[1st Edition] 9780128124727, 9780128124710

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages ix-x
Chapter One - Metabolic Alterations at the Crossroad of Aging and OncogenesisPages 1-42L. Raffaghello, V. Longo
Chapter Two - Cellular and Molecular Mechanisms of Autoimmunity and Lupus NephritisPages 43-154S.K. Devarapu, G. Lorenz, O.P. Kulkarni, H.-J. Anders, S.R. Mulay
Chapter Three - Old and Novel Functions of Caspase-2Pages 155-212M.A. Miles, T. Kitevska-Ilioski, C.J. Hawkins
Chapter Four - Metabolic Reprogramming and Oncogenesis: One Hallmark, Many OrganellesPages 213-231A.S.H. Costa, C. Frezza
Chapter Five - Molecular Biology Digest of Cell MitophagyPages 233-258I. Matic, D. Strobbe, F. Di Guglielmo, M. Campanella
Chapter Six - Regulation of Cell Calcium and Role of Plasma Membrane Calcium ATPasesPages 259-296T. Calì, M. Brini, E. Carafoli
Chapter Seven - Emerging Mechanisms and Roles for Asymmetric CytokinesisPages 297-345C. Thieleke-Matos, D.S. Osório, A.X. Carvalho, E. Morais-de-Sá

Citation preview

INTERNATIONAL REVIEW OF CELL AND MOLECULAR BIOLOGY Series Editors GEOFFREY H. BOURNE JAMES F. DANIELLI KWANG W. JEON MARTIN FRIEDLANDER JONATHAN JARVIK LORENZO GALLUZZI Editorial Advisory Board KEITH BURRIDGE AARON CIECHANOVER SANDRA DEMARIA SILVIA FINNEMANN KWANG JEON

1949–1988 1949–1984 1967–2016 1984–1992 1993–1995 2016–

CARLOS LOPEZ-OTIN WALLACE MARSHALL SHIGEKAZU NAGATA MOSHE OREN ANNE SIMONSEN

Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States 525 B Street, Suite 1800, San Diego, CA 92101-4495, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 125 London Wall, London, EC2Y 5AS, United Kingdom First edition 2017 Copyright © 2017 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-12-812471-0 ISSN: 1937-6448 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Zoe Kruze Acquisition Editor: Alex White Editorial Project Manager: Fenton Coulthurst Production Project Manager: Magesh Kumar Mahalingam Cover Designer: Vicky Pearson Esser Typeset by SPi Global, India

CONTRIBUTORS H.-J. Anders Medizinische Klinik und Poliklinik IV, Klinikum der Universit€at M€ unchen, Munich, Germany M. Brini University of Padova, Padova, Italy T. Calı` University of Padova, Padova, Italy M. Campanella University of Rome Tor Vergata; Regina Elena-National Cancer Institute, Rome, Italy; RVC, University of London; UCL Consortium for Mitochondrial Research, London, United Kingdom E. Carafoli Venetian Institute of Molecular Medicine, Padova, Italy A.X. Carvalho i3S—Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de; Cytoskeletal Dynamics, IBMC, Instituto de Biologia Molecular e Celular, and i3S, Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de, Universidade do Porto, Porto, Portugal A.S.H. Costa Medical Research Council Cancer Unit, University of Cambridge, Hutchison/MRC Research Centre, Cambridge, ENG, United Kingdom S.K. Devarapu Medizinische Klinik und Poliklinik IV, Klinikum der Universit€at M€ unchen, Munich, Germany F. Di Guglielmo University of Rome Tor Vergata, Rome, Italy C. Frezza Medical Research Council Cancer Unit, University of Cambridge, Hutchison/MRC Research Centre, Cambridge, ENG, United Kingdom C.J. Hawkins La Trobe Institute for Molecular Science, La Trobe University, Bundoora, VIC, Australia T. Kitevska-Ilioski La Trobe Institute for Molecular Science, La Trobe University, Bundoora, VIC, Australia O.P. Kulkarni BITS-Pilani Hyderabad Campus, Hyderabad, India

ix

x

Contributors

V. Longo Longevity Institute, Davis School of Gerontology, University of Southern California, Los Angeles, CA, United States; IFOM, FIRC Institute of Molecular Oncology, Milano, Italy G. Lorenz Klinikum rechts der Isar, Abteilung f€ ur Nephrologie, Technische Universit€at M€ unchen, Munich, Germany I. Matic University of Rome Tor Vergata, Rome, Italy M.A. Miles La Trobe Institute for Molecular Science, La Trobe University, Bundoora, VIC, Australia E. Morais-de-Sa´ i3S—Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de; Cell Division and Genomic stability, IBMC, Instituto de Biologia Molecular e Celular, and i3S, Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de, Universidade do Porto, Porto, Portugal S.R. Mulay Medizinische Klinik und Poliklinik IV, Klinikum der Universit€at M€ unchen, Munich, Germany D.S. Oso´rio i3S—Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de; Cytoskeletal Dynamics, IBMC, Instituto de Biologia Molecular e Celular, and i3S, Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de, Universidade do Porto, Porto, Portugal L. Raffaghello Laboratory of Oncology, Istituto Giannina Gaslini, Genova, Italy D. Strobbe Regina Elena-National Cancer Institute, Rome, Italy C. Thieleke-Matos i3S—Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de; Cell Division and Genomic stability, IBMC, Instituto de Biologia Molecular e Celular, and i3S, Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de, Universidade do Porto, Porto, Portugal

CHAPTER ONE

Metabolic Alterations at the Crossroad of Aging and Oncogenesis L. Raffaghello*, V. Longo†,{,1 *Laboratory of Oncology, Istituto Giannina Gaslini, Genova, Italy † Longevity Institute, Davis School of Gerontology, University of Southern California, Los Angeles, CA, United States { IFOM, FIRC Institute of Molecular Oncology, Milano, Italy 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Dysregulation of Metabolism in Cancer and Aging 2.1 Mitochondrial Homeostasis in Cancer 2.2 Geroncogenesis and Gerometabolites: The Pseudohypoxic Aging Side of Oncometabolites 2.3 Sirtuins: Regulators of Metabolism of Cancer and Aging 2.4 Nutrient-Sensing Pathways: A Common Signaling in Aging and Cancer 2.5 Inflammation and Cancer 3. Metabolic Interventions With Effects on Aging and Cancer 3.1 Calorie Restriction 3.2 Protein Restriction 3.3 Fasting and Fasting-Mimicking Diet 3.4 Pharmacological Interventions Mimicking CR 4. Geroscience as a Strategy to Optimize Cancer Therapy 4.1 Fasting and Fasting-Mimicking Diet 4.2 Glycolysis Blockade 4.3 Nutrient-Sensing Pathway Interventions 5. Conclusions Acknowledgments References

2 3 3 6 9 10 13 13 14 15 17 19 24 25 27 27 28 29 29

Abstract Aging represents the major risk factor for cancer. Cancer and aging are characterized by a similar dysregulated metabolism consisting in upregulation of glycolysis and downmodulation of oxidative phosphorylation. In this respect, metabolic interventions can be viewed as promising strategies to promote longevity and to prevent or delay age-related disorders including cancer. In this review, we discuss the most

International Review of Cell and Molecular Biology, Volume 332 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2017.01.003

#

2017 Elsevier Inc. All rights reserved.

1

2

L. Raffaghello and V. Longo

promising metabolic approaches including chronic calorie restriction, periodic fasting/fasting-mimicking diets, and pharmacological interventions mimicking calorie restriction. Metabolic interventions can also be viewed as adjuvant anticancer strategies to be combined to standard cancer therapy (chemotherapeutic agents, ionizing radiation, and drugs with specific molecular target), whose major limiting factors are represented by toxicity against healthy cells but also limited efficacy easily circumvented by tumor cells. In fact, conventional cancer therapy is unable to distinguish normal and cancerous cells and thus causes toxic side effects including secondary malignancies, cardiovascular and respiratory complications, endocrinopathies, and other chronic conditions, that resemble and, in some cases, accelerate the age-related disorders and profoundly affect the quality of life. In this scenario, geroscience contributes to the understanding of the mechanisms of protection of normal cells against a cytotoxic agent and finding strategies focused on the preserving healthy cells while enhancing the efficacy of the treatment against malignant cells.

1. INTRODUCTION Aging represents the major risk factor for cancer. In fact, the incidence of cancer increases with age due to accumulation of mutations. This association can be explained not only by the multihit or Knudson hypothesis, according to which tumor cells need to accumulate the mutations responsible for genome instability and carcinogenesis, but also by a decline of homeostasis occurring during aging (Knudson, 1971; Wu et al., 2014b). The discovery of the role of protooncogenes in accelerating aging and promoting cellular sensitization to stress has encouraged the investigation of metabolic interventions able to promote longevity but also useful in generating differential protection and sensitization of normal and cancer cells, respectively. In this review, we will discuss the most promising treatments including chronic calorie and protein restriction, fasting and fasting-mimicking diets (FMD), and pharmacological interventions mimicking calorie restriction (CR) relevant to both aging and cancer. In addition to their antiaging effects, the above-mentioned strategies can also function as adjuvant approaches to be combined with standard cancer treatments in order to protect healthy cells and tissues and sensitize malignant cells to cytotoxic agents. In fact, one of the major limitations of cancer therapy, including chemotherapy, ionizing radiation, and target-specific drugs, is their inability or limited ability to distinguish normal and cancerous cells. This aspect is associated with the appearance of long-term and often serious

Cancer and Aging Metabolism

3

side effects that not only impair the quality of life but may also accelerate the aging process. For this reason, geroscience, whose purpose is to understand the mechanisms of damage and protection during aging, could lead to novel interventions able to protect the host from the standard cancer treatment while enhancing its efficacy.

2. DYSREGULATION OF METABOLISM IN CANCER AND AGING 2.1 Mitochondrial Homeostasis in Cancer Dysregulation of metabolism represents one of the hallmarks of tumorigenesis. In particular, cancer cells reprogram their carbon metabolism by upregulating glycolysis (Warburg effect) and glutaminolysis and by downregulating oxidative phosphorylation (OXPHOS). These events lead to the increase of biosynthetic intermediates such as nucleotides, amino acids, and lipids which are necessary for cell proliferation and survival (Vander Heiden et al., 2009). Moreover, disruption of mitochondrial homeostasis causes accumulation of reactive oxygen species (ROS) which can contribute to cancer progression (Hamanaka and Chandel, 2010). In accordance with the normal function of their encoded proteins, oncogenes or tumor suppressors regulate cellular metabolism (Cairns et al., 2012). For example, Myc and phosphatidyl inositol 3-kinase (PI3K) are potent inducers of glutaminolysis and glucose uptake, respectively (Elstrom et al., 2004; Wise et al., 2008), while the tumor suppressor p53 inhibits glucose transporters (GLUT), glycolysis, pentose phosphate pathway, de novo fatty acid synthesis, and enhances mitochondrial OXPHOS and tricarboxylic acid (TCA) cycle rate (Wang et al., 2014). A similar effect on the regulation of metabolism is caused by the tumor suppressor liver kinase B1 (LKB1) that is frequently mutated in Peutz-Jegher’s syndrome, characterized by an increased risk of gastrointestinal cancer. LKB1 encodes for a serine-threonine kinase that activates the energetic sensor AMP-dependent kinase (AMPK) and downmodulates the regulator of growth and proliferation known as mammalian target of rapamycin (mTOR) (Shackelford and Shaw, 2009). Von Hippel Lindau disease is another example of disorders associated to cancers and caused by mutations in the Vhl gene, which inhibits the hypoxia-inducible factor α (HIF-α) (Kapitsinou and Haase, 2008; Maxwell et al., 1999). HIF-α is a transcriptional factor which suppresses oxidative metabolism and promotes glycolysis (Semenza, 2010).

4

L. Raffaghello and V. Longo

In the last 10 years, several genetic defects affecting TCA enzymes associated to oncogenesis have been described. In particular, loss-of-function mutations or changes in the amino acid residues have been identified in the cytoplasmic and mitochondrial isoforms of the enzymes succinate dehydrogenase (SDH), isocitrate dehydrogenase (IDH), and fumarate hydratase (FH) (Cardaci and Ciriolo, 2012). The SDH complex is an highly conserved heterotetrameric tumor suppressor, composed by two catalytic subunits (SDHA and SDHB) and two hydrophobic subunits (SDHC and SDHD) (Bardella et al., 2011), which have been found mutated in patients affected by different malignancies including hereditary paragangliomas, pheochromocytomas, gastrointestinal stromal, thyroid and renal tumors, neuroblastoma, and testicular seminoma (Astuti et al., 2001; Bardella et al., 2011; Baysal et al., 2000). FH is a homotetrameric TCA cycle enzyme, which catalyzes the hydration of fumarate to L-malate. Heterozygous FH mutations predispose to multiple cutaneous and uterine leiomyomas, hereditary leiomyomatosis, renal cell cancer, breast, bladder, as well as Leydig cell tumors (Carvajal-Carmona et al., 2006; Lehtonen et al., 2006; Tomlinson et al., 2002). IDH is a member of the β-decarboxylating dehydrogenase family, which catalyzes the oxidative decarboxylation of isocitrate to produce 2-oxoglutarate (α-KG). Three isoforms have been identified so far: the cytosolic IDH1, and the mitochondrial IDH2 and IDH3. Mutations associated to IDH1 and IDH2 have been identified in 70% of grade II–III gliomas, secondary glioblastomas, acute myeloid leukemia, angioimmunoblastic T-cell lymphomas, thyroid, colorectal, and prostate cancer (Abbas et al., 2010; Cairns et al., 2012; De Carli et al., 2009; Yen et al., 2010). As a result of gain-of-function mutations, IDH1 and IDH2 are unable to efficiently convert isocitrate into α-KG and acquire a neomorphic catalytic activity that allows a NADPH-dependent reduction of α-KG into the oncometabolite (R)-2-hydroxyglutaric acid ((R)-2HG) (Dang et al., 2010; Ward and Thompson, 2012). The mechanisms underlying tumorigenesis in cancers characterized by TCA cycle enzyme mutations involve the accumulation of metabolites (succinate, fumarate, and (R)-2-HG) that convey oncogenic signals (oncometabolites) (Yang et al., 2013). In particular, the abnormal accumulation of (R)-2HG mediates its potential tumorigenic effects via several mechanisms: (i) inhibition of ten-eleven translocation family (TET) of dioxygenases and histone lysine demethylase (KDM) which results into enhanced CpG island and histone methylation and the consequent

Cancer and Aging Metabolism

5

remodeling of the cancer cell epigenome toward an undifferentiated and aggressive phenotype, (ii) inhibition of collagen prolyl and lysyl hydroxylases causing impaired collagen maturation and disrupted basement membrane formation, and (iii) inhibition of HIF-α prolyl hydroxylase (PHD) interactions causing a decrease of HIF-1α degradation and an enhancement of pseudohypoxic condition. In addition, accumulation of fumarate and succinate participates in oncogenic signaling through: (i) modification of cysteine residues in proteins which confers a state of constitutive activation of the antioxidant defense pathway mediated by NF-E2-related factor (NRF2), that generates a reductive milieu and promotes cell proliferation; (ii) inhibition of the reactions involved in arginine and purine synthesis; (iii) epigenetic alterations by inhibition of TET and KDM proteins; and (iv) accumulation of HIF-1α which in its turn promotes aerobic glycolysis and angiogenesis (Selak et al., 2005). Furthermore, succinate has recently emerged as a key player in the promotion of inflammation which is functionally associated to cancer development and progression. In particular, proinflammatory macrophages shift their metabolism from OXPHOS to glycolysis resulting in succinate accumulation and oxidation by SDH, which drives ROS production that, finally, leads to increase of HIF-1α and proinflammatory cytokines including IL-1β (Mills et al., 2016). A recent study highlighted the importance of the oncometabolite fumarate as a driver of tumorigenesis (Sciacovelli et al., 2016) by discovering that accumulation of fumarate, associated to FH loss, induces epithelial mesenchymal transition (EMT), a well-known process involved in cancer initiation, dissemination, and metastasis. Specifically, fumarate has been shown to inhibit TET-mediated demethylation of a regulatory region of the antimetastatic miRNA cluster mir-200ba429, leading to the expression of EMT-related transcription factors and enhanced migratory properties (Sciacovelli et al., 2016). Beside the above-mentioned oncometabolites, ADP-ribose, which is synthesized by poly-ADP-ribose polymerases (PARPs) from NAD(+) and is responsible for protein posttranslational modifications, can be considered as an oncometabolite as well. A recent study demonstrated that nuclear pyruvate kinase M2 (PK)M2 binds directly to ADP-ribose, and this poly-ADP-ribose-binding capability is critical for its nuclear localization. Accordingly, PARP inhibition prevents nuclear retention of PKM2 and therefore suppresses cell proliferation and tumor growth (Li et al., 2016).

6

L. Raffaghello and V. Longo

The general mechanism underlying tumor metabolism dysregulation is still under investigation. The prevailing point of view is that the reprogramming of tumor metabolism (Warburg-like effect) occurs after cancer cells accumulate key mutations, promoting additional genome instability (Vander Heiden et al., 2009). According to alternative hypothesis, known as geroncogenesis, during age, normal cells undergo natural alterations in oxidative metabolism, with the consequent increased generation of ROS, which promote additional mutations and tumorigenesis. The latter hypothesis may help explain why aging is the major risk factor for most tumors (Wu et al., 2014b).

2.2 Geroncogenesis and Gerometabolites: The Pseudohypoxic Aging Side of Oncometabolites Consistent with geroncogenesis, aging in mammals is characterized by metabolic alterations similar to those associated with cancer cells, and by a reduction of OXPHOS as a result of alteration of specific electron transport chain (ETC) complexes as well as an increase in aerobic glycolysis in different tissues (Bowling et al., 1993; Hagen et al., 1997; Trounce et al., 1989). For example, in humans, age leads to a decrease in cytochrome oxidase activity in brain and heart (Muller-Hocker, 1989; Ojaimi et al., 1999). Along this line, the activity of complex V decreases in the heart of the Fischer 344 rats and is accompanied by structural changes during aging. In contrast, complexes I and III activity remain unaltered in the heart, liver, and skeletal muscle of mice during aging (Kwong and Sohal, 1998, 2000). As a result of alterations, aging is associated with a higher production of ROS due to a decreased flux through the ETC. This event reduces the activity of upstream complexes, especially complexes I and III, enhancing “electron leak” that generates ROS (Chen and Lesnefsky, 2006). The increased production of ROS by mitochondria leads to greater oxidative damage within mitochondria, including protein sulfhydryl oxidation, lipid peroxidation, and mitochondrial (mt)DNA damage (Floyd et al., 2001; Van Remmen and Richardson, 2001). In support of the importance of mitochondrial-derived ROS to the aging process, it has been shown that interventions and mutations that prolong survival tend to decrease the production of ROS from mitochondria (Chen et al., 2007; de Cabo et al., 2014; Lagouge and Larsson, 2013). Similar metabolic dysregulations have been observed in type 2 diabetes: an age-related disorder characterized by accumulation of HIF-1α and a decline of OXPHOS (Petersen et al., 2004; Ptitsyn et al., 2006).

Cancer and Aging Metabolism

7

An additional possible link between aging and tumorigenesis is supported by the suggestive theory according to which the so-called “gerometabolites” (defined as small-molecule components of normal metabolism whose depletion drives aging) promote the chronic accumulation of oncometabolites which in turn are responsible for pathological metabolic reprogramming. In this scenario, nuclear NAD+, a central metabolic cofactor that decreases during aging (Verdin, 2015), plays a pivotal role. NAD+ is cosubstrate of sirtuins, a family of NAD+-dependent deacetylases that are regulators of metabolism in cancer and aging. Specifically, sirtuin 1 deacetylases HIF1α and promotes its interaction with the ubiquitin ligase VHL which leads to the degradation of HIF-1α (Gomes et al., 2013; Haase, 2009). The decline of NAD+ during aging represents the causal inducer of the accumulation of HIF-1α which promotes a pseudohypoxic state that reduces the carboxylation of α-ketoglutarate. This event leads to the production of the oncometabolite (R)-2HG (Gomes et al., 2013), that, together with the other oncometabolites fumarate and succinate, creates a pseudohypoxic state. The latter condition promotes the activation of c-Myc which is responsible for repressing the transcription of mitochondrial genes. As a consequence, a reprogramming of metabolism characterized by promotion of glycolysis and inhibition of OXPHOS occurs. This scenario, as depicted in Fig. 1, offers the possible connection between aging and cancer where the conjunction ring is represented by pseudohypoxia that is induced by gerometabolites. Pseudohypoxia is then associated with the generation of oncometabolites which block the differentiation and promote stemness through epigenetic mechanisms (Menendez et al., 2014). In this context, several report indicate that hypoxia favors the increase in the cancer stem cell (CSC) pool through HIF-1α and HIF-2α which, in turn, amplify the CSC pool and induce additional dedifferentiation of tumor cells (Carnero and Lleonart, 2016; Keith and Simon, 2007; Li et al., 2009). In particular, hypoxia and HIFs induce stemness in differentiated progenitors and non-CSCs by inducing the expression of genes such as OCT4, SOX2, and NANOG, or the activation of the Notch signaling pathway that regulates cell self-renewal and differentiation (Bennewith and Durand, 2004). The molecular mechanisms that underlie the impact of hypoxic conditions on the CSC compartment require either HIF-1α expression or the inactivation of the hydroxylase activity of PHD domain-containing protein 3 (PHD3) whose loss prevents CSC differentiation and induces dedifferentiation of mature tumor cells (Iriondo et al., 2015). Finally, HIF-1α mediates telomerase transcription in hypoxic cancer cells, maintaining the immortal

8

L. Raffaghello and V. Longo

Gerometabolites

Oncometabolites +

NAD

(R)-2-Hydroxyglutarate

Succinate

Fumarate

g Agin

NADH

pVHL OH

HIF-a

Normoxia

HIF-α proteosomal degradation

OH

HIF-PHDs

HIF-a

Pseudohypoxia C-Myc

Transcription mitochondrial genes

Glycolysis Aging

OXPHOS Cancer

Fig. 1 The link between metabolites and oncometabolites in aging and cancer. Metabolites, such as NAD+, which decline as we age, cause the accumulation of oncometabolites through the induction of a pseudohypoxia condition (Gomes et al., 2013). This latter event occurs as a consequence of an increased HIF-1α stabilization due to the ubiquitin ligase pVHL inhibition. Oncometabolites inhibit the interaction between HIF-1α and prolyl hydroxylases (PHDs) resulting in a decreased HIF-1α degradation. As a consequence, oncometabolites generate a pseudohypoxia condition as other metabolites do. This event converges on c-Myc activation, which represses the transcription of mitochondrial genes resulting in glycolysis promotion and oxidative phosphorylation (OXPHOS) inhibition.

life span of the tumor mass. In fact, 90% of tumors show increased telomerase activity, suggesting that it is an important factor in the maintenance of CSC properties (Blanco et al., 2007). HIF-α and pseudohypoxia have been also implicated in aging although in Caenorhabditis elegans this issue is a matter of debate because HIF-α has been shown to function as both a positive and negative modulator of aging (Leiser and Kaeberlein, 2010). There is also evidence that HIF-α plays a relevant role in mammalian aging. In particular, activation of HIF-1α, which is accompanied by increased HIF-1α DNA binding and activation of transcription of HIF-1α-dependent genes, has been observed in aged rat liver (Kang et al., 2005). Another study examining PHD3 in rats found that PHD3 levels increase with age in liver, heart, and skeletal muscle, and this increase correlates with a decrease in HIF-1α

Cancer and Aging Metabolism

9

activity (Rohrbach et al., 2008). Finally, HIF-1α seems to have a contradictory role in age-related disorders; in fact, studies examining Aβ accumulation in Alzheimer’s disease suggest that increase of HIF-1α tends to be protective in disease-free states, but that increased HIF-1α levels are also a sign of advanced disease progression (Ogunshola and Antoniou, 2009). These seemly conflicting results might be due to the fact that in stress response HIF-1α upregulation can be either protective or reactive in nature (Ogunshola and Antoniou, 2009).

2.3 Sirtuins: Regulators of Metabolism of Cancer and Aging Sirtuins are mono-ADP-ribosyltransferase and beta-nicotinamide adenine dinucleotide (NAD+)-dependent lysine deacetylase enzymes that play key roles in the regulation of metabolism, inflammation, and DNA repair and are critical regulators at the crossroads between cancer and aging (Chalkiadaki and Guarente, 2015; Saunders and Verdin, 2007). The mechanisms underlying the protective effect of sirtuins in aging and cancer include: (1) protection against DNA damage and oxidative stress and (2) protection against accumulation of mutations and genomic instability (Saunders and Verdin, 2007). In fact, the loss of sirtuin expression, activity, or regulation allows cell division to proceed without the proper repair of DNA, resulting in accumulation of mutations and genomic instability which can lead to tumor development. More recently, sirtuin proteins have also been found to finely regulate energy metabolism. In particular, sirtuin 3 facilitates TCA cycle, OXPHOS, and fatty acid metabolism (Hirschey et al., 2010). For example, deletion of Sirtuin 3 causes spontaneous formation of mammary tumors and a metabolic reprogramming characterized by increased glucose uptake and decreased ATP generation (Kim et al., 2010). This metabolic switch is accompanied by an enhanced production of ROS that stabilize HIF-1α, which in its turn promotes glycolysis and reduces OXPHOS (Bell et al., 2011; Finley et al., 2011). The role of sirtuin 3 as a tumor suppressor is however controversial since other studies indicated that sirtuin 3 is overexpressed in breast cancer compared to normal tissues (Alhazzazi et al., 2011; Ashraf et al., 2006). Another sirtuin protein involved in tumor metabolism regulation is sirtuin 6, whose deletion is associated with an increase in HIF-1α and c-Myc, leading to activation of glycolysis (Sebastian et al., 2012; Zhong et al., 2010). Similarly, sirtuin 1, which was the first family member identified as a tumor suppressor that delays lymphoma and protects mice against

10

L. Raffaghello and V. Longo

carcinogen-mediated hepatocellular carcinoma (Herranz et al., 2010; Oberdoerffer et al., 2008), regulates the transcriptional activity of HIF-1α and activates LKB1 through deacetylation (Lan et al., 2008; Lim et al., 2010). Sirtuin 2 is another tumor suppressor involved in maintaining genome stability and suppressing tumorigenesis by deacetylating and activating Cadherin-1 (CDH1), a protein that limits glycolysis (Almeida et al., 2010; Kim et al., 2011). More recently, sirtuin 4 and sirtuin 7 have also been implicated in tumorigenesis with an effect of sirtuin 4 in repressing the mitochondrial glutamine metabolism, and a role of sirtuin 7 in negatively regulating HIF-1α and HIF-2α (Hubbi et al., 2013; Jeong et al., 2013). Although all the above studies indicate that sirtuins can function as tumor suppressors, their role in tumorigenesis is still controversial since different reports indicated that sirtuins, including sirtuin 1, sirtuin 2, and sirtuin 3, also promote cancer (Bosch-Presegue and Vaquero, 2011). In this regard, sirtuin 1 has been shown to inhibit p53 and to stabilize c-Myc, which may explain why its overexpression can increase tumor growth (Menssen et al., 2012; Suh et al., 2011; Villeda et al., 2011).

2.4 Nutrient-Sensing Pathways: A Common Signaling in Aging and Cancer Several studies in yeast, worms, flies, and mice indicate that conserved nutrient-sensing pathways governed by insulin/insulin growth factor (IGF), glucose, and amino acids regulate aging and affect genomic integrity, DNA repair, ROS generation, and cellular apoptosis, highlighting their role in the promotion of cancer (Fontana et al., 2010; Longo and Fontana, 2010). As depicted in Fig. 2, a signaling pathway is triggered when a specific ligand binds to its cognate receptor. These ligands are mainly represented by glucose and amino acids in yeasts; Ins/IGF-1-like peptides in worms and flies; and insulin, IGF-1, and IGF-2 in mammals. In yeast, Ras is involved in the activation of adenylate cyclase (AC) which in its turn activates protein kinase A (PKA), leading to the phosphorylation and inactivation of transcription factors including MSN2/4 and GIS1. The cognate receptors include the insulin-like receptor DAF-2 in worms, InR in flies and IGF-1R, IR-A, and IR-B in mice. Upon the interaction between ligands and their receptors, the signal is transduced through adaptor proteins such as Ras in yeasts and mice, IST-1 in worms, CHICO in flies, and insulin receptor substrate 1 (IRS1-4) in mammals. These proteins are involved in the activation of PI3K which generates phosphatidyl inositol (3,4,5)-trisphosphate (PIP3). In animals, PIP3 activates AKT/PKB which phosphorylates and inactivates

11

Cancer and Aging Metabolism

Yeast

Worms

Flies

Mammals

Dietary restriction

Dietary restriction

Dietary restriction

Dietary restriction

TOR

Insulin/IGF-1 like

Insulin/IGF-1 like

Glucose, amino acids

Grp-1

DAF-2

RAS

IST-1

IGF-1

IGF-R

INR

TOR

TOR

PI3K AGE-1

AC

GHR

GH signaling

CHICO

TOR

GH

RAS

PI3K

(PI3K)

Sch9 (S6K)

AC

S6K

PKA

AKT

RSK1 (S6K)

RIM15

PKA

FOXO

16

F DA

GIS1

MSN2/4

AKT

S6K

AKT

?

FOXO

P

4E-B

-1

HIF

?

?

Antioxidant enzymes (SOD, catalase, and heat shock proteins) autophagy, translation, ER stress

Antiageing

Fig. 2 The nutrient-sensing pathways in aging. Certain nutrients sensing pathways are evolutionarily conserved in different organisms including yeasts, worms, flies, and mammals. These pathways are triggered by ligands (glucose and amino acids in yeast, insulin and insulin growth factor-1 like (IGF-1) in worms and flies, IGF-1 in mammals) which activate a receptor (G protein-coupled receptor 1 (Gpr1) in yeasts; DAF-2 in worms; insulin-like receptor (InR) in flies; and IGF-1R, IR-A, and IR-B in mammals). The signal is then transduced through adaptor proteins which include Ras in yeasts, IST-1 in worms, CHICO in flies, and insulin receptor substrate 1 (IRS1–4) and Ras in mice. In yeast, Ras activates adenylate cyclase and protein kinase A (PKA). In worms, flies, and mice, the adaptors proteins mediate the activation of phosphatidylinositol-4,5-bisphosphate 3-kinase (PI3K) (AGE-1 in worms) which generates phosphatidylinositol (3,4,5)trisphosphate (PIP3). PIP3 activates AKT/PKB, or PKA in yeast, phosphorylates, and inactivates specific stress resistance transcription factors (MSN2/4 and GIS1 in yeasts, DAF-16 in worms, Forkhead box protein O (FOXO) in flies and mice) involved in the upregulation of protective stress resistance genes such as superoxide dismutase (SOD), catalase, and heat shock proteins (HSPs). A parallel pathway involved in the inactivation of the above-mentioned transcription factors is governed by the target of rapamycin (TOR) and its substrate S6 kinase (S6K). The nutrient-sensing pathways can be downmodulated by diet restriction, which reduces the levels of glucose and IGF-1.

specific antiaging transcription factors (DAF-16 in worms, and Forkhead box protein O (FOXO) in flies and mice). This event causes the downregulation of protective stress resistance genes including superoxide dismutase (SOD), catalase, and heat shock proteins (HSPs) which contribute

12

L. Raffaghello and V. Longo

to the protection against oxidative damage contributing to aging as well as cancer. An additional pathway involved in the promotion of aging is controlled by mTOR and the serine kinase 6 (S6K). Dampening these nutrient-sensing pathways by specific mutations, as it occurs in the yeast sch9 null mutants, CHICO mutant flies, Ames and Snell dwarf mice lacking growth hormone (GH) and IGF-1 as well as “Laron dwarf mice” produced by targeted disruption of the GH receptor/GH-binding protein gene (GHR-KO mice), significantly increases the life span (Longo and Finch, 2003). Similar effects can be obtained by strategies based on dietary restriction, which not only extend longevity but also reduce or delay age-related disorders, particularly spontaneous tumors. The effects of these mutations on aging may be explained in part by the finding that fibroblasts from long-lived adult Ames or Snell dwarf mice and GH receptor knockout mice are better protected against oxidative stress, suggesting that IGF-1 has a key role in regulating protection against toxins (Fontana et al., 2010). In agreement with the latter observation, individuals who carry mutations in the GH receptor gene that lead to severe GHR and IGF-1 deficiencies (Laron population) are protected against cancer and diabetes (Guevara-Aguirre et al., 2011). The explanation for these results is provided in part by in vitro studies in which mammary epithelial cells incubated with serum from the Laron individuals were shown to be more protected from DNA damage caused by hydrogen peroxide but are also more likely to die as a consequence of damage compared to those incubated with serum from unaffected relatives. Nutrient signaling pathways play also a relevant role in the regulation of cell metabolism. Upon binding to its receptor, IGF-1/2 can activate Ras which, in turn, can mediate the activation of the PI3K/AKT and MAPK pathways, both of which increase glycolysis through different mechanisms. AKT increases the expression and membrane localization of the glucose transporter GLUT1, which stimulates phosphofructokinase activity and induces the translocation of hexokinases 1 and 2 into the mitochondria where they drive the first reaction of glycolysis (Barthel et al., 1999; DeBerardinis et al., 2008; Robey and Hay, 2006). Downstream of AKT, mTOR stimulates the switch to the glycolytic pathway through upregulation of HIF-1α, protein translation, and lipogenesis by inducing the SRBPE-dependent transcription of different enzymes such as ATP citrate lyase and fatty acid synthase (Krycer et al., 2010; Yang et al., 2002). Interestingly, in an HIF-1α- and Myc-dependent manner, mTOR upregulates the activity of PKM2, a crucial glycolytic enzyme involved in

Cancer and Aging Metabolism

13

promoting the pro-Warburg effect (Sun et al., 2011). PKM2 interacts with HIF-1α and enhances the expression of HIF-1α target genes as well as it upregulates c-Myc transcription (Luo et al., 2011). In addition, mTOR as well as MAPK activate Myc whose overexpression causes the conversion of pyruvate into lactate thereby contributing to the Warburg effect (Dang et al., 2009). Myc is also involved in glutaminolysis induction through the upregulation of glutamate synthesis (Gao et al., 2009).

2.5 Inflammation and Cancer One of the most typical feature and also a probable cause of aging are represented by a chronic and low level state of systemic inflammation in the absence of any infection that has been referred to as inflammaging (Franceschi and Campisi, 2014). Chronic inflammation increases cancer risk and promotes all cancer stages, including cancer initiation, progression, and metastatic diffusion (Mantovani et al., 2008). Thus, inflammation is likely to contribute to age-dependent cancer. Inflammation can be derived from different sources including: (i) damaged macromolecules, organelles, and cells (self-debris) that accumulate with age and behave as “damage”-associated molecular targets that activate innate immunity and the Nlrp3 inflammasome; (ii) products derived from the microbial constituents of the human body, such as oral or gut microbiota, that elicit an inflammatory response (Biagi et al., 2011); (iii) senescent cells which accumulate with age and secrete proinflammatory cytokines; (iv) the coagulation system which is activated and increases during aging; (v) changes in immunosenescence; and (vi) defective or inappropriate regulation of the complement pathway that can lead to local inflammatory reactions (Franceschi and Campisi, 2014). All of these stimuli can result in the activation of nuclear transcription factor NF-κB which is considered as a hub in carcinogenesis, linking inflammation, cellular senescence, and cancer (Ostan et al., 2015).

3. METABOLIC INTERVENTIONS WITH EFFECTS ON AGING AND CANCER Since nutrient-sensing pathways have been demonstrated to regulate longevity and to be modulated by dietary interventions, in the following sections, we will discuss different metabolic approaches demonstrated to have prolongevity effects and to prevent or delay multiple age-related diseases and improve health span. These include: (1) CR, (2) periodic fasting

14

L. Raffaghello and V. Longo

(PF), (3) protein restriction, and (4) pharmacological interventions mimicking CR such as inhibitors of mTOR/S6K signaling, of glycolysis, and of GH or IGF-1 signaling, activators of sirtuins, and pharmacological inhibition of inflammation (Longo et al., 2015).

3.1 Calorie Restriction CR is defined as a 20%–40% reduction in caloric intake without causing malnutrition, and while, in most cases, maintaining meal frequency. CR represents the most potent physiological intervention able to delay and prevent aging and age-associated diseases in single-celled, invertebrate, and vertebrate animals (Fontana and Partridge, 2015). In yeast, starvation obtained by switching the cells from a medium containing nutrients to one containing only water extends the chronological life span (Wei et al., 2008). Similarly, life span extension has been also observed in worms and flies under CR (Fontana et al., 2010). More than 80 years ago, different studies demonstrated that mice and rats under CR live longer and healthier in part by delaying the occurrence of several chronic diseases such as cancer, diabetes, atherosclerosis, and cardiomyopathy (Fontana et al., 2010). In term of cancer prevention, CR has been shown to inhibit the occurrence and growth of spontaneous, chemically induced, and radiation-induced tumors in various animal models (Cheney et al., 1983; Tannenbaum and Silverstone, 1949). Moreover, CR in mice greatly increases insulin sensitivity (Patel et al., 2005). Accordingly, young adult Rhesus monkeys under CR present a significant reduction of age-related pathologies including type 2 diabetes, cardiovascular disease, sarcopenia, and cancer (Colman et al., 2014; Mattison et al., 2012). In addition, immunosenescence and atrophy of the brain’s gray matter are attenuated (Colman et al., 2014). In humans, long-term CR causes metabolic, molecular, and cellular adaptations which are responsible for its protective effect against type 2 diabetes, hypertension, cardiovascular disease, dementia, and cancer (Cava and Fontana, 2013). In particular, overweight women and elderly subjects under 3–4 months of CR present significantly improved cognitive function (Kretsch et al., 1997). The mechanism underlying the antiaging effect of CR include three levels of adaptations: (1) metabolic adaptations associated to a decrease of insulin, IGF-1, sex hormones, oxidative stress, inflammation, and to an increase of cortisol and adiponectin; (2) molecular adaptations which result in downregulation of PI3K/AKT/S6K, mTOR, RAS/MAPK pathways and upregulate NRF2, sirtuins, AMPK, FOXO, and PTEN; and (3) cellular adaptations

Cancer and Aging Metabolism

15

including a decrease in cellular proliferation, oxidative stress, and enhancement of apoptosis, autophagy, DNA repair, genome stability, and immunosurveillance (Longo and Fontana, 2010). For example, CR-mediated inhibition of AKT activates FOXOs transcription factors that, in turn, activate protective systems controlling cell proliferation, autophagy, stress resistance, and DNA repair (Webb and Brunet, 2014). Inhibition of mTORC1 increases autophagy and enhances stem cell function (Johnson et al., 2013; Kapahi et al., 2010). Moreover, genome stability and antioxidant defenses can be increased by the overexpression of certain sirtuins such as sirtuins 1, 3, and 6 and by the activation of heat shock factor 1 (HSF1) and of the NRF2 transcription factor (Akerfelt et al., 2010; Martin-Montalvo et al., 2011).

3.2 Protein Restriction CR, describes the reduction of calories, rather than referring to both the quantity and composition of the diet, which research is now showing to be the major factor responsible of its health span effects in rodents. In fact, it has been observed that neither carbohydrate nor lipid restriction seems to have major effects on longevity alone. In contrast, several studies performed in yeast, worms, flies, and rodents point to protein restriction without a decrease of calorie as a nutritional modification able to increase the life and health span. There is a clear evidence that the reduction of specific amino acids including serine, valine, threonine, asparagine, glutamate, or methionine extends the life span in yeast Saccharomyces cerevisiae, Drosophila melanogaster, and rodents (Ables et al., 2012; Miller et al., 2005; Richie et al., 1994; Wu et al., 2013). Particular interest has been focused on methionine restriction that exerts healthy effects by decreasing mitochondrial oxidative stress and consequently oxidative damage to mitochondrial DNA (Sanchez-Roman and Barja, 2013). The mechanisms underlying the effect of amino acids on aging are associated to their effect on the activation of mTOR and the control of the nonderepressible 2 (GCN2) gene. mTOR is modulated by different essential amino acids, mainly leucine, in a tissue-specific manner, while GCN2 is a serine/threonine-protein kinase that, once activated by amino acid deficiency, stabilizes the transcription factor ATF4 which is essential for the integrated stress response (Li et al., 2014). In yeast, methionine restriction promotes longevity in a GNC2-dependent manner (Wu et al., 2013), but other mechanisms including induction of autophagy

16

L. Raffaghello and V. Longo

(Ruckenstuhl et al., 2014) and mitochondrial retrograde response (Johnson and Johnson, 2014) have been proposed. The importance of other amino acids such as serine, valine, and threonine in promoting aging in yeast has been recently demonstrated (Mirisola et al., 2014). Specifically, threonine and valine promote cellular sensitization and aging primarily by activating the TOR/S6K pathway, while serine induces sensitization via phosphoinositide-dependent protein kinase 1 (PDK1) orthologs Pkh1/2. These events cause intracellular relocalization and transcriptional inhibition of the stress resistance protein kinase Rim15 (Mirisola et al., 2014). The extension of the health span in D. melanogaster is attributable to a restriction of protein-containing yeasts or sugar (Mair et al., 2005). According to these results, in mice, a low-protein and high-carbohydrate diet exerts life-extending effects through downregulation of the hepatic mTOR pathway (Solon-Biet et al., 2014). Protein restriction has been also associated to decrease the prevalence and the severity of age-related diseases. For example, in mice, a low-protein and high-carbohydrate diet reduces the blood pressure, low-density lipoproteins, and triglycerides; improves glucose tolerance; and increases the levels of high-density lipoprotein (Solon-Biet et al., 2014). These results are in agreement with data performed on human subjects in which high-protein and low-carbohydrate diets are associated with an increase of cardiovascular diseases (Floegel and Pischon, 2012; Lagiou et al., 2012). Thus, the balance of protein to carbohydrate, rather than energy intake, may be the driver of a healthy cardiometabolic profile. The serum IGF-1 reduction and mTOR downregulation by an isocaloric restriction of protein has been also associated to a marked inhibition of prostate and breast cancer growth in experimental models (Fontana et al., 2013; Levine et al., 2014). In humans little is known about the effects of protein restricted diets on cancer. However, an interesting study reported that low-protein intake during middle age (between 50 and 65 years old) is associated with decreased risk of cancer and diabetes mortality as well as overall mortality. Conversely, low-protein intake is associated with increased risk of cancer and overall mortality in respondents over 65 (Levine et al., 2014). In agreement with the observation that an increased protein intake and the resulting increase in IGF-1 may prove beneficial for the skeletal muscle metabolism of older adults (Heaney et al., 1999), a very low-protein diet may be detrimental for older adults in whom weight begins to decline thus making them more susceptible to protein malnourishment (Levine et al., 2014).

Cancer and Aging Metabolism

17

3.3 Fasting and Fasting-Mimicking Diet Fasting represents the most extreme version of CR where nutrients are totally eliminated. Two major forms of fasting can be practiced: (i) an intermittent fasting (IF) also known as alternate day fasting that involves a 24-h fast followed by a 24-h nonfasting period and (ii) prolonged and PF in which absence of food lasts two or more consecutive days every 2 or more weeks (Longo and Mattson, 2014). The first experiments performed on the yeast S. cerevisiae demonstrated that the shift from standard growth medium to only water causes a significant increase of life span as well as the resistance to multiple stress (Longo et al., 1997). This intervention is associated to downregulation of mTOR/S6K (Sch9) and RAS-AC-PKA pathways resulting in increased transcription of: (i) stress resistance genes such as SOD, catalase, and HSPs (Madia et al., 2009) and (ii) DNA repair genes including Rev1 (Wei et al., 2008) (Fig. 2). As a result, fasting exerts a protective effect against DNA damage and promotes longevity (Longo and Fontana, 2010). Similar observations have been made in the nematode C. elegans and D. melanogaster in which food deprivation increases the life span through the downmodulation of mTOR/S6K and IGF-1-like/ PI3K/AKT pathways resulting in the activation of transcription factors DAF16 and FOXO (Fontana et al., 2010; Greer et al., 2007; Piper and Partridge, 2007) (Fig. 2). In mice, IF reduces the incidence of spontaneous tumors including lymphoma and sarcoma (Berrigan et al., 2002; Descamps et al., 2005), while PF can delay cancer progression as effectively as chemotherapy (Lee et al., 2012). In addition, IF ameliorates age-related behavioral deficits in the triple-transgenic mouse model of Alzheimer’s disease (Halagappa et al., 2007), reduces degeneration of dopaminergic neurons in models of Parkinson’s disease (Duan and Mattson, 1999), and slows disease progression in huntingtin mutant mice by normalizing glucose metabolism and brain-derived neurotrophic factor levels (Duan et al., 2003). In agreement with the latter results, a recent study demonstrated that IF, when imposed in the middle age, delays or prevents the age-associated impairment of brain functions and promotes healthy aging by improving the motor coordination and learning response recovery (Singh et al., 2012, 2015). Furthermore, IF protects the heart against ischemic injury in myocardial infarction models (Ahmet et al., 2005) and reduces the age-induced inflammation and fibrosis by inhibiting oxidative damage and NF-κB activation (Castello et al., 2010). IF also decreases the diabetes incidence in BB rats (Pedersen et al., 1999) and prevents/reverses different aspects associated to metabolic syndrome by

18

L. Raffaghello and V. Longo

increasing insulin and leptin sensitivity, suppressing inflammation, and stimulating autophagy (Wan et al., 2010). Interestingly, it has been shown that the effects of IF on life span depend on genotype and age of initiation (Goodrick et al., 1990; Kendrick, 1973). In contrast, the use of a FMD for 4 days every 2 weeks started at middle age extends health span, reduces tumor incidence, decreases inflammation, and delays the age-dependent cognitive decline as well as bone loss decline (Brandhorst et al., 2015) In this respect, severe dietary restriction or fasting have been hypothesized to exert beneficial effects in young and middle-age mice, but to be detrimental in older animals that begin to lose weight, according to what has been observed in humans (Brandhorst et al., 2015; Levine et al., 2014). In humans, fasting has been shown to affect certain factors implicated in aging such as insulin and IGF-1, and to cause a fivefold increase of IGF-1 binding protein 1 (IGFBP1) which sequesters IGF-1 from the circulation thus reducing its bioavailability (Fontana et al., 2010; Harvie et al., 2011; Thissen et al., 1994). Notably, the effect of fasting on IGF-1 is due to the combination of protein and CR, but protein intake represents the major regulating factor (Thissen et al., 1994). One of the most evident effects of fasting in humans has been observed in patients affected by rheumatoid arthritis who have clinically significant beneficial long-term effects in term of reduction of pain and inflammation (Muller et al., 2001). However, when normal diet is resumed, most patients relapse unless fasting is followed by a vegetarian diet (Kjeldsen-Kragh et al., 1991). The beneficial effects of fasting have also been observed in subjects affected by hypertension with an improvement of systolic blood pressure below 120 after 13 days of water-only fasting (Goldhamer et al., 2002). Interestingly, a recent case report revealed that a medically supervised 21 days of water-only fasting followed by a diet of minimally processed plant foods free of added sugar, oil, and salt causes tumor regression in a woman affected by lymphoma (Goldhamer et al., 2015). Since fasting is often challenging for individuals affected by pathology, Branhorst et al. developed a FMD to be administered periodically that is low in calories, protein, and sugar (Brandhorst et al., 2015) and which mimics the effects of fasting on markers associated with the stress resistance or longevity, including low levels of glucose and IGF-1 and high levels of ketone bodies and IGFBP-1 (Longo and Mattson, 2014). Bimonthly FMD cycles started at middle age extend the longevity of mice by lowering visceral fat, reducing cancer incidence and skin lesions, rejuvenating the immune system, and

Cancer and Aging Metabolism

19

retarding bone mineral density loss. In old mice, FMD cycles promoted hippocampal neurogenesis, lowered IGF-1 levels and PKA activity, elevated NeuroD1, and improved cognitive performance. In a pilot clinical trial performed in generally healthy adults, three FMD cycles decreased risk factors/biomarkers for aging, diabetes, cardiovascular disease, and cancer without major adverse effects, providing support for the use of FMD to promote health span (Brandhorst et al., 2015). Moreover, preliminary data from a randomized pilot clinical trial conducted to assess the safety and feasibility of the FMD on patients affected by multiple sclerosis showed that the FMD is safe, feasible, and potentially effective by reducing the number of autoimmune lymphocytes (Choi et al., 2016). Among fasting mimetic compounds, polyamine spermidine deserves particular attention since it has been demonstrated to: (i) markedly extend the life span of yeast, flies and worms, and human immune cells; (ii) decrease the oxidative stress; and (iii) exert cardioprotective effects by reducing cardiac hypertrophy and by preserving diastolic function in old mice (Eisenberg et al., 2009, 2016). Interestingly, autophagy, which is able to minimize the functional decline of aged cardiomyocytes by degrading and recycling long-lived damaged proteins as well as dysfunctional mitochondria (Nakai et al., 2007; Taneike et al., 2010), has been shown to be required for spermidinemediated cardioprotection (Eisenberg et al., 2016). In agreement with the latter results, the dietary consumption of sperimidine in humans correlates with reduced blood pressure and a lower incidence of cardiovascular disease (Eisenberg et al., 2016). Of note, spermidine has been also implicated in cancer prevention since it is able to reduce tumor incidence through inhibition of age-associated alteration in DNA methylation status (Soda et al., 2013). Moreover, an elegant recent study demonstrated that spermidine as well as hydroxycitrate, both defined as CR mimetics, improves the inhibition of in vivo tumor growth by chemotherapy (Pietrocola et al., 2016). This effect was only observed for autophagy-competent tumors, depended on the presence of T lymphocytes, and was accompanied by the depletion of regulatory T cells from the tumor bed (Pietrocola et al., 2016).

3.4 Pharmacological Interventions Mimicking CR 3.4.1 Inhibitors of Nutrient-Sensing Pathways Reduced nutrient-sensing pathways including mTOR/S6K, PI3K/AKT, Ras, or AC/PKA signaling, through genetic or pharmacological interventions lead to life span extension in yeast, worm, flies, and mice (Johnson

20

L. Raffaghello and V. Longo

et al., 2013). mTOR kinase is formed by two subunits called TORC1 and TORC2, where the former induces activation of S6K and 4E-BP1. The core components of mTORC1 consist of mTOR, mammalian lethal with sec-13 protein 8 (mLST8), and regulatory-associated protein of TOR (raptor), to which additional proteins including a DEP-domain-containing mTOR-interacting protein (DEPTOR) and proline-rich Akt substrate 40 kDa (PRAS40) are associated. The mTOR complex 2 (mTORC2) core is composed of mTOR, the rapamycin insensitive companion of mTOR (rictor), stress-activated protein kinase-interacting protein 1 (mSIN1), and mLST8. One of the best-known mTOR inhibitors is rapamycin, a pharmacological agent which was initially discovered as an antifungal metabolite and subsequently found to be immunosuppressive by inhibiting S6K1 activation (Chung et al., 1992). In the context of mTOR complex, only mTORC1, which integrates different signals governed by nutrients, growth factors, oxygen, and energy and activates anabolic processes required for cell proliferation and growth, is acutely sensitive to inhibition by rapamycin. Rapamycin binds to the intracellular protein FKBP12 to generate a complex that binds to and destabilized mTORC1 at all times after drug addition, consistent with the capacity of FKBP12-rapamycin to bind to it and weaken the raptor–mTOR interaction (Kim et al., 2002). In contrast, the modulation of mTORC2 by rapamcin is more complex and controversial. Since acute treatment with rapamycin does not perturb mTORC2 signaling and FKBP12-rapamycin cannot bind to intact mTORC2, this complex was originally thought to be rapamycin insensitive (Jacinto et al., 2004; Sarbassov et al., 2004). However, prolonged treatment with rapamycin has been shown to reduce mTORC2 signaling in some, but not all cell types and does so by suppressing mTORC2 assembly (Laplante and Sabatini, 2012; Sarbassov et al., 2006). Rapamycin extends the life span in various model organisms including mice (Harrison et al., 2009; Miller et al., 2005). However, its serious side effects such as metabolic dysregulation (i.e., hyperglycemia, hyperinsulinemia, and insulin resistance) and blockade of hematopoietic cell lineage proliferation hampered its clinical use as an antiaging therapeutic (Soefje et al., 2011). Current clinical trials on healthy older subjects will help to understand whether inhibition of mTOR could be a safe and feasible antiaging intervention. 3.4.2 Inhibitors of Glycolysis Consistent with the metabolic dysregulation associated with aging, i.e., reduction of OXPHOS and increase of glycolysis, there is currently great

Cancer and Aging Metabolism

21

interest in applying pharmacological interventions aimed at inhibiting glucose catabolism and generation. Among these drugs, specific attention has been focused on 2-deoxy-D-glucose (2DG), a glucose analog which, once it is phosphorylated by hexokinase, cannot be further metabolized and consequently blocks glycolysis. Administration of 2DG to rats improves glucose and insulin regulation and increases recovery from stress (Minor et al., 2010). Another compound is an avocado extract called mannoheptulose that inhibits hexokinase and increases the life span in nematodes and mice (Minor et al., 2010). Acarbose, which limits glucose supply to cells by inhibiting α-glucosidase in the intestine, increases the life span of mice (Harrison et al., 2014), and exerts cardioprotective effects in patients affected by diabetes type 2 (Chiasson et al., 2002). 3.4.3 Inhibitors of the GH/IGF-1 Axis Several reports indicate that downregulation of GH/IGF-1 pathway can extend life span in different species (Longo and Finch, 2003). For example, human IGF-1 receptor gene mutations have been found in centenarians (Suh et al., 2008) and decreased serum levels of IGF-1 predict survival in humans with exceptional longevity (Milman et al., 2014). Accordingly, GHR/IGF-1-deficient mice present a lower incidence and delayed occurrence of fatal neoplastic lesions compared with their wild-type littermates (Ikeno et al., 2009; Zhou et al., 1997). Similarly to what it was observed in (GHR/BP) knockout mice, patients affected by GHR-deficient Laron syndrome are characterized by a significant decrease of cancer and diabetes risk (Guevara-Aguirre et al., 2011; Ikeno et al., 2009; Shevah and Laron, 2007; Zhou et al., 1997). According to these results, pharmacological interventions aimed at inhibiting GH/IGF-1 such as monoclonal antibodies and drugs directed against IGF-1R have been used in cancer patients (Carboni et al., 2009). Moreover, several classes of compounds that inhibit GH/IGF-1 axis have been used in patients affected by acromegaly (Giustina et al., 2014). Among the latter drugs, particular attention has been given to somatostatin analogs which lower serum GH and IGF-1 but, unfortunately also reduce other endocrine hormones and can cause serious side effects such as anorexia, diarrhea, and gallstones. For these reasons, the clinical antiaging use of somatostatin analogs is currently poorly understood and pursued. A GHR antagonist currently used in patients affected by acromegaly is pegvisomant which specifically binds to and inhibits GHR (Kopchick et al., 2002; Trainer et al., 2000; van der Lely et al., 2001b). Interestingly, this compound lowers serum IGF-1 levels and blocks the diabetogenic

22

L. Raffaghello and V. Longo

action of GH (Trainer et al., 2000; van der Lely et al., 2001a). However, the very high cost and the need for frequent injections (van der Lely et al., 2012) provide a very tall obstacle for its application as a prolongevity and health span intervention. Finally, an interesting strategy to reduce IGF-1 action is to decrease its availability. In this respect, loss of the protease pregnancy-associated plasma protein-A (PAPP-A) reduces IGF-1 signaling and extends life span in mice while also reducing age-related diseases (Conover, 2013). 3.4.4 Activators of Sirtuins Given their beneficial effects in promoting longevity, sirtuin family proteins are a very interesting drug target. Sirtuin-activating compounds (STACs) including plant-derived metabolites and the well-known resveratrol, represent the first and most potent sirtuin activator and have been shown to extend life span in various organisms (Hubbard and Sinclair, 2014; Pearson et al., 2008). Synthetic activators such as SRT1720 and SRT2104 improve the metabolic profile and extend life span and health span of mice under a high-fat and normal diet (Mitchell et al., 2014; Minor et al., 2011). Interestingly, SRT1720 improves insulin sensitivity, lowers plasma glucose, and increases mitochondrial capacity in experimental diabetes models, thus representing a promising new therapeutic approach for treating age-related diseases such as type 2 diabetes (Milne et al., 2007). Moreover, in rhesus monkeys, under a high-fat and high-sugar diet, resveratrol exerts antiinflammatory effects in visceral white adipose tissue (Jimenez-Gomez et al., 2013). In mice and nonhuman primates fed a high-fat diet, resveratrol protects against the effects of obesity and age-related metabolic decline, increases insulin sensitivity and mitochondrial functions, and prevents liver steatosis (Baur et al., 2006; Fiori et al., 2013; Jimenez-Gomez et al., 2013). In addition, resveratrol delays the onset of neurodegeneration and improves learning and memory in aged mice (Graff et al., 2013; Zhao et al., 2013). The promising results in preclinical models led the clinicians to test resveratrol in humans where it exerts beneficial effects on elderly and obese subjects (Timmers et al., 2011). Furthermore, resveratrol provides positive effects for systolic blood pressure, hemoglobin A1c, and creatinine in patients affected by type 2 diabetes (Hausenblas et al., 2015). Synthetic STACS have also been demonstrated to exert beneficial cardiovascular effects on healthy cigarettes smokers (Venkatasubramanian et al., 2013). The mechanism underlying the beneficial effects of resveratrol is controversial since it has been proposed that the direct activation of sirtuin 1 by

Cancer and Aging Metabolism

23

resveratrol is an in vitro artifact (Beher et al., 2009; Kaeberlein et al., 2005; Pacholec et al., 2010) and that resveratrol works primarily by activating AMPK (Canto et al., 2009), potentially by inhibition of phosphodiesterases (PDE). AMPK may then activate sirtuin 1 indirectly by elevating intracellular levels of its cosubstrate NAD+ (Canto et al., 2009). Alternatively, resveratrol may first activate sirtuin 1 in vivo, leading to AMPK activation via deacetylation and activation of the AMPK kinase LKB1 (Hou et al., 2008; Lan et al., 2008). 3.4.5 Activators of AMPK Pathway AMPK is a serine/threonine kinase which, upon activation by low cellular energy levels, exerts insulin-sensitizing effects resulting in increased glucose uptake in skeletal muscle and fatty acid oxidation in several tissues as well as decreased hepatic glucose production (Ruderman et al., 2013). Different AMPK activators have been developed including biguanides, thiazolidinediones, agonists of glucagon-like peptide-1 receptor, salicylates, and resveratrol (Coughlan et al., 2014). Among biguanides, metformin is a drug used in the first line therapy for type 2 diabetes (Rena et al., 2013) that reduces the risk of cancer and overall mortality (Evans et al., 2005; Franciosi et al., 2013), cardiovascular disease, and possibly cognitive decline (Ng et al., 2014; No author, 1998; Wu et al., 2014a). However, before introducing metformin as an antiaging agent in generally healthy people, further studies to understand the mechanism of action of this compound are necessary. In fact, metformin is also able to inhibit gluconeogenesis, suggesting that, for people under diet-restricted or ketogenic diet, who depend on gluconeogenesis, it might be toxic. According to its prolongevity effects, metformin has been demonstrated to extend the life span of worms and rodents by targeting the folate cycle and methionine metabolism (Anisimov, 2010; Cabreiro et al., 2013; Martin-Montalvo et al., 2013). 3.4.6 Inhibitors of Inflammation Pathways Strategies aimed at decreasing inflammatory pathways can be viewed as candidate target to combat and prevent aging-associated diseases. Various factors contribute to the generation of inflammatory conditions. Some of them are exogenous such as persistent cytomegalovirus infection (Sansoni et al., 2014), while some others are endogeneously produced. The latter include circulating mitochondrial DNA (mtDNA) that is a potent inflammatory stimulus increasing with aging (Pinti et al., 2014), galactosylated N-glycans and proinflammatory micro-RNA (inflammaMIR), all of them potent

24

L. Raffaghello and V. Longo

inflammatory stimuli which increase in circulation with age (Dall’Olio et al., 2013; Olivieri et al., 2013). Approaches targeting one of the pathways that promote age-dependent inflammation, including the de-activation of inflammasomes (Youm et al., 2013), elimination of senescent cells (Tchkonia et al., 2013), and specific dietary restriction regimens have been developed (Berendsen et al., 2013). Initial evidence for the potential of antiinflammatory drugs in retarding aging comes from the effect of nordihydroguaiaretic drugs such as aspirin on life span increase in mice (Strong et al., 2008).

