 Olympus digital camera. Sections of prostate and lung adenocarcinoma were used as positive controls. Quantification of the stained areas was performed using Image Pro Plus software (Media Cybernetics).

Luciferase Assays

To investigate the promoter activity, RAW-264.7 cells (1 × 10^5 cells per well) was plated in 48-well polystyrene plates and transfected with 1 µg of reporter plasmids using LIPOFECTAMINE 2000 reagent (Invitrogen, Carlsbad, CA, USA). THP-1 cells (2 × 10^6) were transfected with 0.5 µg of luciferase reporter plasmids using Nucleofector™ Technology (Lonza, Basel, Switzerland) according to the manufacturer’s instructions. The following plasmids were employed in the assays: Sod1-basal, Sod1-ΔARE, Sod1-WT, 3xARE, and Nrf2-WT. For normalization of the luciferase readout, the plasmid pRL-CMV (Promega) was used. After infection and treatment, the cells were washed with PBS, lysed according to the Dual Luciferase System protocol (Promega), and analyzed using the GloMax®-Multi detection system (Promega Corp., Madison, WI, USA).

Chromatin Immunoprecipitation Assay (ChIP)

Chromatin immunoprecipitation assay analysis was carried out according to the Simple ChIP Enzymatic Chromatin IP kit protocol (Cell Signaling). RAW 264.7 (WT-PKR and DN-PKR) cells or the human monocytic leukemia cell line THP-1 (ATCC:TIB202TM) were plated to confluence in 15 cm dishes. After infection, the cells were fixed with 1% formaldehyde for 10 min at room temperature, followed by the addition of glycine to a final concentration of 125 mM for 5 min at room temperature prior to cell lysis. One unit of micrococcal nuclease was added to the sample and incubated for 20 min at 37°C to digest DNA to the length of approximately 150–800 base-pairs. The chromatin was immunoprecipitated with 5 µg/mL anti-Nrf2 antibody (D1Z9C-XP—Cell Signaling Technology, Danvers, MA, USA) at 4°C under rotation for 16 h. The DNA isolated from the immunoprecipitated material was amplified by real-time PCR using SybrGreen, and the DNA sequences of the primers used were Sod1-ARE.chip-F: 5′-AAGTCCGGGTCCCAGCTCAGAG-3′ and Sod1-ARE.chip-R: 5′-TTGGTGCAAGCACACCGGGAG-3′; p62-ARE.chip-F: 5′-CCCCACAGTTCCCCATTGGC-3′ and p62-ARE.chip-R: 5′-GACAGTGGGGACGCAAAGGC-3′; and Nrf2-AREL2chip-F: 5′-AAGTCCGGGTCCCAGCTCAGAG-3′ and Nrf2-AREL2chip-R: 5′-TTGGTGCAAGCACACCGGGAG-3′. As a control, 1/50 of the digested input chromatin was similarly processed and analyzed in the absence of immunoprecipitation. To calculate the input percentage of the samples, the input was adjusted to 100% (average Ct of input − Log_2 of 50), followed by the application of the 100 × 2(adjusted input − average Ct(IP)) formula.

Cloning and Generation of Luciferase Reporter Plasmids

Total DNA was extracted from THP-1 cells using a Wizard® Genomic DNA Purification kit (Promega) and measured using a BioPhotometer (Eppendorf). One PCR was carried out with primers spanning different regions of the Sod1 and Nrf2 promoters, yielding different fragment sizes, in the following conditions: 20 ng of genomic DNA and 35 cycles of 95°C for 1 min, 58°C for 1 min, and 72°C for 1 min. The DNA sequences of the primers used were Sod1.wt-F: 5′- GTCTCGAGCTGTAGGGTTGTGGCCTTGCCAAA-3′, Sod1.ΔARE-F: 5′-GTCTCGAGGCCAATTTCGCGTACTGCAACCG-3′, Sod1.basal-F: 5′-GTCTCGAGCTCGCGACCCGAGGCTG-3′ and Sod1-R: 5′-GTAGATCTCAGGAGACTACGACGCAAACCAGC-3′; and Nrf2-F: 5’ AAGTCCGGGTCCCAGCTCAGAG 3′ and Nrf2-R: 5′-TGGGGGCGGAACAAGGACCTAG-3′. A 1.8% agarose gel was run for 50 min at 100 V, and the amplicons were extracted from the gel and purified with the Zymoclean Gel DNA Recovery kit ™ (Zymo Research). The amplicons were ligated into a pJet-Blunt plasmid (Fermentas) with T4 ligase (Promega) for the first selection of positive colonies. After confirming positivity through PCR and a digestion assay, a colony was selected and grown, and a new plasmid extraction was performed. Digestion of the pJet-Blunt vector containing subcloned amplicons was performed with the Bgl-II enzyme (Promega), and the products were subjected to electrophoresis on a 2% agarose gel to extract the gel fragments. The pGL2-basic plasmid was also digested with the Bgl-II enzyme for the subsequent binding of the amplicons with T4 ligase enzyme (Promega). The cloned fragments and final vectors were then transformed into DH5α bacteria, and colonies were selected for further confirmation by sequencing. To obtain a luciferase-expressing pGL2-basic plasmid containing three copies of the sequence regulatory region ARE (3xARE), two oligos (5′-ATGCCGCTCGAGAATGACATTTCTAGAATGACATTTCTAGAATGACATTTCTAGAGATCTCGGCCG-3′ and 3′-TACGGCGAGCTCTTACTGTAAAGATCTTACTGTAAAGATCTTACTGTAAAGATCTCTAGAGCCGGC-5′) were designed and annealed to serve as templates for a PCR under the following conditions: 20 ng of DNA oligo and 35 cycles of 95°C for 1 min, 52°C for 1 min, and 72°C for 1 min with the primers 3xARE-F: 5′-ATGCCGCTCGAGAATG 3′, and 3xARE-R: 5′-CGGCCGAGATCTCTAGA 3′. The binding reactions and digestion with the Bgl-II enzyme followed the same protocol as described above.

Lentiviral Production and THP-1 Transduction

HEK-293T cells were used for shNrf2 lentiviral production. Initially, we co-transfected the cells with two packaging plasmids (pΔ8.9 and pVSVG) containing accessory proteins for the generation of the virus and capsid, respectively, along with the plasmid pLKO.1-shMission-Nrf2 (Sigma-Aldrich). For HEK-293T transfection, 60 µL of FuGENE HD reagent (Promega) was used in a 100 mm dish containing approximately 4 × 10^6 cells, along with 10 µg of target plasmid, 6 µg of pVSVG and 4 µg of pΔ8.9. After 24 h of transfection, the culture medium was changed and, over the next 2 days, the supernatants were collected at 10 mL/day. The 20 mL of supernatant was ultracentrifuged at 16,000 rpm for 90 min at 4°C, and the pellet was resuspended in 1 mL of DMEM without serum. Viral transduction in THP-1 cells was accomplished in 2 × 10^6 cells incubated with 1 mL of virus preparation for 48 h.

Fluorimetric Assays

The production of reactive oxygen species (ROS), nitric oxide (NO), and peroxynitrite (OONO) was performed by fluorimetry. For this, 10^5 cells were seeded in black 96-well plates and maintained for 24 h in DMEM containing 10% fetal bovine serum. The day after, the cells were washed three times with PBS, and HBSS medium without serum was added and incubated for 1 h at 37°C and 5% CO_2. The cells were incubated with different fluorescent probes for 1 h. Then, the cells were washed with PBS and treated with medium or infected with L. amazonensis. Fluorescence counting was monitored after incubation at 1-h intervals for up to 6 h (GloMax™). The production of ROS was detected using the probe CM-H2DCFDA (5 mM, Molecular Probes), with excitation at 495 nm and emission at 525 nm. For NO production, the DAF-FM probe (5 mM, Molecular Probes) was used, with excitation at 495 nm and emission at 515 nm. For the production of OONO-, the probe HPF (5 mM, molecular probes) was used, with excitation at 490 nm and emission at 515 nm.