4. GEROSCIENCE AS A STRATEGY TO OPTIMIZE CANCER THERAPY Conventional cancer therapy is mainly based on the use of chemotherapeutics, ionizing radiation, and novel drugs with specific molecular targets, whose major limitation is represented by the toxicity toward healthy cells. Thus, normal cells are damaged and acquire phenotypes similar to those observed during aging. For example, chemotherapy causes mutations and DNA alterations and induces oxidative stress in agreement with what occurs during aging. Moreover, epidemiologic studies have also observed that long-term cancer survivors treated with chemotherapy or radiotherapy have good response in term of tumor regression but often suffer of delayed effects related to cancer treatment including secondary tumors, cardiac pathologies, respiratory complications, infertility, and other chronic disorders. Thus, the discovery of strategies able to generate differential stress resistance (DSR) and sensitization conditions in which tumor cells are sensitized while normal cells are protected against the cytotoxicity of a chemotherapeutic drug is of central importance, particularly as many therapies include multiple chemotherapy drugs or the combination of chemotherapy and non-chemotherapy drugs which can also have adverse effects. Geroscience is a multidisciplinary field whose purpose is the understanding of the association between aging and age-related diseases and the elucidation of the mechanisms underlying cellular damage, protection, and death. Thus, geroscience offers the opportunity to find strategies focused on the protection of healthy cells without interfering with the efficacy of the treatment against malignant cells. In this respect, geroscience and cancer therapy could be considered complementary to each other since the former aims to protect the patient while the second to kill tumor cells (Fig. 3). In the following sections, we will discuss some of the geroscience-based approaches

25

Cancer and Aging Metabolism

em

ot

he

ra py

Radioth erapy

ed get Tar rapy the

ria nd s ho ion toc nt Mi er ve int

Ch

Nutr ientpath sensing way s

Geroscience-based interventions

Cancer therapy

t

Blo gly ck of coly sis

Optimization -Host protection -Tumor sensitization to cytotoxic drug

ng ie sti g d Fa kin ic im m Fasting

Delay/prevention Aging-related diseases

Fig. 3 Geroscience and cancer therapy. Cancer therapy is mainly based on the use of chemotherapeutics, ionizing radiation, and novel drugs with specific molecular targets, whose major limitation is represented by the toxicity toward healthy cells. This is mainly due to their inability to distinguish between healthy and cancer cells. Geroscience, defined as the science of healthy aging, can provide not only strategies able to prevent or delay aging and age-related disorders but also to optimize conventional cancer therapy by protecting the host and sensitizing the cancer cells to the cytotoxic agent. These approaches include fasting and fasting-mimicking diet, mitochondrial interventions, glycolysis blockers, and inhibitors of the nutrient-sensing pathways.

used to generate DSR and stress sensitization conditions in normal and malignant cells, in order to improve the killing of cancer cells, but also to preserve the patient’s healthspan posttreatment.

4.1 Fasting and Fasting-Mimicking Diet Multiple cycles of fasting in tumor-bearing mice can selectively sensitize cancer cells to chemotherapy while protecting normal cells from the associated toxicity (Lee et al., 2012; Raffaghello et al., 2008). These phenomena are known as differential stress sensitization and DSR. Specifically, in healthy cells, fasting was found to activate protective metabolic pathways that confer resistance to a variety of chemotherapeutics (Raffaghello et al., 2008). In contrast, starved cancer cells were found to be unable to turn on such a protective response due to uncontrolled activation of growth promoting signaling cascades by oncogenes and to become even more sensitive to DNA

26

L. Raffaghello and V. Longo

damaging agents (Lee et al., 2012; Safdie et al., 2009, 2012; Shi et al., 2012). The protection of normal cells against chemotherapy-dependent damage was found to be associated to a reduction of IGF-1 and downregulation of protooncogene signals (Lee et al., 2010). In addition to protecting hematopoietic cells from chemotoxicity, multiple cycles of fasting promote hematopoietic stem cell self-renewal to alleviate or reverse the immunosuppression or immunosenescence caused by chemotherapy treatment and aging (Cheng et al., 2014). The latter effect was associated to inhibition of IGF-1 and PKA signaling. More recently, fasting has also been shown to enhance the therapeutic index of chemotreatments by exerting an anti-Warburg effect. In this way, cancer cells are shifted from a glycolytic mode into an uncoupled OXPHOS which promotes increased ROS generation and apoptosis (Bianchi et al., 2015). Preliminary clinical data indicate that fasting in cancer patients is not associated with major adverse effects and may reduce several of the toxic effects of chemotherapy (de Groot et al., 2015; Safdie et al., 2009) including a decreased toxicity to lymphocytes (Cheng et al., 2014). These preliminary clinical data have been collected as part of different pilot Phase I and Phase II clinical trials (NCT01304251, NCT01175837, NCT00936364, and NCT01175837) that demonstrated the safety and efficacy of fasting cycles in combination with chemotherapy in adult oncologic patients. A recent study demonstrated that a FMD is as effective as fasting in killing different cancer cell types (Di Biase et al., 2016). In particular, the combination of chemotherapy and FMD increases the levels of lymphoid progenitor cells and cytotoxic CD8+ tumor-infiltrating lymphocytes, leading to a major delay in breast cancer and melanoma progression. In breast tumors, the mechanism underlying the latter effect involves the downregulation of the stress-protecting enzyme hemeoxygenase-1 (HO-1). These data indicate that cycles of FMD in combination with chemotherapy can enhance T cell-dependent killing of cancer cells through the stimulation of the hematopoietic system and the enhancement of CD8+-dependent tumor cytotoxicity (Di Biase et al., 2016), although it is likely that the initial damage of cancer cells caused by the FMD (Shim et al., 2015) is a key step in the activation of the T cell-dependent immune response. According to these results, back-to-back studies reported that fasting enhances chemotherapy-induced immunosurveillance (Pietrocola et al., 2016). Fasting has also been associated to tyrosine kinase inhibitors (TKI), which represent the most broadly applied cancer therapeutics. However,

Cancer and Aging Metabolism

27

a major limitation of these agents is that their efficacy is short lived since the great majority of patients unavoidably relapse (Gridelli et al., 2014a,b). Thus, strategies that help increase the efficacy of these agents making them more powerful and capable of effectively eradicating cancer cells are warranted. In this respect, fasting has been shown to potentiate the in vivo anticancer effects of various TKI including erlotinib, gefitinib, lapatinib, crizotinib, and regorafenib through inhibition of MAPK kinase signaling and E2F factor transcription (Caffa et al., 2015).

4.2 Glycolysis Blockade As mentioned in the previous chapter, 2-DG is a glucose analog that is phosphorylated by hexokinase to 2-DG-phosphate, which causes a glycolysis blockade. While early studies demonstrated that 2-DG exerts promising anticancer effects in experimental models (Maher et al., 2004), more recent reports show that the mechanisms underlying the activity of 2-DG is heterogeneous and sometimes 2-DG was shown to activate prosurvival pathways in tumor cells (Zhong et al., 2009). Furthermore, the clinical use of 2-DG has been limited by its toxicity upon long-term application (Dwarakanath and Jain, 2009; Landau et al., 1958; Vijayaraghavan et al., 2006). A more promising approach is to combine 2-DG with cytotoxic agents such as chemotherapeutic drugs and ionizing radiation in order to enhance their efficacy (Maschek et al., 2004). In this respect, 2-DG increases the therapeutic index of chemotherapeutic agents in mice-bearing human osteosarcoma and nonsmall cell lung cancer (Maschek et al., 2004). Moreover 2-DG synergizes with etoposide through the induction of immunogenic cell death and in this way increases the life span of immunocompetent tumor-bearing mice (Beneteau et al., 2012). Clearly, the combination of 2-DG with chemotherapy will result in a even higher degree of toxicity, which may be life threatening.

4.3 Nutrient-Sensing Pathway Interventions As described in a previous section, one of the most relevant nutrient-sensing pathway involved in longevity is governed by GH/IGF-1 and glucose which trigger the activation of downstream mTOR/S6K, PI3K/AKT, Ras, and AC/PKA axes, most of them highly conserved from yeast to humans (Fontana et al., 2010). Rapamycin and its analogs (rapalogs) including temsirolimus and everolimus are mTOR inhibitors that have been approved by the FDA for treating different cancers. In addition to their

28

L. Raffaghello and V. Longo

anticancer effects as single agents, rapamycin and rapalogs have been shown to potentiate the efficacy of various cytotoxic agents such chemotherapeutic drugs, ionizing radiation, and proteasome inhibitors against malignant cells through the activation of caspase-dependent apoptosis (Mondesire et al., 2004). In addition, rapamycin and rapalogs prevent epithelial stem cell senescence and protect mice from ionizing radiation-induced side effects (Iglesias-Bartolome et al., 2012). Accordingly, in vitro data indicate that also the combination of rapamacyin and metformin protects normal fibroblasts and epithelial cells against the toxicity of cell-cycle specific chemotherapeutic agents such as mitotic inhibitors (Apontes et al., 2011). However, in vivo rapamycin and other m-TOR inhibitors also cause hyperglycemia which may potentially promote the progression of certain cancers and the sensitization of normal cells to chemotherapy. Metformin has recently emerged as an adjuvant anticancer drug to be used in combination with chemotherapy in order to increase its efficacy and lower the doses. Specifically, metformin has been shown to increase the therapeutic index of various standard chemotherapeutic agents such as paclitaxel, carboplatin, and doxorubicin. This combinatorial effect includes tumor regression and prevention of relapse (Hirsch et al., 2009; Iliopoulos et al., 2011). In agreement with these results, a retrospective analysis of esophageal adenocarcinoma patients under metformin treatment indicated a better response to chemotherapy in subjects treated with metformin compared to those treated with chemotherapy alone (Skinner et al., 2013). The proposed mechanism underlying the chemosensiting effect of metformin against tumor cells is based on downregulation of mTOR pathway and CSC genes (Honjo et al., 2014). In addition to enhancing the efficacy of chemotherapy, metformin has been shown to protect the host against the chemotoxicity by reducing the chemo-induced peripheral neuropathy and sensory deficits (Mao-Ying et al., 2014).

5. CONCLUSIONS Cancer and aging are closely related to each other since they are both associated with damage to DNA and loss of function, but also because aging is a major risk factor for tumorigenesis. In turn, many cancer treatments are contributors to the aging process. From this perspective, geroscience-based approaches appear to be urgent since they provide a potential solution not only for preventing or delaying age-related disorders but also for the optimization and enhancement of conventional cancer treatments. In this

Cancer and Aging Metabolism

29

context, there is a need to educate patients of the long-term side effects caused by standard cancer therapy and to inform them about the strategies used to prevent or reduce such toxicity. In this respect, FMD but also a few drugs including metformin targeting antiaging pathways, or the combination of dietary and pharmacological interventions represent one the most promising approaches that could be incorporated into therapeutic protocols for oncologic patients in order to ameliorate the clinical outcome for these patients with an overall impact on the costs of medical care.

ACKNOWLEDGMENTS L.R. is funded by “Cinque per mille dell’IRPEF–Finanziamento della ricerca sanitaria,” Finanziamento Ricerca Corrente, Ministero Salute. V.L. is funded by NIH grant P01 AG034906 and by Associazione Italiana per la Ricerca sul Cancro (AIRC) 2016 (IG-17605).

REFERENCES Abbas, S., Lugthart, S., Kavelaars, F.G., Schelen, A., Koenders, J.E., Zeilemaker, A., et al., 2010. Acquired mutations in the genes encoding IDH1 and IDH2 both are recurrent aberrations in acute myeloid leukemia: prevalence and prognostic value. Blood 116, 2122–2126. Ables, G.P., Perrone, C.E., Orentreich, D., Orentreich, N., 2012. Methionine-restricted C57BL/6J mice are resistant to diet-induced obesity and insulin resistance but have low bone density. PLoS One 7, e51357. Ahmet, I., Wan, R., Mattson, M.P., Lakatta, E.G., Talan, M., 2005. Cardioprotection by intermittent fasting in rats. Circulation 112, 3115–3121. Akerfelt, M., Morimoto, R.I., Sistonen, L., 2010. Heat shock factors: integrators of cell stress, development and lifespan. Nat. Rev. Mol. Cell Biol. 11, 545–555. Alhazzazi, T.Y., Kamarajan, P., Verdin, E., Kapila, Y.L., 2011. SIRT3 and cancer: tumor promoter or suppressor? Biochim. Biophys. Acta 1816, 80–88. Almeida, A., Bolanos, J.P., Moncada, S., 2010. E3 ubiquitin ligase APC/C-Cdh1 accounts for the Warburg effect by linking glycolysis to cell proliferation. Proc. Natl. Acad. Sci. U.S.A. 107, 738–741. Anisimov, V.N., 2010. Metformin for aging and cancer prevention. Aging (Albany NY) 2, 760–774. Apontes, P., Leontieva, O.V., Demidenko, Z.N., Li, F., Blagosklonny, M.V., 2011. Exploring long-term protection of normal human fibroblasts and epithelial cells from chemotherapy in cell culture. Oncotarget 2, 222–233. Ashraf, N., Zino, S., Macintyre, A., Kingsmore, D., Payne, A.P., George, W.D., et al., 2006. Altered sirtuin expression is associated with node-positive breast cancer. Br. J. Cancer 95, 1056–1061. Astuti, D., Latif, F., Dallol, A., Dahia, P.L., Douglas, F., George, E., et al., 2001. Gene mutations in the succinate dehydrogenase subunit SDHB cause susceptibility to familial pheochromocytoma and to familial paraganglioma. Am. J. Hum. Genet. 69, 49–54. Bardella, C., Pollard, P.J., Tomlinson, I., 2011. SDH mutations in cancer. Biochim. Biophys. Acta 1807, 1432–1443. Barthel, A., Okino, S.T., Liao, J., Nakatani, K., Li, J., Whitlock Jr., J.P., et al., 1999. Regulation of GLUT1 gene transcription by the serine/threonine kinase Akt1. J. Biol. Chem. 274, 20281–20286.

30

L. Raffaghello and V. Longo

Baur, J.A., Pearson, K.J., Price, N.L., Jamieson, H.A., Lerin, C., Kalra, A., et al., 2006. Resveratrol improves health and survival of mice on a high-calorie diet. Nature 444, 337–342. Baysal, B.E., Ferrell, R.E., Willett-Brozick, J.E., Lawrence, E.C., Myssiorek, D., Bosch, A., et al., 2000. Mutations in SDHD, a mitochondrial complex II gene, in hereditary paraganglioma. Science 287, 848–851. Beher, D., Wu, J., Cumine, S., Kim, K.W., Lu, S.C., Atangan, L., et al., 2009. Resveratrol is not a direct activator of SIRT1 enzyme activity. Chem. Biol. Drug Des. 74, 619–624. Bell, E.L., Emerling, B.M., Ricoult, S.J., Guarente, L., 2011. SirT3 suppresses hypoxia inducible factor 1alpha and tumor growth by inhibiting mitochondrial ROS production. Oncogene 30, 2986–2996. Beneteau, M., Zunino, B., Jacquin, M.A., Meynet, O., Chiche, J., Pradelli, L.A., et al., 2012. Combination of glycolysis inhibition with chemotherapy results in an antitumor immune response. Proc. Natl. Acad. Sci. U.S.A. 109, 20071–20076. Bennewith, K.L., Durand, R.E., 2004. Quantifying transient hypoxia in human tumor xenografts by flow cytometry. Cancer Res. 64, 6183–6189. Berendsen, A., Santoro, A., Pini, E., Cevenini, E., Ostan, R., Pietruszka, B., et al., 2013. A parallel randomized trial on the effect of a healthful diet on inflammageing and its consequences in European elderly people: design of the NU-AGE dietary intervention study. Mech. Ageing Dev. 134, 523–530. Berrigan, D., Perkins, S.N., Haines, D.C., Hursting, S.D., 2002. Adult-onset calorie restriction and fasting delay spontaneous tumorigenesis in p53-deficient mice. Carcinogenesis 23, 817–822. Biagi, E., Candela, M., Franceschi, C., Brigidi, P., 2011. The aging gut microbiota: new perspectives. Ageing Res. Rev. 10, 428–429. Bianchi, G., Martella, R., Ravera, S., Marini, C., Capitanio, S., Orengo, A., et al., 2015. Fasting induces anti-Warburg effect that increases respiration but reduces ATP-synthesis to promote apoptosis in colon cancer models. Oncotarget 6, 11806–11819. Blanco, R., Munoz, P., Flores, J.M., Klatt, P., Blasco, M.A., 2007. Telomerase abrogation dramatically accelerates TRF2-induced epithelial carcinogenesis. Genes Dev. 21, 206–220. Bosch-Presegue, L., Vaquero, A., 2011. The dual role of sirtuins in cancer. Genes Cancer 2, 648–662. Bowling, A.C., Mutisya, E.M., Walker, L.C., Price, D.L., Cork, L.C., Beal, M.F., 1993. Age-dependent impairment of mitochondrial function in primate brain. J. Neurochem. 60, 1964–1967. Brandhorst, S., Choi, I.Y., Wei, M., Cheng, C.W., Sedrakyan, S., Navarrete, G., et al., 2015. A periodic diet that mimics fasting promotes multi-system regeneration, enhanced cognitive performance, and health span. Cell Metab. 22, 86–99. Cabreiro, F., Au, C., Leung, K.Y., Vergara-Irigaray, N., Cocheme, H.M., Noori, T., et al., 2013. Metformin retards aging in C. elegans by altering microbial folate and methionine metabolism. Cell 153, 228–239. Caffa, I., D’Agostino, V., Damonte, P., Soncini, D., Cea, M., Monacelli, F., et al., 2015. Fasting potentiates the anticancer activity of tyrosine kinase inhibitors by strengthening MAPK signaling inhibition. Oncotarget 6, 11820–11832. Cairns, R.A., Iqbal, J., Lemonnier, F., Kucuk, C., de Leval, L., Jais, J.P., et al., 2012. IDH2 mutations are frequent in angioimmunoblastic T-cell lymphoma. Blood 119, 1901–1903. Canto, C., Gerhart-Hines, Z., Feige, J.N., Lagouge, M., Noriega, L., Milne, J.C., et al., 2009. AMPK regulates energy expenditure by modulating NAD+ metabolism and SIRT1 activity. Nature 458, 1056–1060. Carboni, J.M., Wittman, M., Yang, Z., Lee, F., Greer, A., Hurlburt, W., et al., 2009. BMS754807, a small molecule inhibitor of insulin-like growth factor-1R/IR. Mol. Cancer Ther. 8, 3341–3349.

Cancer and Aging Metabolism

31

Cardaci, S., Ciriolo, M.R., 2012. TCA cycle defects and cancer: when metabolism tunes redox state. Int. J. Cell Biol. 2012, 161837. Carnero, A., Lleonart, M., 2016. The hypoxic microenvironment: a determinant of cancer stem cell evolution. Bioessays 38 (Suppl. 1), S65–S74. Carvajal-Carmona, L.G., Alam, N.A., Pollard, P.J., Jones, A.M., Barclay, E., Wortham, N., et al., 2006. Adult leydig cell tumors of the testis caused by germline fumarate hydratase mutations. J. Clin. Endocrinol. Metab. 91, 3071–3075. Castello, L., Froio, T., Maina, M., Cavallini, G., Biasi, F., Leonarduzzi, G., et al., 2010. Alternate-day fasting protects the rat heart against age-induced inflammation and fibrosis by inhibiting oxidative damage and NF-kB activation. Free Radic. Biol. Med. 48, 47–54. Cava, E., Fontana, L., 2013. Will calorie restriction work in humans? Aging (Albany NY) 5, 507–514. Chalkiadaki, A., Guarente, L., 2015. The multifaceted functions of sirtuins in cancer. Nat. Rev. Cancer 15, 608–624. Chen, Q., Lesnefsky, E.J., 2006. Depletion of cardiolipin and cytochrome c during ischemia increases hydrogen peroxide production from the electron transport chain. Free Radic. Biol. Med. 40, 976–982. Chen, J.H., Hales, C.N., Ozanne, S.E., 2007. DNA damage, cellular senescence and organismal ageing: causal or correlative? Nucleic Acids Res. 35, 7417–7428. Cheney, K.E., Liu, R.K., Smith, G.S., Meredith, P.J., Mickey, M.R., Walford, R.L., 1983. The effect of dietary restriction of varying duration on survival, tumor patterns, immune function, and body temperature in B10C3F1 female mice. J. Gerontol. 38, 420–430. Cheng, C.W., Adams, G.B., Perin, L., Wei, M., Zhou, X., Lam, B.S., et al., 2014. Prolonged fasting reduces IGF-1/PKA to promote hematopoietic-stem-cell-based regeneration and reverse immunosuppression. Cell Stem Cell 14, 810–823. Chiasson, J.L., Josse, R.G., Gomis, R., Hanefeld, M., Karasik, A., Laakso, M., 2002. Acarbose for prevention of type 2 diabetes mellitus: the STOP-NIDDM randomised trial. Lancet 359, 2072–2077. Choi, I.Y., Piccio, L., Childress, P., Bollman, B., Ghosh, A., Brandhorst, S., et al., 2016. A diet mimicking fasting promotes regeneration and reduces autoimmunity and multiple sclerosis symptoms. Cell Rep. 15, 2136–2146. Chung, J., Kuo, C.J., Crabtree, G.R., Blenis, J., 1992. Rapamycin-FKBP specifically blocks growth-dependent activation of and signaling by the 70 kd S6 protein kinases. Cell 69, 1227–1236. Colman, R.J., Beasley, T.M., Kemnitz, J.W., Johnson, S.C., Weindruch, R., Anderson, R.M., 2014. Caloric restriction reduces age-related and all-cause mortality in rhesus monkeys. Nat. Commun. 5, 3557. Conover, C.A., 2013. Role of PAPP-A in aging and age-related disease. Exp. Gerontol. 48, 612–613. Coughlan, K.A., Valentine, R.J., Ruderman, N.B., Saha, A.K., 2014. AMPK activation: a therapeutic target for type 2 diabetes? Diabetes Metab. Syndr. Obes. 7, 241–253. Dall’Olio, F., Vanhooren, V., Chen, C.C., Slagboom, P.E., Wuhrer, M., Franceschi, C., 2013. N-glycomic biomarkers of biological aging and longevity: a link with inflammaging. Ageing Res. Rev. 12, 685–698. Dang, C.V., Le, A., Gao, P., 2009. MYC-induced cancer cell energy metabolism and therapeutic opportunities. Clin. Cancer Res. 15, 6479–6483. Dang, L., White, D.W., Gross, S., Bennett, B.D., Bittinger, M.A., Driggers, E.M., et al., 2010. Cancer-associated IDH1 mutations produce 2-hydroxyglutarate. Nature 465, 966. de Cabo, R., Carmona-Gutierrez, D., Bernier, M., Hall, M.N., Madeo, F., 2014. The search for antiaging interventions: from elixirs to fasting regimens. Cell 157, 1515–1526. De Carli, E., Wang, X., Puget, S., 2009. IDH1 and IDH2 mutations in gliomas. N. Engl. J. Med. 360, 2248, author reply 2249.

32

L. Raffaghello and V. Longo

de Groot, S., Vreeswijk, M.P., Welters, M.J., Gravesteijn, G., Boei, J.J., Jochems, A., et al., 2015. The effects of short-term fasting on tolerance to (neo) adjuvant chemotherapy in HER2-negative breast cancer patients: a randomized pilot study. BMC Cancer 15, 652. DeBerardinis, R.J., Lum, J.J., Hatzivassiliou, G., Thompson, C.B., 2008. The biology of cancer: metabolic reprogramming fuels cell growth and proliferation. Cell Metab. 7, 11–20. Descamps, O., Riondel, J., Ducros, V., Roussel, A.M., 2005. Mitochondrial production of reactive oxygen species and incidence of age-associated lymphoma in OF1 mice: effect of alternate-day fasting. Mech. Ageing Dev. 126, 1185–1191. Di Biase, S., Lee, C., Brandhorst, S., Manes, B., Buono, R., Cheng, C.W., et al., 2016. Fasting-mimicking diet reduces HO-1 to promote T cell-mediated tumor cytotoxicity. Cancer Cell 30, 136–146. Duan, W., Mattson, M.P., 1999. Dietary restriction and 2-deoxyglucose administration improve behavioral outcome and reduce degeneration of dopaminergic neurons in models of Parkinson’s disease. J. Neurosci. Res. 57, 195–206. Duan, W., Guo, Z., Jiang, H., Ware, M., Li, X.J., Mattson, M.P., 2003. Dietary restriction normalizes glucose metabolism and BDNF levels, slows disease progression, and increases survival in huntingtin mutant mice. Proc. Natl. Acad. Sci. U.S.A. 100, 2911–2916. Dwarakanath, B., Jain, V., 2009. Targeting glucose metabolism with 2-deoxy-D-glucose for improving cancer therapy. Future Oncol. 5, 581–585. Eisenberg, T., Knauer, H., Schauer, A., Buttner, S., Ruckenstuhl, C., Carmona-Gutierrez, D., et al., 2009. Induction of autophagy by spermidine promotes longevity. Nat. Cell Biol. 11, 1305–1314. Eisenberg, T., Abdellatif, M., Schroeder, S., Primessnig, U., Stekovic, S., Pendl, T., et al., 2016. Cardioprotection and lifespan extension by the natural polyamine spermidine. Nat. Med. 22, 1428–1438. Elstrom, R.L., Bauer, D.E., Buzzai, M., Karnauskas, R., Harris, M.H., Plas, D.R., et al., 2004. Akt stimulates aerobic glycolysis in cancer cells. Cancer Res. 64, 3892–3899. Evans, J.M., Donnelly, L.A., Emslie-Smith, A.M., Alessi, D.R., Morris, A.D., 2005. Metformin and reduced risk of cancer in diabetic patients. BMJ 330, 1304–1305. Finley, L.W., Carracedo, A., Lee, J., Souza, A., Egia, A., Zhang, J., et al., 2011. SIRT3 opposes reprogramming of cancer cell metabolism through HIF1alpha destabilization. Cancer Cell 19, 416–428. Fiori, J.L., Shin, Y.K., Kim, W., Krzysik-Walker, S.M., Gonzalez-Mariscal, I., Carlson, O.D., et al., 2013. Resveratrol prevents beta-cell dedifferentiation in nonhuman primates given a high-fat/high-sugar diet. Diabetes 62, 3500–3513. Floegel, A., Pischon, T., 2012. Low carbohydrate-high protein diets. BMJ 344, e3801. Floyd, R.A., West, M., Hensley, K., 2001. Oxidative biochemical markers; clues to understanding aging in long-lived species. Exp. Gerontol. 36, 619–640. Fontana, L., Partridge, L., 2015. Promoting health and longevity through diet: from model organisms to humans. Cell 161, 106–118. Fontana, L., Partridge, L., Longo, V.D., 2010. Extending healthy life span—from yeast to humans. Science 328, 321–326. Fontana, L., Adelaiye, R.M., Rastelli, A.L., Miles, K.M., Ciamporcero, E., Longo, V.D., et al., 2013. Dietary protein restriction inhibits tumor growth in human xenograft models. Oncotarget 4, 2451–2461. Franceschi, C., Campisi, J., 2014. Chronic inflammation (inflammaging) and its potential contribution to age-associated diseases. J. Gerontol. A Biol. Sci. Med. Sci. 69 (Suppl. 1), S4–S9. Franciosi, M., Lucisano, G., Lapice, E., Strippoli, G.F., Pellegrini, F., Nicolucci, A., 2013. Metformin therapy and risk of cancer in patients with type 2 diabetes: systematic review. PLoS One 8, e71583.

Cancer and Aging Metabolism

33

Gao, P., Tchernyshyov, I., Chang, T.C., Lee, Y.S., Kita, K., Ochi, T., et al., 2009. c-Myc suppression of miR-23a/b enhances mitochondrial glutaminase expression and glutamine metabolism. Nature 458, 762–765. Giustina, A., Chanson, P., Kleinberg, D., Bronstein, M.D., Clemmons, D.R., Klibanski, A., et al., 2014. Expert consensus document: a consensus on the medical treatment of acromegaly. Nat. Rev. Endocrinol. 10, 243–248. Goldhamer, A.C., Lisle, D.J., Sultana, P., Anderson, S.V., Parpia, B., Hughes, B., et al., 2002. Medically supervised water-only fasting in the treatment of borderline hypertension. J. Altern. Complement. Med. 8, 643–650. Goldhamer, A.C., Klaper, M., Foorohar, A., Myers, T.R., 2015. Water-only fasting and an exclusively plant foods diet in the management of stage IIIa, low-grade follicular lymphoma. BMJ Case Rep. 2015. Gomes, A.P., Price, N.L., Ling, A.J., Moslehi, J.J., Montgomery, M.K., Rajman, L., et al., 2013. Declining NAD(+) induces a pseudohypoxic state disrupting nuclear-mitochondrial communication during aging. Cell 155, 1624–1638. Goodrick, C.L., Ingram, D.K., Reynolds, M.A., Freeman, J.R., Cider, N., 1990. Effects of intermittent feeding upon body weight and lifespan in inbred mice: interaction of genotype and age. Mech. Ageing Dev. 55, 69–87. Graff, J., Kahn, M., Samiei, A., Gao, J., Ota, K.T., Rei, D., et al., 2013. A dietary regimen of caloric restriction or pharmacological activation of SIRT1 to delay the onset of neurodegeneration. J. Neurosci. 33, 8951–8960. Greer, E.L., Dowlatshahi, D., Banko, M.R., Villen, J., Hoang, K., Blanchard, D., et al., 2007. An AMPK-FOXO pathway mediates longevity induced by a novel method of dietary restriction in C. elegans. Curr. Biol. 17, 1646–1656. Gridelli, C., de Marinis, F., Cappuzzo, F., Di Maio, M., Hirsch, F.R., Mok, T., et al., 2014a. Treatment of advanced non-small-cell lung cancer with epidermal growth factor receptor (EGFR) mutation or ALK gene rearrangement: results of an international expert panel meeting of the Italian Association of Thoracic Oncology. Clin. Lung Cancer 15, 173–181. Gridelli, C., Peters, S., Sgambato, A., Casaluce, F., Adjei, A.A., Ciardiello, F., 2014b. ALK inhibitors in the treatment of advanced NSCLC. Cancer Treat. Rev. 40, 300–306. Guevara-Aguirre, J., Balasubramanian, P., Guevara-Aguirre, M., Wei, M., Madia, F., Cheng, C.W., et al., 2011. Growth hormone receptor deficiency is associated with a major reduction in pro-aging signaling, cancer, and diabetes in humans. Sci. Transl. Med. 3, 70ra13. Haase, V.H., 2009. The VHL tumor suppressor: master regulator of HIF. Curr. Pharm. Des. 15, 3895–3903. Hagen, T.M., Yowe, D.L., Bartholomew, J.C., Wehr, C.M., Do, K.L., Park, J.Y., et al., 1997. Mitochondrial decay in hepatocytes from old rats: membrane potential declines, heterogeneity and oxidants increase. Proc. Natl. Acad. Sci. U.S.A. 94, 3064–3069. Halagappa, V.K., Guo, Z., Pearson, M., Matsuoka, Y., Cutler, R.G., Laferla, F.M., et al., 2007. Intermittent fasting and caloric restriction ameliorate age-related behavioral deficits in the triple-transgenic mouse model of Alzheimer’s disease. Neurobiol. Dis. 26, 212–220. Hamanaka, R.B., Chandel, N.S., 2010. Mitochondrial reactive oxygen species regulate cellular signaling and dictate biological outcomes. Trends Biochem. Sci. 35, 505–513. Harrison, D.E., Strong, R., Sharp, Z.D., Nelson, J.F., Astle, C.M., Flurkey, K., et al., 2009. Rapamycin fed late in life extends lifespan in genetically heterogeneous mice. Nature 460, 392–395. Harrison, D.E., Strong, R., Allison, D.B., Ames, B.N., Astle, C.M., Atamna, H., et al., 2014. Acarbose, 17-alpha-estradiol, and nordihydroguaiaretic acid extend mouse lifespan preferentially in males. Aging Cell 13, 273–282.

34

L. Raffaghello and V. Longo

Harvie, M.N., Pegington, M., Mattson, M.P., Frystyk, J., Dillon, B., Evans, G., et al., 2011. The effects of intermittent or continuous energy restriction on weight loss and metabolic disease risk markers: a randomized trial in young overweight women. Int. J. Obes. (Lond.) 35, 714–727. Hausenblas, H.A., Schoulda, J.A., Smoliga, J.M., 2015. Resveratrol treatment as an adjunct to pharmacological management in type 2 diabetes mellitus—systematic review and meta-analysis. Mol. Nutr. Food Res. 59, 147–159. Heaney, R.P., McCarron, D.A., Dawson-Hughes, B., Oparil, S., Berga, S.L., Stern, J.S., et al., 1999. Dietary changes favorably affect bone remodeling in older adults. J. Am. Diet. Assoc. 99, 1228–1233. Herranz, D., Munoz-Martin, M., Canamero, M., Mulero, F., Martinez-Pastor, B., Fernandez-Capetillo, O., et al., 2010. Sirt1 improves healthy ageing and protects from metabolic syndrome-associated cancer. Nat. Commun. 1, 3. Hirsch, H.A., Iliopoulos, D., Tsichlis, P.N., Struhl, K., 2009. Metformin selectively targets cancer stem cells, and acts together with chemotherapy to block tumor growth and prolong remission. Cancer Res. 69, 7507–7511. Hirschey, M.D., Shimazu, T., Goetzman, E., Jing, E., Schwer, B., Lombard, D.B., et al., 2010. SIRT3 regulates mitochondrial fatty-acid oxidation by reversible enzyme deacetylation. Nature 464, 121–125. Honjo, S., Ajani, J.A., Scott, A.W., Chen, Q., Skinner, H.D., Stroehlein, J., et al., 2014. Metformin sensitizes chemotherapy by targeting cancer stem cells and the mTOR pathway in esophageal cancer. Int. J. Oncol. 45, 567–574. Hou, X., Xu, S., Maitland-Toolan, K.A., Sato, K., Jiang, B., Ido, Y., et al., 2008. SIRT1 regulates hepatocyte lipid metabolism through activating AMP-activated protein kinase. J. Biol. Chem. 283, 20015–20026. Hubbard, B.P., Sinclair, D.A., 2014. Small molecule SIRT1 activators for the treatment of aging and age-related diseases. Trends Pharmacol. Sci. 35, 146–154. Hubbi, M.E., Hu, H., Kshitiz, Gilkes, D.M., Semenza, G.L., 2013. Sirtuin-7 inhibits the activity of hypoxia-inducible factors. J. Biol. Chem. 288, 20768–20775. Iglesias-Bartolome, R., Patel, V., Cotrim, A., Leelahavanichkul, K., Molinolo, A.A., Mitchell, J.B., et al., 2012. mTOR inhibition prevents epithelial stem cell senescence and protects from radiation-induced mucositis. Cell Stem Cell 11, 401–414. Ikeno, Y., Hubbard, G.B., Lee, S., Cortez, L.A., Lew, C.M., Webb, C.R., et al., 2009. Reduced incidence and delayed occurrence of fatal neoplastic diseases in growth hormone receptor/binding protein knockout mice. J. Gerontol. A Biol. Sci. Med. Sci. 64, 522–529. Iliopoulos, D., Hirsch, H.A., Struhl, K., 2011. Metformin decreases the dose of chemotherapy for prolonging tumor remission in mouse xenografts involving multiple cancer cell types. Cancer Res. 71, 3196–3201. Iriondo, O., Rabano, M., Domenici, G., Carlevaris, O., Lopez-Ruiz, J.A., Zabalza, I., et al., 2015. Distinct breast cancer stem/progenitor cell populations require either HIF1alpha or loss of PHD3 to expand under hypoxic conditions. Oncotarget 6, 31721–31739. Jacinto, E., Loewith, R., Schmidt, A., Lin, S., Ruegg, M.A., Hall, A., et al., 2004. Mammalian TOR complex 2 controls the actin cytoskeleton and is rapamycin insensitive. Nat. Cell Biol. 6, 1122–1128. Jeong, S.M., Xiao, C., Finley, L.W., Lahusen, T., Souza, A.L., Pierce, K., et al., 2013. SIRT4 has tumor-suppressive activity and regulates the cellular metabolic response to DNA damage by inhibiting mitochondrial glutamine metabolism. Cancer Cell 23, 450–463. Jimenez-Gomez, Y., Mattison, J.A., Pearson, K.J., Martin-Montalvo, A., Palacios, H.H., Sossong, A.M., et al., 2013. Resveratrol improves adipose insulin signaling and reduces the inflammatory response in adipose tissue of rhesus monkeys on high-fat, high-sugar diet. Cell Metab. 18, 533–545.

Cancer and Aging Metabolism

35

Johnson, J.E., Johnson, F.B., 2014. Methionine restriction activates the retrograde response and confers both stress tolerance and lifespan extension to yeast, mouse and human cells. PLoS One 9, e97729. Johnson, S.C., Rabinovitch, P.S., Kaeberlein, M., 2013. mTOR is a key modulator of ageing and age-related disease. Nature 493, 338–345. Kaeberlein, M., McDonagh, T., Heltweg, B., Hixon, J., Westman, E.A., Caldwell, S.D., et al., 2005. Substrate-specific activation of sirtuins by resveratrol. J. Biol. Chem. 280, 17038–17045. Kang, M.J., Kim, H.J., Kim, H.K., Lee, J.Y., Kim, D.H., Jung, K.J., et al., 2005. The effect of age and calorie restriction on HIF-1-responsive genes in aged liver. Biogerontology 6, 27–37. Kapahi, P., Chen, D., Rogers, A.N., Katewa, S.D., Li, P.W., Thomas, E.L., et al., 2010. With TOR, less is more: a key role for the conserved nutrient-sensing TOR pathway in aging. Cell Metab. 11, 453–465. Kapitsinou, P.P., Haase, V.H., 2008. The VHL tumor suppressor and HIF: insights from genetic studies in mice. Cell Death Differ. 15, 650–659. Keith, B., Simon, M.C., 2007. Hypoxia-inducible factors, stem cells, and cancer. Cell 129, 465–472. Kendrick, D.C., 1973. The effects of infantile stimulation and intermittent fasting and feeding on life span in the black-hooded rat. Dev. Psychobiol. 6, 225–234. Kim, D.H., Sarbassov, D.D., Ali, S.M., King, J.E., Latek, R.R., Erdjument-Bromage, H., et al., 2002. mTOR interacts with raptor to form a nutrient-sensitive complex that signals to the cell growth machinery. Cell 110, 163–175. Kim, H.S., Patel, K., Muldoon-Jacobs, K., Bisht, K.S., Aykin-Burns, N., Pennington, J.D., et al., 2010. SIRT3 is a mitochondria-localized tumor suppressor required for maintenance of mitochondrial integrity and metabolism during stress. Cancer Cell 17, 41–52. Kim, H.S., Vassilopoulos, A., Wang, R.H., Lahusen, T., Xiao, Z., Xu, X., et al., 2011. SIRT2 maintains genome integrity and suppresses tumorigenesis through regulating APC/C activity. Cancer Cell 20, 487–499. Kjeldsen-Kragh, J., Haugen, M., Borchgrevink, C.F., Laerum, E., Eek, M., Mowinkel, P., et al., 1991. Controlled trial of fasting and one-year vegetarian diet in rheumatoid arthritis. Lancet 338, 899–902. Knudson Jr., A.G., 1971. Mutation and cancer: statistical study of retinoblastoma. Proc. Natl. Acad. Sci. U.S.A. 68, 820–823. Kopchick, J.J., Parkinson, C., Stevens, E.C., Trainer, P.J., 2002. Growth hormone receptor antagonists: discovery, development, and use in patients with acromegaly. Endocr. Rev. 23, 623–646. Kretsch, M.J., Green, M.W., Fong, A.K., Elliman, N.A., Johnson, H.L., 1997. Cognitive effects of a long-term weight reducing diet. Int. J. Obes. Relat. Metab. Disord. 21, 14–21. Krycer, J.R., Sharpe, L.J., Luu, W., Brown, A.J., 2010. The Akt-SREBP nexus: cell signaling meets lipid metabolism. Trends Endocrinol. Metab. 21, 268–276. Kwong, L.K., Sohal, R.S., 1998. Substrate and site specificity of hydrogen peroxide generation in mouse mitochondria. Arch. Biochem. Biophys. 350, 118–126. Kwong, L.K., Sohal, R.S., 2000. Age-related changes in activities of mitochondrial electron transport complexes in various tissues of the mouse. Arch. Biochem. Biophys. 373, 16–22. Lagiou, P., Sandin, S., Lof, M., Trichopoulos, D., Adami, H.O., Weiderpass, E., 2012. Low carbohydrate-high protein diet and incidence of cardiovascular diseases in Swedish women: prospective cohort study. BMJ 344, e4026. Lagouge, M., Larsson, N.G., 2013. The role of mitochondrial DNA mutations and free radicals in disease and ageing. J. Intern. Med. 273, 529–543.

36

L. Raffaghello and V. Longo

Lan, F., Cacicedo, J.M., Ruderman, N., Ido, Y., 2008. SIRT1 modulation of the acetylation status, cytosolic localization, and activity of LKB1. Possible role in AMP-activated protein kinase activation. J. Biol. Chem. 283, 27628–27635. Landau, B.R., Laszlo, J., Stengle, J., Burk, D., 1958. Certain metabolic and pharmacologic effects in cancer patients given infusions of 2-deoxy-D-glucose. J. Natl. Cancer Inst. 21, 485–494. Laplante, M., Sabatini, D.M., 2012. mTOR signaling in growth control and disease. Cell 149, 274–293. Lee, C., Safdie, F.M., Raffaghello, L., Wei, M., Madia, F., Parrella, E., et al., 2010. Reduced levels of IGF-I mediate differential protection of normal and cancer cells in response to fasting and improve chemotherapeutic index. Cancer Res. 70, 1564–1572. Lee, C., Raffaghello, L., Brandhorst, S., Safdie, F.M., Bianchi, G., Martin-Montalvo, A., et al., 2012. Fasting cycles retard growth of tumors and sensitize a range of cancer cell types to chemotherapy. Sci. Transl. Med. 4, 124ra27. Lehtonen, H.J., Kiuru, M., Ylisaukko-Oja, S.K., Salovaara, R., Herva, R., Koivisto, P.A., et al., 2006. Increased risk of cancer in patients with fumarate hydratase germline mutation. J. Med. Genet. 43, 523–526. Leiser, S.F., Kaeberlein, M., 2010. The hypoxia-inducible factor HIF-1 functions as both a positive and negative modulator of aging. Biol. Chem. 391, 1131–1137. Levine, M.E., Suarez, J.A., Brandhorst, S., Balasubramanian, P., Cheng, C.W., Madia, F., et al., 2014. Low protein intake is associated with a major reduction in IGF-1, cancer, and overall mortality in the 65 and younger but not older population. Cell Metab. 19, 407–417. Li, Z., Bao, S., Wu, Q., Wang, H., Eyler, C., Sathornsumetee, S., et al., 2009. Hypoxia-inducible factors regulate tumorigenic capacity of glioma stem cells. Cancer Cell 15, 501–513. Li, W., Li, X., Miller, R.A., 2014. ATF4 activity: a common feature shared by many kinds of slow-aging mice. Aging Cell 13, 1012–1018. Li, N., Feng, L., Liu, H., Wang, J., Kasembeli, M., Tran, M.K., et al., 2016. PARP inhibition suppresses growth of EGFR-mutant cancers by targeting nuclear PKM2. Cell Rep. 15, p843–p856. Lim, J.H., Lee, Y.M., Chun, Y.S., Chen, J., Kim, J.E., Park, J.W., 2010. Sirtuin 1 modulates cellular responses to hypoxia by deacetylating hypoxia-inducible factor 1alpha. Mol. Cell 38, 864–878. Longo, V.D., Finch, C.E., 2003. Evolutionary medicine: from dwarf model systems to healthy centenarians? Science 299, 1342–1346. Longo, V.D., Fontana, L., 2010. Calorie restriction and cancer prevention: metabolic and molecular mechanisms. Trends Pharmacol. Sci. 31, 89–98. Longo, V.D., Mattson, M.P., 2014. Fasting: molecular mechanisms and clinical applications. Cell Metab. 19, 181–192. Longo, V.D., Ellerby, L.M., Bredesen, D.E., Valentine, J.S., Gralla, E.B., 1997. Human Bcl-2 reverses survival defects in yeast lacking superoxide dismutase and delays death of wild-type yeast. J. Cell Biol. 137, 1581–1588. Longo, V.D., Antebi, A., Bartke, A., Barzilai, N., Brown-Borg, H.M., Caruso, C., et al., 2015. Interventions to slow aging in humans: are we ready? Aging Cell 14, 497–510. Luo, W., Hu, H., Chang, R., Zhong, J., Knabel, M., O’Meally, R., et al., 2011. Pyruvate kinase M2 is a PHD3-stimulated coactivator for hypoxia-inducible factor 1. Cell 145, 732–744. Madia, F., Wei, M., Yuan, V., Hu, J., Gattazzo, C., Pham, P., et al., 2009. Oncogene homologue Sch9 promotes age-dependent mutations by a superoxide and Rev1/ Polzeta-dependent mechanism. J. Cell Biol. 186, 509–523.

Cancer and Aging Metabolism

37

Maher, J.C., Krishan, A., Lampidis, T.J., 2004. Greater cell cycle inhibition and cytotoxicity induced by 2-deoxy-D-glucose in tumor cells treated under hypoxic vs aerobic conditions. Cancer Chemother. Pharmacol. 53, 116–122. Mair, W., Piper, M.D., Partridge, L., 2005. Calories do not explain extension of life span by dietary restriction in Drosophila. PLoS Biol. 3, e223. Mantovani, A., Allavena, P., Sica, A., Balkwill, F., 2008. Cancer-related inflammation. Nature 454, 436–444. Mao-Ying, Q.L., Kavelaars, A., Krukowski, K., Huo, X.J., Zhou, W., Price, T.J., et al., 2014. The anti-diabetic drug metformin protects against chemotherapy-induced peripheral neuropathy in a mouse model. PLoS One 9, e100701. Martin-Montalvo, A., Villalba, J.M., Navas, P., de Cabo, R., 2011. NRF2, cancer and calorie restriction. Oncogene 30, 505–520. Martin-Montalvo, A., Mercken, E.M., Mitchell, S.J., Palacios, H.H., Mote, P.L., Scheibye-Knudsen, M., et al., 2013. Metformin improves healthspan and lifespan in mice. Nat. Commun. 4, 2192. Maschek, G., Savaraj, N., Priebe, W., Braunschweiger, P., Hamilton, K., Tidmarsh, G.F., et al., 2004. 2-Deoxy-D-glucose increases the efficacy of adriamycin and paclitaxel in human osteosarcoma and non-small cell lung cancers in vivo. Cancer Res. 64, 31–34. Mattison, J.A., Roth, G.S., Beasley, T.M., Tilmont, E.M., Handy, A.M., Herbert, R.L., et al., 2012. Impact of caloric restriction on health and survival in rhesus monkeys from the NIA study. Nature 489, 318–321. Maxwell, P.H., Wiesener, M.S., Chang, G.W., Clifford, S.C., Vaux, E.C., Cockman, M.E., et al., 1999. The tumour suppressor protein VHL targets hypoxia-inducible factors for oxygen-dependent proteolysis. Nature 399, 271–275. Menendez, J.A., Alarcon, T., Joven, J., 2014. Gerometabolites: the pseudohypoxic aging side of cancer oncometabolites. Cell Cycle 13, 699–709. Menssen, A., Hydbring, P., Kapelle, K., Vervoorts, J., Diebold, J., Luscher, B., et al., 2012. The c-MYC oncoprotein, the NAMPT enzyme, the SIRT1-inhibitor DBC1, and the SIRT1 deacetylase form a positive feedback loop. Proc. Natl. Acad. Sci. U.S.A. 109, E187–E196. Miller, R.A., Buehner, G., Chang, Y., Harper, J.M., Sigler, R., Smith-Wheelock, M., 2005. Methionine-deficient diet extends mouse lifespan, slows immune and lens aging, alters glucose, T4, IGF-I and insulin levels, and increases hepatocyte MIF levels and stress resistance. Aging Cell 4, 119–125. Mills, E.L., Kelly, B., Logan, A., Costa, A.S., Varma, M., Bryant, C.E., et al., 2016. Succinate dehydrogenase supports metabolic repurposing of mitochondria to drive inflammatory macrophages. Cell 167, 457–470, e13. Milman, S., Atzmon, G., Huffman, D.M., Wan, J., Crandall, J.P., Cohen, P., et al., 2014. Low insulin-like growth factor-1 level predicts survival in humans with exceptional longevity. Aging Cell 13, 769–771. Milne, J.C., Lambert, P.D., Schenk, S., Carney, D.P., Smith, J.J., Gagne, D.J., et al., 2007. Small molecule activators of SIRT1 as therapeutics for the treatment of type 2 diabetes. Nature 450, 712–716. Minor, R.K., Smith Jr., D.L., Sossong, A.M., Kaushik, S., Poosala, S., Spangler, E.L., et al., 2010. Chronic ingestion of 2-deoxy-D-glucose induces cardiac vacuolization and increases mortality in rats. Toxicol. Appl. Pharmacol. 243, 332–339. Minor, R.K., Baur, J.A., Gomes, A.P., Ward, T.M., Csiszar, A., Mercken, E.M., et al., 2011. SRT1720 improves survival and healthspan of obese mice. Sci. Rep. 1, 70. Mirisola, M.G., Taormina, G., Fabrizio, P., Wei, M., Hu, J., Longo, V.D., 2014. Serine- and threonine/valine-dependent activation of PDK and Tor orthologs converge on Sch9 to promote aging. PLoS Genet. 10, e1004113.

38

L. Raffaghello and V. Longo

Mitchell, S.J., Martin-Montalvo, A., Mercken, E.M., Palacios, H.H., Ward, T.M., Abulwerdi, G., et al., 2014. The SIRT1 activator SRT1720 extends lifespan and improves health of mice fed a standard diet. Cell Rep. 6, 836–843. Mondesire, W.H., Jian, W., Zhang, H., Ensor, J., Hung, M.C., Mills, G.B., et al., 2004. Targeting mammalian target of rapamycin synergistically enhances chemotherapy-induced cytotoxicity in breast cancer cells. Clin. Cancer Res. 10, 7031–7042. Muller, H., de Toledo, F.W., Resch, K.L., 2001. Fasting followed by vegetarian diet in patients with rheumatoid arthritis: a systematic review. Scand. J. Rheumatol. 30, 1–10. Muller-Hocker, J., 1989. Cytochrome-c-oxidase deficient cardiomyocytes in the human heart—an age-related phenomenon. A histochemical ultracytochemical study. Am. J. Pathol. 134, 1167–1173. Nakai, A., Yamaguchi, O., Takeda, T., Higuchi, Y., Hikoso, S., Taniike, M., et al., 2007. The role of autophagy in cardiomyocytes in the basal state and in response to hemodynamic stress. Nat. Med. 13, 619–624. Ng, T.P., Feng, L., Yap, K.B., Lee, T.S., Tan, C.H., Winblad, B., 2014. Long-term metformin usage and cognitive function among older adults with diabetes. J. Alzheimers Dis. 41, 61–68. No author, 1998. Effect of intensive blood-glucose control with metformin on complications in overweight patients with type 2 diabetes (UKPDS 34). UK Prospective Diabetes Study (UKPDS) Group. Lancet 352, 854–865. Oberdoerffer, P., Michan, S., McVay, M., Mostoslavsky, R., Vann, J., Park, S.K., et al., 2008. SIRT1 redistribution on chromatin promotes genomic stability but alters gene expression during aging. Cell 135, 907–918. Ogunshola, O.O., Antoniou, X., 2009. Contribution of hypoxia to Alzheimer’s disease: is HIF-1alpha a mediator of neurodegeneration? Cell. Mol. Life Sci. 66, 3555–3563. Ojaimi, J., Masters, C.L., McLean, C., Opeskin, K., McKelvie, P., Byrne, E., 1999. Irregular distribution of cytochrome c oxidase protein subunits in aging and Alzheimer’s disease. Ann. Neurol. 46, 656–660. Olivieri, F., Rippo, M.R., Monsurro, V., Salvioli, S., Capri, M., Procopio, A.D., et al., 2013. MicroRNAs linking inflamm-aging, cellular senescence and cancer. Ageing Res. Rev. 12, 1056–1068. Ostan, R., Lanzarini, C., Pini, E., Scurti, M., Vianello, D., Bertarelli, C., et al., 2015. Inflammaging and cancer: a challenge for the Mediterranean diet. Nutrients 7, 2589–2621. Pacholec, M., Bleasdale, J.E., Chrunyk, B., Cunningham, D., Flynn, D., Garofalo, R.S., et al., 2010. SRT1720, SRT2183, SRT1460, and resveratrol are not direct activators of SIRT1. J. Biol. Chem. 285, 8340–8351. Patel, N.V., Gordon, M.N., Connor, K.E., Good, R.A., Engelman, R.W., Mason, J., et al., 2005. Caloric restriction attenuates Abeta-deposition in Alzheimer transgenic models. Neurobiol. Aging 26, 995–1000. Pearson, K.J., Baur, J.A., Lewis, K.N., Peshkin, L., Price, N.L., Labinskyy, N., et al., 2008. Resveratrol delays age-related deterioration and mimics transcriptional aspects of dietary restriction without extending life span. Cell Metab. 8, 157–168. Pedersen, C.R., Hagemann, I., Bock, T., Buschard, K., 1999. Intermittent feeding and fasting reduces diabetes incidence in BB rats. Autoimmunity 30, 243–250. Petersen, K.F., Dufour, S., Befroy, D., Garcia, R., Shulman, G.I., 2004. Impaired mitochondrial activity in the insulin-resistant offspring of patients with type 2 diabetes. N. Engl. J. Med. 350, 664–671. Pietrocola, F., Pol, J., Vacchelli, E., Rao, S., Enot, D.P., Baracco, E.E., et al., 2016. Caloric restriction mimetics enhance anticancer immunosurveillance. Cancer Cell 30, 147–160.

Cancer and Aging Metabolism

39

Pinti, M., Cevenini, E., Nasi, M., De Biasi, S., Salvioli, S., Monti, D., et al., 2014. Circulating mitochondrial DNA increases with age and is a familiar trait: implications for “inflamm-aging” Eur. J. Immunol. 44, 1552–1562. Piper, M.D., Partridge, L., 2007. Dietary restriction in Drosophila: delayed aging or experimental artefact? PLoS Genet. 3, e57. Ptitsyn, A., Hulver, M., Cefalu, W., York, D., Smith, S.R., 2006. Unsupervised clustering of gene expression data points at hypoxia as possible trigger for metabolic syndrome. BMC Genomics 7, 318. Raffaghello, L., Lee, C., Safdie, F.M., Wei, M., Madia, F., Bianchi, G., et al., 2008. Starvation-dependent differential stress resistance protects normal but not cancer cells against high-dose chemotherapy. Proc. Natl. Acad. Sci. U.S.A. 105, 8215–8220. Rena, G., Pearson, E.R., Sakamoto, K., 2013. Molecular mechanism of action of metformin: old or new insights? Diabetologia 56, 1898–1906. Richie Jr., J.P., Leutzinger, Y., Parthasarathy, S., Malloy, V., Orentreich, N., Zimmerman, J.A., 1994. Methionine restriction increases blood glutathione and longevity in F344 rats. FASEB J. 8, 1302–1307. Robey, R.B., Hay, N., 2006. Mitochondrial hexokinases, novel mediators of the antiapoptotic effects of growth factors and Akt. Oncogene 25, 4683–4696. Rohrbach, S., Teichert, S., Niemann, B., Franke, C., Katschinski, D.M., 2008. Caloric restriction counteracts age-dependent changes in prolyl-4-hydroxylase domain (PHD) 3 expression. Biogerontology 9, 169–176. Ruckenstuhl, C., Netzberger, C., Entfellner, I., Carmona-Gutierrez, D., Kickenweiz, T., Stekovic, S., et al., 2014. Lifespan extension by methionine restriction requires autophagy-dependent vacuolar acidification. PLoS Genet. 10, e1004347. Ruderman, N.B., Carling, D., Prentki, M., Cacicedo, J.M., 2013. AMPK, insulin resistance, and the metabolic syndrome. J. Clin. Invest. 123, 2764–2772. Safdie, F.M., Dorff, T., Quinn, D., Fontana, L., Wei, M., Lee, C., et al., 2009. Fasting and cancer treatment in humans: a case series report. Aging (Albany NY) 1, 988–1007. Safdie, F., Brandhorst, S., Wei, M., Wang, W., Lee, C., Hwang, S., et al., 2012. Fasting enhances the response of glioma to chemo- and radiotherapy. PLoS One 7, e44603. Sanchez-Roman, I., Barja, G., 2013. Regulation of longevity and oxidative stress by nutritional interventions: role of methionine restriction. Exp. Gerontol. 48, 1030–1042. Sansoni, P., Vescovini, R., Fagnoni, F.F., Akbar, A., Arens, R., Chiu, Y.L., et al., 2014. New advances in CMV and immunosenescence. Exp. Gerontol. 55, 54–62. Sarbassov, D.D., Ali, S.M., Kim, D.H., Guertin, D.A., Latek, R.R., Erdjument-Bromage, H., et al., 2004. Rictor, a novel binding partner of mTOR, defines a rapamycin-insensitive and raptor-independent pathway that regulates the cytoskeleton. Curr. Biol. 14, 1296–1302. Sarbassov, D.D., Ali, S.M., Sengupta, S., Sheen, J.H., Hsu, P.P., Bagley, A.F., et al., 2006. Prolonged rapamycin treatment inhibits mTORC2 assembly and Akt/PKB. Mol. Cell 22, 159–168. Saunders, L.R., Verdin, E., 2007. Sirtuins: critical regulators at the crossroads between cancer and aging. Oncogene 26, 5489–5504. Sciacovelli, M., Goncalves, E., Johnson, T.I., Zecchini, V.R., da Costa, A.S., Gaude, E., et al., 2016. Fumarate is an epigenetic modifier that elicits epithelial-to-mesenchymal transition. Nature 537, 544–547. Sebastian, C., Zwaans, B.M., Silberman, D.M., Gymrek, M., Goren, A., Zhong, L., et al., 2012. The histone deacetylase SIRT6 is a tumor suppressor that controls cancer metabolism. Cell 151, 1185–1199. Selak, M.A., Armour, S.M., MacKenzie, E.D., Boulahbel, H., Watson, D.G., Mansfield, K.D., et al., 2005. Succinate links TCA cycle dysfunction to oncogenesis by inhibiting HIF-alpha prolyl hydroxylase. Cancer Cell 7, 77–85.