Patient Characteristics

Diffuse cutaneous leishmaniasis patients (n = 4) were recruited at our reference clinic in São Luiz, Maranhão, Brazil. DCL patients exhibited chronic progression of the disease with several remissions, multiple nodular and highly parasitized lesions throughout the skin, and a negative DTH response. LCL patients (n = 5), recruited at our reference clinic in Jiquiriçá, Bahia, Brazil, had a single or a few ulcerated lesions present for up to 2 months and a positive DTH response (30). The clinical and epidemiological data from patients with DCL and those with LCL are summarized in Table S3 in Supplementary Material. Skin biopsies were preserved as paraffin-embedded specimens.

Patient Recruitment and Diagnosis for Transcriptomic Analysis

This study was approved by the Ethics Committee of the Gonçalo Moniz Research Center (FioCruz-Bahia). Informed consent was obtained from all patients and healthy controls. CL patients were diagnosed according to characteristic lesion morphology, positive skin test, seropositivity toward Leishmania antigen and/or the presence of parasites in the lesion. LCL patients infected with Leishmania braziliensis (n = 18, 10 male, 29.6 ± 2.3 years) were recruited at diagnosis (before treatment) in two outpatient clinics (Jequié and Jiquiriçá-BA, NE Brazil) covering the same rural area.

Ethics Statement

Written informed consent was obtained from all participants or legal guardians, and all of the data analyzed were anonymized. The project was approved by the Institutional Review Board of Centro de Pesquisas Gonçalo Moniz, FIOCRUZ–BA (license number 136/2007) and complies with the guidelines of the Declaration of Helsinki.

Microarray Analysis

PBMCs from LCL patients and healthy controls were processed in parallel and immediately frozen in Trizol to preserve RNA integrity. Following Trizol extraction, total RNA was further purified using an RNeasy kit according to the manufacturer’s protocol (QIAGEN, Venlo, Netherlands). Affymetrix Whole Genome microarray analysis was performed by the VIB MicroArray Facility (Leuven, Belgium) using a GeneChip® Human Gene 1.0 ST Array with the WT PLUS reagent kit (Affymetrix, Santa Clara, CA, USA) according to the manufacturer’s specifications. Data preprocessing (RMA) was performed using the Bioconductor xps package. Microarray data were deposited in GEO (accession number: GEO Submission (GSE80008) (NCBI tracking system #17832057)).

nCounter Digital Transcriptomics

RNA extraction from skin biopsies was performed as above. Digital quantification of selected genes (NRF2, PKR, SOD1, SOD2, KEAP1, HMOX1) was performed by nCounter (Nanostring). Two-step normalization using internal positive and negative control RNAs, as well as PTPRC (CD45) normalization to correct for differences in tissue leukocyte infiltration, was performed as previously described (31).

Enrichment Analysis

The ingenuity pathway analysis (IPA) program was used to perform the initial pathway/function level analysis on genes determined to be differentially expressed in the microarray analysis (IPA version 9.0, Build 116623, Content version 3211, Ingenuity Systems, Red Wood City, CA, USA). Uncorrected p-values and absolute fold-changes were used with cut-offs of p < 0.05. Based on a scientific literature database, the genes were sorted into gene networks and canonical pathways, and significantly overrepresented pathways were identified. Further enrichment analysis was performed, including Gene Ontology (GO) term enrichment using the WEB-based GEne SeT AnaLysis Toolkit (WebGestalt), KEGG pathway enrichment using the pathway database from the Kyoto Encyclopedia of Genes, and Genomes and transcription factor target enrichment using data from the Broad Institute Molecular Signatures Database (MSigDB). Genesets from the GO, KEGG pathways, WikiPathways, and Pathway Commons databases, as well as transcription factors, were considered overrepresented if their corrected p-value was smaller than 0.05.

Statistical Analysis

The data were analyzed by one-way ANOVA for independent samples or Mann–Whitney (two-sided t-test) using Prism 5 software. The data represent the mean ± SD of the mean. The data are expressed as the average of three independent determinations, and significant differences were indicated by *p < 0.05.

RESULTS

Leishmania Induces Nrf2 via PKR

The oxidative stress response plays a determinant role in the control of intracellular pathogens such as Leishmania (32). L. amazonensis dampens some macrophage functions, including the induction of oxidative stress (33, 34). Importantly, Nrf2 activation may promote infection tolerance, thus favoring pathogen survival. We sought to investigate whether L. amazonensis would induce Nrf2 via PKR. Figure 1A and Figure S1A in Supplementary Material shows that Nrf2 translocated to the nuclei of macrophages during the initial phase of interaction with the parasite. Importantly, Nrf2 translocation was not observed in infected pkr-ko and DN-PKR macrophages, respectively. Nrf2 levels were augmented in 6 h of infection and were induced by PKR signaling (Figure 1B; Figure S1B in Supplementary Material). The main target of PKR, eIf2α, is also not activated by phosphorylation in PKR-deficient cells (Figures S1C,D in Supplementary Material). PKR activation by inducers, such as PolyI:C or IFN-I added to macrophages also induced Nrf2 translocation and expression (Figure 1C). Next, we investigated the binding of Nrf2 to cognate Nrf2 promoter (Figure 1D). Our data show that Nrf2 only occupied the ARE sequences in infected wild-type macrophages by ChIP. To address whether ARE genes are activated in Leishmania infection, we constructed two luciferase reporter plasmids. The 3xARE construct contains the canonical ARE promoter response element, while the other construct contains the Nrf2 promoter (also spanning an ARE-like element). Figure 1E shows that the 3xARE regulatory sequence drove luciferase expression in infected wild-type macrophages, while luciferase expression was abrogated in DN-PKR cells. Importantly, the Nrf2 promoter was also induced in infected wild-type macrophages. In summary, our results show that L. amazonensis induces Nrf2 in a PKR-dependent manner.

FIGURE 1 | Leishmania amazonensis induced nuclear factor erythroid 2-related factor 2 (Nrf2) expression and nuclear translocation in a protein kinase R (PKR)-dependent manner. Peritoneal macrophages from wild-type or PKR-ko 129/sv mice were infected with stationary promastigotes forms of L. amazonensis for 2 h (A) or 6 h (B). Western-blot was carried out for nuclear or total protein extract, respectively, and then assay was performed using Nrf2 antibody. (C) THP-1 cells were infected with Leishmania amazonensis or treated with IFN-α or PolyI:C for 2 h for nuclear extract or 6 h for total protein extract, before western-blot analysis with Nrf2 antibody. (D) RAW-WT-PKR and RAW-DN-PKR cells were infected with stationary promastigotes forms of L. amazonensis for 4 h and then submitted to chromatin immunoprecipitation assay (ChIP) using Nrf2 ChIP-antibody. (E) RAW 264.7 cells were transiently transfected with p3xARE- or pNrf2-promoter-luciferase reporter plasmids constructs and infected with L. amazonensis 24 h post-transfection. Whole-cell lysates were analyzed for luciferase activity 24 h later. Results are representative of three independent experiments. *p < 0.05.

Nrf2 and PKR Signaling Control SOD1 Gene Expression

Recent reports have demonstrated that L. amazonensis activates the classical antiviral response mediated by PKR, leading to Sod1 expression, favoring parasite growth in infected macrophages (35, 36). To address the role of Nrf2 on Sod1 expression in infected macrophages, we cloned the Sod1 promoter and deleted the regulatory regions in the Luciferase vector (pGL2) (Figure 2A). The Sod1 promoter was induced in wild-type infected macrophages, while the deletion of the ARE sequence disrupted Luc expression. Accordingly, Sod1 expression, which is controlled by Nrf2, was only increased in wild-type infected macrophages (Figure 2B). Our data show that ARE element on Sod1 promoter was occupied by Nrf2 only in infected wild-type macrophages by ChIP (Figure 2C). We aimed to test the hypothesis that Sod1 dependence of Nrf2 activity, we developed a macrophage shNrf2 knockdown cell line. In only wild-type infected macrophages, the parasites induce Sod1 expression (Figure 2D). The quantification of infection index show a decrease on proliferation of Leishmania in Nrf2 knockdown cells (Figure 2E). These data support the link between Sod1 and two major signaling pathways represent by Nrf2 and PKR.

FIGURE 2 | Sod1 regulation through nuclear factor erythroid 2-related factor 2 (Nrf2) and protein kinase R (PKR) signaling. (A) RAW 264.7 WT or DN-PKR cells were transiently transfected with Sod1-Luc plasmids and infected with stationary promastigotes of Leishmania amazonensis for additional 24 h before luciferase activity assay. (B) The same cells were also infected with stationary promastigotes of L. amazonensis for 18 h before total protein extract for western-blot analyzes with Sod1 and α-tubulin antibodies. (C) RAW 264.7 cells were infected with stationary promastigotes forms of L. amazonensis for 4 h and then submitted to chromatin immunoprecipitation assay (ChIP) using Nrf2 ChIP-antibody. shNrf2 or shControl THP-1 cells were infected with stationary promastigotes of L. amazonensis, (D) Sod1 protein expression was analyzed, and (E) infection index was evaluated. Results are representative of three independent experiments. *p < 0.05.

Akt1 Controls Nrf2 Induction in Infected Macrophages

Nrf2 activation is controlled at different levels, including indirect phosphorylation by Akt1 (11). Because L. amazonensis promotes Akt1 activation (12), we aimed to investigate its role in Nrf2 induction. Initially, we examined whether the induction of Akt1 by L. amazonensis relied on PKR expression. Figure 3A shows that GSK3 phosphorylation due to Akt1 depended on PKR. The phosphorylation of Akt depends on PKR during Leishmania infection (Figures S3A,B in Supplementary Material). Of note, Nrf2 induction required Akt signaling, as shown in infected shAkt1 macrophages (Figures 3B,C). In macrophages treated with pharmacological inhibitors of Akt1/2 and PI3K (Figures 3D,E), we also observed the same pattern of Nrf2 repression in nucleus translocation and protein expression. As predicted, ARE element, Nrf2 Luciferase and Nrf2 promoter occupancy in ChIP assay were induced by L. amazonensis infection in an Akt1-dependent manner (Figures 3F,G). Likewise, Sod1 expression followed the same PI3K/Akt1 dependence pattern. ChIP assays corroborated these findings, where the occupancy of ARE in the Sod1 promoter by Nrf2 depended on Akt1 (Figure 3H).

FIGURE 3 | Protein kinase R (PKR)-dependent PIK3/Akt signaling activation controls positively the nuclear factor erythroid 2-related factor 2 (Nrf2) functions in Leishmania-infected macrophages. (A) Peritoneal macrophages from wild-type or PKR-ko 129/sv mice were infected with stationary promastigotes forms of Leishmania amazonensis at indicated times. Western-blot was carried out for total protein extract with anti-phospho-GSK3 and anti-GSK3. THP-1 cells stably knocked-down for Akt1 expression (B,C) and treated with PI3K/Akt inhibitors (LY294002, Wortmannin and Akt-inhibitor-VIII) (D,E) were infected with stationary promastigotes forms of Leishmania amazonensis at indicate times. Nuclear and total protein extracts were analyzed using Nrf2 antibody. (F) THP-1 cells were transiently transfected with p3xARE- or pNrf2-promoter Luciferase reporter plasmids. Twenty-four hours post-transfection, cells were differentiated into macrophages with phorbol-12 myristate-13 acetate (PMA) treatment for 6 days. The cells were infected with stationary promastigotes forms of L. amazonensis and/or treated with PI3K/Akt inhibitors for additional 24 h. Whole-cell lysates were analyzed for luciferase activity 24 h later. THP-1 cells were infected with stationary promastigotes forms of L. amazonensis and/or treated with Akt-inhibitor-VIII for 4 h and then submitted for chromatin immunoprecipitation assay (ChIP) using Nrf2 ChIP-antibody and primers for Nrf2 (G) and Sod1 (I) promoters. (H) Western-blot for total protein extract analyses with Sod1 antibody was performed at same conditions of infection and treatment. Results are representative of three independent experiments. *p < 0.05.

Nrf2 Knockdown Promotes Oxidative Stress and Impairs Parasite Survival in Macrophages

We aimed to test the hypothesis that Nrf2 knockdown would favor oxidative stress, leading to the reduction of the parasite load in macrophages. We measured the production of OONO, NO, and ROS as components of the oxidative stress pathway in Nrf2-knockdown infected macrophages (Figure 4A). As expected, the production of ROS and the formation of OONO and NO were enhanced in infected Nrf2-knockdown macrophages. Figure 4B shows that PKR or Akt1 inhibition leads to a similar oxidative stress profile upon infection. Silencing of Nrf2 decreased the infection index, whereas the parasite load was rescued when infected Nrf2-knockdown macrophages were treated with the antioxidant NAC compound (Figure 4C). Notably, the Nrf2 inducer sulforaphane augmented the infection index.

FIGURE 4 | The reactive oxygen species (ROS) were enhanced upon Leishmania amazonensis infection in nuclear factor erythroid 2-related factor 2 (NRF2)/protein kinase R (PKR)/Akt-deficient macrophages. (A) shNrf2 or shControl and (B) wild-type THP-1 cells treated with PKR-inhibitor or Akt-inhibitor-VIII were infected with stationary promastigotes forms of L. amazonensis at indicated times together with probes for quantifying peroxynitrite (OONO), nitric oxide (NO), and ROS, and then analyzed as described in material and methods. (C) THP-1 transiently knocked-down for Nrf2 expression or shControl cells were infected with stationary promastigotes forms of L. amazonensis for 24 h and treated for additional 24 h with sulforaphane (SFN) or NAC (N-acetylcysteine) before the analysis of infection index. The asterisk means the statistic significant differences between the groups. Results are representative of three independent experiments. *p < 0.05.