40

L. Raffaghello and V. Longo

Semenza, G.L., 2010. HIF-1: upstream and downstream of cancer metabolism. Curr. Opin. Genet. Dev. 20, 51–56. Shackelford, D.B., Shaw, R.J., 2009. The LKB1-AMPK pathway: metabolism and growth control in tumour suppression. Nat. Rev. Cancer 9, 563–575. Shevah, O., Laron, Z., 2007. Patients with congenital deficiency of IGF-I seem protected from the development of malignancies: a preliminary report. Growth Horm. IGF Res. 17, 54–57. Shi, Y., Felley-Bosco, E., Marti, T.M., Orlowski, K., Pruschy, M., Stahel, R.A., 2012. Starvation-induced activation of ATM/Chk2/p53 signaling sensitizes cancer cells to cisplatin. BMC Cancer 12, 571. Shim, H.S., Wei, M., Brandhorst, S., Longo, V.D., 2015. Starvation promotes REV1 SUMOylation and p53-dependent sensitization of melanoma and breast cancer cells. Cancer Res. 75, 1056–1067. Singh, R., Lakhanpal, D., Kumar, S., Sharma, S., Kataria, H., Kaur, M., et al., 2012. Late-onset intermittent fasting dietary restriction as a potential intervention to retard age-associated brain function impairments in male rats. Age (Dordr.) 34, 917–933. Singh, R., Manchanda, S., Kaur, T., Kumar, S., Lakhanpal, D., Lakhman, S.S., et al., 2015. Middle age onset short-term intermittent fasting dietary restriction prevents brain function impairments in male Wistar rats. Biogerontology 16, 775–788. Skinner, H.D., McCurdy, M.R., Echeverria, A.E., Lin, S.H., Welsh, J.W., O’Reilly, M.S., et al., 2013. Metformin use and improved response to therapy in esophageal adenocarcinoma. Acta Oncol. 52, 1002–1009. Soda, K., Kano, Y., Chiba, F., Koizumi, K., Miyaki, Y., 2013. Increased polyamine intake inhibits age-associated alteration in global DNA methylation and 1,2dimethylhydrazine-induced tumorigenesis. PLoS One 8, e64357. Soefje, S.A., Karnad, A., Brenner, A.J., 2011. Common toxicities of mammalian target of rapamycin inhibitors. Target. Oncol. 6, 125–129. Solon-Biet, S.M., McMahon, A.C., Ballard, J.W., Ruohonen, K., Wu, L.E., Cogger, V.C., et al., 2014. The ratio of macronutrients, not caloric intake, dictates cardiometabolic health, aging, and longevity in ad libitum-fed mice. Cell Metab. 19, 418–430. Strong, R., Miller, R.A., Astle, C.M., Floyd, R.A., Flurkey, K., Hensley, K.L., et al., 2008. Nordihydroguaiaretic acid and aspirin increase lifespan of genetically heterogeneous male mice. Aging Cell 7, 641–650. Suh, Y., Atzmon, G., Cho, M.O., Hwang, D., Liu, B., Leahy, D.J., et al., 2008. Functionally significant insulin-like growth factor I receptor mutations in centenarians. Proc. Natl. Acad. Sci. U.S.A. 105, 3438–3442. Suh, D.H., Kim, M.K., No, J.H., Chung, H.H., Song, Y.S., 2011. Metabolic approaches to overcoming chemoresistance in ovarian cancer. Ann. N. Y. Acad. Sci. 1229, 53–60. Sun, Q., Chen, X., Ma, J., Peng, H., Wang, F., Zha, X., et al., 2011. Mammalian target of rapamycin up-regulation of pyruvate kinase isoenzyme type M2 is critical for aerobic glycolysis and tumor growth. Proc. Natl. Acad. Sci. U.S.A. 108, 4129–4134. Taneike, M., Yamaguchi, O., Nakai, A., Hikoso, S., Takeda, T., Mizote, I., et al., 2010. Inhibition of autophagy in the heart induces age-related cardiomyopathy. Autophagy 6, 600–606. Tannenbaum, A., Silverstone, H., 1949. The influence of the degree of caloric restriction on the formation of skin tumors and hepatomas in mice. Cancer Res. 9, 724–727. Tchkonia, T., Zhu, Y., van Deursen, J., Campisi, J., Kirkland, J.L., 2013. Cellular senescence and the senescent secretory phenotype: therapeutic opportunities. J. Clin. Invest. 123, 966–972. Thissen, J.P., Ketelslegers, J.M., Underwood, L.E., 1994. Nutritional regulation of the insulin-like growth factors. Endocr. Rev. 15, 80–101.

Cancer and Aging Metabolism

41

Timmers, S., Konings, E., Bilet, L., Houtkooper, R.H., van de Weijer, T., Goossens, G.H., et al., 2011. Calorie restriction-like effects of 30 days of resveratrol supplementation on energy metabolism and metabolic profile in obese humans. Cell Metab. 14, 612–622. Tomlinson, I.P., Alam, N.A., Rowan, A.J., Barclay, E., Jaeger, E.E., Kelsell, D., et al., 2002. Germline mutations in FH predispose to dominantly inherited uterine fibroids, skin leiomyomata and papillary renal cell cancer. Nat. Genet. 30, 406–410. Trainer, P.J., Drake, W.M., Katznelson, L., Freda, P.U., Herman-Bonert, V., van der Lely, A.J., et al., 2000. Treatment of acromegaly with the growth hormone-receptor antagonist pegvisomant. N. Engl. J. Med. 342, 1171–1177. Trounce, I., Byrne, E., Marzuki, S., 1989. Decline in skeletal muscle mitochondrial respiratory chain function: possible factor in ageing. Lancet 1, 637–639. van der Lely, A.J., Hutson, R.K., Trainer, P.J., Besser, G.M., Barkan, A.L., Katznelson, L., et al., 2001a. Long-term treatment of acromegaly with pegvisomant, a growth hormone receptor antagonist. Lancet 358, 1754–1759. van der Lely, A.J., Muller, A., Janssen, J.A., Davis, R.J., Zib, K.A., Scarlett, J.A., et al., 2001b. Control of tumor size and disease activity during cotreatment with octreotide and the growth hormone receptor antagonist pegvisomant in an acromegalic patient. J. Clin. Endocrinol. Metab. 86, 478–481. van der Lely, A.J., Biller, B.M., Brue, T., Buchfelder, M., Ghigo, E., Gomez, R., et al., 2012. Long-term safety of pegvisomant in patients with acromegaly: comprehensive review of 1288 subjects in ACROSTUDY. J. Clin. Endocrinol. Metab. 97, 1589–1597. Van Remmen, H., Richardson, A., 2001. Oxidative damage to mitochondria and aging. Exp. Gerontol. 36, 957–968. Vander Heiden, M.G., Cantley, L.C., Thompson, C.B., 2009. Understanding the Warburg effect: the metabolic requirements of cell proliferation. Science 324, 1029–1033. Venkatasubramanian, S., Noh, R.M., Daga, S., Langrish, J.P., Joshi, N.V., Mills, N.L., et al., 2013. Cardiovascular effects of a novel SIRT1 activator, SRT2104, in otherwise healthy cigarette smokers. J. Am. Heart Assoc. 2, e000042. Verdin, E., 2015. NAD(+) in aging, metabolism, and neurodegeneration. Science 350, 1208–1213. Vijayaraghavan, R., Kumar, D., Dube, S.N., Singh, R., Pandey, K.S., Bag, B.C., et al., 2006. Acute toxicity and cardio-respiratory effects of 2-deoxy-D-glucose: a promising radio sensitiser. Biomed. Environ. Sci. 19, 96–103. Villeda, S.A., Luo, J., Mosher, K.I., Zou, B., Britschgi, M., Bieri, G., et al., 2011. The ageing systemic milieu negatively regulates neurogenesis and cognitive function. Nature 477, 90–94. Wan, R., Ahmet, I., Brown, M., Cheng, A., Kamimura, N., Talan, M., et al., 2010. Cardioprotective effect of intermittent fasting is associated with an elevation of adiponectin levels in rats. J. Nutr. Biochem. 21, 413–417. Wang, R., Wang, L., Li, Y., Hu, H., Shen, L., Shen, X., et al., 2014. FGFR1/3 tyrosine kinase fusions define a unique molecular subtype of non-small cell lung cancer. Clin. Cancer Res. 20, 4107–4114. Ward, P.S., Thompson, C.B., 2012. Metabolic reprogramming: a cancer hallmark even Warburg did not anticipate. Cancer Cell 21, 297–308. Webb, A.E., Brunet, A., 2014. FOXO transcription factors: key regulators of cellular quality control. Trends Biochem. Sci. 39, 159–169. Wei, M., Fabrizio, P., Hu, J., Ge, H., Cheng, C., Li, L., et al., 2008. Life span extension by calorie restriction depends on Rim15 and transcription factors downstream of Ras/PKA, Tor, and Sch9. PLoS Genet. 4, e13. Wise, D.R., DeBerardinis, R.J., Mancuso, A., Sayed, N., Zhang, X.Y., Pfeiffer, H.K., et al., 2008. Myc regulates a transcriptional program that stimulates mitochondrial

42

L. Raffaghello and V. Longo

glutaminolysis and leads to glutamine addiction. Proc. Natl. Acad. Sci. U.S.A. 105, 18782–18787. Wu, Z., Liu, S.Q., Huang, D., 2013. Dietary restriction depends on nutrient composition to extend chronological lifespan in budding yeast Saccharomyces cerevisiae. PLoS One 8, e64448. Wu, J.W., Boudreau, D.M., Park, Y., Simonds, N.I., Freedman, A.N., 2014a. Commonly used diabetes and cardiovascular medications and cancer recurrence and cancer-specific mortality: a review of the literature. Expert Opin. Drug Saf. 13, 1071–1099. Wu, L.E., Gomes, A.P., Sinclair, D.A., 2014b. Geroncogenesis: metabolic changes during aging as a driver of tumorigenesis. Cancer Cell 25, 12–19. Yang, Y.A., Han, W.F., Morin, P.J., Chrest, F.J., Pizer, E.S., 2002. Activation of fatty acid synthesis during neoplastic transformation: role of mitogen-activated protein kinase and phosphatidylinositol 3-kinase. Exp. Cell Res. 279, 80–90. Yang, M., Soga, T., Pollard, P.J., 2013. Oncometabolites: linking altered metabolism with cancer. J. Clin. Invest. 123, 3652–3658. Yen, K.E., Bittinger, M.A., Su, S.M., Fantin, V.R., 2010. Cancer-associated IDH mutations: biomarker and therapeutic opportunities. Oncogene 29, 6409–6417. Youm, Y.H., Grant, R.W., McCabe, L.R., Albarado, D.C., Nguyen, K.Y., Ravussin, A., et al., 2013. Canonical Nlrp3 inflammasome links systemic low-grade inflammation to functional decline in aging. Cell Metab. 18, 519–532. Zhao, Y.N., Li, W.F., Li, F., Zhang, Z., Dai, Y.D., Xu, A.L., et al., 2013. Resveratrol improves learning and memory in normally aged mice through microRNA-CREB pathway. Biochem. Biophys. Res. Commun. 435, 597–602. Zhong, D., Xiong, L., Liu, T., Liu, X., Liu, X., Chen, J., et al., 2009. The glycolytic inhibitor 2-deoxyglucose activates multiple prosurvival pathways through IGF1R. J. Biol. Chem. 284, 23225–23233. Zhong, L., D’Urso, A., Toiber, D., Sebastian, C., Henry, R.E., Vadysirisack, D.D., et al., 2010. The histone deacetylase Sirt6 regulates glucose homeostasis via Hif1alpha. Cell 140, 280–293. Zhou, Y., Xu, B.C., Maheshwari, H.G., He, L., Reed, M., Lozykowski, M., et al., 1997. A mammalian model for Laron syndrome produced by targeted disruption of the mouse growth hormone receptor/binding protein gene (the Laron mouse). Proc. Natl. Acad. Sci. U.S.A. 94, 13215–13220.

CHAPTER TWO

Cellular and Molecular Mechanisms of Autoimmunity and Lupus Nephritis S.K. Devarapu*, G. Lorenz†, O.P. Kulkarni{, H.-J. Anders*, S.R. Mulay*,1 *Medizinische Klinik und Poliklinik IV, Klinikum der Universit€at M€ unchen, Munich, Germany † Klinikum rechts der Isar, Abteilung f€ ur Nephrologie, Technische Universit€at M€ unchen, Munich, Germany { BITS-Pilani Hyderabad Campus, Hyderabad, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. The Phenomenon of Immune Tolerance 2.1 Central Tolerance 2.2 Peripheral Tolerance 3. Factors That Influence the Loss of Immune Tolerance During Autoimmunity 3.1 Genetic Factors and Autoimmunity 3.2 Environmental Factors and Autoimmunity 4. Factors That Induce Autoimmunity 4.1 Epigenetics and Transcription Factors 4.2 Extracellular Vesicles 4.3 Neutrophil Extracellular Traps 4.4 Ion Channels 4.5 Lipids 5. Costimulatory and Coinhibitory Pathways in Autoimmunity 5.1 Costimulatory Pathways 5.2 Coinhibitory Pathways 6. PRRs in Autoimmunity 6.1 PRRs and Autoimmunity 7. Tissue Inflammation and Injury in Autoimmunity 7.1 Immune Complexes 7.2 Lymphocytes 7.3 Monocytes and Macrophages 7.4 Tertiary Lymphoid Organs 8. Genetic Risk Factors for Organ Manifestations in Human Autoimmune Diseases 9. Lupus Nephritis 9.1 Systemic Autoimmunity in SLE 9.2 Autoimmunity and Tissue Inflammation Inside the Kidney 9.3 Animal Models for SLE

International Review of Cell and Molecular Biology, Volume 332 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2016.12.001

#

2017 Elsevier Inc. All rights reserved.

44 46 46 48 53 53 56 59 59 66 70 74 82 84 84 88 90 91 97 97 99 101 102 103 105 106 109 111

43

44

S.K. Devarapu et al.

10. Summary Acknowledgments References

114 115 115

Abstract Autoimmunity involves immune responses directed against self, which are a result of defective self/foreign distinction of the immune system, leading to proliferation of self-reactive lymphocytes, and is characterized by systemic, as well as tissue-specific, inflammation. Numerous mechanisms operate to ensure the immune tolerance to self-antigens. However, monogenetic defects or genetic variants that weaken immune tolerance render susceptibility to the loss of immune tolerance, which is further triggered by environmental factors. In this review, we discuss the phenomenon of immune tolerance, genetic and environmental factors that influence the immune tolerance, factors that induce autoimmunity such as epigenetic and transcription factors, neutrophil extracellular trap formation, extracellular vesicles, ion channels, and lipid mediators, as well as costimulatory or coinhibitory molecules that contribute to an autoimmune response. Further, we discuss the cellular and molecular mechanisms of autoimmune tissue injury and inflammation during systemic lupus erythematosus and lupus nephritis.

1. INTRODUCTION Autoimmunity implies immune responses that are directed against the self. It is usually considered a pathological process that should be avoided by a clear self/foreign distinction of the immune system. Historically, this perspective originates from clinical syndromes of organ destruction by noninfectious triggers for which certain autoantigens and autoantibodies could be identified. However, bioassays used to detect such autoantibodies often display their persistent presence also in healthy people or their transient presence upon infections that provide an unspecific stimulus to clonal lymphocyte expansion—for example, a transient increase of cryoglobulins after mycoplasma infection or persistent levels of low-affinity antinuclear antibodies. Indeed, autoimmunity is a common biological phenomenon that does not always cause a disease. Complex organisms need to maintain their integrity in response to all sorts of threats. For example, threats by infectious organisms require particular host defense mechanisms such as intact barriers, secretory molecules, or local inflammatory responses, referred to as innate immunity. The molecular

Mechanisms of Autoimmunity

45

mechanisms of innate immunity have raised considerable attention since the discovery of the Toll-like receptors (TLRs), and the last decade has much increased our knowledge about how infectious organisms alert the immune system in an antigen-independent manner. It is also of note that the vast majority of the past and present species on this planet entirely rely on the innate immune system for host defense. The evolution of the adaptive immune system introduced a completely new way of immune activation that relies on “antigens,” small supramolecular structures of peptides, and lipid or nucleic acid complexes that are presented to the host’s effector cell repertoire. The way in which priming of adaptive immune responses and imprinting of immune memory evolved it holds the risk for misinterpretations in terms of self-foreign discrimination. This was not a new problem. Errors in self-foreign discrimination also exist at the level of the innate immune system and can contribute to considerable tissue destruction, e.g., in sterile forms of inflammation, where danger-associated molecular patterns (DAMPs) activate TLRs to initiate unnecessary inflammation causing additional tissue injury. However, innate immunity-related errors in self-foreign discrimination do not imprint any immune memory. The numerous mechanisms of immune tolerance assure that potentially autoreactive elements of the adaptive immune system are kept to a minimum and hardly activated. However, the genetic variability of the population implies that some people are able to maintain immune tolerance better than others. In the end, autoimmunity presents like most other noncommunicable diseases. Most people do not experience autoimmune diseases during a lifetime. Very few individuals suffer from monogenetic defects of immune tolerance and experience autoimmune disease early in life. However, a small part of the population carries unfortunate combinations of genetic variants that considerably weaken immune tolerance at different levels, which, eventually triggered by environmental factors, primes an immune response and potentially immune memory upon presentation of an autoantigen. This chapter will describe in detail the molecular and cellular mechanisms of immune tolerance and autoimmunity. The presentation is focused on the understanding of diseases in general and may show additional features in specific autoimmune diseases. A detailed description of the pathogenesis of all the different kinds of autoimmune diseases is beyond the scope of this chapter, but it should prepare the reader well for further studying more specific literature.

46

S.K. Devarapu et al.

2. THE PHENOMENON OF IMMUNE TOLERANCE The immune system identifies and mounts a prompt response to eliminate foreign/nonself-antigens while abstaining the harmful response to self-antigens. This inherent feature of the immune system has been termed as immune tolerance (Burnet and Fenner, 1949; Jerne, 2004). Broadly, immune tolerance can be divided into two categories, viz., natural or self-tolerance and inducible tolerance. Natural or self-tolerance is further subclassified based on the anatomical sites into central and peripheral tolerance (Fig. 1).

2.1 Central Tolerance Of the various immune tolerance methods, central tolerance is the primary event occurring at developmental stages of both T and B cells in the thymus and bone marrow, respectively. The mechanisms of central tolerance involve clonal deletion, clonal anergy, and receptor editing to eliminate autoreactive lymphocytes at maximal efficacy (Fig. 1). 2.1.1 T-Cell Tolerance To achieve tolerance to self-antigens, T lymphocytes undergo two levels of selection processes. After completion of their antigen receptor editing in the thymus, immature T cells with the ability to recognize self-MHC molecule are exposed to self-peptides bound to MHC molecule. Cells that show high affinity for self-peptides MHC complexes are eliminated by clonal deletion, thus negatively selecting the cells with no or low affinity. Thymic dendritic cells (DCs) and cortical and medullary epithelial cells can induce elimination of self-reactive T cells until they reach either CD4+ or CD8+ single-cell stage resulting over 95% of T cells generated in the thymus die by apoptosis (Kappler et al., 1987). The self-antigen-driven thymic B-cell class switching promotes T-cell central tolerance by presenting cognate self-antigens to support the negative selection of CD4+ T cells (Perera et al., 2016). Efficient negative selection is achieved by the display of self-peptides and expression of responsible genes, which are controlled by a transcriptional factor called the autoimmune regulator (AIRE) (Peterson et al., 2008). Medullary thymic epithelial cells (mTECs) selectively express AIRE and drive tissue-specific antigen (TSA) expression to induce negative selection of TSA autoreactive T cells (Anderson et al., 2002). Moreover, CD80hi MHC-IIhi mTECs expressed AIRE and controlled ectopic antigen

Fig. 1 Mechanisms of immune tolerance. The natural immune tolerance is divided into either central or peripheral immune tolerance depending on the anatomical sites in the body. The central tolerance occurs at developmental stages of both T and B cells in the thymus and bone marrow, respectively. The mechanisms of central tolerance involve clonal deletion and either TCR or BCR receptor editing to achieve maximum efficiency to eliminate the autoreactive lymphocytes. The peripheral tolerance primarily occurs in the secondary lymphoid organs such spleens and lymph nodes. The mechanisms of peripheral tolerance involve anergy, immune deviation, immune regulation, and network-mediated suppression, as well as immune privilege.

48

S.K. Devarapu et al.

expression (Gray et al., 2007). AIRE-deficient humans and mice developed autoimmune polyendocrinopathy-candidiasis-ectodermal dystrophy (APS-1 or APECED), suggesting the involvement of AIRE in the central tolerance (Kyewski and Derbinski, 2004). Furthermore, expression of AIRE is also observed in peripheral lymphoid organs such as spleen and lymph nodes, known as extrathymic AIRE-expressing cells (eTACs) (Gardner et al., 2008). High expression levels of MHC-II molecules by eTACs allow them to interact with CD4+ T cells in an intrinsic and extrinsic manner as well as to maintain central and peripheral immune tolerance (Bour-Jordan et al., 2011; Nurieva et al., 2011). 2.1.2 B-Cell Tolerance In the bone marrow, B cells develop tolerance to self-antigens by means of receptor editing and clonal deletion in order to minimize the risk of autoimmune diseases. In general, B cells upon encountering antigens undergo affinity maturation and produce high-affinity antibodies that eventually avoid the possibility of cross-reactivity with self-antigens. Receptor editing is an essential mechanism that contributes to central tolerance (Tiegs et al., 1993). It is achieved by inducing VDJ recombinase and rearrangements, which produce new Ig light chains (edited) with the ability to modify the specificity of the receptor so that it can no longer recognize self-antigens (Tiegs et al., 1993) (Fig. 1). How does the autoreactive human B cells undergo tolerance was recently addressed by Lang et al. using a humanized mouse model, i.e., mice expressing an antihuman Igκ membrane protein to serve as a ubiquitous neo-self-antigen were transplanted with a human immune system (Lang et al., 2016). They followed the fate of self-reactive human κ + B cells relative to nonautoreactive λ + cells and showed that tolerance of human B cells occurs in the bone marrow via combination of receptor editing and clonal deletion. In addition, they reported that a number of available self-antigens and the genetics of the cord blood donor dictate the levels of central tolerance of the autoreactive B cells in the periphery (Lang et al., 2016).

2.2 Peripheral Tolerance The mechanisms of central immune tolerance operate at the early developmental stages of both T and B cells. However, in certain circumstances, self-antigens are not expressed during early life but rather during the adult life. In addition, certain antigens that are produced in nonlymphoid organs

Mechanisms of Autoimmunity

49

are not exposed sufficiently at the primary lymphoid organs and, therefore, are missed during primary tolerance. Such self-antigens might pose a potential threat to the host if not restricted and, hence, must be controlled outside the primary lymphoid organs. These mechanisms are referred to as peripheral tolerance. Peripheral tolerance is maintained primarily in the secondary lymphoid organs such spleens and lymph nodes. Several mechanisms are involved in the regulation of peripheral tolerance such as immune deviation, immune regulation and immune privilege, network-mediated suppression, and co-receptor modulation (Fig. 1). 2.2.1 B- and T-Cell Anergy Although receptor editing and clonal deletion are the primary mechanisms of central tolerance, the maintenance of tolerance is a much more complicated process. The theory of clonal anergy has been evaluated in the maintenance of tolerance. Clonal anergy was first observed in B cells that functionally rest in the nonresponsive state (Nossal and Pike, 1980). In the tolerant animals, anergic lymphocytes have a lower capacity to proliferate and secrete only low levels of antibody upon antigenic or mitogenic stimuli. Moreover, anergic B cells display a reduced life span with delayed deletion state (Goodnow et al., 2009) and a reduced expression of surface immunoglobulin (Ig) M. In addition, impaired signal transduction is attributed as one of the main reasons for the B-cell anergy (Healy et al., 1997). Such anergic B cells have elevated basal levels of intracellular calcium; however, failure to increase it further upon antigenic stimulation due to the increased requirement of B cell-activating factor of TNF family (BAFF) is for the survival of these cells. Hence, anergic B cells are compromised in mounting immune responses owing to the limited/reduced lifespan and impaired signal transduction. Furthermore, T cells also display anergy, although involving different mechanisms compared to B cells. T cells that lack the secondary costimulatory signal undergo anergic state even in the presence of an antigenic signal. Self-antigens presented by the thymic epithelial cells lack secondary signal compelling the reactive T cells into the nonresponsive anergic state. These anergic T cells are incapable of proliferating as well as producing growth factors such as interleukin (IL)-2. As a central theme, cell cycle mediators fail to activate the progression from G1 to S phase in anergic T cells (Subramanian et al., 2006). The biochemical substances such as diacylglycerol, as well as IL-2, play a major role in T-cell anergy (Olenchock et al., 2006; Spitaler et al., 2006; Zha et al., 2006).

50

S.K. Devarapu et al.

2.2.2 Immune Deviation Upon activation by antigen-presenting cells (APCs), naı¨ve T helper (TH) cells in the peripheral lymphoid tissues differentiate into either TH1 or TH2 effector helper cells, which can be distinguished by the cytokines they secrete. For example, TH1 cells secrete IL-2, interferon (IFN) γ, and tumor necrosis factor (TNF) α. They are involved in the macrophage activation, defend intracellular pathogens, and stimulate B cells to secrete specific subclasses of IgG antibodies that can coat extracellular microbes and activate complement (Alberts et al., 2002). TH2 cells secrete IL-4, IL-5, IL-10, and IL-13. They are involved in the macrophage inactivation, respond to extracellular pathogens, and stimulate B cells to secrete antibodies including IgE and some subclasses of IgG that bind to mast cells, basophils, and eosinophils (Alberts et al., 2002). Recent evidence indicates that preferential activation of T-cell subsets can be a mechanism of tolerance induction. For example, administration of bee venom to NZB/W F1 mice increased CD4+CD25+ Treg cells and delayed the development of lupus nephritis (LN) (Lee et al., 2011), whereas exogenous IL-10 administration inhibited corneal allograft rejection by inducing TGFβ and TH1/TH2 deviation (Li et al., 2014). Furthermore, the TH1/TH2 immune deviation was demonstrated to facilitate islet allograft tolerance in mice (Zhang et al., 2010). 2.2.3 Immune Regulation/Suppression The adoptive transfer of lymphocytes from tolerant animals to naive recipients can induce tolerance in recipients. Such an inducible tolerance was thought to be mediated by suppressor and cytotoxic T cells (Bloom et al., 1992); however, eventually, these cells were identified as regulatory T cells (Tregs). 2.2.3.1 Regulatory T Cells

Tregs are produced in the thymus to maintain immune homeostasis. In addition, they also contribute to immune tolerance due to their ability to suppress the function of other T cells. Tregs are essential in monitoring the immune system and compromising their number or function could result in several autoimmune diseases—for example, multiple sclerosis, rheumatoid arthritis, or type 1 diabetes (Chinen et al., 2010; Kim et al., 2007; Kravchenko et al., 2016; Long and Buckner, 2011; Parackova et al., 2016). Phenotypically, Tregs are identified as CD4+CD25+FoxP3+ T cells, of which Foxp3 is a master regulator shown to drive the regulatory

Mechanisms of Autoimmunity

51

activity of Tregs in both humans and mice (Hadaschik et al., 2015). Several studies have explored the functions of Foxp3+ T cells in the immune response. Adoptive transfer of CD4+CD25+FoxP3+ T cells protected FoxP3-deficient mice from the development of autoimmune disease (Hori et al., 2003), in vivo IL-2/anti-IL-2 complex-induced Tregs protected against chronic kidney disease (Polhill et al., 2012). Furthermore, piperlongumine treatment increased the proportion of Tregs and protected MRL/lpr mice from LN (Yao et al., 2014). Additionally, FoxP3
regulatory T cells has also been reported to contribute to immune tolerance—for example, Zohar et al. reported that CXCL-11-dependent induction of FoxP3
regulatory T cells suppressed murine autoimmune encephalomyelitis (Karin and Wildbaum, 2015; Zohar et al., 2014). FoxP3+ Treg cells exert their regulatory function by secreting various immunoregulatory cytokines including IL-9, IL-35, IL-10, and TGFβ that suppress the activity of nearby T cells and APCs by downregulating surface expression of CD80 or CD86 (Wing and Sakaguchi, 2014). Tregs induce the upregulation intracellular cyclic AMP, leading to the inhibition of T-cell proliferation and IL-2 production. Tregs are classified into two groups based on their origin and function into natural Tregs (nTregs) and inducible Tregs (iTregs). The nTregs are generated in the thymus, whereas the iTreg cells develop outside the thymus in various mucosa-associated lymphoid tissues (MALTs). 2.2.3.2 Natural Tregs

Thymus-derived CD4+ cells with high levels of CD25 expression together with the transcription factor FoxP3 (CD4+CD25+FoxP3+) were identified as nTregs. They form approximately 5%–10% of the total CD4+ T-cell population (Walker et al., 2003) and are positively selected thymocytes with a relatively high affinity for self-antigens. The T-cell receptor (TCR) and MHC II with self-peptide interaction signal the development of nTreg in the thymic stroma. Briefly, thymocytes with high affinity for self-antigens undergo clonal deletion, while a small fraction of these thymocytes escape the selection barrier and appear as nTregs moving toward the periphery to patrol along the secondary lymphoid organs. Multiple mechanisms are involved in mediating the immune regulation by nTregs—for example, inhibition of APC’s ability to activate T cells, direct cytotoxicity of T cells, as well as secretion of antiinflammatory cytokines such as IL-10 and TGFβ, which further inhibits T-cell activity. Several recent studies describe the involvement of nTregs and iTregs in various

52

S.K. Devarapu et al.

disease conditions—for example, lung allergic responses and autoimmune thyroiditis in mice (Joetham et al., 2016; Kong et al., 2015; Lin et al., 2013; Metzker et al., 2016). 2.2.3.3 Induced Tregs

Thymic single-positive CD4+ T cells differentiate into CD25+- and FoxP3+-expressing iTregs (CD4+FoxP3+) in the presence of TGFβ, IL-10, and IL-4. The iTregs mature in peripheral sites—for example, MALT, and exert suppressive function by secreting IL-10 and TGFβ, inducing cell cycle arrest or apoptosis in effector T cells, and blocking costimulation and maturation of DCs. Depending on the cytokine secretion, iTregs identified as T regulatory 1 (Tr1) cells that secrete IL-10 and T helper 3 (Th3) cells that secrete TGFβ. Though Tr1 cells do not express FoxP3, their properties in vitro are very similar to those of FoxP3+ Tregs (Yao et al., 2015). It is reported that in multiple sclerosis patients, memory T cells can be induced to iTreg phenotype and their development and function are precursor dependent (Mohiuddin et al., 2016). 2.2.4 Immune Privilege Sites Certain anatomical regions in the body are more favorable for grafting than others since they are able to tolerate the presence of allogeneic and xenogeneic tissues (antigens) without eliciting an inflammatory immune response. Such sites are called immune privileged sites, implying that in these regions when foreign antigens are introduced, the zone does not mount an autoimmune response. One of the reasons for this nonresponsive state is the effective sequestering of the immune cells in these zones. Several areas have been identified to function as immune privileged sites in the body—for example, the eye, brain, uterus, testis, and the fetus in the pregnant females. Classical experiments done by Medawar PB proved this concept, in which it was believed that tolerance is due to the limited access of lymphocytes to these sites and failure of foreign antigen transportation to secondary lymphoid organs preventing immune response initiation (Medawar, 1948). Immune privilege is achieved by several mechanisms. The major mechanisms operating the immune tolerance are lymphatic drainage and physical barrier between blood and tissues. In addition, the absence or minimal expression of MHC class Ia protein on cells of the eye and brain, which is required for the cytotoxic cells activity, also plays an important role in immune tolerance (Niederkorn, 2012). The constitutive expression of CD95/Fas ligand (FasL) and TRAIL (Griffith et al., 1995) in these sites

Mechanisms of Autoimmunity

53

contributes to the immune tolerance by inducing apoptosis of infiltrating lymphocytes, as well as other inflammatory cells, upon their entry into these sites. Furthermore, high expression levels of the antiinflammatory cytokines such as TNFβ and migration inhibitory factor (MIF) contribute to the immune tolerance by inhibiting NK cell-mediated cytolytic activities (Solomos and Rall, 2016; Taylor, 2016).

3. FACTORS THAT INFLUENCE THE LOSS OF IMMUNE TOLERANCE DURING AUTOIMMUNITY 3.1 Genetic Factors and Autoimmunity Genetic variation influences the immune tolerance and autoimmune disease outcomes. The mechanisms that induce genetic variations include sexual reproduction, mutation, migration, random genetic drift, recombination, and natural selection (Ramos et al., 2015). The recombination events mimic the natural selection shaping the diversity of human genome as well as increasing the risk of genetic diseases. For example, lymphocytes achieve their surface receptor diversity by the genomic alterations that occur either(1) at the primary lymphoid organs via somatic VDJ recombination of both B-cell receptors (BCRs) and TCRs or (2) at the secondary lymphoid organs via somatic hypermutations substituting single nucleotides of BCR during the late phase of immune response (Ignatowicz et al., 1996; Laufer et al., 1996; Wardemann et al., 2003). Aberrations in these recombination events lead to autoimmune diseases. Thus, by shaping the diversity of human genome natural selection consistently checks immune function genes and pathways. Many genes have been reported to be associated with autoimmune diseases, for example, Bim, Zap70, Cblb, Ctla4, Fas, and Roquin. How these genes contribute to the loss of immune tolerance and induce autoimmune diseases? When the interaction of immature B cells with self-antigens exceed certain thresholds, they internalize the BCR and halt the maturation process (Hartley et al., 1993). This results in downregulation of CD62 ligand (CD62L) (Hartley et al., 1993) and BAFF (B cell-activating factor) receptors (Mackay et al., 2003) and, however, continues the expression of Rag1 and Rag2, which are required for VDJ recombination for receptor editing (Jankovic et al., 2004; Nemazee and Hogquist, 2003). Failure of receptor editing leads to apoptosis of B cells with self-reactive BCR in either bone marrow or spleen (Hartley et al., 1993). BCL-2-interacting mediator (BIM) of cells death induces B-cell deletion; hence, Bim-deficient mice

54

S.K. Devarapu et al.

show higher autoantibody production as well as spontaneous autoimmunity (Strasser and Bouillet, 2003). Furthermore, TCR activation by self-peptide/ MHC complex in thymic cortical regions induces positive selection. The positively selected T cells move to the thymic medulla to test TCRs for self-reactivity. Medullary thymic epithelial cells and DCs express B7.1 (CD80) and other costimulatory molecules along with self-peptide/MHC (Palmer, 2003). The TCRs with a strong affinity for self-peptide/MHC molecule undergo cell death. Several molecules have been identified that involve the regulation of self-reactive TCR—for example, ZAP70 (ζ-chain-associated protein kinase of 70 kDa) (Sakaguchi et al., 2003), GRB2 (growth-factor-receptor-bound protein 2) (Gong et al., 2001), and MINK (misshapen-Nck-interacting kinase-related kinase) (McCarty et al., 2005). In addition, BIM, FAS, and Nur77 play essential roles in the autoreactive TCR regulation (Strasser and Bouillet, 2003; Zhou et al., 1996). Furthermore, limitations in central tolerance (of both clonal deletion and receptor editing) are backed by peripheral tolerance with various intrinsic genetic elements. These elements increase BCR threshold irrespective of BCR specificity as well as induce apoptosis of those self-reactive B cells. This involves the recruitment of tyrosine phosphatase SHP1 (SH2-domain-containing protein tyrosine phosphatase 1) and SHIP (SH2-domain-containing inositol-5-phosphatase) to the activated BCR and induction of CD5 receptor that regulates B-cell anergy (Healy and Goodnow, 1998; Hippen et al., 2000; Ravetch and Lanier, 2000). In T cells, CTLA4 (cytotoxic T-lymphocyte antigen 4) induces a high threshold of TCR self-reactivity and inhibits T-cell activation and CTLA4 deficiency or variants of the CTLA4 resulted in autoimmunity in humans and mice (Inobe and Schwartz, 2004; Ueda et al., 2003; Walker and Abbas, 2002). Moreover, TCR internalization is enhanced by ubiquitin ligases such as CBL-B, GRAIL, and ITCH tag (Anandasabapathy et al., 2003; Jeon et al., 2004). Therefore, deficiency of any of these ligases resulted in an autoimmune disease in mice (Liu, 2004; Naramura et al., 2002; Yokoi et al., 2002). These mechanisms are summarized in Fig. 2. Although single-gene defects very rarely lead to autoimmune diseases, single-gene mutations in murine models were employed to understand how such defects in immune system cause autoimmunity. For example, alike experimental DNAse-1 deletion, analysis of Dnase-1-knockout mice revealed that normal DNAse-1 activity protects mice from the anti-DNA autoimmune response (Jacob et al., 2002; Wilber et al., 2003). Similarly, deficiency of caspase-activated DNase resulted in pristane-induced murine

Mechanisms of Autoimmunity

55

Fig. 2 Factors that influence the immune tolerance–autoimmunity. Breakdown of immune tolerance results in autoimmunity that involves multiple factors ranging from genetic to the environmental origin. For example, mutations and/or polymorphisms in genes such as Aire, Bim, Zap70, Cblb, Ctla4, Fas, and Roquin are linked to the development of autoimmune diseases in animal models as well as in humans. Other factors that contribute to the breakdown of immune tolerance include upregulation in the surface expression of costimulatory molecules, molecular mimicry by antigens/ adjuvants, the presence of superantigens or modification of proteins into self-antigens, and defective clearance of dead cells exposing the intracellular autoantigens that promote an immune response and, thus, induce autoimmunity.

lupus-like disease owing to the impaired clearance of dead cells in these mice (Frisoni et al., 2007). Deficiency of DNAse-2, that degrades extracellular chromatin, developed a chronic polyarthritis resembling human rheumatoid arthritis in mice (Kawane et al., 2006). Null mutations and hypomorphic variants of the secreted deoxyribonuclease DNASE1L3 are also linked to systemic lupus erythematosus (SLE) since they digest the extracellular microparticle-associated chromatin (Sisirak et al., 2016). Moreover, mice deficient in either NADPH oxidase 2 or rubicon, molecules involved in

56

S.K. Devarapu et al.

LC3-associated phagocytosis (LAP), a form of noncanonical autophagy that remove dying cells, displayed symptoms of autoinflammatory, lupus-like diseases with kidney pathology (Martinez et al., 2016). In addition, protein kinase C delta deficiency was observed in autoimmune lymphoproliferative syndrome as well as SLE in humans (Belot et al., 2013; Kuehn et al., 2013; Salzer et al., 2013). NADPH oxidase deficiency also renders susceptibility to experimental allergic encephalomyelitis by regulating TH lineage commitment to TH17 in mice (Tse et al., 2010). Defects in the NADPH oxidase enzyme also lead to chronic granulomatous disease that involves recurrent life-threatening infections with bacteria and fungi as well as dysregulated inflammatory mechanisms (Rosenzweig, 2008). Moreover, patients with selective IgA deficiency have a greater risk of concomitant autoimmune diseases (Abolhassani et al., 2015). In addition, combined immunodeficiencies are associated with autoimmune phenomena (Schuetz et al., 2010). For example, (1) common variable immunodeficiency (CVID), which is caused by mutations in the transmembrane activator and calcium-modulating cyclophilin ligand interactor (TACI), is associated with X-linked Agammaglobulinemia and autoimmune cytopenia (Cunningham-Rundles, 2008); (2) Wiskott–Aldrich syndrome, which is caused by mutations in the gene encoding for WASP, is associated with glomerulonephritis and autoimmune hemolytic anemia (Bosticardo et al., 2009); (3) chronic granulomatous disease, which is caused by a defect in NADPH oxidase, is associated with chorioretinitis (Rosenzweig, 2008); and (4) hyper-IgE syndrome is associated with autoimmune cytopenias, glomerulonephritis, and SLE (Yamazaki-Nakashimada et al., 2006). Furthermore, numerous Mendelian disorders are associated with an upregulation of type I IFN, collectively referred as type 1 interferonopathies (Crow, 2011). The common pathomechanism involved in type 1 interferonopathies is the upregulation of IFNα, resulting from either inappropriate stimulation or defective negative regulation of type 1 IFN pathway (Crow, 2011). The genes implicated and resulting type 1 interferonopathies are listed in Table 1.

3.2 Environmental Factors and Autoimmunity 3.2.1 Infection and Tissue Injury In spite of being well endowed with mechanisms of immune tolerance, susceptible individuals still develop autoimmune diseases, suggesting the presence of factors that are capable of breaking tolerance by compromising

57

Mechanisms of Autoimmunity

Table 1 Genes Implicated in Type 1 Interferonopathies Gene Autoimmune Disease References

Trex1

Aicardi–Goutie`res syndrome

Livingston and Crow (2016)

Familial chilblain lupus

Rice et al. (2007)

Systemic lupus erythematosus

Fredi et al. (2015)

Rnaseh2 Aicardi–Goutie`res syndrome

Livingston and Crow (2016)

Systemic lupus erythematosus

Gunther et al. (2015)

Aicardi–Goutie`res syndrome

Livingston and Crow (2016)

Familial chilblain lupus

Ravenscroft et al. (2011)

Systemic lupus erythematosus

Ramantani et al. (2011)

Adar1

Aicardi–Goutie`res syndrome

Livingston and Crow (2016)

Ifih1

Aicardi–Goutie`res syndrome

Livingston and Crow (2016)

Singleton–Merten syndrome

Rutsch et al. (2015)

STING-associated vasculopathy

Clarke et al. (2016)

Infantile-onset Familial chilblain lupus

Konig et al. (2017)

Acp5

Spondyloenchrondrodysplasia

Girschick et al. (2015)

Rig1

Singleton–Merten syndrome

Jang et al. (2015)

Isg15

ISG15 deficiency

Zhang et al. (2015)

Usp18

USP18 deficiency

Meuwissen et al. (2016)

Psmb

CANDLE syndrome

Kunimoto et al. (2013)

Pola1

X-linked reticulate pigmentary disorder Starokadomskyy et al. (2016)

Samhd1

Sting

Acp5, acid phosphatase 5; Adar1, adenosine deaminase acting on RNA 1; CANDLE, chronic atypical neutrophilic dermatosis with lipodystrophy and elevated temperature; Ifih1, interferon induced with helicase C domain 1; Isg15, interferon-stimulated gene 15; Pola1, polymerase (DNA) alpha 1; Psmb, proteasome subunit beta; Rig1, retinoic acid-inducible gene 1; Rnaseh2, ribonuclease H2; Samhd1, SAM domain and HD domain 1; Sting, stimulator of interferon genes; Trex1, three-prime repair exonuclease 1; Usp18, ubiquitin-specific peptidase 18.

certain checkpoints of immune tolerance (Goodnow, 2007). Indeed, many factors have been identified that can induce a loss of immune tolerance. 1. Superantigens: Superantigens are virulence proteins produced by a variety of pathogens. They bind to MHC II and TCR to stimulate proliferation of autoreactive T cells and cytokine production (Fraser and Proft, 2008). Exposure to superantigens as a result of infection contribute to initiation, as well as exacerbation, of autoimmune diseases including SLE,

58

S.K. Devarapu et al.

rheumatoid arthritis, multiple sclerosis, psoriasis, and autoimmune type 1 diabetes (Cole and Griffiths, 1993; Conrad et al., 1997; Dar et al., 2016; Kumar et al., 1997; Leung et al., 1995). 2. Molecular mimicry by antigens: Resembles of self-antigens with those of viruses and bacterial epitopes can lead to cross-reacting immune responses known as molecular antigen mimicry. For example, antistreptococcal immunity can cross-react with certain cardiac tissue antigens (Cusick et al., 2012), and the viral RNA recognition receptor TLR7 accelerates murine lupus (Anders et al., 2008). T cells involved in molecular mimicry respond to self-antigens and help autoantibodyproducing B cells to elicit an autoimmune response (Kain et al., 2008; McClain et al., 2005; Ray et al., 1996). 3. Molecular mimicry by adjuvants: During infection and trauma, dying cells release nuclear material such as nucleosomes or U1sn ribonucleoprotein that mimics the structure of viruses. These nuclear materials upon release from dying cell act as autoadjuvants by mimicking viruses and elicit an antiviral-like type I interferon-based immune response, a process typical for SLE (Anders, 2009). 4. Epitope spreading or modification of self-antigen: Modifications of self-antigens such as citrullination increase the potency of self-antigens to induce an immune response (Fig. 2). For example, citrullination of different antigens including fibrinogen, fibronectin, α-enolase, collagen type II, and histones leads to the generation of anticyclic citrullinated peptide antibodies that serve as a biomarker for rheumatoid arthritis (Gavrila et al., 2016; Lipinska et al., 2016; Sakkas et al., 2014). It is proposed that the citrullination of antigens is either a result of smoking, which is associated with increased levels of extracellular peptidyl arginine deiminase 2 (PAD2) in the lungs (Damgaard et al., 2015; Klareskog and Catrina, 2015), or autophagy in synoviocytes in rheumatoid arthritis (Sorice et al., 2016). The molecular mechanisms that assist epitope spreading include endocytic processing, antigen presentation, and somatic hypermutation (Floreani et al., 2016). 3.2.2 Environmental Agents Exposures to physical and chemical environmental agents are also considered as putative causes for the development of autoimmune diseases. For example, exposure to ultraviolet light is associated with SLE since it induces apoptosis of dermal cells resulting in autoantigen production and

Mechanisms of Autoimmunity

59

systemic inflammation (Barbhaiya and Costenbader, 2014; Caricchio et al., 2003). Furthermore, drug categories have been associated with drug-induced autoimmunity—for example, procainamide and hydralazine are associated with SLE (Rubin, 2005; Yokogawa and Vivino, 2009), whereas minocycline and nitrofurantoin are associated with autoimmune hepatitis (Hatoff et al., 1979; Lawrenson et al., 2000). Silica, tropospheric pollutants, and solvent/pesticides have also been implicated in autoimmunity; however, the precise mechanisms are not known (Floreani et al., 2016). Moreover, pristine and naturally occurring hydrocarbons can also induce autoimmunity by inducing the formation of tertiary lymphoid organs (TLOs). Pristane-induced lupus is used as a murine model of SLE and LN (Lech et al., 2010; Savarese et al., 2008).

4. FACTORS THAT INDUCE AUTOIMMUNITY 4.1 Epigenetics and Transcription Factors Gene transcription is an essential process for cellular functions and is regulated by epigenetic modifications. Numerous studies reported that epigenetic modifications occur on the gene loci that encode transcription factors and thus act as an additional regulatory factor for biological functions and disease pathogenesis. Epigenetics is one of the promising areas of investigation in the pathogenesis of autoimmune diseases such as SLE, rheumatoid arthritis, and autoimmune diabetes (Ballestar, 2010; Brown and Wedderburn, 2015; Jeffries and Sawalha, 2011, 2015). DNA methylation, histone modification, and microRNA (miRNA) are some of the known important epigenetic mechanisms. The methylation of DNA is a process in which methyl group is added to a cytosine or an adenine at the 50 position of a CpG dinucleotide. DNA methylation represses gene expression and is regulated by a specific set of enzymes like DNA methyltransferase 1 (DNMT1), DNMT3a, and DNMT3b (Denis et al., 2011). Contrary to the DNA methylation, DNA demethylation reactivates gene expression and is regulated by ten–eleven translocation methylcytosine dioxygenase 1 (TET1), TET2, and TET3 (Abdel-Wahab et al., 2009). Methylation influences the function of transcription factors by four different mechanisms, viz.: 1. Transcription factors do not bind to methylated DNA (Jin et al., 2016). 2. Methylated transcription factors cannot bind to DNA (Ivascu et al., 2007).

60

S.K. Devarapu et al.

3. Transcription factors recruit DNMT1 and repress transcription (Hervouet et al., 2010). 4. Transcription factors regulate transcriptions of methyltransferases and demethyltransferases (Zhang et al., 2006b). Histone modifications are other important epigenetic mechanisms that regulate gene expression and transcriptions factor functions. Histones within nucleosomes can undergo various modifications—for example, methylation, acetylation, deacetylation, and demethylation (Rothbart and Strahl, 2014). The acetylation process is regulated by histone acetyltransferases (HATs) and histone deacetylases (HDACs), and the methylation process is regulated by histone methyltransferases and histone demethylases. There are four known interactions between transcription factors and histone modifications, viz.: 1. Histone modifications influence binding of transcription factors to their target DNA (Gregory et al., 2001). 2. Enzymes that are involved in histone modification also regulate transcription factors (Chuang et al., 2006). 3. Transcription factors recruit HATs and HDACs to their target DNA loci (Yao et al., 2001), and 4. Transcription factors regulate the DNA modification enzymes (Katto et al., 2013). The other important epigenetic modifications are driven by miRNAs, which function as posttranscriptional and posttranslational regulators of gene expression (Chen et al., 2011). miRNAs are, in fact, one of the most important cooperators of transcription factors to various cell functions (Chen et al., 2011). miRNAs and transcription factors regulate the expression of each other in a unilateral negative feedback loop—for example, the expression of transcription factors is negatively regulated by miRNAs, and on the other hand, miRNAs are positively regulated by transcription factors (Krol et al., 2010). Nevertheless, in double-negative feedback loops, the transcription factors regulated miRNAs are directly responsible for the transcriptional activation and inactivation, while the miRNAs themselves are regulated by transcription factors (Arora et al., 2013). Transcription factors and epigenetic modifications that regulate autoimmunity are listed in Table 2. SLE. The transcription factor, regulatory factor X-box 1 (RFX1), recruits DNMT1 and HDAC1 to target gene promoter in CD4+ T cells. During the development of SLE in both humans and mice, RFX1 is found

61

Mechanisms of Autoimmunity

Table 2 Transcription Factors and Epigenetic Mechanisms in Autoimmune Diseases Transcription Factor Expression Epigenetic Mechanisms

Disease

References

RFX1

Decreased Recruitment of DNMT1, Systemic Zhao et al. HDAC, and SUV39H1 lupus (2010a,b) to the promoter regions erythematosus

E4BP4/ NFIL3

Increased

Histone acetylation and methylation

CREMα

Increased

Recruitment of DNMT3a Systemic Hedrich et al. to IL-2 promoter lupus (2012) erythematosus

EBF1

Increased

Regulation by miR-1246 Systemic Luo et al. lupus (2015) erythematosus

RelA

Increased

Regulation by SIRT6

DR3

Systemic Zhao et al. lupus (2013b) erythematosus

Rheumatoid arthritis

Klein and Gay (2015)

Decreased Regulation by DNA hypermethylation

Rheumatoid arthritis

Bull et al. (2008); Takami et al. (2006)

HOXA9

Increased

Regulation by DNA hypomethylation

Autoimmune Miao et al. type 1 (2008) diabetes

NF-κB

Increased

Regulation by H3K4 methyltransferase

Autoimmune Brasacchio type 1 et al. (2009) diabetes

FoxP3

Decreased Regulation by hypermethylation

IRF1

Decreased Deacetylation by Sirtuin I Multiple sclerosis

Yang et al. (2013)

FoxP3

Decreased Regulation by hypermethylation

Multiple sclerosis

Guan et al. (2011)

STAT5

Decreased miR-155

Multiple sclerosis

Lu et al. (2009a)

Autoimmune Tan et al. type 1 (2014); Wang diabetes et al. (2013)

CREMα, cAMP-responsive element modulator α; DR, death receptor; EBF, early B-cell factor; Foxp3, forkhead box P3; IRF, interferon regulatory factor; NFIL3, nuclear factor interleukin-3-regulated protein; NF-κB, nuclear factor kappa B; RFX, regulatory factor X; STAT, signal transducer and activator of transcription.

62

S.K. Devarapu et al.

to be downregulated, which is responsible for DNA hypomethylation and histone H3 hyperacetylation in CD11a and CD70 promoter region in CD4+ T cells, further leading to CD11a and CD70 overexpression (Zhao et al., 2010a,b). Moreover, the transcription factor E4BP4 regulates the expression of CD40L and is overexpressed in CD4+ T cells in SLE patients (Zhao et al., 2013b). Another transcription factor, CREMα, recruits DNMT3a to the IL-2 promoter, regulates chromatin conformation at the IL-17A locus, and contributes to the increased expression of IL-2 and IL-17 in CD4+ T cell in SLE patients (Hedrich et al., 2012). In addition, the transcription factor, Early B-cell factor 1 (EBF1), activates AKT pathway, regulates B-cell functions, and is known to play an important role in B-cell regulation during the development of SLE (Luo et al., 2015). miR-1246 regulates EBF1 expression and contributes the disease pathogenesis in patients (Luo et al., 2015). Rheumatoid arthritis. Epigenetic mechanisms like DNA hypermethylation (Kuchen et al., 2004), aberrant histone modification (Grabiec et al., 2008), and differentially expressed miRNAs (Nakamachi et al., 2009) are also associated with the pathogenesis and progression of rheumatoid arthritis. For example, in mice, SIRT6 suppresses NF-κB-dependent gene expression by deacetylating H3K9 (Kawahara et al., 2009) and thus suppresses the activity of NF-κB target gene-related immune responses that may contribute to the development and progression of rheumatoid arthritis (Klein and Gay, 2015). The death receptor-3 (DR-3) promoter is hypermethylated in experimental rheumatoid arthritis in mice, rendering the synovial cells resistant to apoptosis (Bull et al., 2008; Takami et al., 2006). Autoimmune diabetes. In autoimmune type 1 diabetes, T cells cannot differentiate between self-pancreatic cells from dangerous pathogens and contribute to the disease development (Xie et al., 2014). Epigenetic modifications have been shown to be associated with the pathogenesis of T1D—for example, hypomethylation of the transcription factor HOXA9 in lymphocytes from T1D patients (Miao et al., 2008). The H3K4 methyltransferase upregulates the transcription factor NF-κB and causes an increase in inflammatory gene expression in diabetic mice (Li et al., 2008). Further, an increased H3K4Me1 and a reduced H3K9 methylation also contribute to inflammation by enhanced NF-κB-p65 gene expression in patients (Brasacchio et al., 2009). In addition, the DNA methylation blocked the binding of IRF7 to FoxP3 and thus reduced the number of regulatory T cells, which contributed to the pathogenesis of autoimmune diabetes in patients (Tan et al., 2014; Wang et al., 2013).

Mechanisms of Autoimmunity

63

Multiple sclerosis. In multiple sclerosis, autoimmune responses target myelin sheet in CNS and lead to the progressive degeneration. Yang et al. reported that the histone deacetylase sirtuin I deacetylates IRF1 and contributes to the reduction in a number of TH17 cells (Yang et al., 2013). Further, increased DNA methylation is associated with decreased Treg activity in the pathogenesis of multiple sclerosis (Liu et al., 2010). This reduced Treg activity is attributed to a deficiency of miRNA-155 in Tregs (Lu et al., 2009a). Hypomethylation at IL-17A/ifng loci and increased methylation at the IL-4/FoxP3 loci contribute to the TH1/TH17 imbalance, which is also reported in the development of multiple sclerosis in mice (Guan et al., 2011). 4.1.1 MicroRNAs MicroRNAs are short endogenous noncoding RNAs that are evolutionary conserved and regulate gene expression at posttranscriptional level. Along with the wide range of cellular and developmental process, miRNAs also regulate immune tolerance mechanisms as well as pathogenesis of autoimmune diseases (Garo and Murugaiyan, 2016). For example, overexpression of miR-17–92 in lymphocytes induced lymphoproliferation and autoimmunity in mice (Xiao et al., 2008), as well as strong upregulation of miR-155 characterizes murine and human multiple sclerosis and rheumatoid arthritis (Kurowska-Stolarska et al., 2011; Murugaiyan et al., 2011; Paraboschi et al., 2011). miRNAs that regulate autoimmunity are listed in Table 3. 4.1.1.1 Central Tolerance

Central tolerance is induced during the development of T and B cells. During T-cell development, members of the miR-181 family are abundantly expressed and are involved in the regulation of T-cell selection (Li et al., 2007). The miR-181a binds to dual specificity phosphatases-6 and increases TCR signaling promoting clonal deletion of moderate affinity T cells. Thus, it prevents self-reactive T-cell clones evasion to the periphery and promoting autoimmunity (Li et al., 2012). Further, loss of miR-181a-1/b-1 reduced the basal TCR signaling in peripheral T cells and affected their migration from lymph nodes to sites of tissue inflammation, thereby dampening the development of experimental autoimmune encephalomyelitis (EAE), a murine model for multiple sclerosis (Schaffert et al., 2015). Similar to T cells, miRNAs also regulate receptor editing and B-cell clonal deletion to maintain B-cell tolerance. For example, Dicer-deficient mice, which lack

Table 3 MicroRNAs in Autoimmunity Immune Cells miRNA

Function

References

T cells

miR-181a

Clonal selection

Li et al. (2007, 2012)

miR-181a-1/b-1

TCR signaling and migration from lymph node

Schaffert et al. (2015)

Let-7

Costimulation-independent IL-2 production

Marcais et al. (2014)

miR-16

Costimulation-independent IL-2 production

Marcais et al. (2014)

miR-182

Clonal expansion of helper T cells

Stittrich et al. (2010)

miR-155

Treg differentiation

Lu et al. (2009a)

miR-21

Induction of Th-17 differentiation

Murugaiyan et al. (2015)

miR-326

Induction of Th-17 differentiation

Du et al. (2009)

miR-301a

Induction of Th-17 differentiation

Mycko et al. (2012); Nakahama et al. (2013)

miR-132/212

Induction of Th-17 differentiation

Mycko et al. (2012); Nakahama et al. (2013)

miR-20b

Suppression of Th-17 differentiation

Zhu et al. (2014)

miR-185

BCR signaling

Belver et al. (2010)

miR-155

IgG-class switching

Thai et al. (2007); Vigorito et al. (2007)

miR-150

B-cell differentiation

Xiao et al. (2007)

miR-148a

Inhibition of apoptotic death of immature B cells

Gonzalez-Martin et al. (2016)

miR-155

Regulation of Th1- and Th17-polarizing cytokine expression

Murugaiyan et al. (2011)

miR-21

Induction of T helper cells differentiation to Th2 phenotype

Lu et al. (2009b)

B cells

Dendritic cells

Macrophages miR-146a miR-124

Induction of myeloproliferation and leading to loss of peripheral tolerance Boldin et al. (2011) Induction of systemic deactivation of macrophages

Ponomarev et al. (2011)

BCR, B-cell receptor; IgG, immunoglobulin; IL, interleukin; miR, microRNA; TCR, T-cell receptor; Th, T helper cell; Treg, regulatory T cell.

Mechanisms of Autoimmunity

65

the dicer enzyme involved in miRNA biogenesis, had high autoantibody titers and immune complex deposition in kidneys (Belver et al., 2010) because the Dicer-deficient B cell had skewed BCR repertoire with increased BCR signaling (Koralov et al., 2008). Dicer-deficient mice have impaired follicular B-cell generation and more transitional and marginal zone B cells (Belver et al., 2010). Further, miRNA analysis in these mice identified loss of miR-185, overexpressed in follicular B cells, responsible for the altered BCR signaling (Belver et al., 2010). The B-cell intrinsic miR-155 regulates the germinal center response by promoting the generation of immunoglobulin class-switched plasma cells (Thai et al., 2007; Vigorito et al., 2007). Mature B cells expressed miR-150 that target the expression of a transcription factor c-Myb and thus control B-cell differentiation (Xiao et al., 2007). Moreover, miR-148a suppresses the expression of Gad45a, Bcl2l11, and Pten to protect immature B cells from apoptosis induced by engagement of the BCR, leading to an acceleration of autoimmune disease development (Gonzalez-Martin et al., 2016). These studies highlight the involvement of miRNAs in the T- and B-cell development, selection, and tolerance to self-antigens. 4.1.1.2 Peripheral Tolerance

Peripheral tolerance is mainly a backup mechanism to control autoreactivity of the cells that have escaped central tolerance. The mechanisms of peripheral tolerance include T-cell anergy and regulatory T cells, as described earlier. Dicer-deficient CD4+ T cells have demonstrated to not distinguish activating and anergic stimuli and produce IL-2 even in the absence of costimulation (Marcais et al., 2014). Further studies revealed that in Dicer-deficient CD4+ T cells, miRNA Let-7 and miR-16 target mTOR and Rictor, which are responsible for costimulation-independent IL-2 production (Marcais et al., 2014). In addition, IL-2-induced miR-182 also promotes clonal expansion of activated helper T cells by posttranscriptionally regulating FOXO1 (Stittrich et al., 2010). Inhibition of miR-182 inhibited T-cell expansion in vitro and ameliorated OVA-induced arthritis in vivo (Stittrich et al., 2010). Tregs also play a crucial role in maintaining peripheral tolerance. miRNAs regulate both kinds of Tregs, i.e., nTregs and iTregs. Selective miRNA disruption in Tregs, e.g., Dicer-deficient Tregs, leads to uncontrolled autoimmunity in vivo and the mice rapidly developed the fatal systemic autoimmune disease, which resembled Foxp3 deficiency in vivo (Liston et al., 2008; Zhou et al., 2008). Further, T cell-intrinsic Foxp3-dependent miR-155 promoted

66

S.K. Devarapu et al.