Leishmania Down-Regulates the Nrf2 Negative Regulator Keap1 and Induces Autophagy

Nuclear factor erythroid 2-related factor 2 is sequestered in the cytosol by a homodimer of Keap1, which limits its nuclear translocation. Nrf2 associated with Keap1 is directed to proteasomal degradation by Cul3-mediated poly-ubiquitination (37). However, Keap1 is degraded through p62-mediated autophagy, releasing Nrf2 into the nucleus (38). Given that Leishmania induces autophagy in infected macrophages (39), we sought to investigate the levels of Keap1 in Leishmania infection. Figure 5A shows the prompt decrease in Keap1 levels in infected macrophages and demonstrates that PKR inactivation prevented Keap1 degradation. Moreover, Keap1 reduction was prevented by chloroquine, an autophagy inhibitor (Figure 5B). Given that Nrf2 released via Keap1 degradation promotes the antioxidant response, we addressed whether the inhibition of autophagy would increase the oxidative stress of infected macrophages. As observed in Figure 5C, the levels of ROS, OONO, and NO increased in infected cells treated with chloroquine. We also confirmed that L. amazonensis triggers LC3-I conversion to LC3-II, a marker of autophagy (Figure 5D). Given that the formation of the LC3-p62-Keap1 ternary complex on the autophagosome membrane directs Keap1 to degradation (40), we investigated the induction of p62 in the infection. We show that p62 was induced in infected macrophages, and this effect relied on PKR and Akt1 (Figure 5E and Figures S5A in Supplementary Material, respectively). Moreover, the ablation of Nrf2 expression prevented p62 induction due to infection (Figure 5F). Given that our data indicate that PKR and Akt control the induction of Nrf2, we tested the occupancy of the p62 promoter by Nrf2 in the context of infection. Our data show that Leishmania promoted Nrf2 occupancy, and the inhibition of either PKR or Akt signaling prevented this effect (Figure 5G and Figure S5B in Supplementary Material, respectively).

FIGURE 5 | The nuclear factor erythroid 2-related factor 2 (Nrf2)-inhibitor Kelch-like ECH-associated protein 1 (Keap1) is modulated negatively through protein kinase R (PKR) signaling and p62 autophagy-dependent manner in Leishmania infection. (A) RAW-WT-PKR and RAW-DN-PKR cells were infected with stationary promastigotes forms of Leishmania amazonensis for 2 or 4 h and then western-blot assay were performed with total protein extract using Keap1 antibody. (B) THP-1 cells were infected with stationary promastigotes forms of L. amazonensis at indicate times and/or treated with chloroquine and the total or nuclear protein extracts were analyzed using Keap1 and Nrf2 antibodies. (C) THP-1 cells treated with chloroquine were infected with stationary promastigotes forms of L. amazonensis at indicated times together with probes for quantifying OONO, NO, and ROS. (D) THP-1 cells were infected with stationary promastigotes forms of L. amazonensis and western-blot for LC3-I/II protein was performed. RAW-WT-PKR and RAW-DN-PKR cells (E), and shNrf2 or shControl THP-1 cells (F) were infected with stationary promastigotes forms of L. amazonensis and then the total protein extract was analyzed by western-blot assay with p62 antibody. (G) RAW-WT-PKR and RAW-DN-PKR were infected with stationary promastigotes forms of L. amazonensis for 4 h and then submitted for ChIP assay using Nrf2 ChIP-antibody and primers for p62 promoter. Results are representative of three independent experiments. *p < 0.05.

Nrf2 Protein Levels Are Elevated in Human CL, and L. braziliensis also Induces Nrf2 In Vitro

We aimed to address whether other L. amazonensis strains isolated from patients with localized cutaneous lesions (LCL) or DCL would induce Nrf2 nuclear translocation and the activation of PKR. Figure 6A shows that all distinct strains ofL. amazonensis activated PKR and Nrf2. Given that most of the cases of human CL in Brazil are caused by L. braziliensis, we decided to address whether this species would induce PKR and Nrf2. Figure 6B shows that L. braziliensis activated PKR and Nrf2. Nrf2 activation depended on PKR function (Figure 6C). Moreover, the expression of the Nrf2 target genes p62 and Sod1 was reduced in Nrf2-silenced L. braziliensis-infected macrophages (Figure 6E), and the growth of amastigotes was impaired in Nrf2-knockdown macrophages (Figure 6D). These results prompted us to investigate the levels of Nrf2 and the negative regulator Keap1 in clinical samples from LCL or DCL patients. Figure 6F shows the marked expression of Nrf2 in DCL samples compared to LCL tissues. Accordingly, Keap1 expression was enriched in LCL samples. Altogether, the data show that Nrf2 induction is triggered by distinct species and strains of L. amazonensis, and high levels of Nrf2 are found in patients with DCL, a severe clinical condition that presents with a high number of parasites and poor prognosis (41).

FIGURE 6 | Leishmania braziliensis and different strains of Leishmania amazonensis infections also modulated positively the protein kinase R (PKR)/nuclear factor erythroid 2-related factor 2 (Nrf2) axis pathway. (A) L. amazonensis strains from LCL or diffuse cutaneous leishmaniasis (DCL) patients were used to infect THP-1 cells. Total or nuclear protein extracts were processed and then analyzed by western-blot with phospho-PKR and Nrf2 antibodies, respectively. (B) THP-1 cells were infected with stationary promastigotes forms of L. braziliensis at indicate times and then performed for western-blot with phospho-PKR and Nrf2 antibodies, and 3xARE-promoter Luciferase assays. Peritoneal macrophages of wild-type or PKR-ko mice were infected with L. braziliensis and infection index assays were then analyzed. (C) RAW-WT-PKR and RAW-DN-PKR cells were infected with L. braziliensis and western-blot with anti-Nrf2 was then analyzed. (D) shNrf2 or shControl THP-1 cells were infected with stationary promastigotes forms of L. amazonensis for 24 h before treatment with NAC or polyI:C for additional 24 h. After this time, the cells were fixed and the infection index was evaluated. (E) THP-1 transiently knocked-down for Nrf2 expression or shControl cells were infected with stationary promastigotes forms of L. amazonensis for 4 or 6 h before total protein extract for western-blot analyzes with p62 and Sod1 antibodies. (F) Histological sections from biopsies obtained from lesions of patients with DCL (n = 4) or with LCL (n = 5) were submitted to immunohistochemical reaction with primary antibodies against Nrf2 or Keap1 as previously described. All sections were counterstained with hematoxylin. Digital images (400× magnification) were captured using a Nikon E600 microscope and an Olympus Q-Color 1 digital camera with the Image Pro Plus program. Bars represent 10 µm. Positive cell density was obtained. Graph represents the analysis of reactive positive cells for Nrf2 and Keap1 compared with isotype controls as percentage of positive stained area per total tissue area. Results are representative of three independent experiments. *p < 0.05.

Transcriptomic Analysis Reveals a Pivotal Role of Nrf2 Signaling in CL Patient Samples

Next, we tested for transcription factor enrichment among the 413 genes composing the systemic LCL disease signature. Only five transcription factor motifs were significantly enriched among the promoters of the 413 genes of the LCL disease signature. After the E4F1 motif, the Nrf2 binding site was the second-most significantly represented, being present in 15 of the 413 genes composing the LCL disease signature (Table 1). Among those, p62 (Sqstm1), in bold, was confirmed, in agreement with our in vitro data. We herein present the first disease signature of LCL using a systems biology analysis of the PBMC transcriptome of LCL patients (n = 18) vs. healthy controls (n = 12). Using Affymetrix microarrays (HuGene 1.0), we found that Nrf2 was significantly overexpressed in patient PBMCs vs. controls (1.8-fold, uncorrected p = 0.0002, p = 0.033 using the Benjamini–Hochberg correction for genome-wide testing). The top 50 upregulated genes in patients vs. controls are shown in Table S1 in Supplementary Material. Next, we used IPA to determine which biological pathways and molecular networks were enriched among the LCL disease signature. As shown in Table S2 in Supplementary Material, three antioxidant pathways, i.e., the thioredoxin pathway, the antioxidant action of Vitamin C and the Nrf2 pathway, were significantly enriched in the LCL disease signature.

TABLE 1 | Gene promoters in LCL disease signature are enriched for nuclear factor erythroid 2-related factor 2 transcription factor binding sites.