IL-2-induced STAT5 signaling and Treg differentiation by targeting SOCS-1 (Lu et al., 2009a). miRNA expression in APCs, e.g., DCs, also regulates the antigen presentation and costimulation (Turner et al., 2011). For example, miR-155 present in DCs positively regulates Th1- and Th17-cytokine expression, which is critical for the development of inflammatory T cells (Murugaiyan et al., 2011). Moreover, miR-155-deficient mice showed a delayed development of an EAE, an animal model of multiple sclerosis, protection from collagen-induced arthritis, as well as reduced autoantibody responses and alleviated lupus-like disease in FAS/lpr mice, suggesting that miR-155 confers susceptibility to multiple sclerosis, arthritis, and SLE (KurowskaStolarska et al., 2011; Murugaiyan et al., 2011; Xin et al., 2015). Furthermore, during an allergic airway inflammation the miR-21, which is upregulated in DCs, induced differentiation of T helper cells to Th2 phenotype by inhibiting p35 subunit of Th1-promoting IL-12 (Lu et al., 2009b). miR-21 also upregulated TH17 cells by inhibiting Smad 7 and mediated development of the EAE (Murugaiyan et al., 2015). Mice deficient in miR-21 were also resistant to SLE, dextran sulfate sodium-induced colitis owing to a defect in TH17 differentiation (Garchow and Kiriakidou, 2016; Shi et al., 2013). Other miRNAs that regulate TH17 differentiation are miR-326, miR-301a, miR-132/212, and miR-20b. The overexpression of miR-326 is demonstrated to be associated with the pathogenesis of multiple sclerosis (Du et al., 2009), whereas the overexpression of miR-301a and miR-132/212 is demonstrated to be associated with the pathogenesis of EAE in mice (Mycko et al., 2012; Nakahama et al., 2013). On the other hand, miR-20b suppresses TH17 differentiation by targeting RAR-related orphan receptor γt and STAT3 and protected mice from multiple sclerosis and EAE (Zhu et al., 2014). Together, miRNAs are implicated in the pathogenesis of many autoimmune diseases. Proper regulation of miRNAs is important in disease prevention and therefore miRNAs serve as a promising approach to treat autoimmune disorders.

4.2 Extracellular Vesicles Extracellular vesicles (EVs) are a group of extracellular structures that are released by all kinds of cells and are found in all body fluids (Raposo and Stoorvogel, 2013). These extracellular structures include nucleic acids from the originating cell within a phospholipid bilayer membrane. EVs are

Mechanisms of Autoimmunity

67

subdivided into the exosomes and microparticles based on their size, composition, and mechanism of formation (Raposo and Stoorvogel, 2013). Further, their function varies depending on the origin and microenvironment and may contribute to inflammation, immune signaling, vascular reactivity, angiogenesis, and tissue repair (Turpin et al., 2016). EVs interact with target cells by multiple mechanisms. For example, they regulate cell-signaling pathways by releasing ligands that activate receptors present on the target cells (Pizzirani et al., 2007). EVs also exert their function via a direct membrane contact to target cells, leading to the transfer of intracellular components after fusion or endocytosis (Morelli et al., 2004). Inside the endolysosomal compartments, EVs control antigen presentation or activate endosomal receptors. 4.2.1 The Source of Self-Antigens EVs express peptide–MHC complexes, costimulatory molecules, as well as self-antigens, on the membrane surface. Therefore, EVs might activate autoreactive T cells in the context of MHC. The number of MHC and costimulatory molecules determines an effective autoimmune response (Turpin et al., 2016). Furthermore, microparticles also are a rich source of extracellular DNA, which can bind to lupus DNA autoantibodies (Pisetsky et al., 2011). 4.2.2 Formation of Immune Complexes The binding of soluble antigens to autoantibodies results in the formation of immune complexes. EVs contain autoantigens and therefore participate in the formation of immune complexes. For example, murine monoclonal anti-DNA and antinucleosomal antibodies readily bound to microparticles that are released by dying cells in vitro (Ullal et al., 2011). A similar analysis of plasma from SLE patients showed that microparticles carry IgG, IgM, and C1q, which are associated with autoantibodies and complement activation and their numbers correlate with the anti-DNA levels (Nielsen et al., 2012; Ullal et al., 2011). Interestingly, only the loading of IgG on the microparticles, and not the number of microparticles themselves, is increased in patients with SLE compared to the healthy subjects (Nielsen et al., 2012). In addition to SLE, microparticles also contribute to the immune complex formation in autoimmune arthritis. The collagen receptor glycoprotein VI, present in the synovial fluids of rheumatoid arthritis patients, triggers platelets to release microparticles (Boilard et al., 2010). The levels of platelet-derived microparticles correlated with the rheumatic arthritis

68

S.K. Devarapu et al.

disease activity (Knijff-Dutmer et al., 2002). These local platelet-derived microparticles interact with the autoantibodies against citrullinated peptides and form proinflammatory immune complexes, which stimulate neutrophils to secrete leukotrienes and induce joint inflammation (Cloutier et al., 2013). Nevertheless, the nature of the association between microparticles and IgG is not studied in detail, raising doubts on their involvement in the formation of immune complexes, and warrants further studies. 4.2.3 Autoantigen Presentation Autoantigen presentation depends on the intracellular components being presented via MHC class I and II molecules on the APCs surface. APCs secrete exosomes that originate from MHC class II peptide compartments of the cell and therefore express very high levels of MHC class II (Clayton et al., 2001). Exosomes can directly activate T cells in the absence of viable APCs (Admyre et al., 2006; Hwang et al., 2003) or indirectly by promoting the exchange of functional peptide/MHC complexes between DCs, therefore increasing the number of a particular peptide bearing DCs (Thery et al., 2002). However, the direct activation of T cells requires that the exosomes coexpress intercellular adhesion molecule-1 (ICAM-1), whereas the mature DCs had to express CD80 and CD86 for such exchanges (Hwang et al., 2003; Thery et al., 2002). Moreover, the exosomes released from immature DCs weakly present the MHC class I peptide to T cells compared to mature DCs, and thus, mature DCs were more efficient in activating T-cell responses (Utsugi-Kobukai et al., 2003). Immature DCs can also internalize and process exosomes in the endocytic compartment to load exosome-derived MHC class II molecules for presentation to T cells (Morelli et al., 2004). 4.2.4 Inflammation and Immunity EVs also contribute to inflammation and immunity—for example, platelet-derived microparticles in the synovial fluids of patients with rheumatoid arthritis induce IL-1β secretion activating synovial fibroblasts, which further secreted IL-6 and IL-8 and contributed to synovial inflammation (Berckmans et al., 2005; Boilard et al., 2010). These platelet-derived microparticles also induce thrombotic events in co-operation with the tissue factor contributing to fibrin deposits in the synovial fluid activating synovial fibroblasts to secrete proinflammatory mediators (Boilard et al., 2010; Muller et al., 2003). In addition to platelet-derived microparticles,

Mechanisms of Autoimmunity

69

macrophage and T cell-derived microparticles also contributed to the matrix metalloproteinase and proinflammatory cytokine production by synovial fibroblasts (Distler et al., 2005). These metalloproteinases are also known to erode the blood–brain barrier, a major pathomechanism in multiple sclerosis (Saenz-Cuesta et al., 2014). On the other hand, synovial fibroblasts secreted exosomes expressing TNFα that bind to autoreactive T cells and render them resistant to activation-induced cell death, therefore promoting their survival (Zhang et al., 2006a). Platelet-derived exosomes also participate in lipid metabolism whereby they deliver arachidonic acid to adjacent platelets and endothelial cells, initiating the production of proinflammatory mediators like thromboxane A2 and cyclooxygenase (Barry et al., 1997). In addition, exosomes derived from macrophages and DCs contain enzymes that regulate leukotriene metabolism. For example, when a leukotriene biosynthesis intermediate LTA4 was incubated with intact macrophages, the major product was LTB4, whereas its incubation with DCs resulted in LTC4 production (Esser et al., 2010). Further, exosomes derived from either cell type produced chemotactic eicosanoids and induced granulocyte migration upon stimulation with Ca2+ ionophore and arachidonic acid (Esser et al., 2010). Several studies reported that the pathogen-infected cells released EVs that carry pathogen-associated molecular patterns (PAMPs), as well as EVs carrying DAMPs that are released by cells during stress conditions (Bhatnagar et al., 2007; Thery et al., 1999). These EVs activate the innate immune response by activating the pattern recognition receptors (PRRs). For example, in SLE EVs carrying RNA, as DAMP, activated endosomal TLR7 in plasmacytoid DCs (pDCs) and induced production of INFα (Pisetsky and Lipsky, 2010). In systemic sclerosis patients, EVs associated with oxidized high mobility group protein B1 (HMGB1) activated neutrophils, leading to microvascular injury and induced inflammation (Maugeri et al., 2014). Furthermore, neutrophils-derived EVs also contribute to the pathophysiology of autoimmune diseases. When human neutrophils are stimulated in vitro with myeloperoxidase (MPO) and proteinase 3 (PR3), antigens involved in ANCA-associated vasculitis, they secrete microparticles that express ANCA antigens (MPO, PR3) and tissue factors (Hong et al., 2012). These microparticles also possessed the capability to activate endothelial cells via ICAM-1 as well as to trigger coagulation cascade involving tissue factor, thus contributing to the pathophysiology of ANCA-associated vasculitis (Hong et al., 2012; Kambas et al., 2014).

70

S.K. Devarapu et al.

Together, EVs contribute to the development and pathogenesis of several autoimmune disorders and, therefore, offer potential new targets for therapy.

4.3 Neutrophil Extracellular Traps Neutrophils can expel partially decondensed chromatin upon activation by bacteria or phorbol 12-myristate 13-acetate (PMA) in net-like structures known as neutrophil extracellular traps (NETs). NET formation is primarily an early and nonspecific immune response to pathogens or foreign particles build by our body. The major components of NETs include the decondensed chromatin decorated with cytosolic proteins such as neutrophil elastase (NE), myeloperoxidase (MPO), and histones that synergize to kill bacteria (Brinkmann et al., 2004). PMA-induced NET formation was associated with neutrophil death and, therefore, called NETosis, which was morphologically different from apoptosis and necrosis (Fuchs et al., 2007). Recently, it is demonstrated that PMA or crystalline particles indeed induced receptor-interacting protein kinase 3 (RIPK3)- and mixed lineage kinase domain-like (MLKL)-mediated neutrophil necroptosis during NET formation, which is sometimes also referred as suicidal NETosis (Desai et al., 2016a,b; Mulay et al., 2016a). Neutrophils might also be able to release NETs without neutrophil death, which was wrongly termed as vital NETosis (Desai et al., 2016a; Yipp and Kubes, 2013; Yipp et al., 2012). NETs are eventually cleared either by degradation via DNase 1 (Hakkim et al., 2010) or by engulfment by macrophages (Farrera and Fadeel, 2013). 4.3.1 NETs in Autoimmune Diseases NET formation has a potential to play an important role in the pathogenesis of autoimmune diseases since they expose the otherwise intracellular antigens such as DNA, proteases, and histones to APCs that prime specific immune response against these autoantigens (Pruchniak et al., 2015) (Fig. 3). For examples, autoantibodies against double-stranded DNA (dsDNA) and histones are often found in patients with SLE (Fattal et al., 2010). In addition, autoantibodies against citrullinated protein antigens (ACPAs) originating from NETs are considered a key pathogenic event in the pathogenesis of rheumatic arthritis (Khandpur et al., 2013). 4.3.1.1 Systemic Lupus Erythematosus

A key event in the pathogenesis of SLE is the increased production of IFNγ. Denny et al. identified the presence of a distinct subset of

Cytotoxicity, vascular injury Endothelial cells

A U

Histones Type I interferon

Macrophages

LL37–DNA complex, HMGB1

NLRP3 t

b,

-1

IL

en plem

T O I

Plamacytoid Dendritic cells

M

Com

8 -1 IL

Tissue factor

Coagulation, thrombosis

M U

NET formation

N

Platelets

Neutrophils

Cytokines

I Activation

T Y

T cells Autoantibody production B cells

Fig. 3 See figure legend on next page.

72

S.K. Devarapu et al.

granulocyte population, called low-density granulocytes (LDGs), in the peripheral blood mononuclear cell fraction by density separation of whole blood from SLE patients (Denny et al., 2010). They demonstrated that these LDGs are highly active; secrete increased amounts of proinflammatory cytokines including INFγ; and possessed increased microbicidal and phagocytic capacities compared to their counterparts from the healthy controls, leading to vascular injury (Denny et al., 2010). Moreover, these LDGs have enhanced capacity to form NETs at baseline in vitro, which remains unchanged even after PMA stimulation (Villanueva et al., 2011). These findings suggested that during the development of SLE, these LDGs remain maximally stimulated in vivo, leading to the enhanced externalization of autoantigens and immunostimulatory molecules, e.g., LL-37, MMP9, and dsDNA (Villanueva et al., 2011). Therefore, NET formation attributes to the pathogenesis of SLE. In addition, a substantial proportion of SLE patients also suffer from impaired NET clearance mechanisms. For example, the presence of DNase 1 inhibitors, as well as anti-NET antibodies in the sera of SLE patients, prevented DNAse 1 access to NETs, leading to impaired NET degradation in these patients that correlated with the development of LN (Hakkim et al., 2010). In addition, oxidization of nucleic acids that are released in NETs, as well as NET-induced complement activation, leads to defects in NET clearance, contributing to SLE pathogenesis

Fig. 3 Neutrophils and NET formation in the development of autoimmunity. The activated neutrophils expel partially decondensed chromatin in net-like structures known as neutrophil extracellular traps (NETs) along with the secretion of proinflammatory cytokines. This NET formation exposes the intracellular antigens, e.g., histones, peptide (LL-37)–DNA complexes, and HMGB1. The extracellular histones activate Nlrp3 inflammasome in macrophages inducing IL-1β, IL-18 release. These cytokines further promote the recruitment of neutrophils. In addition, extracellular histones kill endothelial cells, resulting in vasculopathy that leads to the development of renal disease in systemic lupus erythematosus and ANCA-associated vasculitis. The LL-37–DNA complexes as well as HMGB1 present in the NETs facilitate the uptake and recognition of dsDNA by plasmacytoid dendritic cells, leading to the production of higher levels of IFNα, and thus contribute to SLE pathogenesis. Activated neutrophils also release inflammatory cytokines that activate T and B lymphocytes, which contribute the development of autoimmunity. Activated B lymphocytes secrete autoantibodies that form immune complexes, which contribute to the pathogenesis of multiple autoimmune diseases. In addition, NETs promote the expression of the tissue factor, which promotes thrombosis. Thrombus formation contributes the development of renal disease in systemic lupus erythematosus and ANCA-associated vasculitis.

Mechanisms of Autoimmunity

73

(Leffler et al., 2012; Lood et al., 2016). The oxidized mitochondrial DNA present in the NETs released by LDGs triggered autoimmune responses in SLE by activating the STING pathway to activate type I interferon in myeloid cells (Lood et al., 2016). Furthermore, NETs induce upregulation of CD25 and CD69, activation markers, and phosphorylation of TCR-associated signaling kinase ZAP70 in T cells, leading to lowering of their activation threshold (Tillack et al., 2012). This mechanism also represents an important link between innate and adaptive immune responses. LL-37 and HMGB1 present in the NETs have been demonstrated to facilitate the uptake and recognition of dsDNA by pDCs, leading to the production of higher levels of IFNα in a TLR9-dependent manner, and thus contribute to SLE pathogenesis (Garcia-Romo et al., 2011) (Fig. 3). Moreover, several studies reported that compounds inhibiting in NET formation in vivo—for example, PAD-4 inhibitors and mitochondrial reactive oxygen species (ROS) inhibitors, protect mice from SLE (Knight et al., 2015; Lood et al., 2016). 4.3.1.2 Rheumatoid Arthritis

A key event in the pathogenesis of rheumatoid arthritis is the formation of ACPAs. Neutrophils in the synovial fluid of patients with rheumatoid arthritis are reported to show enhanced NETosis compared to neutrophils from healthy controls. Moreover, the extent of NETosis also correlated with ACPAs levels and systemic inflammatory markers, suggesting that accelerated NETosis plays an important role in the pathogenesis of rheumatoid arthritis (Khandpur et al., 2013). Further, Spengler et al. reported that NETosis leads to the release of active PAD isoforms, viz., PAD2 and PAD4, which generate extracellular autoantigens by citrullinating histones and fibrinogens (Spengler et al., 2015). In addition, NETs also significantly augmented the production of proinflammatory cytokines IL-6, IL-8, chemokines, and adhesion molecules by synovial fibroblasts in rheumatoid arthritis (Khandpur et al., 2013). 4.3.1.3 ANCA-Associated Vasculitis

The presence of autoantibodies against MPO and PR3 is used as serological markers of ANCA-associated vasculitis (Jennette and Falk, 2014). These autoantibodies can also induce NETosis, leading to the release of toxic oxygen radicals and to the development of destructive necrotizing vascular and extravascular inflammation (Falk et al., 1990; Soderberg and Segelmark, 2016). Further, neutrophils from patients with ANCA-associated vasculitis

74

S.K. Devarapu et al.

have been reported to show enhanced NETosis compared to neutrophils from healthy controls (Tang et al., 2015). In addition, increased levels of the components of NETs, MPO/DNA complex, and calprotectin were found in sera of patients with ANCA-associated vasculitis (Kessenbrock et al., 2009; Pepper et al., 2013). Interestingly, DCs internalize MPO and PR3 present in NETs and can induce ANCA against MPO, PR3, and dsDNA, when injected into mice, leading to autoimmune vasculitis (Sangaletti et al., 2012). NETs have also been implicated in the pathogenesis of other autoimmune diseases. For example, the presence of NETs was detected in the pancreatic islets during type 1 diabetes mellitus, suggesting that circulating NET components could serve as biomarkers for diagnosis of the disease (Wang et al., 2014c). In addition, hypoglycemia in diabetes favors NET formation, a process impairing wound-healing responses and increasing chronic inflammation (Wong et al., 2015). Furthermore, increased levels of antibodies against NETs were also found the serum of patients with antiphospholipid antibody syndrome (Leffler et al., 2014). Together, NETs play a very important role in the development and pathogenesis of several autoimmune disorders and, therefore, offer new targets for therapy. However, whether potential aberrant NETosis inhibitors will ameliorate disease pathology without affecting host defense requires further examination.

4.4 Ion Channels Ion channels are the ubiquitous transmembrane proteins that allow the selective transport of ions and solutes across the plasma membrane. A range of stimuli control the opening and closing of the ion channels— for example, transmembrane potential difference, ligands, pH, temperature, and mechanical stimuli. Ion channels are mainly divided into three broad categories, viz., voltage-gated, ligand-gated, and acid-sensing ion channels (RamaKrishnan and Sankaranarayanan, 2016). In the electrically excitable cells like neurons, cardiomyocytes, and skeletal muscle cells, ion channels regulate the generation and propagation of the action potential, whereas in the nonexcitable cells, ion channels regulate cell volume, proliferation, and differentiation, as well as fluid and ion transport (RamaKrishnan and Sankaranarayanan, 2016). Moreover, ion channels control the differentiation of stem cells into the particular lineage as well as safeguard the pluripotency of stem cells (Li et al., 2015; Lo et al., 2016).

Mechanisms of Autoimmunity

75

Hematopoietic stem cells (HSCs) differentiate into both lymphoid and myeloid cells—for example, lymphoid progenitor cells differentiate into T and B lymphocytes, whereas myeloid progenitor cells differentiate into neutrophils, basophils, eosinophils, and macrophages (Pillozzi and Becchetti, 2012). All these cells play an important role in the development and progression of autoimmunity. Ion channels can modulate immune responses indirectly by regulating the differentiation of HSCs into either lymphoid or myeloid cells. In addition, ion channels present in the membrane of the immune cells, driving both innate and adaptive immune responses, can directly regulate their functions and therefore regulate the development and progression of autoimmunity (Table 4). 4.4.1 Role in Innate Immune Response and Autoimmunity Neutrophils and macrophages of the innate immune system provide the first line of defense. As described earlier, NETs released by neutrophils contribute to autoimmunity. Neutrophils undergo a respiratory burst and produce ROS during the process of NET formation (Fuchs et al., 2007). This production of superoxide in neutrophils is highly regulated by the second messenger—calcium, whose intracellular concentration is controlled by the calcium ion channels, e.g., store-operated calcium entry (SOCE), transient receptor potential melastatin subfamily 2 (TRPM2), and transient receptor potential vanilloid 2 (TRPV) (Brechard and Tschirhart, 2008; RamaKrishnan and Sankaranarayanan, 2016). Moreover, recently, it is proposed that NETosis involves neutrophil necroptosis (Desai et al., 2016a,b), and necroptosis involves Ca2+ and Na+ influx via TRPM2/7 and Na+ channels, respectively (Galluzzi et al., 2014; Kunzelmann, 2016). In addition to NETosis, calcium influx also controls other functions of neutrophils like phagocytosis, ROS production, and inflammatory processes involving neutrophils (Burgos et al., 2011). The P2X7 potassium channels, as well as Transient receptor potential (TRP) channels, are known to regulate the neutrophil chemotaxis to the target site (RamaKrishnan and Sankaranarayanan, 2016). Further, TRPM7 mediated the calcium-induced neutrophil chemotaxis, adhesion, and invasiveness, as well as toxicity against bone, and cartilages in rheumatoid arthritis patients (Wang et al., 2014a). The anion channels are also reported to play a role in neutrophil functions—for example, CIC-3 is required for normal neutrophil oxidative function, phagocytosis, and transendothelial migration (Moreland et al., 2006). It is speculated that the activated neutrophils swell due to rapid water entry stimulating the chloride channels (Simchowitz et al., 1993). This leads

76

S.K. Devarapu et al.

Table 4 Ion Channels in Autoimmunity Name of the Ion Immune Cells Ion Channel

Neutrophils

Function

References

ROS

Brechard and Tschirhart (2008)

TRPM2

ROS

Brechard and Tschirhart (2008)

TRPV2

ROS

Brechard and Tschirhart (2008)

TRP

Chemotaxis

RamaKrishnan and Sankaranarayanan (2016)

TRPM2/7

Neutrophil necroptosis or NETosis

Desai et al. (2016a); Galluzzi et al. (2014)

TRPM7

Chemotaxis

RamaKrishnan and Sankaranarayanan (2016)

K+

P2X7

Chemotaxis

RamaKrishnan and Sankaranarayanan (2016)

Cl


CIC-3

ROS, phagocytosis, migration

Moreland et al. (2006)

Ca2+ SOCE

H+

Rapid water NADPH activation entry-stimulating and NETosis Cl
channels

Salmon and Ahluwalia (2009)

VGPC

Fujiwara et al. (2013)

ROS, phagocytosis

Macrophages Na+ VGSC (Nav1.6) Phagocytosis Ca2+ CRAC

Phagocytosis, inflammasome activation, T-cell priming

Craner et al. (2005) Vaeth et al. (2015)

77

Mechanisms of Autoimmunity

Table 4 Ion Channels in Autoimmunity—cont’d Name of the Ion Immune Cells Ion Channel Function

K+

References

TRPM2/4/3

ROS, phagocytosis, inflammation

RamaKrishnan and Sankaranarayanan (2016)

CFTR

ROS, phagocytosis, inflammation

RamaKrishnan and Sankaranarayanan (2016)

K(Ca)3.1

Secretion of IL-6 and Gao et al. (2010); IL-8 Xu et al. (2014)

Kvi

Inhibition of inflammation

Moreno et al. (2013)

P2X7

Inflammasome activation

Prochnicki et al. (2016)

VGPC (Kv1.3, Induction of apoptosis Leanza et al. Kv1.1, and Kv1.5) (2012) Cl


T cells

ROS, phagocytosis, Intracellular chloride channel inflammation (CLIC 1)

Ca2+ TRPM7

RamaKrishnan and Sankaranarayanan (2016)

Inhibition of T-cell development

Jin et al. (2008)

TRPV2

Impaired TCR formation

Santoni et al. (2013)

SOCE

Positive T-cell selection

Oh-Hora et al. (2013)

L-type VGCC

Positive T-cell selection

Oh-Hora et al. (2013)

TRPV1

Immunotolerance of RamaKrishnan Tregs and Sankaranarayanan (2016) Continued

78

S.K. Devarapu et al.

Table 4 Ion Channels in Autoimmunity—cont’d Name of the Ion Immune Cells Ion Channel Function

K+

B cells

References

VGPC (Kv1.3)

Proliferation and cytokine production

RamaKrishnan and Sankaranarayanan (2016)

K(Ca)3.1

Memory T-cell development

RamaKrishnan and Sankaranarayanan (2016)

P2X7

Baricordi et al. Maturation, proliferation, cytokine (1996) production

Na+ VGSC

Positive T-cell selection

Lo et al. (2012)

Ca2+ TRPV2

Inhibition of B-cell development

Santoni et al. (2013)

CRAC

Activation, selection, RamaKrishnan and differentiation and Sankaranarayanan (2016)

T-type calcium channel

Proliferation

RamaKrishnan and Sankaranarayanan (2016)

Mg2+ TRPM6/7

B-cell growth

RamaKrishnan and Sankaranarayanan (2016)

K+

VGPC (Kv1.3)

Proliferation and class Amigorena et al. switching of memory (1990) cells

K(Ca)3.1

Proliferation and class Amigorena et al. switching of memory (1990) cells

P2X7

Proliferation

RamaKrishnan and Sankaranarayanan (2016)

79

Mechanisms of Autoimmunity

Table 4 Ion Channels in Autoimmunity—cont’d Name of the Ion Immune Cells Ion Channel Function

H+ Dendritic cells

References

Kvi

Mitogenesis and DNA Amigorena et al. synthesis (1990)

HVCN1

Antibody production Capasso et al. (2010)

Ca2+ TRPM4

Migration to lymph nodes

RamaKrishnan and Sankaranarayanan (2016)

TRPV2

Migration to lymph nodes

RamaKrishnan and Sankaranarayanan (2016)

L-type calcium Antigen presentation RamaKrishnan channel (Cav1.2) and Sankaranarayanan (2016) CFTR

DC differentiation

RamaKrishnan and Sankaranarayanan (2016)

Na+ VGSC (Nav1.7) Cytokine secretion Zsiros et al. (2009) and T-cell activation K+

Cl


VGPC (Kv1.3)

Cytokine secretion Zsiros et al. (2009) and T-cell activation

K(Ca)3.1

Chemokine-induced Shao et al. (2015) migration

P2X7

Cytokine secretion

RamaKrishnan and Sankaranarayanan (2016)

Chloride channel Chemokine-induced Shao et al. (2015) (CCL3) migration

CFTR, cystic fibrosis transmembrane conductance regulator; CRAC, Ca2+ release-activated Ca2+; HVCN1, hyperpolarized voltage-gated proton channel; K(Ca)3.1, CRAC-activated K+ channel; Kvi, voltage-dependent inward rectifier K+ channels; SOCE, store-operated calcium entry; TRPM, transient receptor potential melastatin; TRPV, transient receptor potential vanilloid; VGCC, voltage-gated calcium channel; VGPC, voltage-gated potassium channels; VGSC, voltage-gated sodium channels.

80

S.K. Devarapu et al.

to activation of NADPH oxidase, an enzyme that regulates neutrophil migration, phagocytosis, and NETosis (Salmon and Ahluwalia, 2009). The nicotinic acetylcholine receptor, as well as cAMP, activated chloride channel, and cystic fibrosis transmembrane conductance regulator (CFTR) also regulate the neutrophil functions (RamaKrishnan and Sankaranarayanan, 2016). In addition, voltage-gated proton channels also controlled the neutrophil phagocytosis by regulating cytosolic acid concentration (pH) and superoxide anion production (Fujiwara et al., 2013). Macrophages are involved in phagocytosis of pathogens as well as clearance of NETs and are responsible for both innate and adaptive immune responses. Ion channels also regulate the functions of macrophages. Craner et al. have reported that the voltage-gated sodium channel, Nav1.6, is important for the phagocytosis activity of microglia, residential macrophages in the brain and spinal cord. Therefore, inhibitors of this channel ameliorated the axonal degeneration and inflammatory reactions in an animal model of EAE and multiple sclerosis (Craner et al., 2005). In contrast, expression of human Nav1.5 in mouse macrophages enhanced the recovery in multiple sclerosis (Rahgozar et al., 2013). Furthermore, Ca2+ release-activated Ca2+ (CRAC)-induced Ca2+ influx is important for many effector functions of macrophages including phagocytosis, inflammasome activation, and priming of T cells (Vaeth et al., 2015). Such CRAC-induced Ca2+ influx regulated calcium-activated K+ channel (K(Ca)3.1)-mediated secretion of proinflammatory cytokines IL-6 and IL-8 by activation of NF-ĸB (Gao et al., 2010; Xu et al., 2014), and therefore, chemical inhibition of these channels attenuated the inflammatory reactions of macrophages (Tsai et al., 2013). The antiinflammatory action of 15-epi-lipoxin A4 is also attributed to the inhibition of voltage-dependent and inward rectifier potassium channels (Kv) in macrophages (Moreno et al., 2013). K+ efflux via P2X7 receptors activates NLRP3 inflammasome in macrophages, leading to secretion of IL-1β and IL-18 (Prochnicki et al., 2016). In addition to phagocytic and inflammatory functions by macrophages, the voltage-gated potassium channels also regulate their apoptosis. The Kv1.3, Kv1.1, and Kv1.5 are demonstrated to interact with Bax to induce apoptosis in macrophages, and subsequently, siRNA knockdown of these channels protected macrophages from undergoing apoptotic cell death (Leanza et al., 2012), as well as the Kv1.3-deficient mice showed decreased susceptibility to the development of EAE (Gocke et al., 2012). The presence of anti-IgG antibodies against Kir4.1 in the sera of patients of multiple sclerosis has been reported (Srivastava et al., 2012). Furthermore, macrophage functions are

Mechanisms of Autoimmunity

81

also regulated by Ca2+ influx through TRPM2, TRPM4, and TRPC3; intracellular chloride channel (CLIC 1); and CFTR (RamaKrishnan and Sankaranarayanan, 2016). CFTR acidified the phagolysosomes in macrophages and therefore contributed to increased oxidative burst as well as bacterial clearance (Di et al., 2006). Along with neutrophils and macrophages, ion channels also regulate the functions of basophils and natural killer cells that are involved in innate immune responses during the development of autoimmunity (RamaKrishnan and Sankaranarayanan, 2016). 4.4.2 Role in Adaptive Immune Response and Autoimmunity T and B lymphocytes are the major arms of adaptive immune responses. Ion channels have been implicated in the T-cell development and proliferation. T cell-specific deletion of TRPM7 caused a block in T-cell development at the double-negative (CD4
CD8
) stage, leading to fewer numbers of double-positive (CD4+CD8+) and single-positive (CD4+) T cells in spleen and thymus (Jin et al., 2008). The TRPV2 orchestrates Ca2+ signal in T-cell activation, proliferation, and effector functions since knockdown of TRPV2 in T cells impaired calcium signaling and TCR formation (Santoni et al., 2013). The rise in the intracellular concentration of calcium activates nuclear factor of activated T cells, leading to lymphocyte activation (Gwack et al., 2007). Accordingly, increased expression of CRAC in T lymphocytes of patients with rheumatoid arthritis was reported recently (Liu et al., 2014). Furthermore, the positive selection of T cells also depends on the SOCE as well as on voltage-gated Na+ channel and L-type voltage-gated calcium channel (Lo et al., 2012; Oh-Hora et al., 2013); thus, any alterations in their functions may lead to the development of autoimmune diseases. In addition to calcium channel, potassium channel also regulates T-cell proliferation and cytokine production—for example, increased expression of Kv1.3 and K(Ca)3.1 has been observed on T lymphocytes associated with autoimmune type 1 diabetes (RamaKrishnan and Sankaranarayanan, 2016), whereas inhibition of Kv1.3 on T lymphocytes protected severe combined immunodeficient mice from psoriasis (Gilhar et al., 2011). Recent evidence also suggests that curcumin inhibits Kv1.3 on T effector lymphocytes isolated from patients with rheumatoid arthritis and multiple sclerosis and, therefore, may serve as a potential treatment option for these patients (Lian et al., 2013). Intracellular Ca2+ concentration also plays an important role in B-cell activation, selection, and differentiation into plasma and memory B cells (Scharenberg et al., 2007). Similar to T cells, TRPV2 is expressed in

82

S.K. Devarapu et al.

B cells where it regulates Ca2+ release during B-cell development and activation (Santoni et al., 2013). In addition, potassium channels are also involved in B-cell proliferation and IgG-class switching associated with SLE (RamaKrishnan and Sankaranarayanan, 2016). Moreover, the influx of K+ activated B-cell mitogenesis and DNA synthesis at G1 and S phases (Amigorena et al., 1990). The voltage-gated proton channel HVCN1 that associated with the BCR is involved in mitochondrial ROS production in B cells (Capasso et al., 2010). 4.4.3 Antigen Presentation DCs process the antigens and present the antigenic peptides to T and B cells. It is demonstrated that the immature DCs express sodium channel Nav1.7, which is downregulated with concomitant upregulation of voltage-gated Kv1.3 potassium channel expression during maturation of DCs, and therefore, Kv1.3 blockers can inhibit various functions of mature DCs such as cytokine secretion and T cell-activating potential (Zsiros et al., 2009). Furthermore, intermediate conductance calcium-activated potassium channel K(Ca)3.1 and chloride channel (CIC-3) regulate the chemokine-induced migration of DCs to the lymph nodes where they stimulate adaptive immune responses (Shao et al., 2015). Other ion channels that regulate the DCs migration include TRPM4, TRPV1, and TRPV2 (RamaKrishnan and Sankaranarayanan, 2016). In addition to this, the water-transporting channels AQP-7, AQP-5, and AQP-3 are involved in antigen presentation (RamaKrishnan and Sankaranarayanan, 2016). The voltage-gated potassium and calcium channels also act as an autoantigen and stimulate immune responses during the development of autoimmune type 1 diabetes (Fierabracci and Saura, 2010; Messinger et al., 2009). Together, ion channels extensively regulate both innate and adaptive immune response, and therefore, an aberration in the expression of ion channel might lead to autoimmune disorders. Understanding their functions in more detail will offer multiple new targets as potential novel therapies autoimmune diseases.

4.5 Lipids Lipid mediators are other important molecules that influence the autoimmunity. They are divided into following three classes—class I, arachidonic acidderived eicosanoids—prostaglandins and leukotriene; class II, lysophospholipids and their derivatives; and class III, omega-3-polyunsaturated fatty acid-derived

Mechanisms of Autoimmunity

83

antiinflammatory mediators (Murakami, 2011). Lipids are important and complex component of the plasma membrane. The lipid raft controls protein interactions following the ligand–receptor binding and, thus, promotes the signaling in immune cells (Katagiri et al., 2001). While the complexity of lipids in the plasma membrane is appreciated for long, only recently the developments in lipidomics have increased our knowledge of their dynamics in plasma membrane (Wu et al., 2016). Aberration in lipid functions has been associated with development and complications of several autoimmune disorders—for example, patients with SLE show increased levels of oxidized lipids, viz., oxidized LDL and phospholipids, leading to generation of antibodies against them, which contribute to thrombosis resulting in increase in atherosclerosis and coronary artery events in SLE (Hahn and McMahon, 2008). Abnormalities in lipid-binding proteins of myelin and sphingolipid content that confer increased immunogenicity have been implicated in the development of multiple sclerosis (Reale and Sanchez-Ramon, 2016). Furthermore, the presence of lipid-specific antibodies against sulfatide, sphingomyelin, and oxidized lipids in cerebrospinal fluid derived from individuals with multiple sclerosis has been identified by a large-scale multiplex microarray analysis of lipids present in the myelin sheath, including ganglioside, sulfatide, cerebroside, sphingomyelin, and total brain lipid fractions (Kanter et al., 2006). Lipids also serve as an antigen and regulate immunity via lipid-reactive T cells. Such lipid antigens are presented to T cells by proteins of CD1 family (Dowds et al., 2014). Such lipid-reactive T cells’ number and functions were reduced in patients with SLE as well as mouse models of SLE (Jacinto et al., 2012). These CD1d-reactive T cells, also called invariant natural killer T (NKT) cells, inhibit autoreactive B cells since they express more CD1d than the nonautoreactive B cells (Yang et al., 2011). Therefore, CD1d deficiency in mice leads to exacerbated autoantibody production and developed a lupus-like disease (Yang et al., 2011). Furthermore, the NKT cell levels and functions are also reduced in SLE patients, which correlated with the disease activity (Cho et al., 2011). In particular, the CD4+ T cells from SLE patients display an altered profile of lipid raft-associated glycosphingolipids (GSLs), e.g., lactosylceramide, globotriaosylceramide, and monosialotetrahexosylgangliosi de (McDonald et al., 2014). Interestingly, normalizing GSLs restored function in CD4+ T cells from SLE patients (McDonald et al., 2014).

84

S.K. Devarapu et al.

Together, these observations indicate that lipids and lipid-reactive T cells contribute to autoimmunity via different mechanisms, and therefore, represent potential therapeutic targets for autoimmune diseases.

5. COSTIMULATORY AND COINHIBITORY PATHWAYS IN AUTOIMMUNITY Activation of naive T cells requires two signals acting simultaneously. Interaction of TCR with MHC-peptide molecules comprises signal 1, while costimulation via costimulatory receptors and their corresponding ligands on APCs requires signal 2 for activation of naı¨ve T cells (Lafferty and Cunningham, 1975; Mueller et al., 1989). These costimulatory mechanisms provide molecular checkpoints to ensure that the immune system produces a controlled response to foreign antigens while avoiding pathology and destruction of the host tissue and provide potential avenues for therapeutic intervention. These costimulatory mechanisms regulate Tregs, which are important for self-tolerance (Sakaguchi et al., 2008; Vignali et al., 2008). In fact, as mentioned earlier, either overactivation of costimulatory pathways or loss of coinhibitory mechanisms leads to loss of the self-tolerance. Apart from APCs, B cells and other immune cells also require costimulation for their activation, maturation, and function (Bretscher and Cohn, 1970). Thus, costimulatory and coinhibitory receptors and their signaling pathways regulate overall immune response. We have reviewed these costimulatory and coinhibitory pathways in four important autoimmune diseases including SLE, rheumatoid arthritis, multiple sclerosis, and type 1 diabetes (Table 5).

5.1 Costimulatory Pathways One of the most important costimulatory receptors is CD28, which is activated by its ligands B7.1 (CD80)/B7.2 (CD86). This CD28 stimulation, along with TCR-MHC-II signaling, promotes the proliferation of T cells, as well as their survival, during T-cell priming. Several studies reported attenuation of autoimmune diseases in Cd28-deficient mice— for example, MRL/lpr mice, an animal model for SLE (Tada et al., 1999a); in DBA/1 mice with collagen-induced arthritis (Tada et al., 1999b); and in Cd28-deficient-MBP1–17 TCR transgenic mice, a model of EAE (Oliveira-dos-Santos et al., 1999). Interestingly, other reports suggest that the B7.1 and B7.2 act in a redundant manner in the development of SLE. For example, the dual blockade of B7.1 and B7.2 using antibodies resulted in a significant decrease in autoantibody production compared to the blockade using either of the antibodies alone in MRL/lpr mice

85

Mechanisms of Autoimmunity

Table 5 Costimulatory and Coinhibitory Pathways in Autoimmunity Autoimmune Pathway Disease References

Costimulatory CD28–B7.1/ B7.1 axis

Systemic lupus erythematosus

Liang et al. (1999); Tada et al. (1999a)

Autoimmune type 1 diabetes

Lenschow et al. (1996); Salomon et al. (2000)

Rheumatoid arthritis

Tada et al. (1999b); Webb et al. (1996)

Encephalomyelitis Chang et al. (1999); Oliveira-dos-Santos et al. (1999) ICOS–ICOSL axis

Systemic lupus erythematosus

Hu et al. (2009); Odegard et al. (2008); Teichmann et al. (2015)

Rheumatoid arthritis

Iwai et al. (2002)

Encephalomyelitis Sporici et al. (2001)

CD40–CD40L axis

Autoimmune type 1 diabetes

Ansari et al. (2008); Hawiger et al. (2008)

Systemic lupus erythematosus

Early et al. (1996); Ma et al. (1996)

Rheumatoid arthritis

Kyburz et al. (1999); MacDonald et al. (1997); Tellander et al. (2000)

Encephalomyelitis Gerritse et al. (1996); Howard et al. (1999) Autoimmune type 1 diabetes OX40–OX40L Systemic lupus axis erythematosus Rheumatoid arthritis

Baker et al. (2008); Bour-Jordan et al. (2004); Green et al. (2000) Cunninghame Graham et al. (2008); Jacquemin et al. (2015); Manku et al. (2013) Gwyer Findlay et al. (2014); Yoshioka et al. (2000)

Encephalomyelitis Carboni et al. (2003); Weinberg et al. (1996) Autoimmune type 1 diabetes

Pakala et al. (2004) Continued

86

S.K. Devarapu et al.

Table 5 Costimulatory and Coinhibitory Pathways in Autoimmunity—cont’d Autoimmune Pathway Disease References

Coinhibitory

CTLA4–B7.1/ B7.2 axis

PD1–PDL1/ PDL2 axis

Systemic lupus erythematosus

Butte et al. (2007); Finck et al. (1994); Mihara et al. (2000)

Rheumatoid arthritis

Ko et al. (2010); Quattrocchi et al. (2000); Webb et al. (1996)

Systemic lupus erythematosus

Butte et al. (2007); Laufer et al. (1996); Okazaki and Honjo (2007)

Rheumatoid arthritis

Raptopoulou et al. (2010)

Encephalomyelitis Ansari et al. (2003); Bodhankar et al. (2013); Carter et al. (2007) Autoimmune type 1 diabetes

Ansari et al. (2003); Liang et al. (2003); Wang et al. (2008)

Rheumatoid Levin et al. (2011) TIGIT– CD112/CD155 arthritis axis Encephalomyelitis Joller et al. (2011); Levin et al. (2011) TIM3–galectin 9 axis

Rheumatoid arthritis

Pierer et al. (2009); Seki et al. (2008)

Encephalomyelitis Podojil et al. (2013); Zhu et al. (2005) Autoimmune type 1 diabetes

Sanchez-Fueyo et al. (2003); Wang et al. (2011)

CD, cluster of differentiation; ICOS, inducible T-cell costimulator; PD, programmed cell death protein; TIGIT, T-cell immunoreceptor with Ig and ITIM domains; TIM3, T-cell immunoglobulin and mucin domain-containing 3.

(Liang et al., 1999). The similar redundancy of B7.1 and B7.2 was observed in DBA/1 mice (Webb et al., 1996) as well as in the MOG35–55-induced EAE model in mice (Chang et al., 1999). Further, O‘Neill et al. showed that expression of B7 molecules on B cells is essential for autoreactive T-cell priming in the development of murine proteoglycan-induced arthritis

Mechanisms of Autoimmunity

87

(O’Neill et al., 2007). In contrast to the above observations, both Cd28-deficient NOD mice and B7.1/B7.2-double deficient NOD mice showed an acceleration in the development of autoimmune diabetes, along with a significant reduction in the number of Tregs and enhanced IFNγ production (Lenschow et al., 1996; Salomon et al., 2000). The Inducible T-cell COStimulator (ICOS) or CD278 also induces T-cell activation and the humoral immunity. The function and generation of follicular and extrafollicular T helper cells are dependent on ICOS in NZB/NZW.F1 and MRL/lpr mouse models of SLE (Hu et al., 2009; Odegard et al., 2008; Teichmann et al., 2015). Furthermore, pharmacological blockade of the ICOS–ICOSL pathway with anti-ICOSL inhibited development of arthritis in a mouse model of collagen-induced arthritis (Iwai et al., 2002). In the EAE, selective ICOS blockade exacerbated disease in the priming phase (1–10 days after immunization), whereas it ameliorated the disease in the efferent phase (9–20 days after immunization) (Sporici et al., 2001). However, the Icos-deficient mice developed more severe disease with enhanced production of Th1 and Th17 cell cytokines (Dong et al., 2001; Galicia et al., 2009). In contrast, the Icos-deficient NOD mice, as well as NOD mice treated with anti-ICOSL, showed a delayed development of autoimmune diabetes (Ansari et al., 2008; Hawiger et al., 2008). In addition to the costimulatory receptors, the CD40–CD40L costimulatory pathway also drives the humoral immune responses during the development of autoimmunity. Studies in NZB/NZW.F1 and MRL/lpr mice with the loss of CD40–CD40L axis demonstrated that the B and T cells interact with each other via CD40–CD40L axis and drive the humoral immunity in SLE (Early et al., 1996; Ma et al., 1996). For example, anti-CD40L blockade in combination with CTLA4 antibody in NZB/ NZW.F1 mice has shown synergistic effects by suppressing B- and T-cell activation (Daikh et al., 1997; Wang et al., 2002). Furthermore, in rheumatoid arthritis patients, CD4+ T cells present in synovial fluid express CD40L, and the CD40–CD40L axis contributes to the production of autoantibodies (MacDonald et al., 1997). The pharmacological blockade of CD40–CD40L axis attenuated the progression of murine collagen-induced arthritis (Kyburz et al., 1999; Tellander et al., 2000), as well as anti-CD40L antibody slowed the progression and development of EAE by inhibiting T-cell activation, as well as IFNγ production (Gerritse et al., 1996; Howard et al., 1999). In addition, CD40–CD40L costimulation plays an essential role in the initiation of insulitis and autoreactive T-cell priming during the development of autoimmune diabetes since both CD40L antagonization and Cd40l

88

S.K. Devarapu et al.

deficiency in NOD mice completely abrogated the development of the disease (Baker et al., 2008; Balasa et al., 1997; Bour-Jordan et al., 2004; Green et al., 2000). Another costimulatory pathway that contributes to the development of autoimmune diseases is the OX40–OX40L axis. Several polymorphisms at Tnfsf4, a gene encoding OX40L, are associated with the development of SLE (Cunninghame Graham et al., 2008; Manku et al., 2013). Further, OX40L was reported to contribute to human lupus pathogenesis by promoting T follicular helper response (Jacquemin et al., 2015). Very high expressions of OX40 and OX40L were observed on activated T cells and APCs, respectively, in the inflamed joints in collagen-induced arthritis mouse model (Gwyer Findlay et al., 2014). Moreover, mice deficient in OX40–OX40L signaling show less IFNγ and autoantibody production in the pathogenesis of rheumatoid arthritis (Gwyer Findlay et al., 2014; Yoshioka et al., 2000). Similarly, OX40 is overexpressed on autoreactive T cells during the development of EAE, and therefore, the selective depletion of T cells attenuated the disease pathology (Carboni et al., 2003; Weinberg et al., 1996). However, although the deficiency of OX40 or OX40L delayed the development, it did not result in complete protection from the disease, suggesting a minor role of this axis in the pathogenesis of EAE (Carboni et al., 2003; Ndhlovu et al., 2001; Nohara et al., 2001; Weinberg et al., 1999). The OX40–OX40L axis also demonstrated to contribute to the development of autoimmune diabetes in NOD mice at a late stage (Pakala et al., 2004).

5.2 Coinhibitory Pathways The coinhibitory molecule B7.1 plays an important role in the regulation of immune response in SLE. For example, the B7.1-deficient mice showed exacerbation of SLE in MRL/lpr (Liang et al., 1999). As mentioned earlier, the B7.1 and B7.2 molecules interact with costimulatory CD28 on T-cell surface to transduce stimulatory signal, whereas they can also interact with CTLA4, as well as PD-L1, to transduce the inhibitory signals (Butte et al., 2007). Therefore, an exacerbation in disease pathology in B7.1-deficient MRL/lpr mice can be attributed to the preferential interaction of CTLA4 and PDL1 with B7.1 over B7.2 (Butte et al., 2007). The important evidence of the coinhibitory role of CTLA4–B7 interaction comes from the observation that treatment of CTLA4 antibody attenuated the disease progression in NZB/NZW.F1 mice (Finck et al., 1994; Mihara et al., 2000) as well as in

Mechanisms of Autoimmunity

89

mouse model of collagen-induced arthritis (Knoerzer et al., 1995; Ko et al., 2010; Quattrocchi et al., 2000; Webb et al., 1996). On the contrary, the NOD mice expressing the CTLA4 antibody or treated with CTLA4 antibody showed exacerbation in autoimmune diabetes (Lenschow et al., 1996; Salomon et al., 2000). Another coinhibitory pathway that contributes to the development of autoimmune diseases is the PD1–PDL axis. Polymorphisms at PDCD1 are associated with SLE, rheumatoid arthritis, multiple sclerosis, and type 1 diabetes (Okazaki and Honjo, 2007). The C57BL/6J mice that are deficient in Pdcd1 have higher expression of IgG2b, IgA, and IgG3 in plasma along with reduced expression of CD5, leading to the spontaneous development of SLE (Nishimura et al., 1998; Nishimura et al., 1999). Moreover, pharmacological blockade of PD1 in NZB/NZW.F1 mice resulted in an expansion in the population of Tregs and therefore protected the mice from the development of SLE and associated renal injury (Kasagi et al., 2010; Wong et al., 2013; Wong et al., 2010). Similarly, the Pdcd1-deficient mice exhibited exacerbated rheumatoid arthritis, and pharmacological activation of PD1–PDL axis attenuated the progression of rheumatoid arthritis (Raptopoulou et al., 2010). Further, the PD1–PDL axis also plays an important role in the development of EAE. Both PDL1 and PDL2 were observed on the infiltrating cells. However, the PD1 interaction with PDL1 but not PDL2 on B cells protected mice from EAE (Bodhankar et al., 2013; Carter et al., 2007). Pancreatic β cells also express PDL1; however, its involvement in the development of autoimmune diabetes is ambiguous (Ansari et al., 2003; Liang et al., 2003). For example, PDL1 overexpression on β cells in NOD mice protected them (Wang et al., 2008), whereas in C57Bl6 mice PDL1 overexpression caused the activation of autoreactive T cells, leading to the aggravation of autoimmune diabetes (Subudhi et al., 2004). Similar to the EAE, pharmacological blockade of PD1–PDL1 axis, but not the blockade of PDL2, leads to aggravation of autoimmune diabetes in NOD mice (Ansari et al., 2003). Apart from the coinhibitory pathways discussed earlier, there are some pathways, which also participate in the regulation of immune response— for example, the signaling pathways associated with TIGIT (VSIG9 or VSTM3). The TIGIT interacts with ligands like CD112 and CD115 and suppresses T-cell responses in rheumatoid arthritis (Levin et al., 2011). Further, both the treatment with TIGIT agonist and mice overexpressing TIGIT inhibited the progression, while mice Tigit-deficient mice exhibited exacerbation of EAE (Joller et al., 2011; Levin et al., 2011). Moreover,

90

S.K. Devarapu et al.

coinhibitory receptors, TIM3 and B7-H4, play an important role in the progression of rheumatoid arthritis (Pierer et al., 2009; Seki et al., 2008), EAE (Monney et al., 2002; Podojil et al., 2013; Prasad et al., 2003; Sabatos et al., 2003; Zhu et al., 2005), as well as autoimmune diabetes (Sanchez-Fueyo et al., 2003; Wang et al., 2011). Together, several costimulatory receptors, as well as coinhibitory pathways, regulate the immune responses during the development of autoimmune diseases. Understanding such pathways in detail will provide novel therapeutic options for autoimmune diseases.

6. PRRs IN AUTOIMMUNITY Pioneers like Beutler, Janeway, Medzhitov, and colleagues initiated scientific interest in the field of innate immunity over the past decades (Beutler, 2000; Janeway and Medzhitov, 1999). It is now clear that immune cells use a set of evolutionarily conserved PRRs to detect foreign microorganisms via PAMPs (Cao, 2016). However, these PRRs can also sense mammalian motifs that are delivered by stressed or dying cells (Cao, 2016). Such signals include the release and intracellular engagement of ATP; endogenous TLR ligands, e.g., modified mammalian nucleic acids, are cumulatively termed DAMPs (Cao, 2016; Park et al., 2004). This so-called danger hypothesis not only implies broadened competence of the innate immune system in terms of host defense but also suggests a role for these PRRs in the pathogenesis of autoimmunity and autoimmune disease (Anders and Fogo, 2014; Lech and Anders, 2013; Lech et al., 2015; Lorenz and Anders, 2015; Marshak-Rothstein, 2006; Nickerson et al., 2010; Pawar et al., 2007). Currently, PRRs can be divided into three main families, namely: TLRs, RIG-I-like receptors (RLRs), and NOD-like receptors (NLRs) with different (sub)cellular localizations (Cao, 2016). TLRs can be subdivided into surface expressed (TLR1, 2, and 4–6) and endosomal receptors (TLR3, 7, 8, and 9). RLRs (e.g., MDA5, RIG-I) and NLRs (e.g., AIM2, NLRP3) on the contrary reside in the cytosolic compartment (Cao, 2016; Lorenz et al., 2017). Surface and endosomal TLRs cover the detection of extracellular bacterial wall components, e.g., LPS. They further sense endolysosomal RNA and DNA derived from pathogens or the host cell itself (Wu and Chen, 2014). Whereas these receptors are practically blind to cytosolic pathogens, RLRs like RIG-I or MDA5 and NLRs like AIM2 can detect viral RNA and DNA in the cytosol, respectively (Wu and

Mechanisms of Autoimmunity

91

Chen, 2014). This ubiquity awareness of nuclear acids and other PAMPs and DAMPs is double edged. Indeed, it allows the detection of a broad variety of pathogens independently of their intrusion pathway (Wu and Chen, 2014). However, under autoimmune conditions, NLR ligation by self-nuclear acids can promote autoimmunity and autoimmune tissue inflammation (Cao, 2016). In the following sections, we will discuss how PRRs can influence the pathogenesis of autoimmunity and autoimmune tissue inflammation and discuss their involvement in clinical autoimmune disease such as SLE systemic sclerosis.

6.1 PRRs and Autoimmunity How PRRs contribute to autoimmunity. The self-nucleic acids become immune stimulatory by engaging endolysosomal TLRs. For example, in SLE failure to clear apoptotic blebs on the surface of dying cells containing nuclear material and associated proteins renders these initially inert particles immune stimulatory (Casciola-Rosen et al., 1994; Lorenz and Anders, 2015). After engulfment, nuclear material containing immune complexes can stimulate autoreactive B cells to secrete autoantibodies and APCs to release proinflammatory cytokines and type I interferons in a TLR7- or a TLR9-dependent manner (Anders, 2010; Demaria et al., 2010; Lau et al., 2005; Leadbetter et al., 2002; Lorenz and Anders, 2015). Hence, it is not surprising that overexpression of RNase in TLR7 transgenic mice confers protection against TLR7-driven lupus-like disease (Sun et al., 2013). Within immune cells, ligation of most TLRs results in the recruitment of MyD88 and consequently the activation of NF-κB and mitogen-activated protein kinases (Cao, 2016). In turn, APCs produce inflammatory cytokines, enhance their antigen-presenting capability, and stimulate T- and B-celldependent adaptive immune responses (Lorenz et al., 2017). Different from this is the signaling via TLR3 and TLR4, which alternatively signal via the adapter protein TRIF to induce the production of type I interferons via interferon regulatory transcription factor (IRF) 3–7 (Cao, 2016). In addition, parenchymal cells—for example, renal mesangial cells, also express a restricted set of TLRs (e.g., TLR1–4, TLR6). Upon TLR engagement of endogenous ligands, these cells can produce chemoattractants and interleukins to recruit immune cells and promote autoimmune tissue inflammation (Lorenz et al., 2017). RLRs, like RIG-I and MDA5, detect invading virus via double-stranded or single-stranded cytosolic RNA. Consequently, type I interferon secretion

92

S.K. Devarapu et al.

boosters the antiviral host response (Cao, 2016). However, increased type I interferon production is also prominent in some autoimmune diseases (Lorenz and Anders, 2015). A gain-of-function mutation in the gene coding for MDA5 triggers the spontaneous onset of autoimmunity in mice, which partially depends on exaggerated type I interferon production by DCs (Funabiki et al., 2014). Interestingly, a gain-of-function mutation of the human MDA5 gene locus IFIH1 has also been identified in patients suffering from Aicardi–Goutie`res syndrome (Oda et al., 2014). The IFIH1 rs1990760 T-allele of the human IFIH1 (MDA5-coding) gene has been associated with SLE, rheumatoid arthritis, and other autoimmune diseases (Cen et al., 2013). These data indicate that aberrant RLR activity is involved in the onset of autoimmunity. In addition, parenchymal cells express MDA5 and RIG-I. After activation by RNA motives, these cells produce proinflammatory mediators and enhance inflammation on the tissue level (Hagele et al., 2009; Imaizumi et al., 2012). Activation of NLRs, like NLRP3 and AIM2, can form large molecular complexes with the adaptor protein ASC to recruit caspase-1, which results in the proteolytic activation and secretion of proinflammatory IL-1β and IL-18. Moreover, inflammasome formation can trigger a programmed proinflammatory cell death with necrotic features in macrophages, which is known as pyroptosis (Fernandes-Alnemri et al., 2009; Lorenz et al., 2014a). Further, AIM2 senses cytoplasmatic double-stranded DNA and mediates macrophage activation in response to apoptotic DNA under autoimmune conditions (Fernandes-Alnemri et al., 2009; Zhang et al., 2013). In contrast, the NLRP3 inflammasome can respond to a much broader spectrum of different DAMPs, thereby acting as a sensor of cellular homeostasis (Martinon et al., 2009). NLRP3 inflammasome activation usually comprises a two-step process. First, there is TLR or death receptormediated transcriptional upregulation of pro-IL-1β and pro-IL-18. Second, cellular potassium efflux, ROS generation, or extracellular ATP increase triggers the assembly of the NLRP3-inflammasome and caspase1-dependent IL-1β and IL-18 secretion (Lorenz et al., 2014a). Interestingly, U1-sn RNP and NETs can also stimulate NLRP3 activation (Kahlenberg et al., 2013; Shin et al., 2012). Inadequate secretion of IL-1β and IL-18 not only fosters local inflammatory responses as seen in arthritis but also skews adaptive immune responses—for example, by promoting Th17 T-cell expansion. These cells have been implicated in several autoimmune diseases (Ji et al., 2016; Zhang et al., 2016).