Nrf2 Transcriptome-Wide Correlations Confirm the Links between IFN-I/PKR, ARE, PIK3, and Autophagy Signaling Pathways In Situ

Then, we performed a transcriptome-wide correlation analysis to further investigate whether the molecular links we described at the protein level in vitro might be confirmed at the transcriptional level ex vivo. The expression of a large number of genes was significantly correlated to Nrf2 transcript levels, even following stringent Benjamini–Hochberg correction for multiple testing. Among those, PKR, PIK3C, Sod1, and p62 (SQSTM1) transcripts were positively correlated, whereas Keap1 was negatively correlated, to Nrf2 transcript levels, with minor differences between LCL patients and controls (Figure 7A), thus confirming our protein data of Nrf2 regulation, both upstream or downstream. To validate these microarray results, we performed a targeted analysis of key genes in the Nrf2/PKR crosstalk using nCounter digital transcriptomic quantification in LCL (n = 6) as well as healthy skin biopsies (n = 4). As shown in Figure 7B, unsupervised hierarchical clustering of in situ transcriptomes revealed two major clusters, which coincided with either LCL patients or normal donors (ND). Thus, LCL skin biopsies could be discriminated from healthy skin by differential expression of only six transcripts (Nrf2, PKR, Sod1, Sod2, Keap1, and Hmox1).

FIGURE 7 | Nuclear factor erythroid 2-related factor 2 (Nrf2) transcriptome-wide correlations confirm the links between IFN/protein kinase R (PKR), antioxidant response element (ARE), PIK3, and autophagy signaling pathways in situ. (A) Transcriptome-wide correlation analysis was applied to Nrf2 transcript levels in microarray data (Illumina) obtained from skin biopsies from LCL patients (n = 20, red) and healthy controls (n = 10, green). Spearman correlation coefficient and p-values are shown for individual transcripts, 95% confidence intervals are shown in gray for patients only. (B) Validation of increased PKR/Nrf2 signaling pathway members by digital mRNA quantification. Heat map and hierarchical cluster analysis (Eucledian distance) of selected genes quantified by nCounter digital transcriptomics (Nanostring) in skin biopsies from normal donors (ND, n = 4) and LCL patients (n = 6), normalized according to PTPRC (CD45) expression levels, to account for differences in tissue leukocyte infiltration.

DISCUSSION

The oxidative burst in infected cells is a key microbicide mechanism exhibited by macrophages. However, Leishmania parasites present a repertoire of adaptive mechanisms to cope with the altered redox state of infected macrophages by expressing antioxidant enzymes or interfering with macrophage signaling pathways (42). A growing number of reports indicate that PKR modulates infections caused by intracellular pathogens (43). Notably, increased levels of Sod1 are expressed in macrophages infected by L. amazonensis due to PKR activation (35). Given that the transcription factor Nrf2 is the main regulator of Sod1 expression (44) among other genes involved in the anti-oxidative response, we studied the regulation of Nrf2 in the context of Leishmania infection and tested the hypothesis that PKR is actually a positive regulator of the ARE via Nrf2.

Our results demonstrated that Nrf2 activation depended on PKR signaling. Simple treatment with PKR inducers such as IFN-α and PolyI:C increased the expression and nuclear translocation of Nrf2, demonstrating that the mechanisms of Nrf2 activation through PKR pathway signaling are not exclusively due to Leishmania infection. Remarkably, PKR activation induces a significant increase in Nrf2 expression. PKR phosphorylates eIF2-α, which reduces protein synthesis while upregulating the expression of some genes such as ATF-4. The 5′untranslated region of Nrf2 mRNA presents an internal ribosomal entry site (IRES), allowing enhanced Nrf2 translation in eIF2-α-mediated protein translation (45, 46).

The control of Nrf2 activation requires different post-translational modifications as well as its repression and subsequent degradation via the proteasome (8–11, 47). The non-canonical PI3K/Akt signaling pathway has been linked to the activation of Nrf2 in a number of models. GSK3-mediated inhibitory phosphorylation induces Nrf2 by inhibiting the phosphorylation signal and sequential ubiquitination in the Neh6-Nrf2 domain, allowing its stability and activity (13). In Leishmania infection, PI3K and Akt inhibition reduced the expression of Nrf2 and Sod1 (Figure 3). Our results led us to conclude that PI3K/Akt activation as a result of Leishmania infection is a positive Nrf2 regulator in host cells.

The rise of ROS seems to be a key regulator of infection by intracellular pathogens (48), and the co-evolution of host cells and parasites results in a shared pattern of subversion in the production of these radicals. For example, Trypanosoma cruzi infection in THP-1 cells requires a level of oxidative stress for successful parasitism, given that the overexpression of Nrf2 reduces parasitism (49). Our data from in vitro Nrf2-knockdown macrophages revealed the spontaneous increase of oxidative stress, measured through the levels of ROS, NO, and OONO. The same change in phenotype occurred when PKR and Akt were inhibited, probably due to the reduction of Sod1 and other targets. However, the infection index increased when the cells were treated with SFN and NAC. Our data suggest that Nrf2 activation induces Sod1, thus counteracting the oxidative boost in the cell milieu in infected macrophages.

Several reports (50) have highlighted the close relationship between oxidative stress and the autophagy process. The autophagy pathway plays an important role in resistance to various infections, although it could be subverted, thus favoring some infections (51). It is conceivable that autophagy induced by L. amazonensis may be controlled by PKR, as revealed in other models, thus regulating Nrf2 levels. Accordingly, some studies have shown the importance of oxidative stress sensing in autophagy (15, 16) and have demonstrated that the degradation of Keap1 via autophagy allows cellular redox homeostasis in liver cells. Our data showed that infected macrophages display an increase in LC3-I to LC3-II conversion, thus corroborating the importance of autophagy through this marker during infection.

Kelch-like ECH-associated protein 1 is a negative regulator of Nrf2, and in the context of infection by Leishmania, we demonstrated that Keap1 is regulated after 18 h of infection in a PKR-independent manner (data not included). However, Keap1 stability is decreased in a PKR-dependent manner between 2 and 4 h of infection. When autophagy was inhibited by chloroquine, we noted a cytoplasmic accumulation of Nrf2 and stabilization of Keap1 levels, which was accompanied by high levels of oxidative stress.

Several studies have shown the involvement of the p62 (Sqstm1) protein as a central regulator between Keap1 and Nrf2. Oxidative stress decreases when cells overexpressing p62 bind to this inhibitory protein, leading to autophagosome formation (19, 20). Other studies have shown that Nrf2 positively regulates the expression of p62 (17). Importantly, TLR2 activation culminates in M2 polarization of macrophages (MOX macrophages), which leads to NF-κB-p65 degradation through p62 and lysosomes, characterized by selective autophagy (52). M2 macrophages exhibit antioxidant properties, as judged by the expression of Cox2, IL1β, HO-1, VEGF, and Nrf2 (53). Considering the cascade of signals, our data support the notion that the PKR–PI3K/Akt↔Nrf2 axis regulates p62 gene expression in Leishmania infection and passively triggers the autophagy pathway that culminates in Keap1 degradation, activating Nrf2 and resulting in oxidative cellular homeostasis.