Mechanisms of Autoimmunity

93

6.1.1 PRRs in SLE TLR involvement in autoimmunity is most apparent in murine models of lupus-like disease. TLR7, which senses single-stranded RNA, is required for the production of anti-RNA-targeted autoantibodies in different murine models (Christensen et al., 2006; Savarese et al., 2008). Abrogation of the signaling molecule MyD88 or simultaneous deletion of TLR7 and TLR9 in MRL/lpr completely abrogates ANA production (Nickerson et al., 2010). Further, TLR7 signaling promotes systemic cytokine production, lymphoproliferation, memory T-cell formation, and severity of glomerulonephritis in these mice (Nickerson et al., 2010). In mice, bearing autoimmune susceptibility mutations such as Sle1 overexpression of TLR7 aggravates pathologies (Subramanian et al., 2006). For humans, SNP in the TLR7 gene or UTR has been associated with risk of SLE in different ethnic populations (Kawasaki et al., 2011; Lee et al., 2016; Shen et al., 2010; Wang et al., 2014b). Moreover, overexpression of TLR7 gene has been found in PBMCs from SLE patients (Lyn-Cook et al., 2014). Normalizing TLR7 expression solely in CD19+ B cells significantly decreases anti-RNA autoantibody production and severity of nephritis (Hwang et al., 2012). In accordance with this, in MRL/lpr mice, B-cell intrinsic TLR/MyD88 is required for ANA formation and full-blown nephritis, whereas DC-intrinsic signaling affects different disease features like autoimmune skin inflammation and type I interferon production by pDCs (Teichmann et al., 2013). Thus, TLR7/MyD88 signaling differentially promotes innate and adaptive immune responses in B cells and DCs. Nevertheless, nuclear acid-sensing TLRs do not promote autoimmunity uniformly and show a complex regulatory interplay. TLR9 is a sensor for CpG rich DNA that is able to sense endogenous DNA-containing immune complexes and to induce type I interferon secretion by pDCs (Leadbetter et al., 2002; Lech and Anders, 2013). TLR9 expression in peripheral blood leukocytes correlates with anti-DNA-targeted autoantibody titers and TLR9 upregulation on B cells has been correlated with increased serum creatinine in humans (Chauhan et al., 2013; Nasr et al., 2015). Although TLR9 inhibition reduced lymphoproliferation and renal inflammation upon CpG challenge in murine lupus, deletion of TLR9 in several murine models goes along with exaggerated disease (Nickerson et al., 2010; Patole et al., 2005; Bossaller et al., 2016). The Tlr9-deficient MRL/lpr mice show reduced titers of antinucleosome antibodies, but produce increased amounts of antiRNA-mediated antibodies and present with increased lymphoproliferation and aggravated renal disease (Nickerson et al., 2010). These effects can be

94

S.K. Devarapu et al.

abrogated by simultaneous deletion of TLR7 (Nickerson et al., 2010). Thus, TLR9 negatively regulates TLR7 in this model of systemic autoimmunity (Nickerson et al., 2010). A possible molecular mechanism for this regulatory function of TLR9 may be that TLR9 and TLR7 compete for endosomal trafficking via Unc93B1 (Fukui et al., 2011). Consequently, in the absence of TLR9, TLR7 trafficking to the endosome may be increased (Fukui et al., 2011; Lorenz et al., 2017). Like TLR9, the role of TLR8 is difficult to predict in SLE and autoimmunity. In 564Igi mice that express an anti-RNA-antibody, TLR8 contributes to autoantibody production and type I interferon production. However, in C57BL/6 mice, loss of TLR8 results in lymphoproliferation, anti-dsDNA autoantibody production, and glomerulonephritis (Demaria et al., 2010; Tran et al., 2015). Interestingly, myeloid cell TLR7 signaling is exaggerated in Tlr8-deficient animals and simultaneous deletion of TLR7 can rescue the phenotype of Tlr8-deficient animals (Demaria et al., 2010; Tran et al., 2015). Thus, both TLR8 and TLR9 differentially regulate TLR7-mediated autoimmunity in murine lupus. Besides that, also TLR2 and TLR4 have been implemented in the pathogenesis of autoimmunity. Pristane-induced nephritis as well as nephritis in C57BL6lpr/lpr mice partially depends on the presence of TLR2 and TLR4 and responsiveness of PBMCs to TLR2 and TLR4, in terms of cytokine production, is increased in SLE patients (Klonowska-Szymczyk et al., 2014; Lartigue et al., 2009; Nasr et al., 2015; Summers et al., 2010). Murine macrophages in lupus-prone MRL/lpr mice display defects in lysosomal maturation and acidification. This goes along with prolonged and heightened ROS production and reduced elimination of internalized IgG immune complexes (Monteith et al., 2016). Internalized IgG immune complexes stimulate endolysosomal TLR7 and TLR9 to increase type I interferon production (Monteith et al., 2016). Most importantly, dsDNA from immune complexes was shown to leak into the cytosol and to drive AIM2 inflammasome assembly in macrophages (Monteith et al., 2016). This might explain how knockdown of AIM2 in an apoptotic DNA-induced lupus model decreases systemic inflammation and nephritis (Zhang et al., 2013). The AIM2 expression in human PBMCs is upregulated in SLE patients and correlated to clinical disease severity (Zhang et al., 2013). Therefore, AIM2 might be an important sensor for self-DNA in SLE and contribute to renal inflammation via aberrant activation of infiltrating macrophages (Monteith et al., 2016; Zhang et al., 2013). However, New Zealand Black (NZB) mice, which spontaneously develop autoimmune

Mechanisms of Autoimmunity

95

hemolytic anemia and lupus-like disease, present defective NLRP3 and AIM2 inflammasome signaling (Sester et al., 2015). This implies that other genetic alterations can substitute for aberrant NLR signaling in order to promote SLE pathogenesis. Further, there is the possibility that like TLR9, NLRs have additional negative regulatory functions via interfering with other signaling pathways. In this context, Nlrp3-deficient B6lpr mice have been shown to display exaggerated lymphoproliferation and autoimmune kidney and lung injury, eventually due to decreased immunosuppressive TGFβ signaling in these mice (Lech et al., 2015). On the contrary, inhibition of NLRP3 inflammasome activation reduced Th17 T-cell polarization and beneficially influenced nephritis and lifespan in MRL/lpr and NZM2328 mice (Zhao et al., 2013a). Intensive research is needed to further dissect the complex regulatory interplay among different PRRs to exploit these receptors as therapeutic targets in the future. SLE patients show elevated production of type I interferons similar to a (pseudo)antiviral immune response (Lorenz et al., 2017). In addition to the endosomal TLR3, TLR7 and TLR9 as well as RLRs, e.g., RIG-I and MDA5, are potent triggers of type I interferon and proinflammatory mediator production (Allam et al., 2008; Lorenz and Anders, 2015). In fact, a gain-of-function mutation in MDA5 has been identified in both murine and human lupus-like disease (Funabiki et al., 2014; Van Eyck et al., 2015). Mice expressing a constitutively active MDA5 mutant develop anti-dsDNA autoantibodies, systemic inflammation, and glomerulonephritis, which in part depends on type I interferons (Funabiki et al., 2014). In brief, there can be no doubt aberrant TLR, NLR, and RLR activations are deeply involved in murine and human SLE pathogenesis. Here, they influence both adaptive and innate immunity. 6.1.2 PRRs in Systemic Sclerosis Systemic sclerosis is another clinical example of autoimmunity. The exact pathogenesis of this condition remains elusive. However, it can be characterized by small vessel abnormalities, aberrant innate, and adaptive autoimmunity ultimately ending in fibrosis of the skin and inner organs (Fuschiotti, 2016). Monocytic cell infiltration in the skin precedes the onset of fibrosis in patients (Ishikawa and Ishikawa, 1992; Kraling et al., 1995). Interestingly, Broen and colleagues demonstrated that a TLR2 polymorphism (Pro631) that renders myeloid DCs prone to proinflammatory mediator production correlated with antitopoisomerase positivity, disease severity, and the development of pulmonary hypertension in systemic sclerosis patients

96

S.K. Devarapu et al.

(Broen et al., 2012). In addition, dermal fibroblasts of systemic sclerosis patients express increased surface TLR2 and, therefore, show increased IL-6 production when exposed to the endogenous TLR2 ligand, serum amyloid A (O’Reilly et al., 2014). In analogy, Bhattacharyya et al. found increased TLR4 expression on fibroblasts and vascular cells within dermal or pulmonary lesions of systemic sclerosis patients. They also showed a simultaneous increase of endogenous TLR4 ligands such as the alternative spliced fibronectin domain A in five skin biopsy speciesism (Bhattacharyya et al., 2013, 2014). In vitro TLR4 activation of systemic sclerosis dermal fibroblasts resulted in an increased TGFβ response and profibrotic capacity (Bhattacharyya et al., 2013). Thus, enhanced TLR4 responsiveness for endogenous DAMPs may tip the scales toward uncontrolled fibrosis in systemic sclerosis (Bhattacharyya et al., 2013). Additional data suggest an involvement of endosomal nucleic acid-sensing TLRs in the pathogenesis of systemic sclerosis. Similar to SLE, an involvement of increased type I interferon-responsive gene expression has been demonstrated for dermal fibroblasts in systemic sclerosis (Farina et al., 2010). In this context, chronic activation of TLR3 via subcutaneous Poly(I:C) application in mice was shown to induce interferon and TGFβ depending on the gene expression in dermal fibroblast. Consequently, Poly(I:C)-treated mice developed inflammation and fibrotic remodeling of the skin, which was in part dependent on the TLR3/TRIF axis (Farina et al., 2010). Moreover, similar to the induction of type I interferons in SLE by RNA-containing immune complexes, systemic sclerosis-specific autoantibodies such as antitopoisomerase can induce type I interferon production in PBMCs (Eloranta et al., 2009; Kim et al., 2008). Combined DNase and RNase treatment of patients’ sera or blockage of Fc gamma receptors (FcyR) II on test PBMCs was able to reduce the type I interferon inducing properties of antitopoisomerase antibodies of patients sera (Kim et al., 2008). Therefore, similar to SLE, interferon induction in systemic sclerosis by autoantibodies requires FcyR-mediated uptake and eventually TLR7 or TLR9 engagement (Kim et al., 2008; Santegoets et al., 2011). In brief, these data indicate that dysregulated TLR signaling is an early event in the pathogenesis of systemic sclerosis promoting local tissue inflammation as well as systemic disease features. Although data on the involvement of NLRs and RLRs in systemic sclerosis are rare, an involvement in this disease seems likely. In fact, the NLR, especially NLRP3, is overexpressed in skin biopsies of systemic sclerosis patients and correlates with skin thickening and profibrotic

Mechanisms of Autoimmunity

97

mediator expression (Martinez-Godinez et al., 2015). Inhibition of downstream caspase-1 activation reduced secretion of the proinflammatory mediators IL-1β and IL-18 as well as the production of collagens by systemic sclerosis fibroblasts (Artlett et al., 2011).

7. TISSUE INFLAMMATION AND INJURY IN AUTOIMMUNITY 7.1 Immune Complexes The binding of autoantibodies to soluble autoantigens results in the formation of immune complexes. Immune complexes are involved in several immune responses—for example, phagocytosis, opsonization, and complement activation. Mononuclear phagocytes or red blood cells that bear the complement and Fc receptors usually efficiently clear the immune complexes from the body. However, impairment in their clearance machinery leads to their deposition and subsequent tissue injuries resulting in autoimmune diseases. Such autoimmune responses are classified as type III autoimmune responses. Immune complex-mediated inflammation and tissue injury primarily depend on the Fcγ receptors (FcγRs) that bind to Fc-binding domain of IgG. They are further classified into activating or inhibitory FcγRs and are present in immune cells, e.g., neutrophils, which upon activation by inflammatory mediators shed inhibitory FcγRs and modulate activating FcγRs (Selvaraj et al., 2004). Therefore, the activities of FcγRs at the site of inflammation determine the magnitude of neutrophil responses. In fact, mice lacking the activating FcγRs were protected from SLE, whereas mice lacking inhibitory FcγRs showed exacerbated SLE disease (Mayadas et al., 2009). In addition to FcγRs, the immune complex deposition in tissues activates the complement cascade, which mediates inflammation and tissue injury. Complement C1Q binds to the Fc part of the antibody, leading to activation of anaphylatoxins C5a and C3a (Mayadas et al., 2009). This results in secretion of proinflammatory cytokines and chemokines that activate endothelial cells. Further, activated endothelial cells increase the expression of adhesion receptors on the surface, subsequently leading to enhanced recruitment of neutrophils at the target site (Kim and Luster, 2015; Mayadas et al., 2009). In addition, the complement factor C3a also possesses the potential to induce NETosis. NETosis results in release of complement components C3a and C5a along with the DAMPs (Guglietta et al., 2016; Yuen et al., 2016). These extracellular DAMPs, along with C3a and C5a,

98

S.K. Devarapu et al.

activate other immune cells to secrete proinflammatory cytokines. Some proinflammatory cytokines, e.g., TNFα, possess the potential to induce regulated necrosis and, therefore, cause further DAMPs release from the dying cells. In addition, neutrophils recruited at the site of inflammation can directly induce cytotoxicity via FcγRs, complement receptors, and ROS generation (Mayadas et al., 2009). This sets up an autoamplification loop between cell death and inflammation, which is referred as necroinflammation (Linkermann et al., 2014; Mulay et al., 2016b,c,d) (Fig. 4). Furthermore, the FcγRs bearing neutrophils signal macrophage recruitment at the inflammatory site (Tsuboi et al., 2008), which also contribute to immune complex-mediated inflammation and tissue injury.

Fig. 4 Schematics of immune complex deposition-mediated tissue injury. The binding of autoantibodies to soluble autoantigens results in the formation of immune complexes. Impairment in their clearance machinery leads to their deposition in tissues, where the Fc part of the antibody binds to complement C1Q, and leads to the activation of C3a as well as C5a. These anaphylatoxins then activate the immune cells to secrete proinflammatory cytokines and chemokines as well as increase the recruitment of immune cells, viz., neutrophils in the injured tissues. In addition, C3a and C5a can induce cell necrosis as well as neutrophil extracellular trap (NET) formation and, therefore, induce DAMP release. DAMPs activate other immune cells to secrete proinflammatory cytokines that may possess the potential to induce regulated necrosis (e.g., TNFα), resulting in a further DAMP release. This sets up an autoamplification loop between inflammation and cell death, known as necroinflammation, which subsequently contributes to the tissue injuries resulting in autoimmune diseases.

Mechanisms of Autoimmunity

99

The more common examples of immune complex-mediated autoimmune diseases are SLE (discussed in detail later), all forms of immune complex glomerulonephritis, e.g., IgA nephropathy (Fabiano et al., 2016), and cryoglobulinemic vasculitis. Cryoglobulinemic vasculitis is a form of vasculitis that is caused by the cryoglobulin-containing immune complexes. Cryoglobulins are immunoglobulins that form solid precipitates at low temperature and redissolve at body temperature (Takada et al., 2012). Cryoglobulinemia, presence of cryoglobulins in serum, is often observed in lymphoproliferative disease (type I cryoglobulinemia), or collagen disease and infections of hepatitis C virus (type II and III cryoglobulinemia) (Takada et al., 2012). The disease affects skin, muscles, nerves, lungs, kidneys, and bone marrow (Takada et al., 2012).

7.2 Lymphocytes The key feature of most of the autoimmune diseases is the presence of autoreactive T and/or B lymphocytes in the periphery. Usually, lymphocytes have less affinity toward antigens as such, and therefore, they are unable to react to the antigen directly. However, they become activated when the APCs, which also express high levels of costimulatory molecules, present the antigen to them. 7.2.1 T Lymphocytes T lymphocytes that are present in the inflamed tissues contribute to the inflammatory processes in tissue damage. For example, their presence in the kidney of SLE patients is linked to the decreased renal function (Tsokos, 2011). T cells sense antigen via the T-cell receptor (TCR) in conjunction with the CD3-defined complex of transmembrane proteins (ε, δ, γ, and ζ) to activate a signaling process that decides the effector cell function (Moulton and Tsokos, 2015). Aberrations in this signaling pathway during autoimmune diseases lead to hyperactivated T-cell responses causing defective gene transcription, increased cytokine production, e.g., IL-17 and IFNγ, as well as excessive help to B cells to produce more autoantibodies, e.g., anti-dsDNA autoantibodies in SLE (Moulton and Tsokos, 2015). Furthermore, T cells also express several costimulatory molecules that promote B-cell differentiation, proliferation, and autoantibody production in autoimmune diseases (Kow and Mak, 2013). The TCR complex as well as the costimulatory molecules is present in the lipid rafts on T-cell membrane. Therefore, alterations in the T-cell lipid rafts in

100

S.K. Devarapu et al.

autoimmune diseases lead to a reduction in their activation threshold (Jury et al., 2004). T cells are subdivided depending on the expression of CD4, CD8, and TCR chains (α, β, γ, δ) into: A. Double-negative (CD4
CD8
) T cells. Although the origin of these cells is still not so clear, the populations of TCRαβ+CD4
CD8
T cells are increased in blood of patients with SLE as well as autoimmune lymphoproliferative syndrome (Konya et al., 2014). They produce proinflammatory IL-17 and augment the production of pathogenic anti-DNA autoantibodies associated with LN (Shivakumar et al., 1989). B. TH17 cells. CD4+ T cells undergo a specific differentiation to become TH17 cells that produce IL-17 (Wilson et al., 2007). IL-17 is considered as an important cytokine that contributes to the development of autoimmune inflammation. Mice deficient in IL-17 are protected from SLE (Amarilyo et al., 2014). In addition, the IL-17 blockade was beneficial in ameliorating inflammation in autoimmune diseases like psoriasis, rheumatoid arthritis, and uveitis (Hueber et al., 2010). C. T follicular helper cells. These are activated T helper cells that migrate from thymic extrafollicular area to germinal center where they stimulate the activated B cells to produce autoantibodies and then differentiate into T follicular helper cells expressing CXCR5 (Konya et al., 2014). They are increased in blood of patients with SLE (Yang et al., 2014) and are demonstrated to contribute to the development of lupus in Fas-deficient mice (Futatsugi-Yumikura et al., 2014). D. Tregs. Regulatory T cells are the CD4+ T cells that express the transcription factor Foxp3. They are important for maintaining the self-tolerance and thereby control the immune system. These CD4+Foxp3+ Tregs are decreased in patients with SLE, which contributes to lack of self-tolerance (Konya et al., 2014). 7.2.2 B Lymphocytes B lymphocytes contribute to the development of autoimmune diseases by different cellular functions—for example, A. Secretion of autoantibodies. B cells secrete autoantibodies against DNA, chromatin peptides, and ribonucleoproteins in SLE (Iwata and Tanaka, 2016); against ribonucleoprotein antigens in Sj€ ogren’s syndrome (Shen et al., 2016); and against DNA topoisomerase I, RNA polymerases, and fibrillin-1 during systemic sclerosis (Sakkas and Bogdanos, 2016).

Mechanisms of Autoimmunity

101

B. Presentation of autoantigens. B cells stimulate antigen-specific CD4+ T-cell proliferation after naı¨ve T-cell priming by DC (Giles et al., 2015; Ronchese and Hausmann, 1993). C. Secretion of inflammatory cytokines. B cells secrete cytokines such as IL-6, TNFα, and IL-10 as well as other immunoregulatory cytokines such as IL-2, IFNγ, IL-12, and IL-4 when stimulated with Th cells and antigens (Harris et al., 2000). D. Generation of ectopic germinal centers. B cells help in the process of ectopic germinal center formation during the chronic inflammatory condition. Such ectopic germinal centers are observed in SLE, rheumatoid arthritis, Sj€ ogren’s syndrome, multiple sclerosis, and type 1 diabetes (Aloisi and Pujol-Borrell, 2006; Hampe, 2012). B cells are activated when the antigen binds to the BCR (Yuseff et al., 2013). Other lymphoid cells that cause cytotoxicity and contribute to inflammation by secreting cytokines include NK cells, CD56+ T cells, NKT cells, gamma/ delta (γδ) T cells, and mucosal-associated invariant T cells (Doherty, 2016).

7.3 Monocytes and Macrophages Monocytes and macrophages are involved in various important functions that regulate innate immunity—for example, cytokine production, antigen presentation to T cells, and phagocytosis. They recognize and remove pathogens as well as dead or damaged host cells. Abnormalities in these functions of monocytes and macrophages are linked to the pathogenesis of several autoimmune diseases (Brunini et al., 2016; Hamerman et al., 2016; Katsiari et al., 2010; Roberts et al., 2015). The immune responses by these cells are governed by the expression of surface markers, which help them to sense the environment and respond to it. For example, abnormalities in the FcγRs, involved in phagocytosis as well as inflammatory cytokine production, contributed to the development of LN in NZB/WF1 murine model of lupus (Bergtold et al., 2006; Clynes et al., 1998). The antibodies that form immune complexes have been demonstrated to interact with the FcγRs present in the immune cells (Bergtold et al., 2006; Nimmerjahn and Ravetch, 2008). Such aberrant activation of monocytes and macrophages causes increased production of inflammatory cytokines in SLE, rheumatoid arthritis, as well as Sj€ ogren’s syndrome (Katsiari et al., 2010; Roberts et al., 2015; Tsokos, 2005). These cytokines further contribute to aggravation of the autoimmune disease—for example, IL-6 and IL-10 promoted IgG production; IL-6 and IFN-1 together can

102

S.K. Devarapu et al.

induce B-cell maturation in SLE patients (Jego et al., 2003); whereas IL-6, IL-23, TNFα, IL-1β, and TGFβ have been implicated in the pathogenesis of rheumatoid arthritis (Estrada-Capetillo et al., 2013; Roberts et al., 2015). Recently, it is reported that in healthy individuals an autoregulatory feedback mechanism exists between DCs and regulatory B cells that are mediated by IFNα, and alteration in this mechanism results in SLE (Menon et al., 2016). Monocytes are instrumental in both inductions and maintenance of the immune tolerance. A phenotypic analysis of monocyte-derived DCs from SLE patients in the presence of IL-10 revealed enhanced levels of HLA-DR, CD80, CD9, and CD151 tetraspanins, FN1 (a class II MHC-tetraspanin epitope), CD85j/ILT2, and CD69, suggesting that these cells have an enhanced capacity of antigen representation (Figueroa-Vega et al., 2006). Furthermore, the expression of costimulatory molecules is critical for antigen presentation. Monocytes from SLE patients express high levels of CD40, ICAM-1, STAT-1, and CD69 at the baseline, suggesting their activation status; they indicate an increased costimulatory potential (Kuroiwa et al., 2003). Monocytes and macrophages also play a crucial role in host defense by phagocytosing pathogens as well as dying cells. Increased apoptosis and deficiency in clearing apoptotic cells are considered an important pathomechanism, leading to autoimmune diseases (Biermann et al., 2014). Impaired clearance of dying cells promotes the abundance of autoantigens derived from necrotic cells and thus increased antigen presentation to T and B lymphocytes resulting in increased autoantibody production, chronic inflammation, and severe tissue damage (Biermann et al., 2014).

7.4 Tertiary Lymphoid Organs The persistence exposure of antigens increases the continuous need for extravasation of leukocytes during an autoimmune attack, leading to the formation of the TLOs (Jones and Jones, 2016). The term TLOs is referred to: A. The anatomically distinct infiltrates those are adjacent to T- and B-cell compartments. B. T-cell compartment rich in fibroblasts reticular cells. C. T-cell compartment rich in PNAd+ or MECA79+ high endothelial venules.

Mechanisms of Autoimmunity

103

D. Evidence of B-cell class switching and germinal center reactions in B cells. E. The presence of activation-induced cytidine deaminase enzyme. F. The presence of follicular DCs (Neyt et al., 2012). The formation of TLOs at the site of chronic inflammation is induced by the cytokines and chemokines. For example, overexpression of IL-22, IL-7, LTα, CCL21, and CXCL13 promoted the formation of TLOs (Barone et al., 2015; Marinkovic et al., 2006; Neyt et al., 2012), whereas deletion of CXCL13, CXCR5, and CCR7 inhibits the formation of TLOs (Rangel-Moreno et al., 2007; Wengner et al., 2007; Winter et al., 2010). TLOs allow the activation of naive T cells and B cells by DCs within the germinal centers as well as induce self-reactive T lymphocytes and antibodies (Lee et al., 2006; Moyron-Quiroz et al., 2004). The formation of TLOs is observed in various autoimmune diseases including type 1 diabetes, SLE, rheumatoid arthritis, as well as autoimmune thyroiditis (Armengol et al., 2001; Chang et al., 2011; Lee et al., 2006; RangelMoreno et al., 2006).

8. GENETIC RISK FACTORS FOR ORGAN MANIFESTATIONS IN HUMAN AUTOIMMUNE DISEASES Loss of organ-specific tolerance can be attributed to either lack of thymic presentation of organ-specific antigen or altered antigenicity within the target organ. The identification of the common genetic risk variants, their frequencies in the population (risk allele frequency), and the risks of disease they confer (odds ratio) by the genome-wide association studies (GWASs) have revealed very important information on the genetic risk factors for human autoimmune diseases (Goris and Liston, 2012; Iles, 2008). We have summarized the genetic loci associated with lack of organ-specific tolerance and development of organ-specific autoimmune diseases later. Genetic defects affecting the negative selection process increase the susceptibility to autoimmune disease. For example, the APS-1 is monogenic autoimmune disease because of mutations in gene AIRE (Finnish-German, 1997; Nagamine et al., 1997). Multiple mechanisms of action for AIRE have been suggested, viz.: (1) Aire protein has a transcriptional activity (Kumar et al., 2001; Pitkanen et al., 2000) and is directly involved in the expression

104

S.K. Devarapu et al.

of self-antigen within thymus (Ruan et al., 2007), (2) Aire may function via intermediates, viz., cofactors (Ilmarinen et al., 2008), or indirectly recruits transcriptional components required for target gene expression (Org et al., 2008; Tao et al., 2006). Aire deficiency interacts with the loci associated with the common autoimmune diseases and alters the disease progression in APS-1 patients (Halonen et al., 2002; Kogawa et al., 2002). The Aire pathway is very sensitive, and a slight reduction in Aire expression leads to reduced expression of Aire-dependent antigens. This indicates that small changes in the thymic expression of Aire-dependent antigens may contribute to autoimmune susceptibility in a disease-specific manner (Kont et al., 2008; Liston et al., 2004). For example, the susceptibility alleles of the INS locus cause a two- to threefold decrease in AIRE-dependent thymic expression of the insulin gene (Taubert et al., 2007; Vafiadis et al., 1997). Furthermore, Chrna1 is an organ-specific gene expressed in an AIRE-dependent manner in the thymus. CHRNA1 has allelic variants with reduce thymic expression of the antigen resulting in the autoimmune disease, myasthenia gravis (Giraud et al., 2007). Other susceptibility gene variants with reduced thymic expression are MBP that is associated with multiple sclerosis, and TSHR and TG that are associated with autoimmune diabetes. The common factor involved in all these cases is the organ-specific expression of target antigens. However, in some cases the putative genes might also be associated with imbalance between thymic and peripheral expression—for example, PADI4 locus with rheumatoid arthritis (Yamada and Yamamoto, 2007) and the TREX1 and DNASE1 loci with SLE (Lee-Kirsch et al., 2007; Shin et al., 2004). The PADI4 encodes for peptidyl citrulline, TREX1, and DNASE1 producing DNA fragments. Association of the PDS locus, which produces pendrin, with autoimmune type 1 diabetes explains the synergy between genetic and environmental factors. Pendrin contributes to the iodination of thyroglobulin and increases its antigenicity (Barin et al., 2005). Apart from the expression and antigenicity of the antigen, the other aspect of the immune tolerance revolves around the efficient antigen presentation and that makes MHC-II gene within the HLA locus an important factor. For many autoimmune diseases, HLA is the most important genetic contributor (Chung et al., 2014; Fernando et al., 2008; Goris and Liston, 2012). Two alternative mechanisms exist for the association of altered peptide presentation to autoimmunity, viz., the reduced thymic presentation of major target autoantigens for tolerogenic purposes, and the enhanced peripheral presentation of major target autoantigens. Substitution of aspartic residue at position 57 of HLA-DQ8 with serine, alanine,

Mechanisms of Autoimmunity

105

or valine increases the binding of insulin peptide to HLA-DQ and may elicit an autoimmune response (Faas and Trucco, 1994). Similarly, HLA-DR2 variants have an increased ability to display a dominant epitope of MBP to CD4+ T cells and contribute to the development of multiple sclerosis (Li et al., 2005; Maynard et al., 2005). In addition, some variants of class II HLA-DRB1 are biased toward recognizing a cartilage-specific protein CII in rheumatoid arthritis (Burkhardt et al., 2006). Furthermore, recent GWASs of LN identified multiple LN-susceptibility loci that are independent of HLA regions. For example, the MHC locus rs9263871 located within HCG27 had strongest association with LN (Chung et al., 2014). Other loci that are associated with LN include rs1364989 on 4q11–q13, upstream of PDGF receptor α; rs274068 on 16p12.1 within an intronic region of sodium-dependent glucose cotransporter SCL5A11; and rs7834765 on 8q24.12 as well as other two regions on 6p22 and 9p21 (Chung et al., 2014). Apart from loss of immune tolerance in an antigen-specific manner, other mechanisms can also participate in organ-specific autoimmune disease, viz., increased immune trafficking in the target organ. Multiple genes associated with Crohn’s disease are suggestive of increased leukocyte trafficking, including Atg16l1, Irgm, Mdr1, Mst1, Ncf4, and Nkx-2.3 (Ellson et al., 2006; Rioux et al., 2007; Singh et al., 2006; Suh et al., 2006). In autoimmune diabetes, gene TNFR2 is responsible for increased damage to the islet cells due to prolonged signaling through TNFR2 (Walter et al., 2000). Moreover, GWASs revealed the contribution of PTPN22 to autoimmune type 1 diabetes and rheumatoid arthritis (Begovich et al., 2004; Bottini et al., 2004), as well as NOD2 and IL-23R to Crohn’s disease (Duerr et al., 2006; Hugot et al., 2001; Ogura et al., 2001). In conclusion, the development of autoimmune disease is, therefore, an additive effect of defects in multiple immune tolerance mechanisms being directed toward a specific target organ by additional organ-specific defects, heavily modified by environmental influences.

9. LUPUS NEPHRITIS SLE and its most common organ manifestation, LN, strikingly reflect the implications of systemic autoimmunity and autoimmune tissue injury in clinical reality. In this chapter, we will recapitulate the factors involved in breaking tolerance against self during SLE and describe the mechanisms acting inside the nephritic kidney.

106

S.K. Devarapu et al.

9.1 Systemic Autoimmunity in SLE Genetic susceptibility predisposes to SLE. SLE is rather a clinical syndrome than a defined disease entity. Patients present with variable symptoms and organ involvement can differ among them. Yet, they share serological features like ANAs that document the loss of tolerance against self-nuclear acids. However, what are the reasons for this? Familial aggregation studies support a genetic predisposition to SLE (Alarcon-Segovia et al., 2005; Ghodke-Puranik and Niewold, 2015). In general, SLE follows a polygenetic inheritance pattern, meaning that many risk alleles of moderate effect size (odds ratio 1, 5-2, 5) are present in one individual prone to developing SLE (Ghodke-Puranik and Niewold, 2015). GWASs have identified over 40 risk loci that correlate with an increased risk to develop SLE (Ghodke-Puranik and Niewold, 2015). Strong associations have been found for HLA loci and SLE (Ghodke-Puranik and Niewold, 2015). In addition, genetic alterations in genes promoting type I IFN responses or reducing clearance of apoptotic materials such as IRF5, DNase I, or complement deficiencies have been identified as risk loci in SLE (Ghodke-Puranik and Niewold, 2015; Niewold et al., 2008; Truedsson et al., 2007; Yasutomo et al., 2001). SLE is a heterogeneous disease. In order to understand the heterogenicity of SLE, Banchereau et al. profiled the blood transcriptome of a longitudinal cohort of pediatric lupus patients and have identified a plasmablast signature as a biomarker of disease activity (Banchereau et al., 2016). They also identified neutrophil-related immune signatures with progression to LN (Banchereau et al., 2016). This approach of personalized immunomonitoring also enabled patient stratification into groups that might facilitate implementation of personalized therapies in SLE (Anders and Kretzler, 2016; Anders et al., 2015; Banchereau et al., 2016). 9.1.1 Apoptotic Material Triggers an Inappropriate Immune Response In order to prevent inappropriate immune reactions against self-nuclear acids and associated proteins, the apoptotic material is usually rapidly cleared from the environment (Lorenz et al., 2014b). DNase and RNase degrade extracellular nuclear material (Lorenz et al., 2014b; Yasutomo et al., 2001). C1q marks apoptotic bodies for SCARF1 receptor-dependent clearance by phagocytes (Ramirez-Ortiz et al., 2013). In SLE patients, genetic alterations weaken these defense mechanisms. As a consequence,

Mechanisms of Autoimmunity

107

nuclear material persists in the tissue compartment and can undergo changes rendering it immune stimulatory (Lorenz et al., 2017). In addition, defects in lysosomal degradation of nuclear material lead to prolonged persistence of nuclear material within phagocytes and may facilitate the recycling of DAMPs to the cell membrane (Monteith et al., 2016). Additional sources of extracellular immunogenic material are netting neutrophils (Leffler et al., 2015; Lindau et al., 2014). Interestingly, SLE patients that present with a reduced ability to degrade those NETs also have a reduced ability to degrade secondary necrotic particles and have an increased likelihood of nephritic involvement (Leffler et al., 2015). Cell necrosis in response to injuries or sunburns also increases the load of necrotic material that has to be cleared. Failure of clearance of these intracellular autoantigens induces SLE (Lorenz et al., 2017; Munoz et al., 2010). The deficiency of the tyrosine kinase c-Mer and the Milk fat globule protein factor 8, which are involved in recognition and clearance of early apoptotic cells, results in anti-DNA autoantibodies and SLE (Cohen et al., 2002; Hanayama et al., 2004). The extracellular chromatin because of cell necrosis as well as neutrophil extracellular trap formation is cleared efficiently by endonucleases such as DNase 1, and therefore, the loss of function of DNAse 1 has been associated with renal manifestations in murine SLE (Seredkina and Rekvig, 2011). Furthermore, defects in LAP, a form of noncanonical autophagy to remove dying cells, also contribute to the pathogenesis of murine SLE (Martinez et al., 2016). 9.1.2 Mistaking Self-Nuclear Components for Invading Virus High titer ANA production implies an inappropriate innate and adaptive immune response against self-nuclear acids and associated proteins. Prolonged presence of nuclear particles in the extracellular matrix can reverse epigenetic modifications that normally inhibit self-nuclear acid recognition by PRRs (Kariko et al., 2004; Lorenz et al., 2014b). Some drugs, e.g., procainamide, were also shown to inhibit DNA methylation, thereby enhancing immune simulative properties of self-DNA (Cornacchia et al., 1988). Consequently, after engulfment into phagocytes, secondary necrotic material gains the ability to stimulate innate immune responses via TLR3, 7, 8, and 9, NLRs like AIM2, or RLRs like MDA5, as discussed earlier (Allam et al., 2008; Lorenz et al., 2014b; Zhang et al., 2013). Similar to an infection with intracellular virus innate immune cells, pDCs produce large amounts of type I interferons and initiate

108

S.K. Devarapu et al.

proinflammatory cell response programs (Lorenz et al., 2014b; Savarese et al., 2006). In this context, DNA-containing immune complexes and U1snRNP were shown to activate TLR9 or TLR7 in pDCs, respectively (Leadbetter et al., 2002; Savarese et al., 2006). Since cellular responses to improperly cleared nuclear material mimic those during viral infection, it is likely that viral infection in SLE patients delivers a potent stimulus of active disease flares (Theofilopoulos et al., 2005). These analogies between SLE and viral infectious diseases have led to the concept of pseudo antiviral immunity as a model for SLE pathogenesis (Lorenz et al., 2017). 9.1.3 Directing Adaptive Immunity Against Autoantigens APCs, i.e., DCs, represent the link between innate and adaptive immunity. Due to persistent activation by endogenous immune stimuli, DC’s life span is prolonged in SLE (Guiducci et al., 2010). The pDCs in SLE exhibit increased expression of the IRF3, which goes along with increased type I IFN production in patients (Santana-de Anda et al., 2014). The DCs further show increased expression of costimulatory molecules like CD40 and CD86, as well as an increased activating/inhibitory FcγR expression ratio, which might contribute to a break in self-tolerance in SLE pathogenesis (Carren˜o et al., 2009; Mackern-Oberti et al., 2015). Although the precise mechanisms in breaking T-cell tolerance remain elusive, reduced ability to induce PD1L expression on DCs, which can mediate T-cell suppression via ligation of PD1 on T cells, could promote autoreactive T-cell expansion in SLE (Mackern-Oberti et al., 2015; Mozaffarian et al., 2008). Activated DCs can directly stimulate B-cell proliferation, maturation, and autoantibody production in murine models of SLE (Wan et al., 2008). Furthermore, in SLE, RNA- and DNA-containing autoantigens can directly activate autoreactive B cells via dual engagement of TLR7 or TLR9 and the BCR (Lau et al., 2005; Leadbetter et al., 2002). TLR7 overexpression solely in B cells is sufficient to trigger autoantibody production and lupus progression in Sle1 lupus-susceptible mice (Hwang et al., 2012). The long-lived autoreactive plasma cells rather than the short-lived plasmablasts produce autoantibodies and contribute to chronic humoral autoimmunity in BZB/W mice (Hoyer et al., 2004; Moser et al., 2006). In summary, aberrant stimulation of B cells by DCs and continuous exposure to endogenous PRR ligands foster the selection of self-reactive B cells and promotes autoantibody production.

Mechanisms of Autoimmunity

109

9.2 Autoimmunity and Tissue Inflammation Inside the Kidney 9.2.1 Immune Complex Formation Inside the Kidney LN manifests because of systemic autoimmunity. The formation of immune complexes rather than their passive deposition in different glomerular compartments is a diagnostic hallmark of LN (Lorenz et al., 2017). Immune complexes can harm glomerular cells in different ways. Resident renal cells and immune cells can bind Immunoglobulins via FcR. Immune complexes have the potential to activate the C1q-dependent classical complement pathway. In turn, the proteolytic product C5a serves as a chemoattractant for infiltrating immune cells that cause inflammation and injury (Lorenz et al., 2014b; Yung and Chan, 2015). However, the contribution of complement pathway in the development of SLE is paradoxical. For example, the classical complement C1q participated in phagocytosis of apoptotic cells and its deficiency leads to the development of SLE (Botto, 2001). The anti-dsDNA antibodies may also cross-react within renal cell antigens, such as Annexin II or α-actinin (Yung and Chan, 2015). Last, nuclear material can ligate innate PRRs on resident renal cells and infiltrating immune cells to stimulate local tissue inflammation and cytokine production (Allam et al., 2009; Flur et al., 2009). Immune complexes contain IgM, IgA, and IgG and seem to form in the kidney after binding to renal matrix components (Krishnan et al., 2012; Tojo et al., 1970). Immune complexes can be located in the mesangium, the subendothelial, or the subepithelial space. Outside the glomerular immune complexes can be found along peritubular capillaries (Yu et al., 2010). The histopathological classification of LN reflects their distribution within the renal compartments. Class I and II LN means the presence of mesangial immune complexes. Subendothelial and subepithelial immune complexes can be found in class III and IV or V LN, respectively (Weening et al., 2004). 9.2.2 Innate Immune Signaling Inside the Nephritic Kidney As mentioned earlier, immune complexes have the potential to activate innate PRRs within renal parenchymal cells and infiltrating immune cells. Mesangial cells are reactive to DNA or RNA antigens that aggravate LN (Allam et al., 2008, 2009). Activation of MDA5 in mesangial by exogenous RNA triggers potent type I interferon responses (Flur et al., 2009). Ligation of TLR3 in mesangial cells can upregulate the expression of CXCL1, a strong neutrophil chemoattractant that is enriched during LN (Imaizumi et al., 2014). Glomerular endothelial cells were shown to depend on RIG-I

110

S.K. Devarapu et al.

rather than MDA5 to produce type I interferons and proinflammatory mediators such as IL-6 in response to double-stranded RNA (Hagele et al., 2009). TLR4 is also expressed on podocytes during nephritis. Ligation of TLR4 results in the production of chemokines by podocytes (Banas et al., 2008). Overall, sensing of DAMPs via PRRs in renal parenchymal cells promotes intrarenal inflammation and tissue injury. Furthermore, Stamatiades et al. demonstrated the presence of kidney-specific macrophages that monitor the transport of proteins and particles into the renal interstitium. These kidney macrophages also detect and scavenge the immune complexes in the interstitium and trigger an FcγRIV-dependent inflammatory response and the recruitment of monocytes and neutrophils (Stamatiades et al., 2016). 9.2.3 Immune Cell Infiltration Into Renal Tissue Inflammatory mediators lure a great variety of cells into the nephritic kidney. In some cases, there even is TLO formation inside the tubulointerstitial compartment. Here, B cells can proliferate and undergo somatic hypermutation stimulated by T cells (Chang et al., 2011). In NZB/W mice, autoreactive long-lived anti-dsDNA-specific plasma cells were shown to reside in the kidneys in large numbers (Espeli et al., 2011; Moser et al., 2006). In humans, intrarenal plasma cells correlate with disease severity (Espeli et al., 2011). Among T cells, TH17 cells are currently under intensive investigation in the context of SLE. Increased numbers of these cells along with a skewed TH17/Treg balance have been demonstrated for SLE patients. Th17 T cells infiltrate the renal tissue and promote tissue damage (Koga et al., 2016). Although LN and interstitial infiltrates can develop in the absence of DCs, they promote the progression of nephritis in MRL/lpr mice via stimulating local and systemic inflammatory responses (Sahu et al., 2014; Teichmann et al., 2010). Several different subtypes of DCs and macrophages have been identified inside the kidney and indicate poor prognosis (Bethunaickan et al., 2011). Murine macrophages during nephritis display neither a classical pro- nor an antiinflammatory phenotype. Sahu et al. suggest that this could indicate the missing resolution of inflammatory responses (Sahu et al., 2014). Further research is needed to exactly define the roles of different DC and macrophage subsets during LN. Finally, neutrophils can promote tissue injury by releasing cytokines or ROS inside the kidney (Worthmann et al., 2014). Additionally, they deliver DAMPs to the extracellular space when undergoing NETosis (Lorenz et al., 2014b). Together, immune complexes, aberrant pattern recognition, and misdirected cellular

Mechanisms of Autoimmunity

111

responses trigger recurrent flares of renal inflammation and consecutive tissue injury. The latter can trigger excessive proliferation of renal progenitor cells and exaggerated extracellular matrix production (Smeets et al., 2009). Ultimately, this leaves sclerosing lesions inside most of the glomeruli, which is defined as class VI LN (Weening et al., 2004).

9.3 Animal Models for SLE Most mechanistic data on the etiology and progression of SLE originate from murine models of lupus-like autoimmunity. Similar to human SLE, murine models reflect different modalities of disease pathogenesis, which result in different phenotypes (Peng, 2012). Animal models have been successfully used to identify genetic aberrations associated with disease development and to test therapeutic strategies (Peng, 2012). Nevertheless, translational scientists have to keep several limitations and features of the murine lupus model in mind. The following section will briefly introduce widely used mouse models of SLE and their resulting phenotype. We will further highlight some similarities and differences to human disease. In general, we distinguish spontaneous models (MRL/lpr, NZB/NZW.F1, BXSB/ Yaa) from pharmacologically induced models (e.g., pristane-induced) (Peng, 2012). The latter can be used to study the onset of disease, whereas spontaneous models reveal insight into how genetic alterations predispose to the development of SLE (Peng, 2012). 9.3.1 NZB/NZW.F1 The F1 hybrid from phenotypically mostly unaffected NZB and New Zealand White (NZW) mice results in NZB/W F1 mice (Peng, 2012). In comparison to their ancestors, these mice develop an exaggerated lupus-like phenotype including lymphoproliferation, splenomegaly, ANA production, and immune complex-mediated glomerulonephritis (Table 6) (Peng, 2012). Life expectancy is reduced due to renal failure. Similar to human SLE, severity of the disease is depending on the gender in these mice. Analysis of similarly diseased NZM2410 mice derived from backcrossed NZB/W F1 with NZW mice revealed several lupus susceptibility loci (Sle1-Sle3) linked to MHC-II-related and -nonrelated genes. Subsequent experiments revealed that introduction of these risk alleles into nonautoimmune background (C57BL6) leads to autoantibody production; however, it failed to trigger nephritis (Peng, 2012). This ancestral history resembles the concept of human SLE, where the combinations of several risk loci rather than a monogenetic aberration are needed to

112

S.K. Devarapu et al.

Table 6 Comparison of Phenotypical Features and Cause of Disease in Animal Models of SLE NZB/W F1 MRL/lpr BXYB/Yaa Pristane Phenotype

Spontaneous

+

+

+




Gender

Female

Male/female

Male

Female > male

Arthritis




+




+

Vasculitis

+

+

+

+

Nephritis

+

+

+

+/
*

Mortality (average)

9 months 6 months

5 months

*

Anti-dsDNA Ab

+

+

+

+

Anti-Smith Ab




+




+

Anti-U1snRNP Ab




+




+

Cause

c LSL

Fas
/
> c LSL

Yaa (TLR7") > LSL

TLR7/type I

Skin involvement

LSL, combined lupus susceptibility loci; * depends on background: + in BALB/c, weak in C57BL/6, strong in lupus-prone backgrounds (e.g., MRL/lpr); +, positive; –, negative.

enable disease development. Nevertheless, NZB/W F1 mice fail to develop humoral immunity against U1snRNP, a common human lupus autoantigene (Peng, 2012). 9.3.2 MRL/lpr The MRL/lpr mice develop lymphoproliferation, accumulation of CD4
CD8
T cells, a broad variety of autoantibodies, and severe immune complex nephritis (Peng, 2012) (Table 6). Unlike other models, both males and females develop the disease (Peng, 2012). Skin involvement is a special characteristic of this model. The main cause of the phenotype relates to deficiency in Fas receptor, which belongs to the TNFR superfamily, and induces lymphocyte apoptosis upon ligation by FasL (Peng, 2012). Defective clearance of autoreactive lymphocytes and innate immune cells is considered the main cause of the phenotype. Interestingly, selective

Mechanisms of Autoimmunity

113

deletion of Fas in DCs was sufficient to trigger autoimmunity in C57BL6 mice (Stranges et al., 2007). Additionally, lupus susceptibility loci have been identified within the MRL background that predispose to autoimmunity and associate with disease features (Vidal et al., 1998). Interestingly, deletion of Fas in humans does not classically result in SLE, but rather induces an autoimmune lymphoproliferative disorder without nephritis (Teachey et al., 2010). In addition, the main mechanism of disease induction is a defect in lymphocyte apoptosis (Reap et al., 1995). However, peripheral T lymphocytes from patients demonstrate increased apoptosis, which has been correlated to disease severity (Dhir et al., 2009). 9.3.3 BXSB/Yaa The BXSB/Yaa strain represents the strongest phenotype. On average male mice die after 5 months due to proliferative GN (Peng, 2012). In this model the BXSB bears several different lupus susceptibility loci (BXS1–6) on chromosomes 1, 2, 13, some of which per se confer lupus phenotypes (Haywood et al., 2004). However, in this background, Y-linked autoimmune accelerator (Yaa) has been shown to be the main driver of disease. Yaa is caused by a translocation of X chromosomal ends to the Y chromosome, thereby increasing the gene dose of several genes including TLR7. In this model, increased TLR7 gene expression accounts for many immune cell aberrations, lymphoproliferation, glomerulonephritis, and mortality (Fairhurst et al., 2008). As expected, only male BXSB/Yaa mice develop full-blown disease. Interestingly, in Chinese and Japanese patients an SNP within the TLR7 gene has first been linked to human SLE selectively in male patients (Shen et al., 2010). Meanwhile, TLR7 risk alleles have also been identified in female patients with SLE (dos Santos et al., 2012). As a limitation, monogenetic Lupus is rare in humans. 9.3.4 Pristane The pristane model is a pharmacologically induced model of SLE. It works within several different backgrounds including Balb/c and C57BL6. After i.p. injection with isoprenoid alkane, pristine, mice develop autoantibodies, including U1snRNP, and depending on the background show renal involvement and develop a mild erosive arthritis (Table 6) (Leiss et al., 2013). As such, pristane represents the influence of environmental triggers on SLE development (Peng, 2012). Further, although C57/BL10 mice injected with pristane developed hemorrhagic pulmonary capillaritis (Chowdhary et al., 2007), whether it mimics rare pulmonary capillaritis

114

S.K. Devarapu et al.

in human SLE remains elusive (Zamora et al., 1997). In its function as an autoadjuvant, pristane-induced SLE depends on type I interferon signaling of peritoneal Ly6Chigh monocytes and on the TLR7 signaling axis since the ablation of any of those factors rescues the phenotype (Lee et al., 2008a,b). A type I interferon signature is a common finding in SLE patients, which generates a value for the pristane model to study this aspect. However, in patients, pDCs are the main source of these mediators (Lorenz and Anders, 2015).

10. SUMMARY Unlike autoinflammation and alloimmunity, autoimmunity originates from a spontaneous loss of tolerance against self-proteins and other structures. A number of environmental factors such as drugs, infections, and acquired epigenetic modifications altering gene regulation determine and enhance the susceptibility to the loss of tolerance. Gene variants can weaken or break the checkpoints that maintain immune tolerance in the immune system. Beyond extremely rare monogenetic forms of autoimmunity, the combinations of genetic and environmental factors need to pass a threshold that compromises the safeguard mechanisms of immune tolerance. Infections are common triggers of breaking this threshold in susceptible patients. Although autoimmunity is often transient and potentially harmless, it holds the risk of persistent autoimmunity whenever autovaccination occurs and immune memory is imprinted, implying a potentially lifelong persistence of autoreactive lymphocyte subsets. Consequently, curing of chronic autoimmune disease is hardly possible unless eradicating immune memory that is laid down in long-lived plasma cells in the bone marrow. Hence, the management of autoimmune diseases focusses largely on the control of lymphocyte proliferation to limit the size of autoreactive lymphocyte clones. Indeed, the driving force of autoimmune tissue injury is underlying autoimmune mechanisms located in lymphoid organs. Clonal expansion of the autoreactive lymphocyte clones leads to humoral and cellular immune responses. Immune complex disease, infiltration of autoantigen-specific T cells, and TLO formation contribute to autoimmune tissue injury, inflammation, atrophy, and subsequently organ dysfunction. Currently, the strategy for therapy includes modulating the systemic and peripheral pathomechanisms of autoimmune disease. Inducing remission, maintaining a sufficient suppression of disease activity, and preventing relapsing disease are three different treatment targets to consider. Another

Mechanisms of Autoimmunity

115

treatment target is tissue inflammation, where the problem is to identify nonredundant elements of tissue injury, e.g., complement factors, FC receptors, or Jak/STAT-mediated cytokine signaling. These are currently addressed by steroid therapy for most autoimmune diseases, but more specific targets that allow selective inactivation of lymphocyte cytokines and proinflammatory mediators are approaching the clinic or have already been implemented in the treatment of rheumatoid and psoriatic arthritis. Indeed, novel therapies also help to better understand the pathophysiology. For example, rheumatoid arthritis was considered as a T cell-driven disease up to the moment where anti-CD20 therapies became available and demonstrated profound effects on clinical outcomes of rheumatoid arthritis. This observation implied an unexpected but a central role of CD20+ B cells in the pathogenesis of rheumatoid arthritis and shifted the research interest in this direction. Therefore, we conclude that both bench-to-bedside and bedside-to-bench researches are important approaches to understand immune tolerance and to unravel the pathophysiology of autoimmune diseases. However, several studies often present variable or contradictory results especially while understanding the involvement of EVs, as well as NETs, in the disease pathology, the most probable reason being in the difficulties in their characterization and standardization of the analytical techniques. Whether EVs, as well as NET components, can be used as biomarkers in patients with autoimmune diseases remains to be studied.

ACKNOWLEDGMENTS S.R.M. is supported by the Deutsche Forschungsgemeinschaft (MU 3906/1-1). H.-J.A. is supported by the European Union’s Horizon 2020 research and innovation program under Grant Agreement No. 668036 (RELENT). The views expressed here are the responsibility of the author(s) only. The EU Commission takes no responsibility for any use made of the information set out. Conflict of interest statement: None.

REFERENCES Abdel-Wahab, O., Mullally, A., Hedvat, C., Garcia-Manero, G., Patel, J., Wadleigh, M., Malinge, S., Yao, J., Kilpivaara, O., Bhat, R., Huberman, K., Thomas, S., Dolgalev, I., Heguy, A., Paietta, E., Le Beau, M.M., Beran, M., Tallman, M.S., Ebert, B.L., Kantarjian, H.M., Stone, R.M., Gilliland, D.G., Crispino, J.D., Levine, R.L., 2009. Genetic characterization of TET1, TET2, and TET3 alterations in myeloid malignancies. Blood 114, 144–147. Abolhassani, H., Gharib, B., Shahinpour, S., Masoom, S.N., Havaei, A., Mirminachi, B., Arandi, N., Torabi-Sagvand, B., Khazaei, H.A., Mohammadi, J., Rezaei, N., Aghamohammadi, A., 2015. Autoimmunity in patients with selective IgA deficiency. J. Investig. Allergol. Clin. Immunol. 25, 112–119.

116

S.K. Devarapu et al.

Admyre, C., Johansson, S.M., Paulie, S., Gabrielsson, S., 2006. Direct exosome stimulation of peripheral human T cells detected by ELISPOT. Eur. J. Immunol. 36, 1772–1781. Alarcon-Segovia, D., Alarcon-Riquelme, M.E., Cardiel, M.H., Caeiro, F., Massardo, L., Villa, A.R., Pons-Estel, B.A., 2005. Familial aggregation of systemic lupus erythematosus, rheumatoid arthritis, and other autoimmune diseases in 1,177 lupus patients from the GLADEL cohort. Arthritis Rheum. 52, 1138–1147. Alberts, B., Johnson, A., Lewis, J., et al., 2002. Molecular Biology of the Cell, fourth ed. Garland Science, New York. Allam, R., Pawar, R.D., Kulkarni, O.P., Hornung, V., Hartmann, G., Segerer, S., Akira, S., Endres, S., Anders, H.J., 2008. Viral 5’-triphosphate RNA and non-CpG DNA aggravate autoimmunity and lupus nephritis via distinct TLR-independent immune responses. Eur. J. Immunol. 38, 3487–3498. Allam, R., Lichtnekert, J., Moll, A.G., Taubitz, A., Vielhauer, V., Anders, H.J., 2009. Viral RNA and DNA trigger common antiviral responses in mesangial cells. J. Am. Soc. Nephrol. 20, 1986–1996. Aloisi, F., Pujol-Borrell, R., 2006. Lymphoid neogenesis in chronic inflammatory diseases. Nat. Rev. Immunol. 6, 205–217. Amarilyo, G., Lourenco, E.V., Shi, F.D., La Cava, A., 2014. IL-17 promotes murine lupus. J. Immunol. 193, 540–543. Amigorena, S., Choquet, D., Teillaud, J.L., Korn, H., Fridman, W.H., 1990. Ion channel blockers inhibit B cell activation at a precise stage of the G1 phase of the cell cycle. Possible involvement of K+ channels. J. Immunol. 144, 2038–2045. Anandasabapathy, N., Ford, G.S., Bloom, D., Holness, C., Paragas, V., Seroogy, C., Skrenta, H., Hollenhorst, M., Fathman, C.G., Soares, L., 2003. GRAIL: an E3 ubiquitin ligase that inhibits cytokine gene transcription is expressed in anergic CD4 + T cells. Immunity 18, 535–547. Anders, H.J., 2009. Pseudoviral immunity—a novel concept for lupus. Trends Mol. Med. 15, 553–561. Anders, H.J., 2010. Toll-like receptors and danger signaling in kidney injury. J. Am. Soc. Nephrol. 21, 1270–1274. Anders, H.J., Fogo, A.B., 2014. Immunopathology of lupus nephritis. Semin. Immunopathol. 36, 443–459. Anders, H.J., Kretzler, M., 2016. Glomerular disease: personalized immunomonitoring in lupus and lupus nephritis. Nat. Rev. Nephrol. 12, 320–321. Anders, H.J., Krug, A., Pawar, R.D., 2008. Molecular mimicry in innate immunity? The viral RNA recognition receptor TLR7 accelerates murine lupus. Eur. J. Immunol. 38, 1795–1799. Anders, H.J., Weidenbusch, M., Rovin, B., 2015. Unmet medical needs in lupus nephritis: solutions through evidence-based, personalized medicine. Clin. Kidney J. 8, 492–502. Anderson, M.S., Venanzi, E.S., Klein, L., Chen, Z., Berzins, S.P., Turley, S.J., von Boehmer, H., Bronson, R., Dierich, A., Benoist, C., Mathis, D., 2002. Projection of an immunological self shadow within the thymus by the aire protein. Science 298, 1395–1401. Ansari, M.J., Salama, A.D., Chitnis, T., Smith, R.N., Yagita, H., Akiba, H., Yamazaki, T., Azuma, M., Iwai, H., Khoury, S.J., Auchincloss Jr., H., Sayegh, M.H., 2003. The programmed death-1 (PD-1) pathway regulates autoimmune diabetes in nonobese diabetic (NOD) mice. J. Exp. Med. 198, 63–69. Ansari, M.J., Fiorina, P., Dada, S., Guleria, I., Ueno, T., Yuan, X., Trikudanathan, S., Smith, R.N., Freeman, G., Sayegh, M.H., 2008. Role of ICOS pathway in autoimmune and alloimmune responses in NOD mice. Clin. Immunol. 126, 140–147.

Mechanisms of Autoimmunity

117

Armengol, M.P., Juan, M., Lucas-Martin, A., Fernandez-Figueras, M.T., Jaraquemada, D., Gallart, T., Pujol-Borrell, R., 2001. Thyroid autoimmune disease: demonstration of thyroid antigen-specific B cells and recombination-activating gene expression in chemokine-containing active intrathyroidal germinal centers. Am. J. Pathol. 159, 861–873. Arora, S., Rana, R., Chhabra, A., Jaiswal, A., Rani, V., 2013. miRNA-transcription factor interactions: a combinatorial regulation of gene expression. Mol. Genet. Genomics 288, 77–87. Artlett, C.M., Sassi-Gaha, S., Rieger, J.L., Boesteanu, A.C., Feghali-Bostwick, C.A., Katsikis, P.D., 2011. The inflammasome activating caspase 1 mediates fibrosis and myofibroblast differentiation in systemic sclerosis. Arthritis Rheum. 63, 3563–3574. Baker, R.L., Wagner Jr., D.H., Haskins, K., 2008. CD40 on NOD CD4 T cells contributes to their activation and pathogenicity. J. Autoimmun. 31, 385–392. Balasa, B., Krahl, T., Patstone, G., Lee, J., Tisch, R., McDevitt, H.O., Sarvetnick, N., 1997. CD40 ligand-CD40 interactions are necessary for the initiation of insulitis and diabetes in nonobese diabetic mice. J. Immunol. 159, 4620–4627. Ballestar, E., 2010. Epigenetics lessons from twins: prospects for autoimmune disease. Clin. Rev. Allergy Immunol. 39, 30–41. Banas, M.C., Banas, B., Hudkins, K.L., Wietecha, T.A., Iyoda, M., Bock, E., Hauser, P., Pippin, J.W., Shankland, S.J., Smith, K.D., Stoelcker, B., Liu, G., Grone, H.J., Kramer, B.K., Alpers, C.E., 2008. TLR4 links podocytes with the innate immune system to mediate glomerular injury. J. Am. Soc. Nephrol. 19, 704–713. Banchereau, R., Hong, S., Cantarel, B., Baldwin, N., Baisch, J., Edens, M., Cepika, A.M., Acs, P., Turner, J., Anguiano, E., Vinod, P., Kahn, S., Obermoser, G., Blankenship, D., Wakeland, E., Nassi, L., Gotte, A., Punaro, M., Liu, Y.J., Banchereau, J., Rossello-Urgell, J., Wright, T., Pascual, V., 2016. Personalized immunomonitoring uncovers molecular networks that stratify lupus patients. Cell 165, 551–565. Barbhaiya, M., Costenbader, K.H., 2014. Ultraviolet radiation and systemic lupus erythematosus. Lupus 23, 588–595. Baricordi, O.R., Ferrari, D., Melchiorri, L., Chiozzi, P., Hanau, S., Chiari, E., Rubini, M., Di Virgilio, F., 1996. An ATP-activated channel is involved in mitogenic stimulation of human T lymphocytes. Blood 87, 682–690. Barin, J.G., Talor, M.V., Sharma, R.B., Rose, N.R., Burek, C.L., 2005. Iodination of murine thyroglobulin enhances autoimmune reactivity in the NOD.H2 mouse. Clin. Exp. Immunol. 142, 251–259. Barone, F., Nayar, S., Campos, J., Cloake, T., Withers, D.R., Toellner, K.M., Zhang, Y., Fouser, L., Fisher, B., Bowman, S., Rangel-Moreno, J., Garcia-Hernandez Mde, L., Randall, T.D., Lucchesi, D., Bombardieri, M., Pitzalis, C., Luther, S.A., Buckley, C.D., 2015. IL-22 regulates lymphoid chemokine production and assembly of tertiary lymphoid organs. Proc. Natl. Acad. Sci. U.S.A. 112, 11024–11029. Barry, O.P., Pratico, D., Lawson, J.A., FitzGerald, G.A., 1997. Transcellular activation of platelets and endothelial cells by bioactive lipids in platelet microparticles. J. Clin. Invest. 99, 2118–2127. Begovich, A.B., Carlton, V.E., Honigberg, L.A., Schrodi, S.J., Chokkalingam, A.P., Alexander, H.C., Ardlie, K.G., Huang, Q., Smith, A.M., Spoerke, J.M., Conn, M.T., Chang, M., Chang, S.Y., Saiki, R.K., Catanese, J.J., Leong, D.U., Garcia, V.E., McAllister, L.B., Jeffery, D.A., Lee, A.T., Batliwalla, F., Remmers, E., Criswell, L.A., Seldin, M.F., Kastner, D.L., Amos, C.I., Sninsky, J.J., Gregersen, P.K., 2004. A missense single-nucleotide polymorphism in a gene encoding a protein tyrosine phosphatase (PTPN22) is associated with rheumatoid arthritis. Am. J. Hum. Genet. 75, 330–337.

118

S.K. Devarapu et al.

Belot, A., Kasher, P.R., Trotter, E.W., Foray, A.P., Debaud, A.L., Rice, G.I., Szynkiewicz, M., Zabot, M.T., Rouvet, I., Bhaskar, S.S., Daly, S.B., Dickerson, J.E., Mayer, J., O’Sullivan, J., Juillard, L., Urquhart, J.E., Fawdar, S., Marusiak, A.A., Stephenson, N., Waszkowycz, B., W Beresford, M., Biesecker, L.G., C M Black, G., Rene, C., Eliaou, J.F., Fabien, N., Ranchin, B., Gaffney, P.M., Rozenberg, F., Lebon, P., Malcus, C., Crow, Y.J., Brognard, J., Bonnefoy, N., 2013. Protein kinase cdelta deficiency causes mendelian systemic lupus erythematosus with B cell-defective apoptosis and hyperproliferation. Arthritis Rheum. 65, 2161–2171. Belver, L., de Yebenes, V.G., Ramiro, A.R., 2010. MicroRNAs prevent the generation of autoreactive antibodies. Immunity 33, 713–722. Berckmans, R.J., Nieuwland, R., Kraan, M.C., Schaap, M.C., Pots, D., Smeets, T.J., Sturk, A., Tak, P.P., 2005. Synovial microparticles from arthritic patients modulate chemokine and cytokine release by synoviocytes. Arthritis Res. Ther. 7, R536–544. Bergtold, A., Gavhane, A., D’Agati, V., Madaio, M., Clynes, R., 2006. FcR-bearing myeloid cells are responsible for triggering murine lupus nephritis. J. Immunol. 177, 7287–7295. Bethunaickan, R., Berthier, C.C., Ramanujam, M., Sahu, R., Zhang, W., Sun, Y., Bottinger, E.P., Ivashkiv, L., Kretzler, M., Davidson, A., 2011. A unique hybrid renal mononuclear phagocyte activation phenotype in murine systemic lupus erythematosus nephritis. J. Immunol. 186, 4994–5003. Beutler, B., 2000. Endotoxin, toll-like receptor 4, and the afferent limb of innate immunity. Curr. Opin. Microbiol. 3, 23–28. Bhatnagar, S., Shinagawa, K., Castellino, F.J., Schorey, J.S., 2007. Exosomes released from macrophages infected with intracellular pathogens stimulate a proinflammatory response in vitro and in vivo. Blood 110, 3234–3244. Bhattacharyya, S., Kelley, K., Melichian, D.S., Tamaki, Z., Fang, F., Su, Y., Feng, G., Pope, R.M., Budinger, G.R.S., Mutlu, G.M., Lafyatis, R., Radstake, T., Feghali-Bostwick, C., Varga, J., 2013. Toll-like receptor 4 signaling augments transforming growth factor-β responses: a novel mechanism for maintaining and amplifying fibrosis in scleroderma. Am. J. Pathol. 182, 192–205. Bhattacharyya, S., Tamaki, Z., Wang, W., Hinchcliff, M., Hoover, P., Getsios, S., White, E.S., Varga, J., 2014. FibronectinEDA promotes chronic cutaneous fibrosis through Toll-like receptor signaling. Sci. Transl. Med. 6, 232ra250. Biermann, M.H., Veissi, S., Maueroder, C., Chaurio, R., Berens, C., Herrmann, M., Munoz, L.E., 2014. The role of dead cell clearance in the etiology and pathogenesis of systemic lupus erythematosus: dendritic cells as potential targets. Expert Rev. Clin. Immunol. 10, 1151–1164. Bloom, B.R., Salgame, P., Diamond, B., 1992. Revisiting and revising suppressor T cells. Immunol. Today 13, 131–136. Bodhankar, S., Galipeau, D., Vandenbark, A.A., Offner, H., 2013. PD-1 interaction with PD-L1 but not PD-L2 on B-cells mediates protective effects of estrogen against EAE. J. Clin. Cell. Immunol. 4, 143. Boilard, E., Nigrovic, P.A., Larabee, K., Watts, G.F., Coblyn, J.S., Weinblatt, M.E., Massarotti, E.M., Remold-O’Donnell, E., Farndale, R.W., Ware, J., Lee, D.M., 2010. Platelets amplify inflammation in arthritis via collagen-dependent microparticle production. Science 327, 580–583. Boldin, M.P., Taganov, K.D., Rao, D.S., Yang, L., Zhao, J.L., Kalwani, M., Garcia-Flores, Y., Luong, M., Devrekanli, A., Xu, J., Sun, G., Tay, J., Linsley, P.S., Baltimore, D., 2011. miR-146a is a significant brake on autoimmunity, myeloproliferation, and cancer in mice. J. Exp. Med. 208, 1189–1201.

Mechanisms of Autoimmunity

119

Bossaller, L., Christ, A., Pelka, K., Nundel, K., Chiang, P.I., Pang, C., Mishra, N., Busto, P., Bonegio, R.G., Schmidt, R.E., Latz, E., Marshak-Rothstein, A., 2016. TLR9 deficiency leads to accelerated renal disease and myeloid lineage abnormalities in pristane-induced murine lupus. J. Immunol. 197 (4), 1044–1053. Bosticardo, M., Marangoni, F., Aiuti, A., Villa, A., Grazia Roncarolo, M., 2009. Recent advances in understanding the pathophysiology of Wiskott-Aldrich syndrome. Blood 113, 6288–6295. Bottini, N., Musumeci, L., Alonso, A., Rahmouni, S., Nika, K., Rostamkhani, M., MacMurray, J., Meloni, G.F., Lucarelli, P., Pellecchia, M., Eisenbarth, G.S., Comings, D., Mustelin, T., 2004. A functional variant of lymphoid tyrosine phosphatase is associated with type I diabetes. Nat. Genet. 36, 337–338. Botto, M., 2001. Links between complement deficiency and apoptosis. Arthritis Res. 3, 207–210. Bour-Jordan, H., Salomon, B.L., Thompson, H.L., Szot, G.L., Bernhard, M.R., Bluestone, J.A., 2004. Costimulation controls diabetes by altering the balance of pathogenic and regulatory T cells. J. Clin. Invest. 114, 979–987. Bour-Jordan, H., Esensten, J.H., Martinez-Llordella, M., Penaranda, C., Stumpf, M., Bluestone, J.A., 2011. Intrinsic and extrinsic control of peripheral T-cell tolerance by costimulatory molecules of the CD28/ B7 family. Immunol. Rev. 241, 180–205. Brasacchio, D., Okabe, J., Tikellis, C., Balcerczyk, A., George, P., Baker, E.K., Calkin, A.C., Brownlee, M., Cooper, M.E., El-Osta, A., 2009. Hyperglycemia induces a dynamic cooperativity of histone methylase and demethylase enzymes associated with gene-activating epigenetic marks that coexist on the lysine tail. Diabetes 58, 1229–1236. Brechard, S., Tschirhart, E.J., 2008. Regulation of superoxide production in neutrophils: role of calcium influx. J. Leukoc. Biol. 84, 1223–1237. Bretscher, P., Cohn, M., 1970. A theory of self-nonself discrimination. Science 169, 1042–1049. Brinkmann, V., Reichard, U., Goosmann, C., Fauler, B., Uhlemann, Y., Weiss, D.S., Weinrauch, Y., Zychlinsky, A., 2004. Neutrophil extracellular traps kill bacteria. Science 303, 1532–1535. Broen, J.C., Bossini-Castillo, L., van Bon, L., Vonk, M.C., Knaapen, H., Beretta, L., Rueda, B., Hesselstrand, R., Herrick, A., Worthington, J., Hunzelman, N., Denton, C.P., Fonseca, C., Riemekasten, G., Kiener, H.P., Scorza, R., Simeon, C.P., Ortego-Centeno, N., Gonzalez-Gay, M.A., Airo, P., Coenen, M.J., Martin, J., Radstake, T.R., 2012. A rare polymorphism in the gene for Toll-like receptor 2 is associated with systemic sclerosis phenotype and increases the production of inflammatory mediators. Arthritis Rheum. 64, 264–271. Brown, C.C., Wedderburn, L.R., 2015. Genetics: mapping autoimmune disease epigenetics: what’s on the horizon? Nat. Rev. Rheumatol. 11, 131–132. Brunini, F., Page, T.H., Gallieni, M., Pusey, C.D., 2016. The role of monocytes in ANCA-associated vasculitides. Autoimmun. Rev. 5 (11), 1046–1053. Bull, M.J., Williams, A.S., Mecklenburgh, Z., Calder, C.J., Twohig, J.P., Elford, C., Evans, B.A., Rowley, T.F., Slebioda, T.J., Taraban, V.Y., Al-Shamkhani, A., Wang, E.C., 2008. The death receptor 3-TNF-like protein 1A pathway drives adverse bone pathology in inflammatory arthritis. J. Exp. Med. 205, 2457–2464. Burgos, R.A., Conejeros, I., Hidalgo, M.A., Werling, D., Hermosilla, C., 2011. Calcium influx, a new potential therapeutic target in the control of neutrophil-dependent inflammatory diseases in bovines. Vet. Immunol. Immunopathol. 143, 1–10. Burkhardt, H., Huffmeier, U., Spriewald, B., Bohm, B., Rau, R., Kallert, S., Engstrom, A., Holmdahl, R., Reis, A., 2006. Association between protein tyrosine phosphatase 22 variant R620W in conjunction with the HLA-DRB1 shared epitope and humoral

120

S.K. Devarapu et al.

autoimmunity to an immunodominant epitope of cartilage-specific type II collagen in early rheumatoid arthritis. Arthritis Rheum. 54, 82–89. Burnet, F.M., Fenner, F., 1949. The Production of Antibodies, second ed. Macmillan and Co., Ltd., Melbourne, Australia. Butte, M.J., Keir, M.E., Phamduy, T.B., Sharpe, A.H., Freeman, G.J., 2007. Programmed death-1 ligand 1 interacts specifically with the B7-1 costimulatory molecule to inhibit T cell responses. Immunity 27, 111–122. Cao, X., 2016. Self-regulation and cross-regulation of pattern-recognition receptor signalling in health and disease. Nat. Rev. Immunol. 16, 35–50. Capasso, M., Bhamrah, M.K., Henley, T., Boyd, R.S., Langlais, C., Cain, K., Dinsdale, D., Pulford, K., Khan, M., Musset, B., Cherny, V.V., Morgan, D., Gascoyne, R.D., Vigorito, E., DeCoursey, T.E., MacLennan, I.C., Dyer, M.J., 2010. HVCN1 modulates BCR signal strength via regulation of BCR-dependent generation of reactive oxygen species. Nat. Immunol. 11, 265–272. Carboni, S., Aboul-Enein, F., Waltzinger, C., Killeen, N., Lassmann, H., Pena-Rossi, C., 2003. CD134 plays a crucial role in the pathogenesis of EAE and is upregulated in the CNS of patients with multiple sclerosis. J. Neuroimmunol. 145, 1–11. Caricchio, R., McPhie, L., Cohen, P.L., 2003. Ultraviolet B radiation-induced cell death: critical role of ultraviolet dose in inflammation and lupus autoantigen redistribution. J. Immunol. 171, 5778–5786. Carren˜o, L.J., Pacheco, R., Gutierrez, M.A., Jacobelli, S., Kalergis, A.M., 2009. Disease activity in systemic lupus erythematosus is associated with an altered expression of low-affinity Fcγ receptors and costimulatory molecules on dendritic cells. Immunology 128, 334–341. Carter, L.L., Leach, M.W., Azoitei, M.L., Cui, J., Pelker, J.W., Jussif, J., Benoit, S., Ireland, G., Luxenberg, D., Askew, G.R., Milarski, K.L., Groves, C., Brown, T., Carito, B.A., Percival, K., Carreno, B.M., Collins, M., Marusic, S., 2007. PD-1/ PD-L1, but not PD-1/PD-L2, interactions regulate the severity of experimental autoimmune encephalomyelitis. J. Neuroimmunol. 182, 124–134. Casciola-Rosen, L.A., Anhalt, G., Rosen, A., 1994. Autoantigens targeted in systemic lupus erythematosus are clustered in two populations of surface structures on apoptotic keratinocytes. J. Exp. Med. 179, 1317–1330. Cen, H., Wang, W., Leng, R.X., Wang, T.Y., Pan, H.F., Fan, Y.G., Wang, B., Ye, D.Q., 2013. Association of IFIH1 rs1990760 polymorphism with susceptibility to autoimmune diseases: a meta-analysis. Autoimmunity 46, 455–462. Chang, T.T., Jabs, C., Sobel, R.A., Kuchroo, V.K., Sharpe, A.H., 1999. Studies in B7-deficient mice reveal a critical role for B7 costimulation in both induction and effector phases of experimental autoimmune encephalomyelitis. J. Exp. Med. 190, 733–740. Chang, A., Henderson, S.G., Brandt, D., Liu, N., Guttikonda, R., Hsieh, C., Kaverina, N., Utset, T.O., Meehan, S.M., Quigg, R.J., Meffre, E., Clark, M.R., 2011. In situ B cell-mediated immune responses and tubulointerstitial inflammation in human lupus nephritis. J. Immunol. 186, 1849–1860. Chauhan, S.K., Singh, V.V., Rai, R., Rai, M., Rai, G., 2013. Distinct autoantibody profiles in systemic lupus erythematosus patients are selectively associated with TLR7 and TLR9 upregulation. J. Clin. Immunol. 33, 954–964. Chen, C.Y., Chen, S.T., Fuh, C.S., Juan, H.F., Huang, H.C., 2011. Coregulation of transcription factors and microRNAs in human transcriptional regulatory network. BMC Bioinformatics 12 (Suppl. 1), S41. Chinen, T., Volchkov, P.Y., Chervonsky, A.V., Rudensky, A.Y., 2010. A critical role for regulatory T cell-mediated control of inflammation in the absence of commensal microbiota. J. Exp. Med. 207, 2323–2330.

Mechanisms of Autoimmunity

121

Cho, Y.N., Kee, S.J., Lee, S.J., Seo, S.R., Kim, T.J., Lee, S.S., Kim, M.S., Lee, W.W., Yoo, D.H., Kim, N., Park, Y.W., 2011. Numerical and functional deficiencies of natural killer T cells in systemic lupus erythematosus: their deficiency related to disease activity. Rheumatology (Oxford) 50, 1054–1063. Chowdhary, V.R., Grande, J.P., Luthra, H.S., David, C.S., 2007. Characterization of haemorrhagic pulmonary capillaritis: another manifestation of Pristane-induced lupus. Rheumatology (Oxford) 46, 1405–1410. Christensen, S.R., Shupe, J., Nickerson, K., Kashgarian, M., Flavell, R.A., Shlomchik, M.J., 2006. Toll-like receptor 7 and TLR9 dictate autoantibody specificity and have opposing inflammatory and regulatory roles in a murine model of lupus. Immunity 25, 417–428. Chuang, H.C., Chang, C.W., Chang, G.D., Yao, T.P., Chen, H., 2006. Histone deacetylase 3 binds to and regulates the GCMa transcription factor. Nucleic Acids Res. 34, 1459–1469. Chung, S.A., Brown, E.E., Williams, A.H., Ramos, P.S., Berthier, C.C., Bhangale, T., Alarcon-Riquelme, M.E., Behrens, T.W., Criswell, L.A., Graham, D.C., Demirci, F.Y., Edberg, J.C., Gaffney, P.M., Harley, J.B., Jacob, C.O., Kamboh, M.I., Kelly, J.A., Manzi, S., Moser-Sivils, K.L., Russell, L.P., Petri, M., Tsao, B.P., Vyse, T.J., Zidovetzki, R., Kretzler, M., Kimberly, R.P., Freedman, B.I., Graham, R.R., Langefeld, C.D., 2014. Lupus nephritis susceptibility loci in women with systemic lupus erythematosus. J. Am. Soc. Nephrol. 25, 2859–2870. Clarke, S.L., Pellowe, E.J., de Jesus, A.A., Goldbach-Mansky, R., Hilliard, T.N., Ramanan, A.V., 2016. Interstitial lung disease caused by STING-associated vasculopathy with onset in infancy. Am. J. Respir. Crit. Care Med. 194, 639–642. Clayton, A., Court, J., Navabi, H., Adams, M., Mason, M.D., Hobot, J.A., Newman, G.R., Jasani, B., 2001. Analysis of antigen presenting cell derived exosomes, based on immuno-magnetic isolation and flow cytometry. J. Immunol. Methods 247, 163–174. Cloutier, N., Tan, S., Boudreau, L.H., Cramb, C., Subbaiah, R., Lahey, L., Albert, A., Shnayder, R., Gobezie, R., Nigrovic, P.A., Farndale, R.W., Robinson, W.H., Brisson, A., Lee, D.M., Boilard, E., 2013. The exposure of autoantigens by microparticles underlies the formation of potent inflammatory components: the microparticle-associated immune complexes. EMBO Mol. Med. 5, 235–249. Clynes, R., Dumitru, C., Ravetch, J.V., 1998. Uncoupling of immune complex formation and kidney damage in autoimmune glomerulonephritis. Science 279, 1052–1054. Cohen, P.L., Caricchio, R., Abraham, V., Camenisch, T.D., Jennette, J.C., Roubey, R.A., Earp, H.S., Matsushima, G., Reap, E.A., 2002. Delayed apoptotic cell clearance and lupus-like autoimmunity in mice lacking the c-mer membrane tyrosine kinase. J. Exp. Med. 196, 135–140. Cole, B.C., Griffiths, M.M., 1993. Triggering and exacerbation of autoimmune arthritis by the Mycoplasma arthritidis superantigen MAM. Arthritis Rheum. 36, 994–1002. Conrad, B., Weissmahr, R.N., Boni, J., Arcari, R., Schupbach, J., Mach, B., 1997. A human endogenous retroviral superantigen as candidate autoimmune gene in type I diabetes. Cell 90, 303–313. Cornacchia, E., Golbus, J., Maybaum, J., Strahler, J., Hanash, S., Richardson, B., 1988. Hydralazine and procainamide inhibit T cell DNA methylation and induce autoreactivity. J. Immunol. 140, 2197–2200. Craner, M.J., Damarjian, T.G., Liu, S., Hains, B.C., Lo, A.C., Black, J.A., Newcombe, J., Cuzner, M.L., Waxman, S.G., 2005. Sodium channels contribute to microglia/macrophage activation and function in EAE and MS. Glia 49, 220–229. Crow, Y.J., 2011. Type I interferonopathies: a novel set of inborn errors of immunity. Ann. N. Y. Acad. Sci. 1238, 91–98.

122

S.K. Devarapu et al.

Cunninghame Graham, D.S., Graham, R.R., Manku, H., Wong, A.K., Whittaker, J.C., Gaffney, P.M., Moser, K.L., Rioux, J.D., Altshuler, D., Behrens, T.W., Vyse, T.J., 2008. Polymorphism at the TNF superfamily gene TNFSF4 confers susceptibility to systemic lupus erythematosus. Nat. Genet. 40, 83–89. Cunningham-Rundles, C., 2008. Autoimmune manifestations in common variable immunodeficiency. J. Clin. Immunol. 28 (Suppl. 1), S42–45. Cusick, M.F., Libbey, J.E., Fujinami, R.S., 2012. Molecular mimicry as a mechanism of autoimmune disease. Clin. Rev. Allergy Immunol. 42, 102–111. Daikh, D.I., Finck, B.K., Linsley, P.S., Hollenbaugh, D., Wofsy, D., 1997. Long-term inhibition of murine lupus by brief simultaneous blockade of the B7/CD28 and CD40/gp39 costimulation pathways. J. Immunol. 159, 3104–3108. Damgaard, D., Friberg Bruun Nielsen, M., Quisgaard Gaunsbaek, M., Palarasah, Y., Svane-Knudsen, V., Nielsen, C.H., 2015. Smoking is associated with increased levels of extracellular peptidylarginine deiminase 2 (PAD2) in the lungs. Clin. Exp. Rheumatol. 33, 405–408. Dar, S.A., Janahi, E.M., Haque, S., Akhter, N., Jawed, A., Wahid, M., Ramachandran, V.G., Bhattacharya, S.N., Banerjee, B.D., Das, S., 2016. Superantigen influence in conjunction with cytokine polymorphism potentiates autoimmunity in systemic lupus erythematosus patients. Immunol. Res. 64, 1001–1012. Demaria, O., Pagni, P.P., Traub, S., de Gassart, A., Branzk, N., Murphy, A.J., Valenzuela, D.M., Yancopoulos, G.D., Flavell, R.A., Alexopoulou, L., 2010. TLR8 deficiency leads to autoimmunity in mice. J. Clin. Invest. 120, 3651–3662. Denis, H., Ndlovu, M.N., Fuks, F., 2011. Regulation of mammalian DNA methyltransferases: a route to new mechanisms. EMBO Rep. 12, 647–656. Denny, M.F., Yalavarthi, S., Zhao, W., Thacker, S.G., Anderson, M., Sandy, A.R., McCune, W.J., Kaplan, M.J., 2010. A distinct subset of proinflammatory neutrophils isolated from patients with systemic lupus erythematosus induces vascular damage and synthesizes type I IFNs. J. Immunol. 184, 3284–3297. Desai, J., Kumar, S.V., Mulay, S.R., Konrad, L., Romoli, S., Schauer, C., Herrmann, M., Bilyy, R., Muller, S., Popper, B., Nakazawa, D., Weidenbusch, M., Thomasova, D., Krautwald, S., Linkermann, A., Anders, H.J., 2016a. PMA and crystal-induced neutrophil extracellular trap formation involves RIPK1-RIPK3-MLKL signaling. Eur. J. Immunol. 46, 223–229. Desai, J., Mulay, S.R., Nakazawa, D., Anders, H.J., 2016b. Matters of life and death. How neutrophils die or survive along NET release and is “NETosis” ¼ necroptosis? Cell. Mol. Life Sci. 73, 2211–2219. Dhir, V., Singh, A.P., Aggarwal, A., Naik, S., Misra, R., 2009. Increased T-lymphocyte apoptosis in lupus correlates with disease activity and may be responsible for reduced T-cell frequency: a cross-sectional and longitudinal study. Lupus 18, 785–791. Di, A., Brown, M.E., Deriy, L.V., Li, C., Szeto, F.L., Chen, Y., Huang, P., Tong, J., Naren, A.P., Bindokas, V., Palfrey, H.C., Nelson, D.J., 2006. CFTR regulates phagosome acidification in macrophages and alters bactericidal activity. Nat. Cell Biol. 8, 933–944. Distler, J.H., Jungel, A., Huber, L.C., Seemayer, C.A., Reich 3rd, C.F., Gay, R.E., Michel, B.A., Fontana, A., Gay, S., Pisetsky, D.S., Distler, O., 2005. The induction of matrix metalloproteinase and cytokine expression in synovial fibroblasts stimulated with immune cell microparticles. Proc. Natl. Acad. Sci. U.S.A. 102, 2892–2897. Doherty, D.G., 2016. Immunity, tolerance and autoimmunity in the liver: a comprehensive review. J. Autoimmun. 66, 60–75. Dong, C., Juedes, A.E., Temann, U.A., Shresta, S., Allison, J.P., Ruddle, N.H., Flavell, R.A., 2001. ICOS co-stimulatory receptor is essential for T-cell activation and function. Nature 409, 97–101.

Mechanisms of Autoimmunity

123

dos Santos, B.P., Valverde, J.V., Rohr, P., Monticielo, O.A., Brenol, J.C., Xavier, R.M., Chies, J.A., 2012. TLR7/8/9 polymorphisms and their associations in systemic lupus erythematosus patients from southern Brazil. Lupus 21, 302–309. Dowds, C.M., Kornell, S.C., Blumberg, R.S., Zeissig, S., 2014. Lipid antigens in immunity. Biol. Chem. 395, 61–81. Du, C., Liu, C., Kang, J., Zhao, G., Ye, Z., Huang, S., Li, Z., Wu, Z., Pei, G., 2009. MicroRNA miR-326 regulates TH-17 differentiation and is associated with the pathogenesis of multiple sclerosis. Nat. Immunol. 10, 1252–1259. Duerr, R.H., Taylor, K.D., Brant, S.R., Rioux, J.D., Silverberg, M.S., Daly, M.J., Steinhart, A.H., Abraham, C., Regueiro, M., Griffiths, A., Dassopoulos, T., Bitton, A., Yang, H., Targan, S., Datta, L.W., Kistner, E.O., Schumm, L.P., Lee, A.T., Gregersen, P.K., Barmada, M.M., Rotter, J.I., Nicolae, D.L., Cho, J.H., 2006. A genome-wide association study identifies IL23R as an inflammatory bowel disease gene. Science 314, 1461–1463. Early, G.S., Zhao, W., Burns, C.M., 1996. Anti-CD40 ligand antibody treatment prevents the development of lupus-like nephritis in a subset of New Zealand black x New Zealand white mice. Response correlates with the absence of an anti-antibody response. J. Immunol. 157, 3159–3164. Ellson, C.D., Davidson, K., Ferguson, G.J., O’Connor, R., Stephens, L.R., Hawkins, P.T., 2006. Neutrophils from p40phox-/- mice exhibit severe defects in NADPH oxidase regulation and oxidant-dependent bacterial killing. J. Exp. Med. 203, 1927–1937. Eloranta, M.L., Lovgren, T., Finke, D., Mathsson, L., Ronnelid, J., Kastner, B., Alm, G.V., Ronnblom, L., 2009. Regulation of the interferon-alpha production induced by RNA-containing immune complexes in plasmacytoid dendritic cells. Arthritis Rheum. 60, 2418–2427. Espeli, M., Bokers, S., Giannico, G., Dickinson, H.A., Bardsley, V., Fogo, A.B., Smith, K.G., 2011. Local renal autoantibody production in lupus nephritis. J. Am. Soc. Nephrol. 22, 296–305. Esser, J., Gehrmann, U., D’Alexandri, F.L., Hidalgo-Estevez, A.M., Wheelock, C.E., Scheynius, A., Gabrielsson, S., Radmark, O., 2010. Exosomes from human macrophages and dendritic cells contain enzymes for leukotriene biosynthesis and promote granulocyte migration. J. Allergy Clin. Immunol. 126, 1032–1040. 1040 e1031–1034. Estrada-Capetillo, L., Hernandez-Castro, B., Monsivais-Urenda, A., Alvarez-Quiroga, C., Layseca-Espinosa, E., Abud-Mendoza, C., Baranda, L., Urzainqui, A., Sanchez-Madrid, F., Gonzalez-Amaro, R., 2013. Induction of Th17 lymphocytes and Treg cells by monocyte-derived dendritic cells in patients with rheumatoid arthritis and systemic lupus erythematosus. Clin. Dev. Immunol. 2013, 584303. Faas, S., Trucco, M., 1994. The genes influencing the susceptibility to IDDM in humans. J. Endocrinol. Invest. 17, 477–495. Fabiano, R.C., Pinheiro, S.V., Simoes, E.S.A.C., 2016. Immunoglobulin A nephropathy: a pathophysiology view. Inflamm. Res. 65, 757–770. Fairhurst, A.M., Hwang, S.H., Wang, A., Tian, X.H., Boudreaux, C., Zhou, X.J., Casco, J., Li, Q.Z., Connolly, J.E., Wakeland, E.K., 2008. Yaa autoimmune phenotypes are conferred by overexpression of TLR7. Eur. J. Immunol. 38, 1971–1978. Falk, R.J., Terrell, R.S., Charles, L.A., Jennette, J.C., 1990. Anti-neutrophil cytoplasmic autoantibodies induce neutrophils to degranulate and produce oxygen radicals in vitro. Proc. Natl. Acad. Sci. U.S.A. 87, 4115–4119. Farina, G.A., York, M.R., Di Marzio, M., Collins, C.A., Meller, S., Homey, B., Rifkin, I.R., Marshak-Rothstein, A., Radstake, T.R., Lafyatis, R., 2010. Poly(I:C) drives type I IFN- and TGFbeta-mediated inflammation and dermal fibrosis simulating altered gene expression in systemic sclerosis. J. Invest. Dermatol. 130, 2583–2593.

124

S.K. Devarapu et al.

Farrera, C., Fadeel, B., 2013. Macrophage clearance of neutrophil extracellular traps is a silent process. J. Immunol. 191, 2647–2656. Fattal, I., Shental, N., Mevorach, D., Anaya, J.M., Livneh, A., Langevitz, P., Zandman-Goddard, G., Pauzner, R., Lerner, M., Blank, M., Hincapie, M.E., Gafter, U., Naparstek, Y., Shoenfeld, Y., Domany, E., Cohen, I.R., 2010. An antibody profile of systemic lupus erythematosus detected by antigen microarray. Immunology 130, 337–343. Fernandes-Alnemri, T., Yu, J.W., Datta, P., Wu, J., Alnemri, E.S., 2009. AIM2 activates the inflammasome and cell death in response to cytoplasmic DNA. Nature 458, 509–513. Fernando, M.M., Stevens, C.R., Walsh, E.C., De Jager, P.L., Goyette, P., Plenge, R.M., Vyse, T.J., Rioux, J.D., 2008. Defining the role of the MHC in autoimmunity: a review and pooled analysis. PLoS Genet. 4, e1000024. Fierabracci, A., Saura, F., 2010. Identification of a common autoantigenic epitope of protein disulfide isomerase, golgin-160 and voltage-gated potassium channel in type 1 diabetes. Diabetes Res. Clin. Pract. 88, e14–16. Figueroa-Vega, N., Galindo-Rodriguez, G., Bajana, S., Portales-Perez, D., Abud-Mendoza, C., Sanchez-Torres, C., Gonzalez-Amaro, R., 2006. Phenotypic analysis of IL-10-treated, monocyte-derived dendritic cells in patients with systemic lupus erythematosus. Scand. J. Immunol. 64, 668–676. Finck, B.K., Linsley, P.S., Wofsy, D., 1994. Treatment of murine lupus with CTLA4Ig. Science 265, 1225–1227. Finnish-German, A.C., 1997. An autoimmune disease, APECED, caused by mutations in a novel gene featuring two PHD-type zinc-finger domains. Nat. Genet. 17, 399–403. Floreani, A., Leung, P.S., Gershwin, M.E., 2016. Environmental basis of autoimmunity. Clin. Rev. Allergy Immunol. 50, 287–300. Flur, K., Allam, R., Zecher, D., Kulkarni, O.P., Lichtnekert, J., Schwarz, M., Beutler, B., Vielhauer, V., Anders, H.J., 2009. Viral RNA induces type I interferon-dependent cytokine release and cell death in mesangial cells via melanoma-differentiation-associated gene-5: implications for viral infection-associated glomerulonephritis. Am. J. Pathol. 175, 2014–2022. Fraser, J.D., Proft, T., 2008. The bacterial superantigen and superantigen-like proteins. Immunol. Rev. 225, 226–243. Fredi, M., Bianchi, M., Andreoli, L., Greco, G., Olivieri, I., Orcesi, S., Fazzi, E., Cereda, C., Tincani, A., 2015. Typing TREX1 gene in patients with systemic lupus erythematosus. Reumatismo 67, 1–7. Frisoni, L., McPhie, L., Kang, S.A., Monestier, M., Madaio, M., Satoh, M., Caricchio, R., 2007. Lack of chromatin and nuclear fragmentation in vivo impairs the production of lupus anti-nuclear antibodies. J. Immunol. 179, 7959–7966. Fuchs, T.A., Abed, U., Goosmann, C., Hurwitz, R., Schulze, I., Wahn, V., Weinrauch, Y., Brinkmann, V., Zychlinsky, A., 2007. Novel cell death program leads to neutrophil extracellular traps. J. Cell Biol. 176, 231–241. Fujiwara, Y., Takeshita, K., Nakagawa, A., Okamura, Y., 2013. Structural characteristics of the redox-sensing coiled coil in the voltage-gated H + channel. J. Biol. Chem. 288, 17968–17975. Fukui, R., Saitoh, S., Kanno, A., Onji, M., Shibata, T., Ito, A., Onji, M., Matsumoto, M., Akira, S., Yoshida, N., Miyake, K., 2011. Unc93B1 restricts systemic lethal inflammation by orchestrating Toll-like receptor 7 and 9 trafficking. Immunity 35, 69–81. Funabiki, M., Kato, H., Miyachi, Y., Toki, H., Motegi, H., Inoue, M., Minowa, O., Yoshida, A., Deguchi, K., Sato, H., Ito, S., Shiroishi, T., Takeyasu, K., Noda, T., Fujita, T., 2014. Autoimmune disorders associated with gain of function of the intracellular sensor MDA5. Immunity 40, 199–212.

Mechanisms of Autoimmunity

125

Fuschiotti, P., 2016. Current perspectives on the immunopathogenesis of systemic sclerosis. Immunotargets Ther. 5, 21–35. Futatsugi-Yumikura, S., Matsushita, K., Fukuoka, A., Takahashi, S., Yamamoto, N., Yonehara, S., Nakanishi, K., Yoshimoto, T., 2014. Pathogenic Th2-type follicular helper T cells contribute to the development of lupus in Fas-deficient mice. Int. Immunol. 26, 221–231. Galicia, G., Kasran, A., Uyttenhove, C., De Swert, K., Van Snick, J., Ceuppens, J.L., 2009. ICOS deficiency results in exacerbated IL-17 mediated experimental autoimmune encephalomyelitis. J. Clin. Immunol. 29, 426–433. Galluzzi, L., Kepp, O., Kroemer, G., 2014. MLKL regulates necrotic plasma membrane permeabilization. Cell Res. 24, 139–140. Gao, Y.D., Hanley, P.J., Rinne, S., Zuzarte, M., Daut, J., 2010. Calcium-activated K(+) channel (K(Ca)3.1) activity during Ca(2 +) store depletion and store-operated Ca(2+) entry in human macrophages. Cell Calcium 48, 19–27. Garchow, B., Kiriakidou, M., 2016. MicroRNA-21 deficiency protects from lupus-like autoimmunity in the chronic graft-versus-host disease model of systemic lupus erythematosus. Clin. Immunol. 162, 100–106. Garcia-Romo, G.S., Caielli, S., Vega, B., Connolly, J., Allantaz, F., Xu, Z., Punaro, M., Baisch, J., Guiducci, C., Coffman, R.L., Barrat, F.J., Banchereau, J., Pascual, V., 2011. Netting neutrophils are major inducers of type I IFN production in pediatric systemic lupus erythematosus. Sci. Transl. Med. 3, 73ra20. Gardner, J.M., Devoss, J.J., Friedman, R.S., Wong, D.J., Tan, Y.X., Zhou, X., Johannes, K.P., Su, M.A., Chang, H.Y., Krummel, M.F., Anderson, M.S., 2008. Deletional tolerance mediated by extrathymic Aire-expressing cells. Science 321, 843–847. Garo, L.P., Murugaiyan, G., 2016. Contribution of microRNAs to autoimmune diseases. Cell. Mol. Life Sci. 73, 2041–2051. Gavrila, B.I., Ciofu, C., Stoica, V., 2016. Biomarkers in rheumatoid arthritis, what is new? J. Med. Life 9, 144–148. Gerritse, K., Laman, J.D., Noelle, R.J., Aruffo, A., Ledbetter, J.A., Boersma, W.J., Claassen, E., 1996. CD40-CD40 ligand interactions in experimental allergic encephalomyelitis and multiple sclerosis. Proc. Natl. Acad. Sci. U.S.A. 93, 2499–2504. Ghodke-Puranik, Y., Niewold, T.B., 2015. Immunogenetics of systemic lupus erythematosus: a comprehensive review. J. Autoimmun. 64, 125–136. Giles, J.R., Kashgarian, M., Koni, P.A., Shlomchik, M.J., 2015. B cell-specific MHC class II deletion reveals multiple nonredundant roles for B cell antigen presentation in murine lupus. J. Immunol. 195, 2571–2579. Gilhar, A., Bergman, R., Assay, B., Ullmann, Y., Etzioni, A., 2011. The beneficial effect of blocking Kv1.3 in the psoriasiform SCID mouse model. J. Invest. Dermatol. 131, 118–124. Giraud, M., Taubert, R., Vandiedonck, C., Ke, X., Levi-Strauss, M., Pagani, F., Baralle, F.E., Eymard, B., Tranchant, C., Gajdos, P., Vincent, A., Willcox, N., Beeson, D., Kyewski, B., Garchon, H.J., 2007. An IRF8-binding promoter variant and AIRE control CHRNA1 promiscuous expression in thymus. Nature 448, 934–937. Girschick, H., Wolf, C., Morbach, H., Hertzberg, C., Lee-Kirsch, M.A., 2015. Severe immune dysregulation with neurological impairment and minor bone changes in a child with spondyloenchondrodysplasia due to two novel mutations in the ACP5 gene. Pediatr. Rheumatol. Online J. 13, 37. Gocke, A.R., Lebson, L.A., Grishkan, I.V., Hu, L., Nguyen, H.M., Whartenby, K.A., Chandy, K.G., Calabresi, P.A., 2012. Kv1.3 deletion biases T cells toward an immunoregulatory phenotype and renders mice resistant to autoimmune encephalomyelitis. J. Immunol. 188, 5877–5886.

126

S.K. Devarapu et al.

Gong, Q., Cheng, A.M., Akk, A.M., Alberola-Ila, J., Gong, G., Pawson, T., Chan, A.C., 2001. Disruption of T cell signaling networks and development by Grb2 haploid insufficiency. Nat. Immunol. 2, 29–36. Gonzalez-Martin, A., Adams, B.D., Lai, M., Shepherd, J., Salvador-Bernaldez, M., Salvador, J.M., Lu, J., Nemazee, D., Xiao, C., 2016. The microRNA miR-148a functions as a critical regulator of B cell tolerance and autoimmunity. Nat. Immunol. 17, 433–440. Goodnow, C.C., 2007. Multistep pathogenesis of autoimmune disease. Cell 130, 25–35. Goodnow, C.C., Crosbie, J., Adelstein, S., Lavoie, T.B., Smith-Gill, S.J., Brink, R.A., Pritchard-Briscoe, H., Wotherspoon, J.S., Loblay, R.H., Raphael, K., Trent, R.J., Basten, A., 2009. Altered immunoglobulin expression and functional silencing of self-reactive B lymphocytes in transgenic mice. J. Immunol. 183, 5442–5448. Goris, A., Liston, A., 2012. The immunogenetic architecture of autoimmune disease. Cold Spring Harb. Perspect. Biol. 4, 1–14. Grabiec, A.M., Tak, P.P., Reedquist, K.A., 2008. Targeting histone deacetylase activity in rheumatoid arthritis and asthma as prototypes of inflammatory disease: should we keep our HATs on? Arthritis Res. Ther. 10, 226. Gray, D., Abramson, J., Benoist, C., Mathis, D., 2007. Proliferative arrest and rapid turnover of thymic epithelial cells expressing Aire. J. Exp. Med. 204, 2521–2528. Green, E.A., Wong, F.S., Eshima, K., Mora, C., Flavell, R.A., 2000. Neonatal tumor necrosis factor alpha promotes diabetes in nonobese diabetic mice by CD154-independent antigen presentation to CD8(+) T cells. J. Exp. Med. 191, 225–238. Gregory, P.D., Wagner, K., Horz, W., 2001. Histone acetylation and chromatin remodeling. Exp. Cell Res. 265, 195–202. Griffith, T.S., Brunner, T., Fletcher, S.M., Green, D.R., Ferguson, T.A., 1995. Fas ligand-induced apoptosis as a mechanism of immune privilege. Science 270, 1189–1192. Guan, H., Nagarkatti, P.S., Nagarkatti, M., 2011. CD44 reciprocally regulates the differentiation of encephalitogenic Th1/Th17 and Th2/regulatory T cells through epigenetic modulation involving DNA methylation of cytokine gene promoters, thereby controlling the development of experimental autoimmune encephalomyelitis. J. Immunol. 186, 6955–6964. Guglietta, S., Chiavelli, A., Zagato, E., Krieg, C., Gandini, S., Ravenda, P.S., Bazolli, B., Lu, B., Penna, G., Rescigno, M., 2016. Coagulation induced by C3aR-dependent NETosis drives protumorigenic neutrophils during small intestinal tumorigenesis. Nat. Commun. 7, 11037. Guiducci, C., Gong, M., Xu, Z., Gill, M., Chaussabel, D., Meeker, T., Chan, J.H., Wright, T., Punaro, M., Bolland, S., Soumelis, V., Banchereau, J., Coffman, R.L., Pascual, V., Barrat, F.J., 2010. TLR recognition of self nucleic acids hampers glucocorticoid activity in lupus. Nature 465, 937–941. Gunther, C., Kind, B., Reijns, M.A., Berndt, N., Martinez-Bueno, M., Wolf, C., Tungler, V., Chara, O., Lee, Y.A., Hubner, N., Bicknell, L., Blum, S., Krug, C., Schmidt, F., Kretschmer, S., Koss, S., Astell, K.R., Ramantani, G., Bauerfeind, A., Morris, D.L., Cunninghame Graham, D.S., Bubeck, D., Leitch, A., Ralston, S.H., Blackburn, E.A., Gahr, M., Witte, T., Vyse, T.J., Melchers, I., Mangold, E., Nothen, M.M., Aringer, M., Kuhn, A., Luthke, K., Unger, L., Bley, A., Lorenzi, A., Isaacs, J.D., Alexopoulou, D., Conrad, K., Dahl, A., Roers, A., Alarcon-Riquelme, M.E., Jackson, A.P., Lee-Kirsch, M.A., 2015. Defective removal of ribonucleotides from DNA promotes systemic autoimmunity. J. Clin. Invest. 125, 413–424. Gwack, Y., Feske, S., Srikanth, S., Hogan, P.G., Rao, A., 2007. Signalling to transcription: store-operated Ca2+ entry and NFAT activation in lymphocytes. Cell Calcium 42, 145–156.

Mechanisms of Autoimmunity

127

Gwyer Findlay, E., Danks, L., Madden, J., Cavanagh, M.M., McNamee, K., McCann, F., Snelgrove, R.J., Shaw, S., Feldmann, M., Taylor, P.C., Horwood, N.J., Hussell, T., 2014. OX40L blockade is therapeutic in arthritis, despite promoting osteoclastogenesis. Proc. Natl. Acad. Sci. U.S.A. 111, 2289–2294. Hadaschik, E.N., Wei, X., Leiss, H., Heckmann, B., Niederreiter, B., Steiner, G., Ulrich, W., Enk, A.H., Smolen, J.S., Stummvoll, G.H., 2015. Regulatory T cell-deficient scurfy mice develop systemic autoimmune features resembling lupus-like disease. Arthritis Res. Ther. 17, 35. Hagele, H., Allam, R., Pawar, R.D., Anders, H.J., 2009. Double-stranded RNA activates type I interferon secretion in glomerular endothelial cells via retinoic acid-inducible gene (RIG)-1. Nephrol. Dial. Transplant. 24, 3312–3318. Hahn, B.H., McMahon, M., 2008. Atherosclerosis and systemic lupus erythematosus: the role of altered lipids and of autoantibodies. Lupus 17, 368–370. Hakkim, A., Furnrohr, B.G., Amann, K., Laube, B., Abed, U.A., Brinkmann, V., Herrmann, M., Voll, R.E., Zychlinsky, A., 2010. Impairment of neutrophil extracellular trap degradation is associated with lupus nephritis. Proc. Natl. Acad. Sci. U.S.A. 107, 9813–9818. Halonen, M., Eskelin, P., Myhre, A.G., Perheentupa, J., Husebye, E.S., Kampe, O., Rorsman, F., Peltonen, L., Ulmanen, I., Partanen, J., 2002. AIRE mutations and human leukocyte antigen genotypes as determinants of the autoimmune polyendocrinopathy-candidiasis-ectodermal dystrophy phenotype. J. Clin. Endocrinol. Metab. 87, 2568–2574. Hamerman, J.A., Pottle, J., Ni, M., He, Y., Zhang, Z.Y., Buckner, J.H., 2016. Negative regulation of TLR signaling in myeloid cells—implications for autoimmune diseases. Immunol. Rev. 269, 212–227. Hampe, C.S., 2012. B cell in autoimmune diseases. Scientifica (Cairo). 2012, 1–18. Hanayama, R., Tanaka, M., Miyasaka, K., Aozasa, K., Koike, M., Uchiyama, Y., Nagata, S., 2004. Autoimmune disease and impaired uptake of apoptotic cells in MFG-E8-deficient mice. Science 304, 1147–1150. Harris, D.P., Haynes, L., Sayles, P.C., Duso, D.K., Eaton, S.M., Lepak, N.M., Johnson, L.L., Swain, S.L., Lund, F.E., 2000. Reciprocal regulation of polarized cytokine production by effector B and T cells. Nat. Immunol. 1, 475–482. Hartley, S.B., Cooke, M.P., Fulcher, D.A., Harris, A.W., Cory, S., Basten, A., Goodnow, C.C., 1993. Elimination of self-reactive B lymphocytes proceeds in two stages: arrested development and cell death. Cell 72, 325–335. Hatoff, D.E., Cohen, M., Schweigert, B.F., Talbert, W.M., 1979. Nitrofurantoin: another cause of drug-induced chronic active hepatitis? A report of a patient with HLA-B8 antigen. Am. J. Med. 67, 117–121. Hawiger, D., Tran, E., Du, W., Booth, C.J., Wen, L., Dong, C., Flavell, R.A., 2008. ICOS mediates the development of insulin-dependent diabetes mellitus in nonobese diabetic mice. J. Immunol. 180, 3140–3147. Haywood, M.E., Rogers, N.J., Rose, S.J., Boyle, J., McDermott, A., Rankin, J.M., Thiruudaian, V., Lewis, M.R., Fossati-Jimack, L., Izui, S., Walport, M.J., Morley, B.J., 2004. Dissection of BXSB lupus phenotype using mice congenic for chromosome 1 demonstrates that separate intervals direct different aspects of disease. J. Immunol. 173, 4277–4285. Healy, J.I., Goodnow, C.C., 1998. Positive versus negative signaling by lymphocyte antigen receptors. Annu. Rev. Immunol. 16, 645–670. Healy, J.I., Dolmetsch, R.E., Timmerman, L.A., Cyster, J.G., Thomas, M.L., Crabtree, G.R., Lewis, R.S., Goodnow, C.C., 1997. Different nuclear signals are activated by the B cell receptor during positive versus negative signaling. Immunity 6, 419–428.

128

S.K. Devarapu et al.

Hedrich, C.M., Crispin, J.C., Rauen, T., Ioannidis, C., Apostolidis, S.A., Lo, M.S., Kyttaris, V.C., Tsokos, G.C., 2012. cAMP response element modulator alpha controls IL2 and IL17A expression during CD4 lineage commitment and subset distribution in lupus. Proc. Natl. Acad. Sci. U.S.A. 109, 16606–16611. Hervouet, E., Vallette, F.M., Cartron, P.F., 2010. Dnmt1/Transcription factor interactions: an alternative mechanism of DNA methylation inheritance. Genes Cancer 1, 434–443. Hippen, K.L., Tze, L.E., Behrens, T.W., 2000. CD5 maintains tolerance in anergic B cells. J. Exp. Med. 191, 883–890. Hong, Y., Eleftheriou, D., Hussain, A.A., Price-Kuehne, F.E., Savage, C.O., Jayne, D., Little, M.A., Salama, A.D., Klein, N.J., Brogan, P.A., 2012. Anti-neutrophil cytoplasmic antibodies stimulate release of neutrophil microparticles. J. Am. Soc. Nephrol. 23, 49–62. Hori, S., Nomura, T., Sakaguchi, S., 2003. Control of regulatory T cell development by the transcription factor Foxp3. Science 299, 1057–1061. Howard, L.M., Miga, A.J., Vanderlugt, C.L., Dal Canto, M.C., Laman, J.D., Noelle, R.J., Miller, S.D., 1999. Mechanisms of immunotherapeutic intervention by anti-CD40L (CD154) antibody in an animal model of multiple sclerosis. J. Clin. Invest. 103, 281–290. Hoyer, B.F., Moser, K., Hauser, A.E., Peddinghaus, A., Voigt, C., Eilat, D., Radbruch, A., Hiepe, F., Manz, R.A., 2004. Short-lived plasmablasts and long-lived plasma cells contribute to chronic humoral autoimmunity in NZB/W mice. J. Exp. Med. 199, 1577–1584. Hu, Y.L., Metz, D.P., Chung, J., Siu, G., Zhang, M., 2009. B7RP-1 blockade ameliorates autoimmunity through regulation of follicular helper T cells. J. Immunol. 182, 1421–1428. Hueber, W., Patel, D.D., Dryja, T., Wright, A.M., Koroleva, I., Bruin, G., Antoni, C., Draelos, Z., Gold, M.H., Psoriasis Study, G., Durez, P., Tak, P.P., Gomez-Reino, J.J., Rheumatoid Arthritis Study, G., Foster, C.S., Kim, R.Y., Samson, C.M., Falk, N.S., Chu, D.S., Callanan, D., Nguyen, Q.D., Uveitis Study, G., Rose, K., Haider, A., Di Padova, F., 2010. Effects of AIN457, a fully human antibody to interleukin-17A, on psoriasis, rheumatoid arthritis, and uveitis. Sci. Transl. Med. 2, 52ra72. Hugot, J.P., Chamaillard, M., Zouali, H., Lesage, S., Cezard, J.P., Belaiche, J., Almer, S., Tysk, C., O’Morain, C.A., Gassull, M., Binder, V., Finkel, Y., Cortot, A., Modigliani, R., Laurent-Puig, P., Gower-Rousseau, C., Macry, J., Colombel, J.F., Sahbatou, M., Thomas, G., 2001. Association of NOD2 leucine-rich repeat variants with susceptibility to Crohn’s disease. Nature 411, 599–603. Hwang, I., Shen, X., Sprent, J., 2003. Direct stimulation of naive T cells by membrane vesicles from antigen-presenting cells: distinct roles for CD54 and B7 molecules. Proc. Natl. Acad. Sci. U.S.A. 100, 6670–6675. Hwang, S.H., Lee, H., Yamamoto, M., Jones, L.A., Dayalan, J., Hopkins, R., Zhou, X.J., Yarovinsky, F., Connolly, J.E., Curotto de Lafaille, M.A., Wakeland, E.K., Fairhurst, A.M., 2012. B cell TLR7 expression drives anti-RNA autoantibody production and exacerbates disease in systemic lupus erythematosus-prone mice. J. Immunol. 189, 5786–5796. Ignatowicz, L., Kappler, J., Marrack, P., 1996. The repertoire of T cells shaped by a single MHC/peptide ligand. Cell 84, 521–529. Iles, M.M., 2008. What can genome-wide association studies tell us about the genetics of common disease? PLoS Genet. 4, e33. Ilmarinen, T., Kangas, H., Kytomaa, T., Eskelin, P., Saharinen, J., Seeler, J.S., Tanhuanpaa, K., Chan, F.Y., Slattery, R.M., Alakurtti, K., Palvimo, J.J., Ulmanen, I., 2008. Functional interaction of AIRE with PIAS1 in transcriptional regulation. Mol. Immunol. 45, 1847–1862.

Mechanisms of Autoimmunity

129

Imaizumi, T., Aizawa-Yashiro, T., Tsuruga, K., Tanaka, H., Matsumiya, T., Yoshida, H., Tatsuta, T., Xing, F., Hayakari, R., Satoh, K., 2012. Melanoma differentiation-associated gene 5 regulates the expression of a chemokine CXCL10 in human mesangial cells: implications for chronic inflammatory renal diseases. Tohoku J. Exp. Med. 228, 17–26. Imaizumi, T., Aizawa, T., Segawa, C., Shimada, M., Tsuruga, K., Kawaguchi, S., Matsumiya, T., Yoshida, H., Joh, K., Tanaka, H., 2014. Toll-like receptor 3 signaling contributes to the expression of a neutrophil chemoattractant, CXCL1 in human mesangial cells. Clin. Exp. Nephrol. 19, 761–770. Inobe, M., Schwartz, R.H., 2004. CTLA-4 engagement acts as a brake on CD4 + T cell proliferation and cytokine production but is not required for tuning T cell reactivity in adaptive tolerance. J. Immunol. 173, 7239–7248. Ishikawa, O., Ishikawa, H., 1992. Macrophage infiltration in the skin of patients with systemic sclerosis. J. Rheumatol. 19, 1202–1206. Ivascu, C., Wasserkort, R., Lesche, R., Dong, J., Stein, H., Thiel, A., Eckhardt, F., 2007. DNA methylation profiling of transcription factor genes in normal lymphocyte development and lymphomas. Int. J. Biochem. Cell Biol. 39, 1523–1538. Iwai, H., Kozono, Y., Hirose, S., Akiba, H., Yagita, H., Okumura, K., Kohsaka, H., Miyasaka, N., Azuma, M., 2002. Amelioration of collagen-induced arthritis by blockade of inducible costimulator-B7 homologous protein costimulation. J. Immunol. 169, 4332–4339. Iwata, S., Tanaka, Y., 2016. B-cell subsets, signaling and their roles in secretion of autoantibodies. Lupus 25, 850–856. Jacinto, J., Kim, P.J., Singh, R.R., 2012. Disparate effects of depletion of CD1d-reactive T cells during early versus late stages of disease in a genetically susceptible model of lupus. Lupus 21, 485–490. Jacob, M., Napirei, M., Ricken, A., Dixkens, C., Mannherz, H.G., 2002. Histopathology of lupus-like nephritis in Dnase1-deficient mice in comparison to NZB/W F1 mice. Lupus 11, 514–527. Jacquemin, C., Schmitt, N., Contin-Bordes, C., Liu, Y., Narayanan, P., Seneschal, J., Maurouard, T., Dougall, D., Davizon, E.S., Dumortier, H., Douchet, I., Raffray, L., Richez, C., Lazaro, E., Duffau, P., Truchetet, M.E., Khoryati, L., Mercie, P., Couzi, L., Merville, P., Schaeverbeke, T., Viallard, J.F., Pellegrin, J.L., Moreau, J.F., Muller, S., Zurawski, S., Coffman, R.L., Pascual, V., Ueno, H., Blanco, P., 2015. OX40 ligand contributes to human lupus pathogenesis by promoting T follicular helper response. Immunity 42, 1159–1170. Janeway Jr., C.A., Medzhitov, R., 1999. Lipoproteins take their toll on the host. Curr. Biol. 9, R879–882. Jang, M.A., Kim, E.K., Now, H., Nguyen, N.T., Kim, W.J., Yoo, J.Y., Lee, J., Jeong, Y.M., Kim, C.H., Kim, O.H., Sohn, S., Nam, S.H., Hong, Y., Lee, Y.S., Chang, S.A., Jang, S.Y., Kim, J.W., Lee, M.S., Lim, S.Y., Sung, K.S., Park, K.T., Kim, B.J., Lee, J.H., Kim, D.K., Kee, C., Ki, C.S., 2015. Mutations in DDX58, which encodes RIG-I, cause atypical Singleton-Merten syndrome. Am. J. Hum. Genet. 96, 266–274. Jankovic, M., Casellas, R., Yannoutsos, N., Wardemann, H., Nussenzweig, M.C., 2004. RAGs and regulation of autoantibodies. Annu. Rev. Immunol. 22, 485–501. Jeffries, M.A., Sawalha, A.H., 2011. Epigenetics in systemic lupus erythematosus: leading the way for specific therapeutic agents. Int. J. Clin. Rheumatol. 6, 423–439. Jeffries, M.A., Sawalha, A.H., 2015. Autoimmune disease in the epigenetic era: how has epigenetics changed our understanding of disease and how can we expect the field to evolve? Expert Rev. Clin. Immunol. 11, 45–58.

130

S.K. Devarapu et al.

Jego, G., Palucka, A.K., Blanck, J.P., Chalouni, C., Pascual, V., Banchereau, J., 2003. Plasmacytoid dendritic cells induce plasma cell differentiation through type I interferon and interleukin 6. Immunity 19, 225–234. Jennette, J.C., Falk, R.J., 2014. Pathogenesis of antineutrophil cytoplasmic autoantibody-mediated disease. Nat. Rev. Rheumatol. 10, 463–473. Jeon, M.S., Atfield, A., Venuprasad, K., Krawczyk, C., Sarao, R., Elly, C., Yang, C., Arya, S., Bachmaier, K., Su, L., Bouchard, D., Jones, R., Gronski, M., Ohashi, P., Wada, T., Bloom, D., Fathman, C.G., Liu, Y.C., Penninger, J.M., 2004. Essential role of the E3 ubiquitin ligase Cbl-b in T cell anergy induction. Immunity 21, 167–177. Jerne, N.K., 2004. The somatic generation of immune recognition. 1971. Eur. J. Immunol. 34, 1234–1242. Ji, J., Xu, J., Zhao, S., Liu, F., Qi, J., Song, Y., Ren, J., Wang, T., Dou, H., Hou, Y., 2016. Myeloid-derived suppressor cells contribute to systemic lupus erythematosus by regulating differentiation of Th17 cells and Tregs. Clin. Sci. (Lond.) 130, 1453–1467. Jin, J., Desai, B.N., Navarro, B., Donovan, A., Andrews, N.C., Clapham, D.E., 2008. Deletion of Trpm7 disrupts embryonic development and thymopoiesis without altering Mg2 + homeostasis. Science 322, 756–760. Jin, J., Lian, T., Gu, C., Yu, K., Gao, Y.Q., Su, X.D., 2016. The effects of cytosine methylation on general transcription factors. Sci. Rep. 6, 29119. Joetham, A., Schedel, M., O’Connor, B.P., Kim, S., Takeda, K., Abbott, J., Gelfand, E.W., 2016. Inducible and naturally occurring T regulatory cells enhance lung allergic responses Via divergent transcriptional pathways. J. Allergy Clin. Immunol. [Epub ahead of print]. Joller, N., Hafler, J.P., Brynedal, B., Kassam, N., Spoerl, S., Levin, S.D., Sharpe, A.H., Kuchroo, V.K., 2011. Cutting edge: TIGIT has T cell-intrinsic inhibitory functions. J. Immunol. 186, 1338–1342. Jones, G.W., Jones, S.A., 2016. Ectopic lymphoid follicles: inducible centres for generating antigen-specific immune responses within tissues. Immunology 147, 141–151. Jury, E.C., Kabouridis, P.S., Flores-Borja, F., Mageed, R.A., Isenberg, D.A., 2004. Altered lipid raft-associated signaling and ganglioside expression in T lymphocytes from patients with systemic lupus erythematosus. J. Clin. Invest. 113, 1176–1187. Kahlenberg, J.M., Carmona-Rivera, C., Smith, C.K., Kaplan, M.J., 2013. Neutrophil extracellular trap-associated protein activation of the NLRP3 inflammasome is enhanced in lupus macrophages. J. Immunol. 190, 1217–1226. Kain, R., Exner, M., Brandes, R., Ziebermayr, R., Cunningham, D., Alderson, C.A., Davidovits, A., Raab, I., Jahn, R., Ashour, O., Spitzauer, S., Sunder-Plassmann, G., Fukuda, M., Klemm, P., Rees, A.J., Kerjaschki, D., 2008. Molecular mimicry in pauci-immune focal necrotizing glomerulonephritis. Nat. Med. 14, 1088–1096. Kambas, K., Chrysanthopoulou, A., Vassilopoulos, D., Apostolidou, E., Skendros, P., Girod, A., Arelaki, S., Froudarakis, M., Nakopoulou, L., Giatromanolaki, A., Sidiropoulos, P., Koffa, M., Boumpas, D.T., Ritis, K., Mitroulis, I., 2014. Tissue factor expression in neutrophil extracellular traps and neutrophil derived microparticles in antineutrophil cytoplasmic antibody associated vasculitis may promote thromboinflammation and the thrombophilic state associated with the disease. Ann. Rheum. Dis. 73, 1854–1863. Kanter, J.L., Narayana, S., Ho, P.P., Catz, I., Warren, K.G., Sobel, R.A., Steinman, L., Robinson, W.H., 2006. Lipid microarrays identify key mediators of autoimmune brain inflammation. Nat. Med. 12, 138–143. Kappler, J.W., Roehm, N., Marrack, P., 1987. T cell tolerance by clonal elimination in the thymus. Cell 49, 273–280. Kariko, K., Ni, H., Capodici, J., Lamphier, M., Weissman, D., 2004. mRNA is an endogenous ligand for Toll-like receptor 3. J. Biol. Chem. 279, 12542–12550.