Patients with localized cutaneous lesion (LCL) exhibit predominant expression of iNOS, IL-1β, IL-6, MCP-1, TNF-α, and IFN-γ, while anergic diffuse cutaneous leishmaniasis (ADCL) lesions are characterized by the presence of IL-4, IL-5, IL-10, and MIP-1α and the low expression of iNOS (54, 55). Our in vitro data showed that L. braziliensis, the prominent causative agent of LCL, also induces Nrf2 in a PKR-dependent fashion. This observation underlines the importance of this signaling pathway in other Leishmania species besides L. amazonensis. However, the immunohistochemistry analysis of LCL vs. DCL lesions revealed a strong Nrf2 reaction in the latter group, while the Keap1 signal was predominant in the former clinical samples. These results indicate that Nrf2 activation may contribute to the poor oxidative response and, consequently, the high parasite burden in DCL patients.

The data obtained in this study confirm and extend our previous finding of an IFN-I/Sod1 axis, linked to increased parasite burden (56) and therapeutic failure in both localized cutaneous leishmaniasis and DCL (31). This study now reveals that this IFN-I/Sod1 link is critically mediated by Nrf2/ARE signaling. Our genome-wide study revealed Nrf2 as a master regulator of the in situ (skin biopsies) transcriptome (Figure 7), both in health and disease, which is in agreement with its central role in proteostasis and ancient molecular networks, conserved in evolution from Drosophila to man (57). There is a negative correlation with the Keap1 transcript skin biopsies, which was corroborated by our findings at the protein level in both LCL and DCL skin biopsies (Figure 6F). These results point to a possible compartmentalization of the pathogen-driven immune response between tissues in CL, where cutaneous ulcers in LCL are exposed to a complex microbiome, which strongly influences the local immune response, in addition to Leishmania antigens (58).

Due to its strong pleiotropic effects and its essential function in normal homeostasis, Nrf2 itself is not yet a target of choice for therapeutic intervention in LCL. However, this study reiterates our previous suggestion that downstream targets of Nrf2, such as Sod1, represent excellent therapeutic targets in LCL. Previous works from our and other groups (59–62) have shown that the Sod1 inhibitor DETC or its precursor molecule, disulfiram, are plausible therapeutic alternatives that have been used extensively in humans for decades with an excellent safety profile. In conclusion, we demonstrate for the first time the prominent role of Nrf2 and the PKR↔PI3K/Akt↔p62/autophagy axis in human and experimental leishmaniasis (Figure 7B). Collectively, our data propose a signaling-based scenario that may reveal a pivotal molecular basis for CL pathogenesis as well as its therapeutic potential. A schematic model based in our results is depicted in Figure 8.

FIGURE 8 | Proposed model for the protein kinase R (PKR)-dependent nuclear factor erythroid 2-related factor 2 (Nrf2) activation in Leishmania infection. Internalized parasite signals through the endosomal compartment via TLR2 and induce activation of PKR by dimerization and subsequent autophosphorylation. Subsequently, we found that GSK3 phosphorylation is dependent of PKR signaling, allowing that not occur inhibition of Nrf2 through Neh6 inhibitory domain. This activation of Nrf2 is also dependent of Keap1 inhibition through of autophagic and PKR pathways. These mechanisms induce nuclear translocation Nrf2, increasing the gene expression of Sod1, Nrf2, and p62. The sequestosome-1 (p62) could be recruiting, together with processed LC3-II and Keap1 for autophagic vacuoles, allowing greater Nrf2 activation and inhibition of oxidative stress through antioxidant enzymes.

ETHICS STATEMENT

Written informed consent was obtained from all participants or legal guardians, and all of the data analyzed were anonymized. The project was approved by the Institutional Review Board of Centro de Pesquisas Gonçalo Moniz, FIOCRUZ–BA (license number 136/2007) and complies with the guidelines of the Declaration of Helsinki.

AUTHOR CONTRIBUTIONS

AV—designed and performed experiments, analyzed data, and wrote the manuscript; TC-S—performed initial experiments for the study; AS—provided reagents, supervised experiments, and critically evaluated the manuscript; VB, JF-C, RK, AB, TD, JW, and VB—carried out experiments with patient samples, analyzed the results, and provided input for experimental design and interpretation; NF—critically reviewed the manuscript and analyzed data, and UL—directed the study, analyzed the data, and wrote the manuscript.

ACKNOWLEDGMENTS

In memoriam to late Prof. Mario Alberto Cardoso da Silva Neto, IBqM—UFRJ, our forever most sincere acknowledgments for his comments and suggestions to this work.

FUNDING

This work was supported by the Brazilian research funding agencies Fundação Carlos Chagas Filho de Amparo a Pesquisa do Estado do Rio de Janeiro (FAPERJ) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) and Belgian research funding agency FWO (grant G0D6817N) and by the FNRS (N.310030-173180) to NF and by the FNRS International cooperation (No IZRJZ3_164176/1) to NF and UGL.

SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at http://journal.frontiersin.org/article/10.3389/fimmu.2017.01127/full#supplementary-material.

REFERENCES

1. Alvar J, Vélez ID, Bern C, Herrero M, Desjeux P, Cano J, et al. Leishmaniasis worldwide and global estimates of its incidence. PLoS One (2012) 7(5):e35671. doi:10.1371/journal.pone.0035671

2. Cecílio P, Pérez-Cabezas B, Santarém N, Maciel J, Rodrigues V, Cordeiro da Silva A. Deception and manipulation: the arms of Leishmania, a successful parasite. Front Immunol (2014) 5:480. doi:10.3389/fimmu.2014.00480

3. Kaye P, Scott P. Leishmaniasis: complexity at the host-pathogen interface. Nat Rev Microbiol (2011) 9(8):604–15. doi:10.1038/nrmicro2608

4. Olive AJ, Sassetti CM. Metabolic crosstalk between host and pathogen: sensing, adapting and competing. Nat Rev Microbiol (2016) 14(4):221–34. doi:10.1038/nrmicro.2016.1

5. Wasserman WW, Fahl WE. Functional antioxidant responsive elements. Proc Natl Acad Sci U S A (1997) 94(10):5361–6. doi:10.1073/pnas.94.10.5361

6. Lee IT, Wang SW, Lee CW, Chang CC, Lin CC, Luo SF, et al. Lipoteichoic acid induces HO-1 expression via the TLR2/MyD88/c-Src/NADPH oxidase pathway and Nrf2 in human tracheal smooth muscle cells. J Immunol (2008) 181(7):5098–110. doi:10.4049/jimmunol.181.7.5098

7. Vijayan V, Baumgart-Vogt E, Naidu S, Qian G, Immenschuh S. Bruton’s tyrosine kinase is required for TLR-dependent heme oxygenase-1 gene activation via Nrf2 in macrophages. J Immunol (2011) 187(2):817–27. doi:10.4049/jimmunol.1003631

8. Cullinan SB, Zhang D, Hannink M, Arvisais E, Kaufman RJ, Diehl JA. Nrf2 is a direct PERK substrate and effector of PERK-dependent cell survival. Mol Cell Biol (2003) 23(20):7198–209. doi:10.1128/MCB.23.20.7198-7209.2003

9. Niture SK, Jain AK, Jaiswal AK. Antioxidant-induced modification of INrf2 cysteine 151 and PKC-delta-mediated phosphorylation of Nrf2 serine 40 are both required for stabilization and nuclear translocation of Nrf2 and increased drug resistance. J Cell Sci (2009) 122(Pt 24):4452–64. doi:10.1242/jcs.058537

10. Chowdhry S, Zhang Y, McMahon M, Sutherland C, Cuadrado A, Hayes JD. Nrf2 is controlled by two distinct β-TrCP recognition motifs in its Neh6 domain, one of which can be modulated by GSK-3 activity. Oncogene (2003) 32(32):3765–81. doi:10.1038/onc.2012.388

11. Rada P, Rojo AI, Evrard-Todeschi N, Innamorato NG, Cotte A, Jaworski T, et al. Structural and functional characterization of Nrf2 degradation by the glycogen synthase kinase 3/β-TrCP axis. Mol Cell Biol (2012) 32(17):3486–99. doi:10.1128/MCB.00180-12

12. Ruhland A, Leal N, Kima PE. Leishmania promastigotes activate PI3K/Akt signalling to confer host cell resistance to apoptosis. Cell Microbiol (2007) 9(1):84–96. doi:10.1111/j.1462-5822.2006.00769.x

13. Chen HH, Chen YT, Huang YW, Tsai HJ, Kuo CC. 4-Ketopinoresinol, a novel naturally occurring ARE activator, induces the Nrf2/HO-1 axis and protects against oxidative stress-induced cell injury via activation of PI3K/AKT signaling. Free Radic Biol Med (2012) 52(6):1054–66. doi:10.1016/j.freeradbiomed.2011.12.012

14. Syklotis GP, Bohmann D. Stress-activated cap’n’collar transcription factors in aging and human disease. Sci Signal (2013) 3(112):re3. doi:10.1126/scisignal.3112re3

15. Taguchi K, Fujikawa N, Komatsu M, Ishii T, Unno M, Akaike T, et al. Keap1 degradation by autophagy for the maintenance of redox homeostasis. Proc Natl Acad Sci U S A (2012) 109(34):13561–6. doi:10.1073/pnas.1121572109

16. Filomeni G, De-Zio D, Cecconi F. Oxidative stress and autophagy: the clash between damage and metabolic needs. Cell Death Differ (2015) 22(3):377–88. doi:10.1038/cdd.2014.150

17. Jain A, Lamark T, Sjøttem E, Larsen KB, Awuh JA, Øvervatn A, et al. p62/SQSTM1 is a target gene for transcription factor NRF2 and creates a positive feedback loop by inducing antioxidant response element-driven gene transcription. J Biol Chem (2010) 285(29):22576–91. doi:10.1074/jbc.M110.118976

18. Geetha T, Wooten MW. Structure and functional properties of the ubiquitin binding protein p62. FEBS Lett (2012) 512(1–3):19–24. doi:10.1016/S0014-5793(02)02286-X

19. Ichimura Y, Waguri S, Sou YS, Kageyama S, Hasegawa J, Ishimura R, et al. Phosphorylation of p62 activates the Keap1-Nrf2 pathway during selective autophagy. Mol Cell (2013) 51(5):618–31. doi:10.1016/j.molcel.2013.08.003

20. Yin S, Cao W. Toll-like receptor signaling induces Nrf2 pathway activation through p62-triggered Keap1 degradation. Mol Cell Biol (2015) 35(15):2673–83. doi:10.1128/MCB.00105-15

21. Nanduri S, Carpick BW, Yang Y, Williams BR, Qin J. Structure of the double-stranded RNA-binding domain of the protein kinase PKR reveals the molecular basis of its dsRNA-mediated activation. EMBO J (1998) 17(18):5458–65. doi:10.1093/emboj/17.18.5458

22. Meurs E, Chong K, Galabru J, Thomas NS, Kerr IM, Williams BR, et al. Molecular cloning and characterization of the human double-stranded RNA-activated protein kinase induced by interferon. Cell (1990) 62(2):379–90. doi:10.1016/0092-8674(90)90374-N

23. Taylor DR, Lee SB, Romano PR, Marshak DR, Hinnebusch AG, Esteban M, et al. Autophosphorylation sites participate in the activation of the double-stranded-RNA-activated protein kinase PKR. Mol Cell Biol (1996) 16:6295–302. doi:10.1128/MCB.16.11.6295

24. Thomis DC, Samuel CE. Mechanism of interferon action: evidence for intermolecular autophosphorylation and autoactivation of the interferon-induced, RNA-dependent protein kinase PKR. J Virol (1993) 67(12):7695–700.

25. Dey M, Mann BR, Anshu A, Mannan MA. Activation of protein kinase PKR requires dimerization-induced cis-phosphorylation within the activation loop. J Biol Chem (2014) 289(9):5747–57. doi:10.1074/jbc.M113.527796

26. Tallóczy Z, Jiang W, Virgin HW IV, Leib DA, Scheuner D, Kaufman RJ, et al. Regulation of starvation- and virus-induced autophagy by the eIF2alpha kinase signaling pathway. Proc Natl Acad Sci U S A (2002) 99(1):190–5. doi:10.1073/pnas.012485299

27. Tallóczy Z, Virgin HW IV, Levine B. PKR-dependent autophagic degradation of herpes simplex virus type 1. Autophagy (2006) 2(1):24–9. doi:10.4161/auto.2176

28. Shen S, Niso-Santano M, Adjemian S, Takehara T, Malik SA, Minoux H, et al. Cytoplasmic STAT3 represses autophagy by inhibiting PKR activity. Mol Cell (2012) 48(5):667–80. doi:10.1016/j.molcel.2012.09.013

29. Ambjorn M, Ejlerskov P, Liu Y, Lees M, Jäättelä M, Issazadeh-Navikas S. IFNB1/interferon-β-induced autophagy in MCF-7 breast cancer cells counteracts its proapoptotic function. Autophagy (2013) 9(3):287–302. doi:10.4161/auto.22831

30. Convit J, Pinardi ME, Rondon AJ. Diffuse cutaneous leishmaniasis: a disease due to an immunological defect of the host. Trans R Soc Trop Med Hyg (1972) 66(4):603–10. doi:10.1016/0035-9203(72)90306-9

31. Khouri R, Santos GS, Soares G, Costa JM, Barral A, Barral-Netto M, et al. SOD1 plasma level as a biomarker for therapeutic failure in cutaneous leishmaniasis. J Infect Dis (2014) 210(2):306–10. doi:10.1093/infdis/jiu087

32. Deschacht M, Van Assche T, Hendrickx S, Bult H, Maes L, Cos P. Role of oxidative stress and apoptosis in the cellular response of murine macrophages upon Leishmania infection. Parasitology (2012) 139(11):1429–37. doi:10.1017/S003118201200073X

33. Soong L. Subversion and utilization of host innate defense by Leishmania amazonensis. Front Immunol (2012) 3:58. doi:10.3389/fimmu.2012.00058

34. Almeida TF, Palma LC, Mendez LC, Noronha-Dutra AA, Veras PS. Leishmania amazonensis fails to induce the release of reactive oxygen intermediates by CBA macrophages. Parasite Immunol (2012) 34(10):492–8. doi:10.1111/j.1365-3024.2012.01384.x

35. Pereira RMS, Teixeira KL, Barreto-de-Souza V, Calegari-Silva TC, De-Melo LD, Soares DC, et al. Novel role for the double-stranded RNA-activated protein kinase PKR: modulation of macrophage infection by the protozoan parasite Leishmania. FASEB J (2010) 24(2):617–26. doi:10.1096/fj.09-140053

36. Vivarini AC, Pereira RMS, Teixeira KL, Calegari-Silva TC, Bellio M, Laurenti MD, et al. Human cutaneous leishmaniasis: interferon-dependent expression of double-stranded RNA-dependent protein kinase (PKR) via TLR2. FASEB J (2011) 25(12):4162–73. doi:10.1096/fj.11-185165

37. Zhang M, An C, Gao Y, Leak RK, Chen J, Zhang F. Emerging roles of Nrf2 and phase II antioxidant enzymes in neuroprotection. Prog Neurobiol (2013) 100:30–47. doi:10.1016/j.pneurobio.2012.09.003