Mechanisms of Autoimmunity

131

Karin, N., Wildbaum, G., 2015. The role of chemokines in adjusting the balance between CD4+ effector T cell subsets and FOXp3-negative regulatory T cells. Int. Immunopharmacol. 28, 829–835. Kasagi, S., Kawano, S., Okazaki, T., Honjo, T., Morinobu, A., Hatachi, S., Shimatani, K., Tanaka, Y., Minato, N., Kumagai, S., 2010. Anti-programmed cell death 1 antibody reduces CD4+PD-1 + T cells and relieves the lupus-like nephritis of NZB/W F1 mice. J. Immunol. 184, 2337–2347. Katagiri, Y.U., Kiyokawa, N., Fujimoto, J., 2001. A role for lipid rafts in immune cell signaling. Microbiol. Immunol. 45, 1–8. Katsiari, C.G., Liossis, S.N., Sfikakis, P.P., 2010. The pathophysiologic role of monocytes and macrophages in systemic lupus erythematosus: a reappraisal. Semin. Arthritis Rheum. 39, 491–503. Katto, J., Engel, N., Abbas, W., Herbein, G., Mahlknecht, U., 2013. Transcription factor NFkappaB regulates the expression of the histone deacetylase SIRT1. Clin. Epigenetics 5, 11. Kawahara, T.L., Michishita, E., Adler, A.S., Damian, M., Berber, E., Lin, M., McCord, R.A., Ongaigui, K.C., Boxer, L.D., Chang, H.Y., Chua, K.F., 2009. SIRT6 links histone H3 lysine 9 deacetylation to NF-kappaB-dependent gene expression and organismal life span. Cell 136, 62–74. Kawane, K., Ohtani, M., Miwa, K., Kizawa, T., Kanbara, Y., Yoshioka, Y., Yoshikawa, H., Nagata, S., 2006. Chronic polyarthritis caused by mammalian DNA that escapes from degradation in macrophages. Nature 443, 998–1002. Kawasaki, A., Furukawa, H., Kondo, Y., Ito, S., Hayashi, T., Kusaoi, M., Matsumoto, I., Tohma, S., Takasaki, Y., Hashimoto, H., Sumida, T., Tsuchiya, N., 2011. TLR7 single-nucleotide polymorphisms in the 3’ untranslated region and intron 2 independently contribute to systemic lupus erythematosus in Japanese women: a case-control association study. Arthritis Res. Ther. 13, R41. Kessenbrock, K., Krumbholz, M., Schonermarck, U., Back, W., Gross, W.L., Werb, Z., Grone, H.J., Brinkmann, V., Jenne, D.E., 2009. Netting neutrophils in autoimmune small-vessel vasculitis. Nat. Med. 15, 623–625. Khandpur, R., Carmona-Rivera, C., Vivekanandan-Giri, A., Gizinski, A., Yalavarthi, S., Knight, J.S., Friday, S., Li, S., Patel, R.M., Subramanian, V., Thompson, P., Chen, P., Fox, D.A., Pennathur, S., Kaplan, M.J., 2013. NETs are a source of citrullinated autoantigens and stimulate inflammatory responses in rheumatoid arthritis. Sci. Transl. Med. 5, 178ra140. Kim, N.D., Luster, A.D., 2015. The role of tissue resident cells in neutrophil recruitment. Trends Immunol. 36, 547–555. Kim, J.M., Rasmussen, J.P., Rudensky, A.Y., 2007. Regulatory T cells prevent catastrophic autoimmunity throughout the lifespan of mice. Nat. Immunol. 8, 191–197. Kim, D., Peck, A., Santer, D., Patole, P., Schwartz, S.M., Molitor, J.A., Arnett, F.C., Elkon, K.B., 2008. Induction of interferon-alpha by scleroderma sera containing autoantibodies to topoisomerase I: association of higher interferon-alpha activity with lung fibrosis. Arthritis Rheum. 58, 2163–2173. Klareskog, L., Catrina, A.I., 2015. Autoimmunity: lungs and citrullination. Nat. Rev. Rheumatol. 11, 261–262. Klein, K., Gay, S., 2015. Epigenetics in rheumatoid arthritis. Curr. Opin. Rheumatol. 27, 76–82. Klonowska-Szymczyk, A., Wolska, A., Robak, T., Cebula-Obrzut, B., Smolewski, P., Robak, E., 2014. Expression of toll-like receptors 3, 7, and 9 in peripheral blood mononuclear cells from patients with systemic lupus erythematosus. Mediators Inflamm. 2014, 381418.

132

S.K. Devarapu et al.

Knight, J.S., Subramanian, V., O’Dell, A.A., Yalavarthi, S., Zhao, W., Smith, C.K., Hodgin, J.B., Thompson, P.R., Kaplan, M.J., 2015. Peptidylarginine deiminase inhibition disrupts NET formation and protects against kidney, skin and vascular disease in lupus-prone MRL/lpr mice. Ann. Rheum. Dis. 74, 2199–2206. Knijff-Dutmer, E.A., Koerts, J., Nieuwland, R., Kalsbeek-Batenburg, E.M., van de Laar, M.A., 2002. Elevated levels of platelet microparticles are associated with disease activity in rheumatoid arthritis. Arthritis Rheum. 46, 1498–1503. Knoerzer, D.B., Karr, R.W., Schwartz, B.D., Mengle-Gaw, L.J., 1995. Collagen-induced arthritis in the BB rat. Prevention of disease by treatment with CTLA-4-Ig. J. Clin. Invest. 96, 987–993. Ko, H.J., Cho, M.L., Lee, S.Y., Oh, H.J., Heo, Y.J., Moon, Y.M., Kang, C.M., Kwok, S.K., Ju, J.H., Park, S.H., Park, K.S., Kim, H.Y., 2010. CTLA4-Ig modifies dendritic cells from mice with collagen-induced arthritis to increase the CD4+CD25+Foxp3 + regulatory T cell population. J. Autoimmun. 34, 111–120. Koga, T., Ichinose, K., Tsokos, G.C., 2016. T cells and IL-17 in lupus nephritis. Clin. Immunol. [Epub ahead of print]. Kogawa, K., Kudoh, J., Nagafuchi, S., Ohga, S., Katsuta, H., Ishibashi, H., Harada, M., Hara, T., Shimizu, N., 2002. Distinct clinical phenotype and immunoreactivity in Japanese siblings with autoimmune polyglandular syndrome type 1 (APS-1) associated with compound heterozygous novel AIRE gene mutations. Clin. Immunol. 103, 277–283. Kong, Y.M., Brown, N.K., Morris, G.P., Flynn, J.C., 2015. The essential role of circulating thyroglobulin in maintaining dominance of natural regulatory T cell function to prevent autoimmune thyroiditis. Horm. Metab. Res. 47, 711–720. Konig, N., Fiehn, C., Wolf, C., Schuster, M., Cura Costa, E., Tungler, V., Alvarez, H.A., Chara, O., Engel, K., Goldbach-Mansky, R., Gunther, C., Lee-Kirsch, M.A., 2017. Familial chilblain lupus due to a gain-of-function mutation in STING. Ann. Rheum. Dis. 76, 468–472. Kont, V., Laan, M., Kisand, K., Merits, A., Scott, H.S., Peterson, P., 2008. Modulation of Aire regulates the expression of tissue-restricted antigens. Mol. Immunol. 45, 25–33. Konya, C., Paz, Z., Tsokos, G.C., 2014. The role of T cells in systemic lupus erythematosus: an update. Curr. Opin. Rheumatol. 26, 493–501. Koralov, S.B., Muljo, S.A., Galler, G.R., Krek, A., Chakraborty, T., Kanellopoulou, C., Jensen, K., Cobb, B.S., Merkenschlager, M., Rajewsky, N., Rajewsky, K., 2008. Dicer ablation affects antibody diversity and cell survival in the B lymphocyte lineage. Cell 132, 860–874. Kow, N.Y., Mak, A., 2013. Costimulatory pathways: physiology and potential therapeutic manipulation in systemic lupus erythematosus. Clin. Dev. Immunol. 2013, 245928. Kraling, B.M., Maul, G.G., Jimenez, S.A., 1995. Mononuclear cellular infiltrates in clinically involved skin from patients with systemic sclerosis of recent onset predominantly consist of monocytes/macrophages. Pathobiology 63, 48–56. Kravchenko, P.N., Zhulai, G.A., Churov, A.V., Oleinik, E.K., Oleinik, V.M., Barysheva, O.Y., Vezikova, N.N., Marusenko, I.M., 2016. Subpopulations of regulatory T-lymphocytes in the peripheral blood of patients with rheumatoid arthritis. Vestn. Ross. Akad. Med. Nauk. 2, 48–153. Krishnan, M.R., Wang, C., Marion, T.N., 2012. Anti-DNA autoantibodies initiate experimental lupus nephritis by binding directly to the glomerular basement membrane in mice. Kidney Int. 82, 184–192. Krol, J., Loedige, I., Filipowicz, W., 2010. The widespread regulation of microRNA biogenesis, function and decay. Nat. Rev. Genet. 11, 597–610. Kuchen, S., Seemayer, C.A., Rethage, J., von Knoch, R., Kuenzler, P., Beat, A.M., Gay, R.E., Gay, S., Neidhart, M., 2004. The L1 retroelement-related p40 protein induces p38delta MAP kinase. Autoimmunity 37, 57–65.

Mechanisms of Autoimmunity

133

Kuehn, H.S., Niemela, J.E., Rangel-Santos, A., Zhang, M., Pittaluga, S., Stoddard, J.L., Hussey, A.A., Evbuomwan, M.O., Priel, D.A., Kuhns, D.B., Park, C.L., Fleisher, T.A., Uzel, G., Oliveira, J.B., 2013. Loss-of-function of the protein kinase C delta (PKCdelta) causes a B-cell lymphoproliferative syndrome in humans. Blood 121, 3117–3125. Kumar, V., Aziz, F., Sercarz, E., Miller, A., 1997. Regulatory T cells specific for the same framework 3 region of the Vβ8.2 chain Are involved in the control of collagen II-induced arthritis and experimental autoimmune encephalomyelitis. J. Exp. Med. 185, 1725–1733. Kumar, P.G., Laloraya, M., Wang, C.Y., Ruan, Q.G., Davoodi-Semiromi, A., Kao, K.J., She, J.X., 2001. The autoimmune regulator (AIRE) is a DNA-binding protein. J. Biol. Chem. 276, 41357–41364. Kunimoto, K., Kimura, A., Uede, K., Okuda, M., Aoyagi, N., Furukawa, F., Kanazawa, N., 2013. A new infant case of Nakajo-Nishimura syndrome with a genetic mutation in the immunoproteasome subunit: an overlapping entity with JMP and CANDLE syndrome related to PSMB8 mutations. Dermatology 227, 26–30. Kunzelmann, K., 2016. Ion channels in regulated cell death. Cell. Mol. Life Sci. 73, 2387–2403. Kuroiwa, T., Schlimgen, R., Illei, G.G., Boumpas, D.T., 2003. Monocyte response to Th1 stimulation and effector function toward human mesangial cells are not impaired in patients with lupus nephritis. Clin. Immunol. 106, 65–72. Kurowska-Stolarska, M., Alivernini, S., Ballantine, L.E., Asquith, D.L., Millar, N.L., Gilchrist, D.S., Reilly, J., Ierna, M., Fraser, A.R., Stolarski, B., McSharry, C., Hueber, A.J., Baxter, D., Hunter, J., Gay, S., Liew, F.Y., McInnes, I.B., 2011. MicroRNA-155 as a proinflammatory regulator in clinical and experimental arthritis. Proc. Natl. Acad. Sci. U.S.A. 108, 11193–11198. Kyburz, D., Corr, M., Brinson, D.C., Von Damm, A., Tighe, H., Carson, D.A., 1999. Human rheumatoid factor production is dependent on CD40 signaling and autoantigen. J. Immunol. 163, 3116–3122. Kyewski, B., Derbinski, J., 2004. Self-representation in the thymus: an extended view. Nat. Rev. Immunol. 4, 688–698. Lafferty, K.J., Cunningham, A.J., 1975. A new analysis of allogeneic interactions. Aust. J. Exp. Biol. Med. Sci. 53, 27–42. Lang, J., Ota, T., Kelly, M., Strauch, P., Freed, B.M., Torres, R.M., Nemazee, D., Pelanda, R., 2016. Receptor editing and genetic variability in human autoreactive B cells. J. Exp. Med. 213, 93–108. Lartigue, A., Colliou, N., Calbo, S., Francois, A., Jacquot, S., Arnoult, C., Tron, F., Gilbert, D., Musette, P., 2009. Critical role of TLR2 and TLR4 in autoantibody production and glomerulonephritis in lpr mutation-induced mouse lupus. J. Immunol. 183, 6207–6216. Lau, C.M., Broughton, C., Tabor, A.S., Akira, S., Flavell, R.A., Mamula, M.J., Christensen, S.R., Shlomchik, M.J., Viglianti, G.A., Rifkin, I.R., Marshak-Rothstein, A., 2005. RNA-associated autoantigens activate B cells by combined B cell antigen receptor/Toll-like receptor 7 engagement. J. Exp. Med. 202, 1171–1177. Laufer, T.M., DeKoning, J., Markowitz, J.S., Lo, D., Glimcher, L.H., 1996. Unopposed positive selection and autoreactivity in mice expressing class II MHC only on thymic cortex. Nature 383, 81–85. Lawrenson, R.A., Seaman, H.E., Sundstrom, A., Williams, T.J., Farmer, R.D., 2000. Liver damage associated with minocycline use in acne: a systematic review of the published literature and pharmacovigilance data. Drug Saf. 23, 333–349. Leadbetter, E.A., Rifkin, I.R., Hohlbaum, A.M., Beaudette, B.C., Shlomchik, M.J., Marshak-Rothstein, A., 2002. Chromatin-IgG complexes activate B cells by dual engagement of IgM and toll-like receptors. Nature 416, 603–607.

134

S.K. Devarapu et al.

Leanza, L., Zoratti, M., Gulbins, E., Szabo, I., 2012. Induction of apoptosis in macrophages via Kv1.3 and Kv1.5 potassium channels. Curr. Med. Chem. 19, 5394–5404. Lech, M., Anders, H.J., 2013. The pathogenesis of lupus nephritis. J. Am. Soc. Nephrol. 24, 1357–1366. Lech, M., Skuginna, V., Kulkarni, O.P., Gong, J., Wei, T., Stark, R.W., Garlanda, C., Mantovani, A., Anders, H.J., 2010. Lack of SIGIRR/TIR8 aggravates hydrocarbon oil-induced lupus nephritis. J. Pathol. 220, 596–607. Lech, M., Lorenz, G., Kulkarni, O.P., Grosser, M.O., Stigrot, N., Darisipudi, M.N., Gunthner, R., Wintergerst, M.W., Anz, D., Susanti, H.E., Anders, H.J., 2015. NLRP3 and ASC suppress lupus-like autoimmunity by driving the immunosuppressive effects of TGF-beta receptor signalling. Ann. Rheum. Dis. 74, 2224–2235. Lee, Y., Chin, R.K., Christiansen, P., Sun, Y., Tumanov, A.V., Wang, J., Chervonsky, A.V., Fu, Y.X., 2006. Recruitment and activation of naive T cells in the islets by lymphotoxin beta receptor-dependent tertiary lymphoid structure. Immunity 25, 499–509. Lee, P.Y., Kumagai, Y., Li, Y., Takeuchi, O., Yoshida, H., Weinstein, J., Kellner, E.S., Nacionales, D., Barker, T., Kelly-Scumpia, K., van Rooijen, N., Kumar, H., Kawai, T., Satoh, M., Akira, S., Reeves, W.H., 2008a. TLR7-dependent and FcγR-independent production of type I interferon in experimental mouse lupus. J. Exp. Med. 205, 2995–3006. Lee, P.Y., Weinstein, J.S., Nacionales, D.C., Scumpia, P.O., Li, Y., Butfiloski, E., van Rooijen, N., Moldawer, L., Satoh, M., Reeves, W.H., 2008b. A novel type I IFN-producing cell subset in murine lupus. J. Immunol. 180, 5101–5108. Lee, H., Lee, E.J., Kim, H., Lee, G., Um, E.J., Kim, Y., Lee, B.Y., Bae, H., 2011. Bee venom-associated Th1/Th2 immunoglobulin class switching results in immune tolerance of NZB/W F1 murine lupus nephritis. Am. J. Nephrol. 34, 163–172. Lee, Y.H., Choi, S.J., Ji, J.D., Song, G.G., 2016. Association between toll-like receptor polymorphisms and systemic lupus erythematosus: a meta-analysis update. Lupus 25, 593–601. Lee-Kirsch, M.A., Gong, M., Chowdhury, D., Senenko, L., Engel, K., Lee, Y.A., de Silva, U., Bailey, S.L., Witte, T., Vyse, T.J., Kere, J., Pfeiffer, C., Harvey, S., Wong, A., Koskenmies, S., Hummel, O., Rohde, K., Schmidt, R.E., Dominiczak, A.F., Gahr, M., Hollis, T., Perrino, F.W., Lieberman, J., Hubner, N., 2007. Mutations in the gene encoding the 30 -50 DNA exonuclease TREX1 are associated with systemic lupus erythematosus. Nat. Genet. 39, 1065–1067. Leffler, J., Martin, M., Gullstrand, B., Tyden, H., Lood, C., Truedsson, L., Bengtsson, A.A., Blom, A.M., 2012. Neutrophil extracellular traps that are not degraded in systemic lupus erythematosus activate complement exacerbating the disease. J. Immunol. 188, 3522–3531. Leffler, J., Stojanovich, L., Shoenfeld, Y., Bogdanovic, G., Hesselstrand, R., Blom, A.M., 2014. Degradation of neutrophil extracellular traps is decreased in patients with antiphospholipid syndrome. Clin. Exp. Rheumatol. 32, 66–70. Leffler, J., Ciacma, K., Gullstrand, B., Bengtsson, A.A., Martin, M., Blom, A.M., 2015. A subset of patients with systemic lupus erythematosus fails to degrade DNA from multiple clinically relevant sources. Arthritis Res. Ther. 17, 205. Leiss, H., Niederreiter, B., Bandur, T., Schwarzecker, B., Bluml, S., Steiner, G., Ulrich, W., Smolen, J.S., Stummvoll, G.H., 2013. Pristane-induced lupus as a model of human lupus arthritis: evolvement of autoantibodies, internal organ and joint inflammation. Lupus 22, 778–792. Lenschow, D.J., Herold, K.C., Rhee, L., Patel, B., Koons, A., Qin, H.Y., Fuchs, E., Singh, B., Thompson, C.B., Bluestone, J.A., 1996. CD28/B7 regulation of Th1 and Th2 subsets in the development of autoimmune diabetes. Immunity 5, 285–293.

Mechanisms of Autoimmunity

135

Leung, D.Y., Travers, J.B., Giorno, R., Norris, D.A., Skinner, R., Aelion, J., Kazemi, L.V., Kim, M.H., Trumble, A.E., Kotb, M., et al., 1995. Evidence for a streptococcal superantigen-driven process in acute guttate psoriasis. J. Clin. Invest. 96, 2106–2112. Levin, S.D., Taft, D.W., Brandt, C.S., Bucher, C., Howard, E.D., Chadwick, E.M., Johnston, J., Hammond, A., Bontadelli, K., Ardourel, D., Hebb, L., Wolf, A., Bukowski, T.R., Rixon, M.W., Kuijper, J.L., Ostrander, C.D., West, J.W., Bilsborough, J., Fox, B., Gao, Z., Xu, W., Ramsdell, F., Blazar, B.R., Lewis, K.E., 2011. Vstm3 is a member of the CD28 family and an important modulator of T-cell function. Eur. J. Immunol. 41, 902–915. Li, Y., Huang, Y., Lue, J., Quandt, J.A., Martin, R., Mariuzza, R.A., 2005. Structure of a human autoimmune TCR bound to a myelin basic protein self-peptide and a multiple sclerosis-associated MHC class II molecule. EMBO J. 24, 2968–2979. Li, Q.J., Chau, J., Ebert, P.J., Sylvester, G., Min, H., Liu, G., Braich, R., Manoharan, M., Soutschek, J., Skare, P., Klein, L.O., Davis, M.M., Chen, C.Z., 2007. miR-181a is an intrinsic modulator of T cell sensitivity and selection. Cell 129, 147–161. Li, Y., Reddy, M.A., Miao, F., Shanmugam, N., Yee, J.K., Hawkins, D., Ren, B., Natarajan, R., 2008. Role of the histone H3 lysine 4 methyltransferase, SET7/9, in the regulation of NF-kappaB-dependent inflammatory genes. Relevance to diabetes and inflammation. J. Biol. Chem. 283, 26771–26781. Li, G., Yu, M., Lee, W.W., Tsang, M., Krishnan, E., Weyand, C.M., Goronzy, J.J., 2012. Decline in miR-181a expression with age impairs T cell receptor sensitivity by increasing DUSP6 activity. Nat. Med. 18, 1518–1524. Li, B., Tian, L., Diao, Y., Li, X., Zhao, L., Wang, X., 2014. Exogenous IL-10 induces corneal transplantation immune tolerance by a mechanism associated with the altered Th1/Th2 cytokine ratio and the increased expression of TGF-beta. Mol. Med. Rep. 9, 2245–2250. Li, W., Li, G., Zhang, Y., Wei, S., Song, M., Wang, W., Yuan, X., Wu, H., Yang, Y., 2015. Role of P2 x 7 receptor in the differentiation of bone marrow stromal cells into osteoblasts and adipocytes. Exp. Cell Res. 339, 367–379. Lian, Y.T., Yang, X.F., Wang, Z.H., Yang, Y., Yang, Y., Shu, Y.W., Cheng, L.X., Liu, K., 2013. Curcumin serves as a human kv1.3 blocker to inhibit effector memory T lymphocyte activities. Phytother. Res. 27, 1321–1327. Liang, B., Gee, R.J., Kashgarian, M.J., Sharpe, A.H., Mamula, M.J., 1999. B7 costimulation in the development of lupus: autoimmunity arises either in the absence of B7.1/B7.2 or in the presence of anti-b7.1/B7.2 blocking antibodies. J. Immunol. 163, 2322–2329. Liang, S.C., Latchman, Y.E., Buhlmann, J.E., Tomczak, M.F., Horwitz, B.H., Freeman, G.J., Sharpe, A.H., 2003. Regulation of PD-1, PD-L1, and PD-L2 expression during normal and autoimmune responses. Eur. J. Immunol. 33, 2706–2716. Lin, X., Chen, M., Liu, Y., Guo, Z., He, X., Brand, D., Zheng, S.G., 2013. Advances in distinguishing natural from induced Foxp3(+) regulatory T cells. Int. J. Clin. Exp. Pathol. 6, 116–123. Lindau, D., Mussard, J., Rabsteyn, A., Ribon, M., Kotter, I., Igney, A., Adema, G.J., Boissier, M.C., Rammensee, H.G., Decker, P., 2014. TLR9 independent interferon alpha production by neutrophils on NETosis in response to circulating chromatin, a key lupus autoantigen. Ann. Rheum. Dis. 73, 2199–2207. Linkermann, A., Stockwell, B.R., Krautwald, S., Anders, H.J., 2014. Regulated cell death and inflammation: an auto-amplification loop causes organ failure. Nat. Rev. Immunol. 14, 759–767. Lipinska, J., Lipinska, S., Kasielski, M., Smolewska, E., 2016. Anti-MCV and anti-CCP antibodies-diagnostic and prognostic value in children with juvenile idiopathic arthritis (JIA). Clin. Rheumatol. 35, 2699–2706.

136

S.K. Devarapu et al.

Liston, A., Gray, D.H., Lesage, S., Fletcher, A.L., Wilson, J., Webster, K.E., Scott, H.S., Boyd, R.L., Peltonen, L., Goodnow, C.C., 2004. Gene dosage—limiting role of Aire in thymic expression, clonal deletion, and organ-specific autoimmunity. J. Exp. Med. 200, 1015–1026. Liston, A., Lu, L.F., O’Carroll, D., Tarakhovsky, A., Rudensky, A.Y., 2008. Dicer-dependent microRNA pathway safeguards regulatory T cell function. J. Exp. Med. 205, 1993–2004. Liu, Y.C., 2004. Ubiquitin ligases and the immune response. Annu. Rev. Immunol. 22, 81–127. Liu, B., Tahk, S., Yee, K.M., Fan, G., Shuai, K., 2010. The ligase PIAS1 restricts natural regulatory T cell differentiation by epigenetic repression. Science 330, 521–525. Liu, S., Watanabe, S., Shudou, M., Kuno, M., Miura, H., Maeyama, K., 2014. Upregulation of store-operated Ca(2 +) entry in the naive CD4(+) T cells with aberrant cytokine releasing in active rheumatoid arthritis. Immunol. Cell Biol. 92, 752–760. Livingston, J.H., Crow, Y.J., 2016. Neurologic phenotypes associated with mutations in TREX1, RNASEH2A, RNASEH2B, RNASEH2C, SAMHD1, ADAR1, and IFIH1: Aicardi-Goutieres syndrome and beyond. Neuropediatrics 47, 355–360. Lo, W.L., Donermeyer, D.L., Allen, P.M., 2012. A voltage-gated sodium channel is essential for the positive selection of CD4(+) T cells. Nat. Immunol. 13, 880–887. Lo, I.C., Chan, H.C., Qi, Z., Ng, K.L., So, C., Tsang, S.Y., 2016. TRPV3 channel negatively regulates cell cycle progression and safeguards the pluripotency of embryonic stem cells. J. Cell. Physiol. 231, 403–413. Long, S.A., Buckner, J.H., 2011. CD4+FOXP3 + T regulatory cells in human autoimmunity: more than a numbers game. J. Immunol. 187, 2061–2066. Lood, C., Blanco, L.P., Purmalek, M.M., Carmona-Rivera, C., De Ravin, S.S., Smith, C.K., Malech, H.L., Ledbetter, J.A., Elkon, K.B., Kaplan, M.J., 2016. Neutrophil extracellular traps enriched in oxidized mitochondrial DNA are interferogenic and contribute to lupus-like disease. Nat. Med. 22, 146–153. Lorenz, G., Anders, H.J., 2015. Neutrophils, dendritic cells, toll-like receptors, and interferon-alpha in lupus nephritis. Semin. Nephrol. 35, 410–426. Lorenz, G., Darisipudi, M.N., Anders, H.J., 2014a. Canonical and non-canonical effects of the NLRP3 inflammasome in kidney inflammation and fibrosis. Nephrol. Dial. Transplant. 29, 41–48. Lorenz, G., Desai, J., Anders, H.J., 2014b. Lupus nephritis: update on mechanisms of systemic autoimmunity and kidney immunopathology. Curr. Opin. Nephrol. Hypertens. 23, 211–217. Lorenz G., Lech M. and Anders H.J., 2017. Toll-like receptor activation in the pathogenesis of lupus nephritis. Clin. Immunol. (in press). Lu, L.F., Thai, T.H., Calado, D.P., Chaudhry, A., Kubo, M., Tanaka, K., Loeb, G.B., Lee, H., Yoshimura, A., Rajewsky, K., Rudensky, A.Y., 2009a. Foxp3-dependent microRNA155 confers competitive fitness to regulatory T cells by targeting SOCS1 protein. Immunity 30, 80–91. Lu, T.X., Munitz, A., Rothenberg, M.E., 2009b. MicroRNA-21 is up-regulated in allergic airway inflammation and regulates IL-12p35 expression. J. Immunol. 182, 4994–5002. Luo, S., Liu, Y., Liang, G., Zhao, M., Wu, H., Liang, Y., Qiu, X., Tan, Y., Dai, Y., Yung, S., Chan, T.M., Lu, Q., 2015. The role of microRNA-1246 in the regulation of B cell activation and the pathogenesis of systemic lupus erythematosus. Clin. Epigenetics 7, 24. Lyn-Cook, B.D., Xie, C., Oates, J., Treadwell, E., Word, B., Hammons, G., Wiley, K., 2014. Increased expression of Toll-like receptors (TLRs) 7 and 9 and other cytokines in systemic lupus erythematosus (SLE) patients: ethnic differences and potential new targets for therapeutic drugs. Mol. Immunol. 61, 38–43.

Mechanisms of Autoimmunity

137

Ma, J., Xu, J., Madaio, M.P., Peng, Q., Zhang, J., Grewal, I.S., Flavell, R.A., Craft, J., 1996. Autoimmune lpr/lpr mice deficient in CD40 ligand: spontaneous Ig class switching with dichotomy of autoantibody responses. J. Immunol. 157, 417–426. MacDonald, K.P., Nishioka, Y., Lipsky, P.E., Thomas, R., 1997. Functional CD40 ligand is expressed by T cells in rheumatoid arthritis. J. Clin. Invest. 100, 2404–2414. Mackay, F., Schneider, P., Rennert, P., Browning, J., 2003. BAFF AND APRIL: a tutorial on B cell survival. Annu. Rev. Immunol. 21, 231–264. Mackern-Oberti, J.P., Llanos, C., Riedel, C.A., Bueno, S.M., Kalergis, A.M., 2015. Contribution of dendritic cells to the autoimmune pathology of systemic lupus erythematosus. Immunology 146, 497–507. Manku, H., Langefeld, C.D., Guerra, S.G., Malik, T.H., Alarcon-Riquelme, M., Anaya, J.M., Bae, S.C., Boackle, S.A., Brown, E.E., Criswell, L.A., Freedman, B.I., Gaffney, P.M., Gregersen, P.A., Guthridge, J.M., Han, S.H., Harley, J.B., Jacob, C.O., James, J.A., Kamen, D.L., Kaufman, K.M., Kelly, J.A., Martin, J., Merrill, J.T., Moser, K.L., Niewold, T.B., Park, S.Y., Pons-Estel, B.A., Sawalha, A.H., Scofield, R.H., Shen, N., Stevens, A.M., Sun, C., Gilkeson, G.S., Edberg, J.C., Kimberly, R.P., Nath, S.K., Tsao, B.P., Vyse, T.J., 2013. Trans-ancestral studies fine map the SLE-susceptibility locus TNFSF4. PLoS Genet. 9, e1003554. Marcais, A., Blevins, R., Graumann, J., Feytout, A., Dharmalingam, G., Carroll, T., Amado, I.F., Bruno, L., Lee, K., Walzer, T., Mann, M., Freitas, A.A., Boothby, M., Fisher, A.G., Merkenschlager, M., 2014. MicroRNA-mediated regulation of mTOR complex components facilitates discrimination between activation and anergy in CD4 T cells. J. Exp. Med. 211, 2281–2295. Marinkovic, T., Garin, A., Yokota, Y., Fu, Y.X., Ruddle, N.H., Furtado, G.C., Lira, S.A., 2006. Interaction of mature CD3+CD4 + T cells with dendritic cells triggers the development of tertiary lymphoid structures in the thyroid. J. Clin. Invest. 116, 2622–2632. Marshak-Rothstein, A., 2006. Toll-like receptors in systemic autoimmune disease. Nat. Rev. Immunol. 6, 823–835. Martinez, J., Cunha, L.D., Park, S., Yang, M., Lu, Q., Orchard, R., Li, Q.Z., Yan, M., Janke, L., Guy, C., Linkermann, A., Virgin, H.W., Green, D.R., 2016. Noncanonical autophagy inhibits the autoinflammatory, lupus-like response to dying cells. Nature 533, 115–119. Martinez-Godinez, M.A., Cruz-Dominguez, M.P., Jara, L.J., Dominguez-Lopez, A., Jarillo-Luna, R.A., Vera-Lastra, O., Montes-Cortes, D.H., Campos-Rodriguez, R., Lopez-Sanchez, D.M., Mejia-Barradas, C.M., Castelan-Chavez, E.E., Miliar-Garcia, A., 2015. Expression of NLRP3 inflammasome, cytokines and vascular mediators in the skin of systemic sclerosis patients. Isr. Med. Assoc. J. 17, 5–10. Martinon, F., Mayor, A., Tschopp, J., 2009. The inflammasomes: guardians of the body. Annu. Rev. Immunol. 27, 229–265. Maugeri, N., Rovere-Querini, P., Baldini, M., Baldissera, E., Sabbadini, M.G., Bianchi, M.E., Manfredi, A.A., 2014. Oxidative stress elicits platelet/leukocyte inflammatory interactions via HMGB1: a candidate for microvessel injury in systemic sclerosis. Antioxid. Redox Signal. 20, 1060–1074. Mayadas, T.N., Tsokos, G.C., Tsuboi, N., 2009. Mechanisms of immune complex-mediated neutrophil recruitment and tissue injury. Circulation 120, 2012–2024. Maynard, J., Petersson, K., Wilson, D.H., Adams, E.J., Blondelle, S.E., Boulanger, M.J., Wilson, D.B., Garcia, K.C., 2005. Structure of an autoimmune T cell receptor complexed with class II peptide-MHC: insights into MHC bias and antigen specificity. Immunity 22, 81–92.

138

S.K. Devarapu et al.

McCarty, N., Paust, S., Ikizawa, K., Dan, I., Li, X., Cantor, H., 2005. Signaling by the kinase MINK is essential in the negative selection of autoreactive thymocytes. Nat. Immunol. 6, 65–72. McClain, M.T., Heinlen, L.D., Dennis, G.J., Roebuck, J., Harley, J.B., James, J.A., 2005. Early events in lupus humoral autoimmunity suggest initiation through molecular mimicry. Nat. Med. 11, 85–89. McDonald, G., Deepak, S., Miguel, L., Hall, C.J., Isenberg, D.A., Magee, A.I., Butters, T., Jury, E.C., 2014. Normalizing glycosphingolipids restores function in CD4 + T cells from lupus patients. J. Clin. Invest. 124, 712–724. Medawar, P.B., 1948. Immunity to homologous grafted skin; the fate of skin homografts transplanted to the brain, to subcutaneous tissue, and to the anterior chamber of the eye. Br. J. Exp. Pathol. 29, 58–69. Menon, M., Blair, P.A., Isenberg, D.A., Mauri, C., 2016. A regulatory feedback between plasmacytoid dendritic cells and regulatory B cells is aberrant in systemic lupus erythematosus. Immunity 44, 683–697. Messinger, R.B., Naik, A.K., Jagodic, M.M., Nelson, M.T., Lee, W.Y., Choe, W.J., Orestes, P., Latham, J.R., Todorovic, S.M., Jevtovic-Todorovic, V., 2009. In vivo silencing of the Ca(V)3.2 T-type calcium channels in sensory neurons alleviates hyperalgesia in rats with streptozocin-induced diabetic neuropathy. Pain 145, 184–195. Metzker, M., Shipkova, M., von Ahsen, N., Andag, R., Abe, M., Canzler, O., Klett, C., Leicht, S., Olbricht, C., Wieland, E., 2016. Analytical evaluation of a real-time PCR-based DNA demethylation assay to assess the frequency of naturally occurring regulatory T cells in peripheral blood. Clin. Biochem. 49, 1173–1180. Meuwissen, M.E., Schot, R., Buta, S., Oudesluijs, G., Tinschert, S., Speer, S.D., Li, Z., van Unen, L., Heijsman, D., Goldmann, T., Lequin, M.H., Kros, J.M., Stam, W., Hermann, M., Willemsen, R., Brouwer, R.W., Van, I.W.F., Martin-Fernandez, M., de Coo, I., Dudink, J., de Vries, F.A., Bertoli Avella, A., Prinz, M., Crow, Y.J., Verheijen, F.W., Pellegrini, S., Bogunovic, D., Mancini, G.M., 2016. Human USP18 deficiency underlies type 1 interferonopathy leading to severe pseudo-TORCH syndrome. J. Exp. Med. 213, 1163–1174. Miao, F., Smith, D.D., Zhang, L., Min, A., Feng, W., Natarajan, R., 2008. Lymphocytes from patients with type 1 diabetes display a distinct profile of chromatin histone H3 lysine 9 dimethylation: an epigenetic study in diabetes. Diabetes 57, 3189–3198. Mihara, M., Tan, I., Chuzhin, Y., Reddy, B., Budhai, L., Holzer, A., Gu, Y., Davidson, A., 2000. CTLA4Ig inhibits T cell-dependent B-cell maturation in murine systemic lupus erythematosus. J. Clin. Invest. 106, 91–101. Mohiuddin, I.H., Pillai, V., Baughman, E.J., Greenberg, B.M., Frohman, E.M., Crawford, M.P., Sinha, S., Karandikar, N.J., 2016. Induction of regulatory T-cells from memory T-cells is perturbed during acute exacerbation of multiple sclerosis. Clin. Immunol. 166–167, 12–18. Monney, L., Sabatos, C.A., Gaglia, J.L., Ryu, A., Waldner, H., Chernova, T., Manning, S., Greenfield, E.A., Coyle, A.J., Sobel, R.A., Freeman, G.J., Kuchroo, V.K., 2002. Th1specific cell surface protein Tim-3 regulates macrophage activation and severity of an autoimmune disease. Nature 415, 536–541. Monteith, A.J., Kang, S., Scott, E., Hillman, K., Rajfur, Z., Jacobson, K., Costello, M.J., Vilen, B.J., 2016. Defects in lysosomal maturation facilitate the activation of innate sensors in systemic lupus erythematosus. Proc. Natl. Acad. Sci. U.S.A. 113, E2142–2151. Moreland, J.G., Davis, A.P., Bailey, G., Nauseef, W.M., Lamb, F.S., 2006. Anion channels, including ClC-3, are required for normal neutrophil oxidative function, phagocytosis, and transendothelial migration. J. Biol. Chem. 281, 12277–12288. Morelli, A.E., Larregina, A.T., Shufesky, W.J., Sullivan, M.L., Stolz, D.B., Papworth, G.D., Zahorchak, A.F., Logar, A.J., Wang, Z., Watkins, S.C., Falo Jr., L.D., Thomson, A.W.,

Mechanisms of Autoimmunity

139

2004. Endocytosis, intracellular sorting, and processing of exosomes by dendritic cells. Blood 104, 3257–3266. Moreno, C., Prieto, P., Macias, A., Pimentel-Santillana, M., de la Cruz, A., Traves, P.G., Bosca, L., Valenzuela, C., 2013. Modulation of voltage-dependent and inward rectifier potassium channels by 15-epi-lipoxin-A4 in activated murine macrophages: implications in innate immunity. J. Immunol. 191, 6136–6146. Moser, K., Muehlinghaus, G., Manz, R., Mei, H., Voigt, C., Yoshida, T., Dorner, T., Hiepe, F., Radbruch, A., 2006. Long-lived plasma cells in immunity and immunopathology. Immunol. Lett. 103, 83–85. Moulton, V.R., Tsokos, G.C., 2015. T cell signaling abnormalities contribute to aberrant immune cell function and autoimmunity. J. Clin. Invest. 125, 2220–2227. Moyron-Quiroz, J.E., Rangel-Moreno, J., Kusser, K., Hartson, L., Sprague, F., Goodrich, S., Woodland, D.L., Lund, F.E., Randall, T.D., 2004. Role of inducible bronchus associated lymphoid tissue (iBALT) in respiratory immunity. Nat. Med. 10, 927–934. Mozaffarian, N., Wiedeman, A.E., Stevens, A.M., 2008. Active systemic lupus erythematosus is associated with failure of antigen-presenting cells to express programmed death ligand-1. Rheumatology (Oxford) 47, 1335–1341. Mueller, D.L., Jenkins, M.K., Schwartz, R.H., 1989. Clonal expansion versus functional clonal inactivation: a costimulatory signalling pathway determines the outcome of T cell antigen receptor occupancy. Annu. Rev. Immunol. 7, 445–480. Mulay, S.R., Desai, J., Kumar, S.V., Eberhard, J.N., Thomasova, D., Romoli, S., Grigorescu, M., Kulkarni, O.P., Popper, B., Vielhauer, V., Zuchtriegel, G., Reichel, C., Brasen, J.H., Romagnani, P., Bilyy, R., Munoz, L.E., Herrmann, M., Liapis, H., Krautwald, S., Linkermann, A., Anders, H.J., 2016a. Cytotoxicity of crystals involves RIPK3-MLKL-mediated necroptosis. Nat. Commun. 7, 10274. Mulay, S.R., Holderied, A., Kumar, S.V., Anders, H.J., 2016b. Targeting Inflammation in so-called acute kidney injury. Semin. Nephrol. 36, 17–30. Mulay, S.R., Kumar, S.V., Lech, M., Desai, J., Anders, H.J., 2016c. How kidney cell death induces renal necroinflammation. Semin. Nephrol. 36, 162–173. Mulay, S.R., Linkermann, A., Anders, H.J., 2016d. Necroinflammation in kidney disease. J. Am. Soc. Nephrol. 27, 27–39. Muller, I., Klocke, A., Alex, M., Kotzsch, M., Luther, T., Morgenstern, E., Zieseniss, S., Zahler, S., Preissner, K., Engelmann, B., 2003. Intravascular tissue factor initiates coagulation via circulating microvesicles and platelets. FASEB J. 17, 476–478. Munoz, L.E., Lauber, K., Schiller, M., Manfredi, A.A., Herrmann, M., 2010. The role of defective clearance of apoptotic cells in systemic autoimmunity. Nat. Rev. Rheumatol. 6, 280–289. Murakami, M., 2011. Lipid mediators in life science. Exp. Anim. 60, 7–20. Murugaiyan, G., Beynon, V., Mittal, A., Joller, N., Weiner, H.L., 2011. Silencing microRNA-155 ameliorates experimental autoimmune encephalomyelitis. J. Immunol. 187, 2213–2221. Murugaiyan, G., da Cunha, A.P., Ajay, A.K., Joller, N., Garo, L.P., Kumaradevan, S., Yosef, N., Vaidya, V.S., Weiner, H.L., 2015. MicroRNA-21 promotes Th17 differentiation and mediates experimental autoimmune encephalomyelitis. J. Clin. Invest. 125, 1069–1080. Mycko, M.P., Cichalewska, M., Machlanska, A., Cwiklinska, H., Mariasiewicz, M., Selmaj, K.W., 2012. MicroRNA-301a regulation of a T-helper 17 immune response controls autoimmune demyelination. Proc. Natl. Acad. Sci. U.S.A. 109, E1248–1257. Nagamine, K., Peterson, P., Scott, H.S., Kudoh, J., Minoshima, S., Heino, M., Krohn, K.J., Lalioti, M.D., Mullis, P.E., Antonarakis, S.E., Kawasaki, K., Asakawa, S., Ito, F., Shimizu, N., 1997. Positional cloning of the APECED gene. Nat. Genet. 17, 393–398.

140

S.K. Devarapu et al.

Nakahama, T., Hanieh, H., Nguyen, N.T., Chinen, I., Ripley, B., Millrine, D., Lee, S., Nyati, K.K., Dubey, P.K., Chowdhury, K., Kawahara, Y., Kishimoto, T., 2013. Aryl hydrocarbon receptor-mediated induction of the microRNA-132/212 cluster promotes interleukin-17-producing T-helper cell differentiation. Proc. Natl. Acad. Sci. U.S.A. 110, 11964–11969. Nakamachi, Y., Kawano, S., Takenokuchi, M., Nishimura, K., Sakai, Y., Chin, T., Saura, R., Kurosaka, M., Kumagai, S., 2009. MicroRNA-124a is a key regulator of proliferation and monocyte chemoattractant protein 1 secretion in fibroblast-like synoviocytes from patients with rheumatoid arthritis. Arthritis Rheum. 60, 1294–1304. Naramura, M., Jang, I.K., Kole, H., Huang, F., Haines, D., Gu, H., 2002. c-Cbl and Cbl-b regulate T cell responsiveness by promoting ligand-induced TCR down-modulation. Nat. Immunol. 3, 1192–1199. Nasr, A.S., Fawzy, S.M., Gheita, T.A., El-Khateeb, E., 2015. Expression of Toll-like receptors 3 and 9 in Egyptian systemic lupus erythematosus patients. Z. Rheumatol. 75, 502–507. Ndhlovu, L.C., Ishii, N., Murata, K., Sato, T., Sugamura, K., 2001. Critical involvement of OX40 ligand signals in the T cell priming events during experimental autoimmune encephalomyelitis. J. Immunol. 167, 2991–2999. Nemazee, D., Hogquist, K.A., 2003. Antigen receptor selection by editing or downregulation of V(D)J recombination. Curr. Opin. Immunol. 15, 182–189. Neyt, K., Perros, F., GeurtsvanKessel, C.H., Hammad, H., Lambrecht, B.N., 2012. Tertiary lymphoid organs in infection and autoimmunity. Trends Immunol. 33, 297–305. Nickerson, K.M., Christensen, S.R., Shupe, J., Kashgarian, M., Kim, D., Elkon, K., Shlomchik, M.J., 2010. TLR9 regulates TLR7- and MyD88-dependent autoantibody production and disease in a murine model of lupus. J. Immunol. 184, 1840–1848. Niederkorn, J.Y., 2012. Ocular immune privilege and ocular melanoma: parallel universes or immunological plagiarism? Front. Immunol. 3, 148. Nielsen, C.T., Ostergaard, O., Stener, L., Iversen, L.V., Truedsson, L., Gullstrand, B., Jacobsen, S., Heegaard, N.H., 2012. Increased IgG on cell-derived plasma microparticles in systemic lupus erythematosus is associated with autoantibodies and complement activation. Arthritis Rheum. 64, 1227–1236. Niewold, T.B., Kelly, J.A., Flesch, M.H., Espinoza, L.R., Harley, J.B., Crow, M.K., 2008. Association of the IRF5 risk haplotype with high serum interferon-alpha activity in systemic lupus erythematosus patients. Arthritis Rheum. 58, 2481–2487. Nimmerjahn, F., Ravetch, J.V., 2008. Fcgamma receptors as regulators of immune responses. Nat. Rev. Immunol. 8, 34–47. Nishimura, H., Minato, N., Nakano, T., Honjo, T., 1998. Immunological studies on PD-1 deficient mice: implication of PD-1 as a negative regulator for B cell responses. Int. Immunol. 10, 1563–1572. Nishimura, H., Nose, M., Hiai, H., Minato, N., Honjo, T., 1999. Development of lupus-like autoimmune diseases by disruption of the PD-1 gene encoding an ITIM motif-carrying immunoreceptor. Immunity 11, 141–151. Nohara, C., Akiba, H., Nakajima, A., Inoue, A., Koh, C.S., Ohshima, H., Yagita, H., Mizuno, Y., Okumura, K., 2001. Amelioration of experimental autoimmune encephalomyelitis with anti-OX40 ligand monoclonal antibody: a critical role for OX40 ligand in migration, but not development, of pathogenic T cells. J. Immunol. 166, 2108–2115. Nossal, G.J., Pike, B.L., 1980. Clonal anergy: persistence in tolerant mice of antigen-binding B lymphocytes incapable of responding to antigen or mitogen. Proc. Natl. Acad. Sci. U.S.A. 77, 1602–1606. Nurieva, R.I., Liu, X., Dong, C., 2011. Molecular mechanisms of T-cell tolerance. Immunol. Rev. 241, 133–144.

Mechanisms of Autoimmunity

141

Oda, H., Nakagawa, K., Abe, J., Awaya, T., Funabiki, M., Hijikata, A., Nishikomori, R., Funatsuka, M., Ohshima, Y., Sugawara, Y., Yasumi, T., Kato, H., Shirai, T., Ohara, O., Fujita, T., Heike, T., 2014. Aicardi-Goutieres syndrome is caused by IFIH1 mutations. Am. J. Hum. Genet. 95, 121–125. Odegard, J.M., Marks, B.R., DiPlacido, L.D., Poholek, A.C., Kono, D.H., Dong, C., Flavell, R.A., Craft, J., 2008. ICOS-dependent extrafollicular helper T cells elicit IgG production via IL-21 in systemic autoimmunity. J. Exp. Med. 205, 2873–2886. Ogura, Y., Bonen, D.K., Inohara, N., Nicolae, D.L., Chen, F.F., Ramos, R., Britton, H., Moran, T., Karaliuskas, R., Duerr, R.H., Achkar, J.P., Brant, S.R., Bayless, T.M., Kirschner, B.S., Hanauer, S.B., Nunez, G., Cho, J.H., 2001. A frameshift mutation in NOD2 associated with susceptibility to Crohn’s disease. Nature 411, 603–606. Oh-Hora, M., Komatsu, N., Pishyareh, M., Feske, S., Hori, S., Taniguchi, M., Rao, A., Takayanagi, H., 2013. Agonist-selected T cell development requires strong T cell receptor signaling and store-operated calcium entry. Immunity 38, 881–895. Okazaki, T., Honjo, T., 2007. PD-1 and PD-1 ligands: from discovery to clinical application. Int. Immunol. 19, 813–824. Olenchock, B.A., Guo, R., Carpenter, J.H., Jordan, M., Topham, M.K., Koretzky, G.A., Zhong, X.P., 2006. Disruption of diacylglycerol metabolism impairs the induction of T cell anergy. Nat. Immunol. 7, 1174–1181. Oliveira-dos-Santos, A.J., Ho, A., Tada, Y., Lafaille, J.J., Tonegawa, S., Mak, T.W., Penninger, J.M., 1999. CD28 costimulation is crucial for the development of spontaneous autoimmune encephalomyelitis. J. Immunol. 162, 4490–4495. O’Neill, S.K., Cao, Y., Hamel, K.M., Doodes, P.D., Hutas, G., Finnegan, A., 2007. Expression of CD80/86 on B cells is essential for autoreactive T cell activation and the development of arthritis. J. Immunol. 179, 5109–5116. O’Reilly, S., Cant, R., Ciechomska, M., Finnigan, J., Oakley, F., Hambleton, S., van Laar, J.M., 2014. Serum amyloid A induces interleukin-6 in dermal fibroblasts via toll-like receptor 2, interleukin-1 receptor-associated kinase 4 and nuclear factor-kappaB. Immunology 143, 331–340. Org, T., Chignola, F., Hetenyi, C., Gaetani, M., Rebane, A., Liiv, I., Maran, U., Mollica, L., Bottomley, M.J., Musco, G., Peterson, P., 2008. The autoimmune regulator PHD finger binds to non-methylated histone H3K4 to activate gene expression. EMBO Rep. 9, 370–376. Pakala, S.V., Bansal-Pakala, P., Halteman, B.S., Croft, M., 2004. Prevention of diabetes in NOD mice at a late stage by targeting OX40/OX40 ligand interactions. Eur. J. Immunol. 34, 3039–3046. Palmer, E., 2003. Negative selection—clearing out the bad apples from the T-cell repertoire. Nat. Rev. Immunol. 3, 383–391. Paraboschi, E.M., Solda, G., Gemmati, D., Orioli, E., Zeri, G., Benedetti, M.D., Salviati, A., Barizzone, N., Leone, M., Duga, S., Asselta, R., 2011. Genetic association and altered gene expression of mir-155 in multiple sclerosis patients. Int. J. Mol. Sci. 12, 8695–8712. Parackova, Z., Kayserova, J., Danova, K., Sismova, K., Dudkova, E., Sumnik, Z., Kolouskova, S., Lebl, J., Stechova, K., Sediva, A., 2016. T regulatory lymphocytes in type 1 diabetes: impaired CD25 expression and IL-2 induced STAT5 phosphorylation in pediatric patients. Autoimmunity 49, 523–531. Park, J.S., Svetkauskaite, D., He, Q., Kim, J.Y., Strassheim, D., Ishizaka, A., Abraham, E., 2004. Involvement of toll-like receptors 2 and 4 in cellular activation by high mobility group box 1 protein. J. Biol. Chem. 279, 7370–7377. Patole, P.S., Zecher, D., Pawar, R.D., Grone, H.J., Schlondorff, D., Anders, H.J., 2005. G-rich DNA suppresses systemic lupus. J. Am. Soc. Nephrol. 16, 3273–3280.

142

S.K. Devarapu et al.

Pawar, R.D., Ramanjaneyulu, A., Kulkarni, O.P., Lech, M., Segerer, S., Anders, H.J., 2007. Inhibition of toll-like receptor-7 (TLR-7) or TLR-7 plus TLR-9 attenuates glomerulonephritis and lung injury in experimental lupus. J. Am. Soc. Nephrol. 18, 1721–1731. Peng, S.L., 2012. Experimental use of mouse models of systemic lupus erythematosus. Methods Mol. Biol. 900, 135–168. Pepper, R.J., Hamour, S., Chavele, K.M., Todd, S.K., Rasmussen, N., Flint, S., Lyons, P.A., Smith, K.G., Pusey, C.D., Cook, H.T., Salama, A.D., 2013. Leukocyte and serum S100A8/S100A9 expression reflects disease activity in ANCA-associated vasculitis and glomerulonephritis. Kidney Int. 83, 1150–1158. Perera, J., Zheng, Z., Li, S., Gudjonson, H., Kalinina, O., Benichou, J.I., Block, K.E., Louzoun, Y., Yin, D., Chong, A.S., Dinner, A.R., Weigert, M., Huang, H., 2016. Self-antigen-driven thymic B cell class switching promotes T cell central tolerance. Cell Rep. 17, 387–398. Peterson, P., Org, T., Rebane, A., 2008. Transcriptional regulation by AIRE: molecular mechanisms of central tolerance. Nat. Rev. Immunol. 8, 948–957. Pierer, M., Schulz, A., Rossol, M., Kendzia, E., Kyburz, D., Haentzschel, H., Baerwald, C., Wagner, U., 2009. Herpesvirus entry mediator-Ig treatment during immunization aggravates rheumatoid arthritis in the collagen-induced arthritis model. J. Immunol. 182, 3139–3145. Pillozzi, S., Becchetti, A., 2012. Ion channels in hematopoietic and mesenchymal stem cells. Stem Cells Int. 2012, 217910. Pisetsky, D.S., Lipsky, P.E., 2010. Microparticles as autoadjuvants in the pathogenesis of SLE. Nat. Rev. Rheumatol. 6, 368–372. Pisetsky, D.S., Gauley, J., Ullal, A.J., 2011. Microparticles as a source of extracellular DNA. Immunol. Res. 49, 227–234. Pitkanen, J., Doucas, V., Sternsdorf, T., Nakajima, T., Aratani, S., Jensen, K., Will, H., Vahamurto, P., Ollila, J., Vihinen, M., Scott, H.S., Antonarakis, S.E., Kudoh, J., Shimizu, N., Krohn, K., Peterson, P., 2000. The autoimmune regulator protein has transcriptional transactivating properties and interacts with the common coactivator CREB-binding protein. J. Biol. Chem. 275, 16802–16809. Pizzirani, C., Ferrari, D., Chiozzi, P., Adinolfi, E., Sandona, D., Savaglio, E., Di Virgilio, F., 2007. Stimulation of P2 receptors causes release of IL-1beta-loaded microvesicles from human dendritic cells. Blood 109, 3856–3864. Podojil, J.R., Liu, L.N., Marshall, S.A., Chiang, M.Y., Goings, G.E., Chen, L., Langermann, S., Miller, S.D., 2013. B7-H4Ig inhibits mouse and human T-cell function and treats EAE via IL-10/Treg-dependent mechanisms. J. Autoimmun. 44, 71–81. Polhill, T., Zhang, G.Y., Hu, M., Sawyer, A., Zhou, J.J., Saito, M., Webster, K.E., Wang, Y., Wang, Y., Grey, S.T., Sprent, J., Harris, D.C., Alexander, S.I., Wang, Y.M., 2012. IL-2/IL-2Ab complexes induce regulatory T cell expansion and protect against proteinuric CKD. J. Am. Soc. Nephrol. 23, 1303–1308. Ponomarev, E.D., Veremeyko, T., Barteneva, N., Krichevsky, A.M., Weiner, H.L., 2011. MicroRNA-124 promotes microglia quiescence and suppresses EAE by deactivating macrophages via the C/EBP-alpha-PU.1 pathway. Nat. Med. 17, 64–70. Prasad, D.V., Richards, S., Mai, X.M., Dong, C., 2003. B7S1, a novel B7 family member that negatively regulates T cell activation. Immunity 18, 863–873. Prochnicki, T., Mangan, M.S., Latz, E., 2016. Recent insights into the molecular mechanisms of the NLRP3 inflammasome activation. F1000Res 5, 1469. Pruchniak, M.P., Kotula, I., Manda-Handzlik, A., 2015. Neutrophil extracellular traps (Nets) impact upon autoimmune disorders. Cent. Eur. J. Immunol. 40, 217–224.

Mechanisms of Autoimmunity

143

Quattrocchi, E., Dallman, M.J., Feldmann, M., 2000. Adenovirus-mediated gene transfer of CTLA-4Ig fusion protein in the suppression of experimental autoimmune arthritis. Arthritis Rheum. 43, 1688–1697. Rahgozar, K., Wright, E., Carrithers, L.M., Carrithers, M.D., 2013. Mediation of protection and recovery from experimental autoimmune encephalomyelitis by macrophages expressing the human voltage-gated sodium channel NaV1.5. J. Neuropathol. Exp. Neurol. 72, 489–504. RamaKrishnan, A.M., Sankaranarayanan, K., 2016. Understanding autoimmunity: the ion channel perspective. Autoimmun. Rev. 15, 585–620. Ramantani, G., Hausler, M., Niggemann, P., Wessling, B., Guttmann, H., Mull, M., Tenbrock, K., Lee-Kirsch, M.A., 2011. Aicardi-Goutieres syndrome and systemic lupus erythematosus (SLE) in a 12-year-old boy with SAMHD1 mutations. J. Child Neurol. 26, 1425–1428. Ramirez-Ortiz, Z.G., Pendergraft 3rd, W.F., Prasad, A., Byrne, M.H., Iram, T., Blanchette, C.J., Luster, A.D., Hacohen, N., El Khoury, J., Means, T.K., 2013. The scavenger receptor SCARF1 mediates the clearance of apoptotic cells and prevents autoimmunity. Nat. Immunol. 14, 917–926. Ramos, P.S., Shedlock, A.M., Langefeld, C.D., 2015. Genetics of autoimmune diseases: insights from population genetics. J. Hum. Genet. 60, 657–664. Rangel-Moreno, J., Hartson, L., Navarro, C., Gaxiola, M., Selman, M., Randall, T.D., 2006. Inducible bronchus-associated lymphoid tissue (iBALT) in patients with pulmonary complications of rheumatoid arthritis. J. Clin. Invest. 116, 3183–3194. Rangel-Moreno, J., Moyron-Quiroz, J.E., Hartson, L., Kusser, K., Randall, T.D., 2007. Pulmonary expression of CXC chemokine ligand 13, CC chemokine ligand 19, and CC chemokine ligand 21 is essential for local immunity to influenza. Proc. Natl. Acad. Sci. U.S.A. 104, 10577–10582. Raposo, G., Stoorvogel, W., 2013. Extracellular vesicles: exosomes, microvesicles, and friends. J. Cell Biol. 200, 373–383. Raptopoulou, A.P., Bertsias, G., Makrygiannakis, D., Verginis, P., Kritikos, I., Tzardi, M., Klareskog, L., Catrina, A.I., Sidiropoulos, P., Boumpas, D.T., 2010. The programmed death 1/programmed death ligand 1 inhibitory pathway is up-regulated in rheumatoid synovium and regulates peripheral T cell responses in human and murine arthritis. Arthritis Rheum. 62, 1870–1880. Ravenscroft, J.C., Suri, M., Rice, G.I., Szynkiewicz, M., Crow, Y.J., 2011. Autosomal dominant inheritance of a heterozygous mutation in SAMHD1 causing familial chilblain lupus. Am. J. Med. Genet. A 155A, 235–237. Ravetch, J.V., Lanier, L.L., 2000. Immune inhibitory receptors. Science 290, 84–89. Ray, S.K., Putterman, C., Diamond, B., 1996. Pathogenic autoantibodies are routinely generated during the response to foreign antigen: a paradigm for autoimmune disease. Proc. Natl. Acad. Sci. U.S.A. 93, 2019–2024. Reale, M., Sanchez-Ramon, S., 2016. Lipids at the cross-road of autoimmunity in multiple sclerosis. Curr. Med. Chem. [Epub ahead of print]. Reap, E.A., Leslie, D., Abrahams, M., Eisenberg, R.A., Cohen, P.L., 1995. Apoptosis abnormalities of splenic lymphocytes in autoimmune lpr and gld mice. J. Immunol. 154, 936–943. Rice, G., Newman, W.G., Dean, J., Patrick, T., Parmar, R., Flintoff, K., Robins, P., Harvey, S., Hollis, T., O’Hara, A., Herrick, A.L., Bowden, A.P., Perrino, F.W., Lindahl, T., Barnes, D.E., Crow, Y.J., 2007. Heterozygous mutations in TREX1 cause familial chilblain lupus and dominant Aicardi-Goutieres syndrome. Am. J. Hum. Genet. 80, 811–815.