38. Ishimura R, Tanaka K, Komatsu M. Dissection of the role of p62/Sqstm1 in activation of Nrf2 during xenophagy. FEBS Lett (2014) 588(5):822–8. doi:10.1016/j.febslet.2014.01.045

39. Pinheiro RO, Nunes MP, Pinheiro CS, D’Avila H, Bozza PT, Takiya CM, et al. A. Induction of autophagy correlates with increased parasite load of Leishmania amazonensis in BALB/c but not C57BL/6 macrophages. Microbes Infect (2009) 11(2):181–90. doi:10.1016/j.micinf.2008.11.006

40. Komatsu M, Kurokawa H, Waguri S, Taguchi K, Kobayashi A, Ichimura Y, et al. The selective autophagy substrate p62 activates the stress responsive transcription factor Nrf2 through inactivation of Keap1. Nat Cell Biol (2010) 12(3):213–23. doi:10.1038/ncb2021

41. França-costa J, Wanderley JL, Deolindo P, Zarattini JB, Costa J, Soong L, et al. Exposure of phosphatidylserine on Leishmania amazonensis isolates is associated with diffuse cutaneous leishmaniasis and parasite infectivity. PLoS One (2012) 7(5):e36595. doi:10.1371/journal.pone.0036595

42. Henard CA, Carlsen ED, Hay C, Kima PE, Soong L. Leishmania amazonensis amastigotes highly express a tryparedoxin peroxidase isoform that increases parasite resistance to macrophage antimicrobial defenses and fosters parasite virulence. PLoS Negl Trop Dis (2014) 8(7):e3000. doi:10.1371/journal.pntd.000300

43. Ogolla PS, Portillo JA, White CL, Patel K, Lamb B, Sen GC, et al. The protein kinase double-stranded RNA-dependent (PKR) enhances protection against disease caused by a non-viral pathogen. PLoS Pathog (2013) 9(8):e1003557. doi:10.1371/journal.ppat.1003557

44. Milani P, Gagliardi S, Cova E, Cereda C. SOD1 transcriptional and posttranscriptional regulation and its potential implications in ALS. Neurol Res Int (2011) 2011:458427. doi:10.1155/2011/458427

45. Li W, Thakor N, Xu EY, Huang Y, Chen C, Yu R, et al. An internal ribosomal entry site mediates redox-sensitive translation of Nrf2. Nucleic Acids Res (2010) 38(3):778–88. doi:10.1093/nar/gkp1048

46. Purdom-Dickinson SE, Sheveleva EV, Sun H, Chen QM. Translational control of nrf2 protein in activation of antioxidant response by oxidants. Mol Pharmacol (2007) 72(4):1074–81. doi:10.1124/mol.107.035360

47. Keum YS. Regulation of the Keap1/Nrf2 system by chemopreventive sulforaphane: implications of posttranslational modifications. Ann N Y Acad Sci (2011) 1229:184–9. doi:10.1111/j.1749-6632.2011.06092.x

48. Fang FC. Antimicrobial reactive oxygen and nitrogen species: concepts and controversies. Nat Rev Microbiol (2004) 2(10):820–32. doi:10.1038/nrmicro1004

49. Paiva CN, Feijó DF, Dutra FF, Carneiro VC, Freitas GB, Alves LS, et al. Oxidative stress fuels Trypanosoma cruzi infection in mice. J Clin Invest (2012) 122(7):2531–42. doi:10.1172/JCI58525

50. Lee J, Giordano S, Zhang J. Autophagy, mitochondria and oxidative stress: cross-talk and redox signalling. Biochem J (2012) 441(2):523–40. doi:10.1172/JCI58525

51. Saitoh T, Akira S. Regulation of innate immune responses by autophagy-related proteins. J Cell Biol (2010) 189(6):925–35. doi:10.1083/jcb.201002021

52. Chang CP, Su YC, Hu C, Lei HY. TLR2-dependent selective autophagy regulates NF-κB lysosomal degradation in hepatoma-derived M2 macrophage differentiation. Cell Death Differ (2013) 20(3):515–23. doi:10.1038/cdd.2012.146

53. Shalhoub J, Falck-Hansen MA, Davies AH, Monaco C. Innate immunity and monocyte-macrophage activation in atherosclerosis. J Inflamm (Lond) (2011) 8:9. doi:10.1186/1476-9255-8-9

54. Cáceres-Dittmar G, Tapia FJ, Sánchez MA, Yamamura M, Uyemura K, Modlin RL, et al. Determination of the cytokine profile in American cutaneous leishmaniasis using the polymerase chain reaction. Clin Exp Immunol (1993) 91(3):500–5. doi:10.1111/j.1365-2249.1993.tb05931.x

55. Qadoumi M, Becker I, Donhauser N, Röllinghoff M, Bogdan C. Expression of inducible nitric oxide synthase in skin lesions of patients with American cutaneous leishmaniasis. Infect Immun (2002) 70(8):4638–42. doi:10.1128/IAI.70.8.4638-4642.2002

56. Khouri R, Bafica A, Silva Mda P, Noronha A, Kolb JP, Wietzerbin J, et al. IFN-β impairs superoxide-dependent parasite killing in human macrophages: evidence for a deleterious role of SOD1 in cutaneous leismaniasis. J Immunol (2009) 182(4):2525–31. doi:10.4049/jimmunol.0802860

57. Lacher SE, Lee JS, Wang X, Campbell MR, Bell DA, Slattery M. Beyond antioxidant genes in the ancient Nrf2 regulatory network. Free Radic Biol Med (2015) 88(Pt B):452–65. doi:10.1016/j.freeradbiomed.2015.06.044

58. Naik S, Bouladoux N, Wilhelm C, Molloy MJ, Salcedo R, Kastenmuller W, et al. Compartmentalized control of skin immunity by resident commensals. Science (2012) 337(6098):1115–9. doi:10.1126/science.1225152

59. Khouri R, Novais F, Santana G, de Oliveira CI, Vannier dos Santos MA, Barral A, et al. DETC induces Leishmania parasite killing in human in vitro and murine in vivo models: a promising therapeutic alternative in leishmaniasis. PLoS One (2010) 5(12):e14394. doi:10.1371/journal.pone.0014394

60. Chavali AK, Blazier AS, Tlaxca JL, Jensen PA, Pearson RD, Papin JA. Metabolic network analysis predicts efficacy of FDA-approved drugs targeting the causative agent of a neglected tropical disease. BMC Syst Biol (2012) 6:27. doi:10.1186/1752-0509-6-27

61. Peniche AG, Renslo AR, Melby PC, Travi BL. Antileishmanial activity of disulfiram and thiuram disulfide analogs in an ex vivo model system is selectively enhanced by the addition of divalent metal ions. Antimicrob Agents Chemother (2015) 59(10):6463–70. doi:10.1128/AAC.05131-14

62. Novais FO, Carvalho LP, Passos S, Roos DS, Carvalho EM, Scott P, et al. Genomic profiling of human Leishmania braziliensis lesions identifies transcriptional modules associated with cutaneous immunopathology. J Invest Dermatol (2015) 135(1):94–101. doi:10.1038/jid.2014.305

Conflict of Interest Statement: The authors declare that they have no conflicts of interest with the contents of this article. The handling editor declared a shared affiliation, though no other collaboration, with several of the authors, AV, TS, KD, and UL.

Copyright © 2017 Vivarini, Calegari-Silva, Saliba, Boaventura, França-Costa, Khouri, Dierckx, Dias-Teixeira, Fasel, Barral, Borges, Van Weyenbergh and Lopes. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.