144

S.K. Devarapu et al.

Rioux, J.D., Xavier, R.J., Taylor, K.D., Silverberg, M.S., Goyette, P., Huett, A., Green, T., Kuballa, P., Barmada, M.M., Datta, L.W., Shugart, Y.Y., Griffiths, A.M., Targan, S.R., Ippoliti, A.F., Bernard, E.J., Mei, L., Nicolae, D.L., Regueiro, M., Schumm, L.P., Steinhart, A.H., Rotter, J.I., Duerr, R.H., Cho, J.H., Daly, M.J., Brant, S.R., 2007. Genome-wide association study identifies new susceptibility loci for Crohn disease and implicates autophagy in disease pathogenesis. Nat. Genet. 39, 596–604. Roberts, C.A., Dickinson, A.K., Taams, L.S., 2015. The interplay between monocytes/macrophages and CD4(+) T cell subsets in rheumatoid arthritis. Front. Immunol. 6, 571. Ronchese, F., Hausmann, B., 1993. B lymphocytes in vivo fail to prime naive T cells but can stimulate antigen-experienced T lymphocytes. J. Exp. Med. 177, 679–690. Rosenzweig, S.D., 2008. Inflammatory manifestations in chronic granulomatous disease (CGD). J. Clin. Immunol. 28 (Suppl. 1), S67–72. Rothbart, S.B., Strahl, B.D., 2014. Interpreting the language of histone and DNA modifications. Biochim. Biophys. Acta 1839, 627–643. Ruan, Q.G., Tung, K., Eisenman, D., Setiady, Y., Eckenrode, S., Yi, B., Purohit, S., Zheng, W.P., Zhang, Y., Peltonen, L., She, J.X., 2007. The autoimmune regulator directly controls the expression of genes critical for thymic epithelial function. J. Immunol. 178, 7173–7180. Rubin, R.L., 2005. Drug-induced lupus. Toxicology 209, 135–147. Rutsch, F., MacDougall, M., Lu, C., Buers, I., Mamaeva, O., Nitschke, Y., Rice, G.I., Erlandsen, H., Kehl, H.G., Thiele, H., Nurnberg, P., Hohne, W., Crow, Y.J., Feigenbaum, A., Hennekam, R.C., 2015. A specific IFIH1 gain-of-function mutation causes Singleton-Merten syndrome. Am. J. Hum. Genet. 96, 275–282. Sabatos, C.A., Chakravarti, S., Cha, E., Schubart, A., Sanchez-Fueyo, A., Zheng, X.X., Coyle, A.J., Strom, T.B., Freeman, G.J., Kuchroo, V.K., 2003. Interaction of Tim-3 and Tim-3 ligand regulates T helper type 1 responses and induction of peripheral tolerance. Nat. Immunol. 4, 1102–1110. Saenz-Cuesta, M., Osorio-Querejeta, I., Otaegui, D., 2014. Extracellular vesicles in multiple sclerosis: what are they telling us? Front. Cell. Neurosci. 8, 100. Sahu, R., Bethunaickan, R., Singh, S., Davidson, A., 2014. Structure and function of renal macrophages and dendritic cells from lupus-prone mice. Arthritis Rheumatol. 66, 1596–1607. Sakaguchi, N., Takahashi, T., Hata, H., Nomura, T., Tagami, T., Yamazaki, S., Sakihama, T., Matsutani, T., Negishi, I., Nakatsuru, S., Sakaguchi, S., 2003. Altered thymic T-cell selection due to a mutation of the ZAP-70 gene causes autoimmune arthritis in mice. Nature 426, 454–460. Sakaguchi, S., Yamaguchi, T., Nomura, T., Ono, M., 2008. Regulatory T cells and immune tolerance. Cell 133, 775–787. Sakkas, L.I., Bogdanos, D.P., 2016. Systemic sclerosis: new evidence re-enforces the role of B cells. Autoimmun. Rev. 15, 155–161. Sakkas, L.I., Bogdanos, D.P., Katsiari, C., Platsoucas, C.D., 2014. Anti-citrullinated peptides as autoantigens in rheumatoid arthritis-relevance to treatment. Autoimmun. Rev. 13, 1114–1120. Salmon, M.D., Ahluwalia, J., 2009. Swell activated chloride channel function in human neutrophils. Biochem. Biophys. Res. Commun. 381, 462–465. Salomon, B., Lenschow, D.J., Rhee, L., Ashourian, N., Singh, B., Sharpe, A., Bluestone, J.A., 2000. B7/CD28 costimulation is essential for the homeostasis of the CD4+CD25+ immunoregulatory T cells that control autoimmune diabetes. Immunity 12, 431–440. Salzer, E., Santos-Valente, E., Klaver, S., Ban, S.A., Emminger, W., Prengemann, N.K., Garncarz, W., Mullauer, L., Kain, R., Boztug, H., Heitger, A., Arbeiter, K., Eitelberger, F., Seidel, M.G., Holter, W., Pollak, A., Pickl, W.F., Forster-Waldl, E.,

Mechanisms of Autoimmunity

145

Boztug, K., 2013. B-cell deficiency and severe autoimmunity caused by deficiency of protein kinase C delta. Blood 121, 3112–3116. Sanchez-Fueyo, A., Tian, J., Picarella, D., Domenig, C., Zheng, X.X., Sabatos, C.A., Manlongat, N., Bender, O., Kamradt, T., Kuchroo, V.K., Gutierrez-Ramos, J.C., Coyle, A.J., Strom, T.B., 2003. Tim-3 inhibits T helper type 1-mediated auto- and alloimmune responses and promotes immunological tolerance. Nat. Immunol. 4, 1093–1101. Sangaletti, S., Tripodo, C., Chiodoni, C., Guarnotta, C., Cappetti, B., Casalini, P., Piconese, S., Parenza, M., Guiducci, C., Vitali, C., Colombo, M.P., 2012. Neutrophil extracellular traps mediate transfer of cytoplasmic neutrophil antigens to myeloid dendritic cells toward ANCA induction and associated autoimmunity. Blood 120, 3007–3018. Santana-de Anda, K., Gomez-Martin, D., Monsivais-Urenda, A.E., Salgado-Bustamante, M., Gonzalez-Amaro, R., Alcocer-Varela, J., 2014. Interferon regulatory factor 3 as key element of the interferon signature in plasmacytoid dendritic cells from systemic lupus erythematosus patients: novel genetic associations in the Mexican mestizo population. Clin. Exp. Immunol. 178, 428–437. Santegoets, K.C., van Bon, L., van den Berg, W.B., Wenink, M.H., Radstake, T.R., 2011. Toll-like receptors in rheumatic diseases: are we paying a high price for our defense against bugs? FEBS Lett. 585, 3660–3666. Santoni, G., Farfariello, V., Liberati, S., Morelli, M.B., Nabissi, M., Santoni, M., Amantini, C., 2013. The role of transient receptor potential vanilloid type-2 ion channels in innate and adaptive immune responses. Front. Immunol. 4, 34. Savarese, E., Chae, O.W., Trowitzsch, S., Weber, G., Kastner, B., Akira, S., Wagner, H., Schmid, R.M., Bauer, S., Krug, A., 2006. U1 small nuclear ribonucleoprotein immune complexes induce type I interferon in plasmacytoid dendritic cells through TLR7. Blood 107, 3229–3234. Savarese, E., Steinberg, C., Pawar, R.D., Reindl, W., Akira, S., Anders, H.J., Krug, A., 2008. Requirement of toll-like receptor 7 for pristane-induced production of autoantibodies and development of murine lupus nephritis. Arthritis Rheum. 58, 1107–1115. Schaffert, S.A., Loh, C., Wang, S., Arnold, C.P., Axtell, R.C., Newell, E.W., Nolan, G., Ansel, K.M., Davis, M.M., Steinman, L., Chen, C.Z., 2015. mir-181a-1/b-1 modulates tolerance through opposing activities in selection and peripheral T cell function. J. Immunol. 195, 1470–1479. Scharenberg, A.M., Humphries, L.A., Rawlings, D.J., 2007. Calcium signalling and cell-fate choice in B cells. Nat. Rev. Immunol. 7, 778–789. Schuetz, C., Niehues, T., Friedrich, W., Schwarz, K., 2010. Autoimmunity, autoinflammation and lymphoma in combined immunodeficiency (CID). Autoimmun. Rev. 9, 477–482. Seki, M., Oomizu, S., Sakata, K.M., Sakata, A., Arikawa, T., Watanabe, K., Ito, K., Takeshita, K., Niki, T., Saita, N., Nishi, N., Yamauchi, A., Katoh, S., Matsukawa, A., Kuchroo, V., Hirashima, M., 2008. Galectin-9 suppresses the generation of Th17, promotes the induction of regulatory T cells, and regulates experimental autoimmune arthritis. Clin. Immunol. 127, 78–88. Selvaraj, P., Fifadara, N., Nagarajan, S., Cimino, A., Wang, G., 2004. Functional regulation of human neutrophil Fc gamma receptors. Immunol. Res. 29, 219–230. Seredkina, N., Rekvig, O.P., 2011. Acquired loss of renal nuclease activity is restricted to DNaseI and is an organ-selective feature in murine lupus nephritis. Am. J. Pathol. 179, 1120–1128. Sester, D.P., Sagulenko, V., Thygesen, S.J., Cridland, J.A., Loi, Y.S., Cridland, S.O., Masters, S.L., Genske, U., Hornung, V., Andoniou, C.E., Sweet, M.J.,

146

S.K. Devarapu et al.

Degli-Esposti, M.A., Schroder, K., Stacey, K.J., 2015. Deficient NLRP3 and AIM2 inflammasome function in autoimmune NZB mice. J. Immunol. 195, 1233–1241. Shao, Z., Gaurav, R., Agrawal, D.K., 2015. Intermediate-conductance calcium-activated potassium channel KCa3.1 and chloride channel modulate chemokine ligand (CCL19/CCL21)-induced migration of dendritic cells. Transl. Res. 166, 89–102. Shen, N., Fu, Q., Deng, Y., Qian, X., Zhao, J., Kaufman, K.M., Wu, Y.L., Yu, C.Y., Tang, Y., Chen, J.Y., Yang, W., Wong, M., Kawasaki, A., Tsuchiya, N., Sumida, T., Kawaguchi, Y., Howe, H.S., Mok, M.Y., Bang, S.Y., Liu, F.L., Chang, D.M., Takasaki, Y., Hashimoto, H., Harley, J.B., Guthridge, J.M., Grossman, J.M., Cantor, R.M., Song, Y.W., Bae, S.C., Chen, S., Hahn, B.H., Lau, Y.L., Tsao, B.P., 2010. Sex-specific association of X-linked Toll-like receptor 7 (TLR7) with male systemic lupus erythematosus. Proc. Natl. Acad. Sci. U.S.A. 107, 15838–15843. Shen, L., Gao, C., Suresh, L., Xian, Z., Song, N., Chaves, L.D., Yu, M., Ambrus Jr., J.L., 2016. Central role for marginal zone B cells in an animal model of Sjogren’s syndrome. Clin. Immunol. 168, 30–36. Shi, C., Liang, Y., Yang, J., Xia, Y., Chen, H., Han, H., Yang, Y., Wu, W., Gao, R., Qin, H., 2013. MicroRNA-21 knockout improve the survival rate in DSS induced fatal colitis through protecting against inflammation and tissue injury. PLoS One 8, e66814. Shin, H.D., Park, B.L., Kim, L.H., Lee, H.S., Kim, T.Y., Bae, S.C., 2004. Common DNase I polymorphism associated with autoantibody production among systemic lupus erythematosus patients. Hum. Mol. Genet. 13, 2343–2350. Shin, M.S., Kang, Y., Lee, N., Kim, S.H., Kang, K.S., Lazova, R., Kang, I., 2012. U1-small nuclear ribonucleoprotein activates the NLRP3 inflammasome in human monocytes. J. Immunol. 188, 4769–4775. Shivakumar, S., Tsokos, G.C., Datta, S.K., 1989. T cell receptor alpha/beta expressing double-negative (CD4-/CD8-) and CD4 + T helper cells in humans augment the production of pathogenic anti-DNA autoantibodies associated with lupus nephritis. J. Immunol. 143, 103–112. Simchowitz, L., Textor, J.A., Cragoe Jr., E.J., 1993. Cell volume regulation in human neutrophils: 2-(aminomethyl)phenols as Cl- channel inhibitors. Am. J. Physiol. 265, C143–155. Singh, S.B., Davis, A.S., Taylor, G.A., Deretic, V., 2006. Human IRGM induces autophagy to eliminate intracellular mycobacteria. Science 313, 1438–1441. Sisirak, V., Sally, B., D’Agati, V., Martinez-Ortiz, W., Ozcakar, Z.B., David, J., Rashidfarrokhi, A., Yeste, A., Panea, C., Chida, A.S., Bogunovic, M., Ivanov, I.I., Quintana, F.J., Sanz, I., Elkon, K.B., Tekin, M., Yalcinkaya, F., Cardozo, T.J., Clancy, R.M., Buyon, J.P., Reizis, B., 2016. Digestion of chromatin in apoptotic cell microparticles prevents autoimmunity. Cell 166, 88–101. Smeets, B., Angelotti, M.L., Rizzo, P., Dijkman, H., Lazzeri, E., Mooren, F., Ballerini, L., Parente, E., Sagrinati, C., Mazzinghi, B., Ronconi, E., Becherucci, F., Benigni, A., Steenbergen, E., Lasagni, L., Remuzzi, G., Wetzels, J., Romagnani, P., 2009. Renal progenitor cells contribute to hyperplastic lesions of podocytopathies and crescentic glomerulonephritis. J. Am. Soc. Nephrol. 20, 2593–2603. Soderberg, D., Segelmark, M., 2016. Neutrophil extracellular traps in ANCA-associated vasculitis. Front. Immunol. 7, 256. Solomos, A.C., Rall, G.F., 2016. Get It through your thick head: emerging principles in neuroimmunology and neurovirology redefine central nervous system “immune privilege”. ACS Chem. Neurosci. 7, 435–441. Sorice, M., Iannuccelli, C., Manganelli, V., Capozzi, A., Alessandri, C., Lococo, E., Garofalo, T., Di Franco, M., Bombardieri, M., Nerviani, A., Misasi, R., Valesini, G.,

Mechanisms of Autoimmunity

147

2016. Autophagy generates citrullinated peptides in human synoviocytes: a possible trigger for anti-citrullinated peptide antibodies. Rheumatology (Oxford) 55, 1374–1385. Spengler, J., Lugonja, B., Ytterberg, A.J., Zubarev, R.A., Creese, A.J., Pearson, M.J., Grant, M.M., Milward, M., Lundberg, K., Buckley, C.D., Filer, A., Raza, K., Cooper, P.R., Chapple, I.L., Scheel-Toellner, D., 2015. Release of active peptidyl arginine deiminases by neutrophils can explain production of extracellular citrullinated autoantigens in rheumatoid arthritis synovial fluid. Arthritis Rheumatol. 67, 3135–3145. Spitaler, M., Emslie, E., Wood, C.D., Cantrell, D., 2006. Diacylglycerol and protein kinase D localization during T lymphocyte activation. Immunity 24, 535–546. Sporici, R.A., Beswick, R.L., von Allmen, C., Rumbley, C.A., Hayden-Ledbetter, M., Ledbetter, J.A., Perrin, P.J., 2001. ICOS ligand costimulation is required for T-cell encephalitogenicity. Clin. Immunol. 100, 277–288. Srivastava, R., Aslam, M., Kalluri, S.R., Schirmer, L., Buck, D., Tackenberg, B., Rothhammer, V., Chan, A., Gold, R., Berthele, A., Bennett, J.L., Korn, T., Hemmer, B., 2012. Potassium channel KIR4.1 as an immune target in multiple sclerosis. N. Engl. J. Med. 367, 115–123. Stamatiades, E.G., Tremblay, M.E., Bohm, M., Crozet, L., Bisht, K., Kao, D., Coelho, C., Fan, X., Yewdell, W.T., Davidson, A., Heeger, P.S., Diebold, S., Nimmerjahn, F., Geissmann, F., 2016. Immune monitoring of trans-endothelial transport by kidney-resident macrophages. Cell 166, 991–1003. Starokadomskyy, P., Gemelli, T., Rios, J.J., Xing, C., Wang, R.C., Li, H., Pokatayev, V., Dozmorov, I., Khan, S., Miyata, N., Fraile, G., Raj, P., Xu, Z., Xu, Z., Ma, L., Lin, Z., Wang, H., Yang, Y., Ben-Amitai, D., Orenstein, N., Mussaffi, H., Baselga, E., Tadini, G., Grunebaum, E., Sarajlija, A., Krzewski, K., Wakeland, E.K., Yan, N., de la Morena, M.T., Zinn, A.R., Burstein, E., 2016. DNA polymerase-alpha regulates the activation of type I interferons through cytosolic RNA:DNA synthesis. Nat. Immunol. 17, 495–504. Stittrich, A.B., Haftmann, C., Sgouroudis, E., Kuhl, A.A., Hegazy, A.N., Panse, I., Riedel, R., Flossdorf, M., Dong, J., Fuhrmann, F., Heinz, G.A., Fang, Z., Li, N., Bissels, U., Hatam, F., Jahn, A., Hammoud, B., Matz, M., Schulze, F.M., Baumgrass, R., Bosio, A., Mollenkopf, H.J., Grun, J., Thiel, A., Chen, W., Hofer, T., Loddenkemper, C., Lohning, M., Chang, H.D., Rajewsky, N., Radbruch, A., Mashreghi, M.F., 2010. The microRNA miR-182 is induced by IL-2 and promotes clonal expansion of activated helper T lymphocytes. Nat. Immunol. 11, 1057–1062. Stranges, P.B., Watson, J., Cooper, C.J., Choisy-Rossi, C.M., Stonebraker, A.C., Beighton, R.A., Hartig, H., Sundberg, J.P., Servick, S., Kaufmann, G., Fink, P.J., Chervonsky, A.V., 2007. Elimination of antigen-presenting cells and autoreactive T cells by Fas contributes to prevention of autoimmunity. Immunity 26, 629–641. Strasser, A., Bouillet, P., 2003. The control of apoptosis in lymphocyte selection. Immunol. Rev. 193, 82–92. Subramanian, S., Tus, K., Li, Q.Z., Wang, A., Tian, X.H., Zhou, J., Liang, C., Bartov, G., McDaniel, L.D., Zhou, X.J., Schultz, R.A., Wakeland, E.K., 2006. A Tlr7 translocation accelerates systemic autoimmunity in murine lupus. Proc. Natl. Acad. Sci. U.S.A. 103, 9970–9975. Subudhi, S.K., Zhou, P., Yerian, L.M., Chin, R.K., Lo, J.C., Anders, R.A., Sun, Y., Chen, L., Wang, Y., Alegre, M.L., Fu, Y.X., 2004. Local expression of B7-H1 promotes organ-specific autoimmunity and transplant rejection. J. Clin. Invest. 113, 694–700. Suh, C.I., Stull, N.D., Li, X.J., Tian, W., Price, M.O., Grinstein, S., Yaffe, M.B., Atkinson, S., Dinauer, M.C., 2006. The phosphoinositide-binding protein p40phox

148

S.K. Devarapu et al.

activates the NADPH oxidase during FcgammaIIA receptor-induced phagocytosis. J. Exp. Med. 203, 1915–1925. Summers, S.A., Hoi, A., Steinmetz, O.M., O’Sullivan, K.M., Ooi, J.D., Odobasic, D., Akira, S., Kitching, A.R., Holdsworth, S.R., 2010. TLR9 and TLR4 are required for the development of autoimmunity and lupus nephritis in pristane nephropathy. J. Autoimmun. 35, 291–298. Sun, X., Wiedeman, A., Agrawal, N., Teal, T.H., Tanaka, L., Hudkins, K.L., Alpers, C.E., Bolland, S., Buechler, M.B., Hamerman, J.A., Ledbetter, J.A., Liggitt, D., Elkon, K.B., 2013. Increased ribonuclease expression reduces inflammation and prolongs survival in TLR7 transgenic mice. J. Immunol. 190, 2536–2543. Tada, Y., Nagasawa, K., Ho, A., Morito, F., Koarada, S., Ushiyama, O., Suzuki, N., Ohta, A., Mak, T.W., 1999a. Role of the costimulatory molecule CD28 in the development of lupus in MRL/lpr mice. J. Immunol. 163, 3153–3159. Tada, Y., Nagasawa, K., Ho, A., Morito, F., Ushiyama, O., Suzuki, N., Ohta, H., Mak, T.W., 1999b. CD28-deficient mice are highly resistant to collagen-induced arthritis. J. Immunol. 162, 203–208. Takada, S., Shimizu, T., Hadano, Y., Matsumoto, K., Kataoka, Y., Arima, Y., Inoue, T., Sorano, S., 2012. Cryoglobulinemia (review). Mol. Med. Rep. 6, 3–8. Takami, N., Osawa, K., Miura, Y., Komai, K., Taniguchi, M., Shiraishi, M., Sato, K., Iguchi, T., Shiozawa, K., Hashiramoto, A., Shiozawa, S., 2006. Hypermethylated promoter region of DR3, the death receptor 3 gene, in rheumatoid arthritis synovial cells. Arthritis Rheum. 54, 779–787. Tan, T., Xiang, Y., Chang, C., Zhou, Z., 2014. Alteration of regulatory T cells in type 1 diabetes mellitus: a comprehensive review. Clin. Rev. Allergy Immunol. 47, 234–243. Tang, S., Zhang, Y., Yin, S.W., Gao, X.J., Shi, W.W., Wang, Y., Huang, X., Wang, L., Zou, L.Y., Zhao, J.H., Huang, Y.J., Shan, L.Y., Gounni, A.S., Wu, Y.Z., Zhang, J.B., 2015. Neutrophil extracellular trap formation is associated with autophagy-related signalling in ANCA-associated vasculitis. Clin. Exp. Immunol. 180, 408–418. Tao, Y., Kupfer, R., Stewart, B.J., Williams-Skipp, C., Crowell, C.K., Patel, D.D., Sain, S., Scheinman, R.I., 2006. AIRE recruits multiple transcriptional components to specific genomic regions through tethering to nuclear matrix. Mol. Immunol. 43, 335–345. Taubert, R., Schwendemann, J., Kyewski, B., 2007. Highly variable expression of tissue-restricted self-antigens in human thymus: implications for self-tolerance and autoimmunity. Eur. J. Immunol. 37, 838–848. Taylor, A.W., 2016. Ocular immune privilege and transplantation. Front. Immunol. 7, 37. Teachey, D.T., Seif, A.E., Grupp, S.A., 2010. Advances in the management and understanding of autoimmune lymphoproliferative syndrome (ALPS). Br. J. Haematol. 148, 205–216. Teichmann, L.L., Ols, M.L., Kashgarian, M., Reizis, B., Kaplan, D.H., Shlomchik, M.J., 2010. Dendritic cells in lupus are not required for activation of T and B cells but promote their expansion, resulting in tissue damage. Immunity 33, 967–978. Teichmann, L.L., Schenten, D., Medzhitov, R., Kashgarian, M., Shlomchik, M.J., 2013. Signals via the adaptor MyD88 in B cells and DCs make distinct and synergistic contributions to immune activation and tissue damage in lupus. Immunity 38, 528–540. Teichmann, L.L., Cullen, J.L., Kashgarian, M., Dong, C., Craft, J., Shlomchik, M.J., 2015. Local triggering of the ICOS coreceptor by CD11c(+) myeloid cells drives organ inflammation in lupus. Immunity 42, 552–565. Tellander, A.C., Michaelsson, E., Brunmark, C., Andersson, M., 2000. Potent adjuvant effect by anti-CD40 in collagen-induced arthritis. Enhanced disease is accompanied by increased production of collagen type-II reactive IgG2a and IFN-gamma. J. Autoimmun. 14, 295–302.

Mechanisms of Autoimmunity

149

Thai, T.H., Calado, D.P., Casola, S., Ansel, K.M., Xiao, C., Xue, Y., Murphy, A., Frendewey, D., Valenzuela, D., Kutok, J.L., Schmidt-Supprian, M., Rajewsky, N., Yancopoulos, G., Rao, A., Rajewsky, K., 2007. Regulation of the germinal center response by microRNA-155. Science 316, 604–608. Theofilopoulos, A.N., Baccala, R., Beutler, B., Kono, D.H., 2005. Type I interferons (alpha/beta) in immunity and autoimmunity. Annu. Rev. Immunol. 23, 307–336. Thery, C., Regnault, A., Garin, J., Wolfers, J., Zitvogel, L., Ricciardi-Castagnoli, P., Raposo, G., Amigorena, S., 1999. Molecular characterization of dendritic cell-derived exosomes. Selective accumulation of the heat shock protein hsc73. J. Cell Biol. 147, 599–610. Thery, C., Duban, L., Segura, E., Veron, P., Lantz, O., Amigorena, S., 2002. Indirect activation of naive CD4 + T cells by dendritic cell-derived exosomes. Nat. Immunol. 3, 1156–1162. Tiegs, S.L., Russell, D.M., Nemazee, D., 1993. Receptor editing in self-reactive bone marrow B cells. J. Exp. Med. 177, 1009–1020. Tillack, K., Breiden, P., Martin, R., Sospedra, M., 2012. T lymphocyte priming by neutrophil extracellular traps links innate and adaptive immune responses. J. Immunol. 188, 3150–3159. Tojo, T., Friou, G.J., Spiegelberg, H.L., 1970. Immunoglobulin G subclass of human antinuclear antibodies. Clin. Exp. Immunol. 6, 145–151. Tran, N.L., Manzin-Lorenzi, C., Santiago-Raber, M.-L., 2015. Toll-like receptor 8 deletion accelerates autoimmunity in a mouse model of lupus through a Toll-like receptor 7-dependent mechanism. Immunology 145, 60–70. Truedsson, L., Bengtsson, A.A., Sturfelt, G., 2007. Complement deficiencies and systemic lupus erythematosus. Autoimmunity 40, 560–566. Tsai, K.L., Chang, H.F., Wu, S.N., 2013. The inhibition of inwardly rectifying K+ channels by memantine in macrophages and microglial cells. Cell. Physiol. Biochem. 31, 938–951. Tse, H.M., Thayer, T.C., Steele, C., Cuda, C.M., Morel, L., Piganelli, J.D., Mathews, C.E., 2010. NADPH oxidase deficiency regulates Th lineage commitment and modulates autoimmunity. J. Immunol. 185, 5247–5258. Tsokos, G.C., 2005. Systemic lupus erythematosus and Sjogren’s syndrome. Curr. Opin. Rheumatol. 17, 511–512. Tsokos, G.C., 2011. Systemic lupus erythematosus. N. Engl. J. Med. 365, 2110–2121. Tsuboi, N., Asano, K., Lauterbach, M., Mayadas, T.N., 2008. Human neutrophil Fcgamma receptors initiate and play specialized nonredundant roles in antibody-mediated inflammatory diseases. Immunity 28, 833–846. Turner, M.L., Schnorfeil, F.M., Brocker, T., 2011. MicroRNAs regulate dendritic cell differentiation and function. J. Immunol. 187, 3911–3917. Turpin, D., Truchetet, M.E., Faustin, B., Augusto, J.F., Contin-Bordes, C., Brisson, A., Blanco, P., Duffau, P., 2016. Role of extracellular vesicles in autoimmune diseases. Autoimmun. Rev. 15, 174–183. Ueda, H., Howson, J.M., Esposito, L., Heward, J., Snook, H., Chamberlain, G., Rainbow, D.B., Hunter, K.M., Smith, A.N., Di Genova, G., Herr, M.H., Dahlman, I., Payne, F., Smyth, D., Lowe, C., Twells, R.C., Howlett, S., Healy, B., Nutland, S., Rance, H.E., Everett, V., Smink, L.J., Lam, A.C., Cordell, H.J., Walker, N.M., Bordin, C., Hulme, J., Motzo, C., Cucca, F., Hess, J.F., Metzker, M.L., Rogers, J., Gregory, S., Allahabadia, A., Nithiyananthan, R., Tuomilehto-Wolf, E., Tuomilehto, J., Bingley, P., Gillespie, K.M., Undlien, D.E., Ronningen, K.S., Guja, C., Ionescu-Tirgoviste, C., Savage, D.A., Maxwell, A.P., Carson, D.J., Patterson, C.C., Franklyn, J.A., Clayton, D.G., Peterson, L.B., Wicker, L.S., Todd, J.A., Gough, S.C., 2003. Association of the T-cell regulatory gene CTLA4 with susceptibility to autoimmune disease. Nature 423, 506–511.

150

S.K. Devarapu et al.

Ullal, A.J., Reich 3rd, C.F., Clowse, M., Criscione-Schreiber, L.G., Tochacek, M., Monestier, M., Pisetsky, D.S., 2011. Microparticles as antigenic targets of antibodies to DNA and nucleosomes in systemic lupus erythematosus. J. Autoimmun. 36, 173–180. Utsugi-Kobukai, S., Fujimaki, H., Hotta, C., Nakazawa, M., Minami, M., 2003. MHC class I-mediated exogenous antigen presentation by exosomes secreted from immature and mature bone marrow derived dendritic cells. Immunol. Lett. 89, 125–131. Vaeth, M., Zee, I., Concepcion, A.R., Maus, M., Shaw, P., Portal-Celhay, C., Zahra, A., Kozhaya, L., Weidinger, C., Philips, J., Unutmaz, D., Feske, S., 2015. Ca2 + signaling but not store-operated Ca2+ entry is required for the function of macrophages and dendritic cells. J. Immunol. 195, 1202–1217. Vafiadis, P., Bennett, S.T., Todd, J.A., Nadeau, J., Grabs, R., Goodyer, C.G., Wickramasinghe, S., Colle, E., Polychronakos, C., 1997. Insulin expression in human thymus is modulated by INS VNTR alleles at the IDDM2 locus. Nat. Genet. 15, 289–292. Van Eyck, L., De Somer, L., Pombal, D., Bornschein, S., Frans, G., Humblet-Baron, S., Moens, L., de Zegher, F., Bossuyt, X., Wouters, C., Liston, A., 2015. Brief report: IFIH1 mutation causes systemic lupus erythematosus with selective IgA deficiency. Arthritis Rheumatol. 67, 1592–1597. Vidal, S., Kono, D.H., Theofilopoulos, A.N., 1998. Loci predisposing to autoimmunity in MRL-Fas lpr and C57BL/6-Faslpr mice. J. Clin. Invest. 101, 696–702. Vignali, D.A., Collison, L.W., Workman, C.J., 2008. How regulatory T cells work. Nat. Rev. Immunol. 8, 523–532. Vigorito, E., Perks, K.L., Abreu-Goodger, C., Bunting, S., Xiang, Z., Kohlhaas, S., Das, P.P., Miska, E.A., Rodriguez, A., Bradley, A., Smith, K.G., Rada, C., Enright, A.J., Toellner, K.M., Maclennan, I.C., Turner, M., 2007. microRNA-155 regulates the generation of immunoglobulin class-switched plasma cells. Immunity 27, 847–859. Villanueva, E., Yalavarthi, S., Berthier, C.C., Hodgin, J.B., Khandpur, R., Lin, A.M., Rubin, C.J., Zhao, W., Olsen, S.H., Klinker, M., Shealy, D., Denny, M.F., Plumas, J., Chaperot, L., Kretzler, M., Bruce, A.T., Kaplan, M.J., 2011. Netting neutrophils induce endothelial damage, infiltrate tissues, and expose immunostimulatory molecules in systemic lupus erythematosus. J. Immunol. 187, 538–552. Walker, L.S., Abbas, A.K., 2002. The enemy within: keeping self-reactive T cells at bay in the periphery. Nat. Rev. Immunol. 2, 11–19. Walker, M.R., Kasprowicz, D.J., Gersuk, V.H., Benard, A., Van Landeghen, M., Buckner, J.H., Ziegler, S.F., 2003. Induction of FoxP3 and acquisition of T regulatory activity by stimulated human CD4+CD25- T cells. J. Clin. Invest. 112, 1437–1443. Walter, U., Franzke, A., Sarukhan, A., Zober, C., von Boehmer, H., Buer, J., Lechner, O., 2000. Monitoring gene expression of TNFR family members by beta-cells during development of autoimmune diabetes. Eur. J. Immunol. 30 (4), 1224–1232. Wan, S., Zhou, Z., Duan, B., Morel, L., 2008. Direct B cell stimulation by dendritic cells in a mouse model of lupus. Arthritis Rheum. 58, 1741–1750. Wang, X., Huang, W., Mihara, M., Sinha, J., Davidson, A., 2002. Mechanism of action of combined short-term CTLA4Ig and anti-CD40 ligand in murine systemic lupus erythematosus. J. Immunol. 168, 2046–2053. Wang, C.J., Chou, F.C., Chu, C.H., Wu, J.C., Lin, S.H., Chang, D.M., Sytwu, H.K., 2008. Protective role of programmed death 1 ligand 1 (PD-L1)in nonobese diabetic mice: the paradox in transgenic models. Diabetes 57, 1861–1869. Wang, X., Hao, J., Metzger, D.L., Mui, A., Ao, Z., Akhoundsadegh, N., Langermann, S., Liu, L., Chen, L., Ou, D., Verchere, C.B., Warnock, G.L., 2011. Early treatment of NOD mice with B7-H4 reduces the incidence of autoimmune diabetes. Diabetes 60, 3246–3255.

Mechanisms of Autoimmunity

151

Wang, Z., Zheng, Y., Hou, C., Yang, L., Li, X., Lin, J., Huang, G., Lu, Q., Wang, C.Y., Zhou, Z., 2013. DNA methylation impairs TLR9 induced Foxp3 expression by attenuating IRF-7 binding activity in fulminant type 1 diabetes. J. Autoimmun. 41, 50–59. Wang, C.H., Rong, M.Y., Wang, L., Ren, Z., Chen, L.N., Jia, J.F., Li, X.Y., Wu, Z.B., Chen, Z.N., Zhu, P., 2014a. CD147 up-regulates calcium-induced chemotaxis, adhesion ability and invasiveness of human neutrophils via a TRPM-7-mediated mechanism. Rheumatology (Oxford) 53, 2288–2296. Wang, C.M., Chang, S.W., Wu, Y.J., Lin, J.C., Ho, H.H., Chou, T.C., Yang, B., Wu, J., Chen, J.Y., 2014b. Genetic variations in Toll-like receptors (TLRs 3/7/8) are associated with systemic lupus erythematosus in a Taiwanese population. Sci. Rep. 4, 3792. Wang, Y., Xiao, Y., Zhong, L., Ye, D., Zhang, J., Tu, Y., Bornstein, S.R., Zhou, Z., Lam, K.S., Xu, A., 2014c. Increased neutrophil elastase and proteinase 3 and augmented NETosis are closely associated with beta-cell autoimmunity in patients with type 1 diabetes. Diabetes 63, 4239–4248. Wardemann, H., Yurasov, S., Schaefer, A., Young, J.W., Meffre, E., Nussenzweig, M.C., 2003. Predominant autoantibody production by early human B cell precursors. Science 301, 1374–1377. Webb, L.M., Walmsley, M.J., Feldmann, M., 1996. Prevention and amelioration of collagen-induced arthritis by blockade of the CD28 co-stimulatory pathway: requirement for both B7-1 and B7-2. Eur. J. Immunol. 26, 2320–2328. Weening, J.J., D’Agati, V.D., Schwartz, M.M., Seshan, S.V., Alpers, C.E., Appel, G.B., Balow, J.E., Bruijn, J.A., Cook, T., Ferrario, F., Fogo, A.B., Ginzler, E.M., Hebert, L., Hill, G., Hill, P., Jennette, J.C., Kong, N.C., Lesavre, P., Lockshin, M., Looi, L.M., Makino, H., Moura, L.A., Nagata, M., 2004. The classification of glomerulonephritis in systemic lupus erythematosus revisited. Kidney Int. 65, 521–530. Weinberg, A.D., Bourdette, D.N., Sullivan, T.J., Lemon, M., Wallin, J.J., Maziarz, R., Davey, M., Palida, F., Godfrey, W., Engleman, E., Fulton, R.J., Offner, H., Vandenbark, A.A., 1996. Selective depletion of myelin-reactive T cells with the anti-OX-40 antibody ameliorates autoimmune encephalomyelitis. Nat. Med. 2, 183–189. Weinberg, A.D., Wegmann, K.W., Funatake, C., Whitham, R.H., 1999. Blocking OX-40/ OX-40 ligand interaction in vitro and in vivo leads to decreased T cell function and amelioration of experimental allergic encephalomyelitis. J. Immunol. 162, 1818–1826. Wengner, A.M., Hopken, U.E., Petrow, P.K., Hartmann, S., Schurigt, U., Brauer, R., Lipp, M., 2007. CXCR5- and CCR7-dependent lymphoid neogenesis in a murine model of chronic antigen-induced arthritis. Arthritis Rheum. 56, 3271–3283. Wilber, A., O’Connor, T.P., Lu, M.L., Karimi, A., Schneider, M.C., 2003. Dnase1l3 deficiency in lupus-prone MRL and NZB/W F1 mice. Clin. Exp. Immunol. 134, 46–52. Wilson, N.J., Boniface, K., Chan, J.R., McKenzie, B.S., Blumenschein, W.M., Mattson, J.D., Basham, B., Smith, K., Chen, T., Morel, F., Lecron, J.C., Kastelein, R.A., Cua, D.J., McClanahan, T.K., Bowman, E.P., de Waal Malefyt, R., 2007. Development, cytokine profile and function of human interleukin 17-producing helper T cells. Nat. Immunol. 8, 950–957. Wing, J.B., Sakaguchi, S., 2014. Foxp3(+) T(reg) cells in humoral immunity. Int. Immunol. 26, 61–69. Winter, S., Loddenkemper, C., Aebischer, A., Rabel, K., Hoffmann, K., Meyer, T.F., Lipp, M., Hopken, U.E., 2010. The chemokine receptor CXCR5 is pivotal for ectopic mucosa-associated lymphoid tissue neogenesis in chronic Helicobacter pylori-induced inflammation. J. Mol. Med. (Berl.) 88, 1169–1180. Wong, M., La Cava, A., Singh, R.P., Hahn, B.H., 2010. Blockade of programmed death-1 in young (New Zealand black x New Zealand white)F1 mice promotes the activity of

152

S.K. Devarapu et al.

suppressive CD8 + T cells that protect from lupus-like disease. J. Immunol. 185, 6563–6571. Wong, M., La Cava, A., Hahn, B.H., 2013. Blockade of programmed death-1 in young (New Zealand Black x New Zealand White)F1 mice promotes the suppressive capacity of CD4 + regulatory T cells protecting from lupus-like disease. J. Immunol. 190, 5402–5410. Wong, S.L., Demers, M., Martinod, K., Gallant, M., Wang, Y., Goldfine, A.B., Kahn, C.R., Wagner, D.D., 2015. Diabetes primes neutrophils to undergo NETosis, which impairs wound healing. Nat. Med. 21, 815–819. Worthmann, K., Gueler, F., von Vietinghoff, S., Davalos-Misslitz, A., Wiehler, F., Davidson, A., Witte, T., Haller, H., Schiffer, M., Falk, C.S., Schiffer, L., 2014. Pathogenetic role of glomerular CXCL13 expression in lupus nephritis. Clin. Exp. Immunol. 178, 20–27. Wu, J., Chen, Z.J., 2014. Innate immune sensing and signaling of cytosolic nucleic acids. Annu. Rev. Immunol. 32, 461–488. Wu, W., Shi, X., Xu, C., 2016. Regulation of T cell signalling by membrane lipids. Nat. Rev. Immunol. 16, 690–701. Xiao, C., Calado, D.P., Galler, G., Thai, T.H., Patterson, H.C., Wang, J., Rajewsky, N., Bender, T.P., Rajewsky, K., 2007. MiR-150 controls B cell differentiation by targeting the transcription factor c-Myb. Cell 131, 146–159. Xiao, C., Srinivasan, L., Calado, D.P., Patterson, H.C., Zhang, B., Wang, J., Henderson, J.M., Kutok, J.L., Rajewsky, K., 2008. Lymphoproliferative disease and autoimmunity in mice with increased miR-17-92 expression in lymphocytes. Nat. Immunol. 9, 405–414. Xie, Z., Chang, C., Zhou, Z., 2014. Molecular mechanisms in autoimmune type 1 diabetes: a critical review. Clin. Rev. Allergy Immunol. 47, 174–192. Xin, Q., Li, J., Dang, J., Bian, X., Shan, S., Yuan, J., Qian, Y., Liu, Z., Liu, G., Yuan, Q., Liu, N., Ma, X., Gao, F., Gong, Y., Liu, Q., 2015. miR-155 deficiency ameliorates autoimmune inflammation of systemic lupus erythematosus by targeting S1pr1 in faslpr/lpr mice. J. Immunol. 194, 5437–5445. Xu, H., Lai, W., Zhang, Y., Liu, L., Luo, X., Zeng, Y., Wu, H., Lan, Q., Chu, Z., 2014. Tumor-associated macrophage-derived IL-6 and IL-8 enhance invasive activity of LoVo cells induced by PRL-3 in a KCNN4 channel-dependent manner. BMC Cancer 14, 330. Yamada, R., Yamamoto, K., 2007. Mechanisms of disease: genetics of rheumatoid arthritis—ethnic differences in disease-associated genes. Nat. Clin. Pract. Rheumatol. 3, 644–650. Yamazaki-Nakashimada, M., Zaltzman-Girshevich, S., Garcia de la Puente, S., De Leon-Bojorge, B., Espinosa-Padilla, S., Saez-de-Ocariz, M., Carrasco-Daza, D., Hernandez-Bautista, V., Perez-Fernandez, L., Espinosa-Rosales, F., 2006. Hyper-IgE syndrome and autoimmunity in Mexican children. Pediatr. Nephrol. 21, 1200–1205. Yang, J.Q., Wen, X., Kim, P.J., Singh, R.R., 2011. Invariant NKT cells inhibit autoreactive B cells in a contact- and CD1d-dependent manner. J. Immunol. 186, 1512–1520. Yang, H., Lee, S.M., Gao, B., Zhang, J., Fang, D., 2013. Histone deacetylase sirtuin 1 deacetylates IRF1 protein and programs dendritic cells to control Th17 protein differentiation during autoimmune inflammation. J. Biol. Chem. 288, 37256–37266. Yang, X., Yang, J., Chu, Y., Xue, Y., Xuan, D., Zheng, S., Zou, H., 2014. T follicular helper cells and regulatory B cells dynamics in systemic lupus erythematosus. PLoS One 9e88441. Yao, Y.L., Yang, W.M., Seto, E., 2001. Regulation of transcription factor YY1 by acetylation and deacetylation. Mol. Cell. Biol. 21, 5979–5991.

Mechanisms of Autoimmunity

153

Yao, L., Chen, H.P., Ma, Q., 2014. Piperlongumine alleviates lupus nephritis in MRLFas(lpr) mice by regulating the frequency of Th17 and regulatory T cells. Immunol. Lett. 161, 76–80. Yao, Y., Vent-Schmidt, J., McGeough, M.D., Wong, M., Hoffman, H.M., Steiner, T.S., Levings, M.K., 2015. Tr1 cells, but Not Foxp3 + regulatory T cells, suppress NLRP3 inflammasome activation via an IL-10-dependent mechanism. J. Immunol. 195, 488–497. Yasutomo, K., Horiuchi, T., Kagami, S., Tsukamoto, H., Hashimura, C., Urushihara, M., Kuroda, Y., 2001. Mutation of DNASE1 in people with systemic lupus erythematosus. Nat. Genet. 28, 313–314. Yipp, B.G., Kubes, P., 2013. NETosis: how vital is it? Blood 122, 2784–2794. Yipp, B.G., Petri, B., Salina, D., Jenne, C.N., Scott, B.N., Zbytnuik, L.D., Pittman, K., Asaduzzaman, M., Wu, K., Meijndert, H.C., Malawista, S.E., de Boisfleury Chevance, A., Zhang, K., Conly, J., Kubes, P., 2012. Infection-induced NETosis is a dynamic process involving neutrophil multitasking in vivo. Nat. Med. 18, 1386–1393. Yokogawa, N., Vivino, F.B., 2009. Hydralazine-induced autoimmune disease: comparison to idiopathic lupus and ANCA-positive vasculitis. Mod. Rheumatol. 19, 338–347. Yokoi, N., Komeda, K., Wang, H.Y., Yano, H., Kitada, K., Saitoh, Y., Seino, Y., Yasuda, K., Serikawa, T., Seino, S., 2002. Cblb is a major susceptibility gene for rat type 1 diabetes mellitus. Nat. Genet. 31, 391–394. Yoshioka, T., Nakajima, A., Akiba, H., Ishiwata, T., Asano, G., Yoshino, S., Yagita, H., Okumura, K., 2000. Contribution of OX40/OX40 ligand interaction to the pathogenesis of rheumatoid arthritis. Eur. J. Immunol. 30, 2815–2823. Yu, F., Wu, L.H., Tan, Y., Li, L.H., Wang, C.L., Wang, W.K., Qu, Z., Chen, M.H., Gao, J.J., Li, Z.Y., Zheng, X., Ao, J., Zhu, S.N., Wang, S.X., Zhao, M.H., Zou, W.Z., Liu, G., 2010. Tubulointerstitial lesions of patients with lupus nephritis classified by the 2003 International Society of Nephrology and Renal Pathology Society system. Kidney Int. 77, 820–829. Yuen, J., Pluthero, F.G., Douda, D.N., Riedl, M., Cherry, A., Ulanova, M., Kahr, W.H., Palaniyar, N., Licht, C., 2016. NETosing neutrophils activate complement both on their own NETs and bacteria via alternative and non-alternative pathways. Front. Immunol. 7, 137. Yung, S., Chan, T.M., 2015. Mechanisms of kidney injury in lupus nephritis—the role of anti-dsDNA antibodies. Front. Immunol. 6, 475. Yuseff, M.I., Pierobon, P., Reversat, A., Lennon-Dumenil, A.M., 2013. How B cells capture, process and present antigens: a crucial role for cell polarity. Nat. Rev. Immunol. 13, 475–486. Zamora, M.R., Warner, M.L., Tuder, R., Schwarz, M.I., 1997. Diffuse alveolar hemorrhage and systemic lupus erythematosus. Clinical presentation, histology, survival, and outcome. Medicine 76, 192–202. Zha, Y., Marks, R., Ho, A.W., Peterson, A.C., Janardhan, S., Brown, I., Praveen, K., Stang, S., Stone, J.C., Gajewski, T.F., 2006. T cell anergy is reversed by active Ras and is regulated by diacylglycerol kinase-alpha. Nat. Immunol. 7, 1166–1173. Zhang, H.G., Liu, C., Su, K., Yu, S., Zhang, L., Zhang, S., Wang, J., Cao, X., Grizzle, W., Kimberly, R.P., 2006a. A membrane form of TNF-alpha presented by exosomes delays T cell activation-induced cell death. J. Immunol. 176, 7385–7393. Zhang, Q., Wang, H.Y., Woetmann, A., Raghunath, P.N., Odum, N., Wasik, M.A., 2006b. STAT3 induces transcription of the DNA methyltransferase 1 gene (DNMT1) in malignant T lymphocytes. Blood 108, 1058–1064. Zhang, D., Sanchez-Fueyo, A., Kawamoto, K., Alexopoulos, S.P., Zhang, W., Zheng, X.X., 2010. Th1 to Th2 immune deviation facilitates, but does not cause, islet allograft tolerance in mice. Cytokine 51, 311–319.

154

S.K. Devarapu et al.

Zhang, W., Cai, Y., Xu, W., Yin, Z., Gao, X., Xiong, S., 2013. AIM2 facilitates the apoptotic DNA-induced systemic lupus erythematosus via arbitrating macrophage functional maturation. J. Clin. Immunol. 33, 925–937. Zhang, X., Bogunovic, D., Payelle-Brogard, B., Francois-Newton, V., Speer, S.D., Yuan, C., Volpi, S., Li, Z., Sanal, O., Mansouri, D., Tezcan, I., Rice, G.I., Chen, C., Mansouri, N., Mahdaviani, S.A., Itan, Y., Boisson, B., Okada, S., Zeng, L., Wang, X., Jiang, H., Liu, W., Han, T., Liu, D., Ma, T., Wang, B., Liu, M., Liu, J.Y., Wang, Q.K., Yalnizoglu, D., Radoshevich, L., Uze, G., Gros, P., Rozenberg, F., Zhang, S.Y., Jouanguy, E., Bustamante, J., Garcia-Sastre, A., Abel, L., Lebon, P., Notarangelo, L.D., Crow, Y.J., Boisson-Dupuis, S., Casanova, J.L., Pellegrini, S., 2015. Human intracellular ISG15 prevents interferon-alpha/beta over-amplification and auto-inflammation. Nature 517, 89–93. Zhang, H., Fu, R., Guo, C., Huang, Y., Wang, H., Wang, S., Zhao, J., Yang, N., 2016. Anti-dsDNA antibodies bind to TLR4 and activate NLRP3 inflammasome in lupus monocytes/macrophages. J. Transl. Med. 14, 156. Zhao, M., Sun, Y., Gao, F., Wu, X., Tang, J., Yin, H., Luo, Y., Richardson, B., Lu, Q., 2010a. Epigenetics and SLE: RFX1 downregulation causes CD11a and CD70 overexpression by altering epigenetic modifications in lupus CD4 + T cells. J. Autoimmun. 35, 58–69. Zhao, M., Wu, X., Zhang, Q., Luo, S., Liang, G., Su, Y., Tan, Y., Lu, Q., 2010b. RFX1 regulates CD70 and CD11a expression in lupus T cells by recruiting the histone methyltransferase SUV39H1. Arthritis Res. Ther. 12, R227. Zhao, J., Wang, H., Dai, C., Wang, H., Zhang, H., Huang, Y., Wang, S., Gaskin, F., Yang, N., Fu, S.M., 2013a. P2X7 blockade attenuates murine lupus nephritis by inhibiting activation of the NLRP3/ASC/caspase 1 pathway. Arthritis Rheum. 65, 3176–3185. Zhao, M., Liu, Q., Liang, G., Wang, L., Luo, S., Tang, Q., Zhao, H., Su, Y., Yung, S., Chan, T.M., Lu, Q., 2013b. E4BP4 overexpression: a protective mechanism in CD4 + T cells from SLE patients. J. Autoimmun. 41, 152–160. Zhou, T., Cheng, J., Yang, P., Wang, Z., Liu, C., Su, X., Bluethmann, H., Mountz, J.D., 1996. Inhibition of Nur77/Nurr1 leads to inefficient clonal deletion of self-reactive T cells. J. Exp. Med. 183, 1879–1892. Zhou, X., Jeker, L.T., Fife, B.T., Zhu, S., Anderson, M.S., McManus, M.T., Bluestone, J.A., 2008. Selective miRNA disruption in T reg cells leads to uncontrolled autoimmunity. J. Exp. Med. 205, 1983–1991. Zhu, C., Anderson, A.C., Schubart, A., Xiong, H., Imitola, J., Khoury, S.J., Zheng, X.X., Strom, T.B., Kuchroo, V.K., 2005. The Tim-3 ligand galectin-9 negatively regulates T helper type 1 immunity. Nat. Immunol. 6, 1245–1252. Zhu, E., Wang, X., Zheng, B., Wang, Q., Hao, J., Chen, S., Zhao, Q., Zhao, L., Wu, Z., Yin, Z., 2014. miR-20b suppresses Th17 differentiation and the pathogenesis of experimental autoimmune encephalomyelitis by targeting RORgammat and STAT3. J. Immunol. 192, 5599–5609. Zohar, Y., Wildbaum, G., Novak, R., Salzman, A.L., Thelen, M., Alon, R., Barsheshet, Y., Karp, C.L., Karin, N., 2014. CXCL11-dependent induction of FOXP3-negative regulatory T cells suppresses autoimmune encephalomyelitis. J. Clin. Invest. 124, 2009–2022. Zsiros, E., Kis-Toth, K., Hajdu, P., Gaspar, R., Bielanska, J., Felipe, A., Rajnavolgyi, E., Panyi, G., 2009. Developmental switch of the expression of ion channels in human dendritic cells. J. Immunol. 183, 4483–4492.

CHAPTER THREE

Old and Novel Functions of Caspase-2 M.A. Miles, T. Kitevska-Ilioski, C.J. Hawkins1 La Trobe Institute for Molecular Science, La Trobe University, Bundoora, VIC, Australia 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Biochemical Activity 2.1 Minimal Specificity 2.2 Tools for Monitoring Activity 2.3 Cellular Substrates 3. Activation 3.1 Cleavage 3.2 Induced Proximity Activates Caspase-2 Within the PIDDosome 4. Caspase-2-Mediated Responses to DNA Damage and Mitotic Stress 4.1 DNA Damage-Induced Caspase-2 PIDDosome Formation 4.2 CHK1 Inhibition of PIDDosome Formation and Its Regulation by p53 4.3 Inconsistencies Regarding Outcomes of PIDDosome Signaling and Its Inhibition by CHK1 4.4 PIDDosome Inhibition by BubR1 4.5 An Alternative PIDD Complex Induces NF-κB 4.6 Phosphorylation in Prodomain or Linker Prevents Caspase-2 Activation 4.7 Caspase-2-Mediated Cell Cycle Arrest vs Apoptosis, Following DNA Damage or Mitotic Stress 4.8 Genomic Instability and Aneuploidy Due to Caspase-2 Inhibition 5. Responses to ER Stress 5.1 Brucella Infection-Mediated ER Stress 5.2 Rhabdovirus Infection-Mediated ER Stress 6. Relieving Oxidative Stress 6.1 Aging 6.2 Amelioration of Oxidative Stress 6.3 Suppression of Autophagy 6.4 Suppression of ROS-Driven Osteoclastogenesis 7. Metabolism 8. Cancer 8.1 Mixed Messages From Human and Mouse Research 8.2 Promoting Neuroblastoma 8.3 Tumor Suppressor

International Review of Cell and Molecular Biology, Volume 332 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2016.12.002

#

2017 Elsevier Inc. All rights reserved.

156 157 157 157 160 167 167 168 170 170 170 171 172 172 173 174 176 177 177 178 179 179 179 180 181 182 184 184 186 186

155

156

M.A. Miles et al.

9. Vision 9.1 Retinal Cell Death 9.2 Loss of Retinal Ganglion Cells Following Optic Nerve Injury 10. Neurological Conditions 10.1 Exacerbating Ischemic/Reperfusion Injury 10.2 Excitotoxicity 10.3 Withdrawal of Nerve Growth Factor or Serum 10.4 Alzheimer’s Disease 10.5 Huntington’s and Motor Neuron Diseases 11. Conclusions 11.1 Why Has Caspase-2 Been Evolutionarily Conserved? 11.2 Manipulating Caspase-2 for Therapeutic Benefit 11.3 Could a Common Mechanism Underlie the Apparently Disparate Cellular Roles of Caspase-2? 11.4 Molecular Pathways Upstream and Downstream of Caspase-2 References

189 189 190 191 191 192 193 193 194 195 195 196 197 198 201

Abstract Although caspase-2 is a highly conserved protease that has received a lot of research attention, consensus about its roles and the molecular mechanisms that underpin them has been elusive. Recent improvements to our understanding of the activities of caspase-2 have been facilitated by the development and refinement of techniques allowing identification of cellular processes instigated by this caspase. Following DNA damage, caspase-2 can be activated in a molecular complex called the “PIDDosome”; however, other stimuli provoke caspase-2-dependent activities that do not appear to involve this complex. Further research is needed into the mechanisms that activate caspase-2, and the substrates that it cleaves to accomplish its functions. Apart from DNA damage, caspase-2 has also been implicated in responses to other cellular stresses including oxidative damage, endoplasmic reticulum stress, and aberrant mitotic signaling. Caspase-2 sensitized animals fed diets high in fat and sugar to glucose intolerance and liver disease, so drugs that target this protease may be useful to prevent or treat metabolic conditions. Caspase-2 loss enhanced the survival of retinal ganglion cells following optic nerve damage, prompting hope that caspase-2 inhibitors may help treat optic nerve injuries. Caspase-2 predisposed animals to neuroblastoma but tended to provide protection against oncogene-driven cancers. Intriguingly, caspase-2 facilitated host cell death following viral or bacterial infection, raising the possibility that its evolutionary retention may reflect its ability to induce defensive apoptosis following intracellular infection.

1. INTRODUCTION The caspases comprise an ancient family of cysteine proteases with aspartate specificities, which includes orthologs from nematodes (Conradt et al., 2016), insects (Cooper et al., 2009), and mammals (Eckhart et al.,

Old and Novel Functions of Caspase-2

157

2008) that share biochemical activities and functional roles. Eighteen distinct caspases arose during mammalian evolution, but multiple duplication, deletion, and mutation events have produced a human genome that contains 12 genes encoding enzymatically competent caspases (Eckhart et al., 2008). Some of these are apparently dispensable for mammalian life, as caspases-4/-5, -10, -14, and -16 are absent from platypus, rodent, dolphin, and guinea pig genomes, respectively (Eckhart et al., 2008; Strasser et al., 2015). Almost every caspase that is shared by all mammals has a clearly defined primary role, either in the maturation of inflammatory cytokines (caspase-1), the initiation of apoptosis (caspases-8 and -9), or its execution (caspases-3, -6, and -7), although ancillary functions are still being assigned to these enzymes (Shalini et al., 2015a). All studied mammalian genomes contain a single conserved caspase-2 gene, implying that some selective pressure drove its retention throughout evolution, yet ascertaining its role(s) has proven difficult. This review summarizes the current state of knowledge regarding the biochemical and biological functions of caspase-2, focusing on recent developments in this field. We also highlight opportunities for future research that may help clarify the functions of this mysterious caspase.

2. BIOCHEMICAL ACTIVITY 2.1 Minimal Specificity Early data from studies employing Positional-Scanning Combinatorial Peptide Libraries and synthetic peptides (Talanian et al., 1997; Thornberry et al., 1997) suggested that caspase-2 would most efficiently cleave sequences bearing aspartate in the P1 position (the residue on the amino side of the cleavage site) and that it could tolerate a variety of residues in P2 and P3 positions (the next two residues toward the amino terminus). Caspase-2 preferred aspartate in P4, a hydrophobic amino acid in P5 (Talanian et al., 1997; Thornberry et al., 1997), and glycine in the P10 position (the residue immediately carboxyl to the cleavage site) (Kitevska et al., 2014).

2.2 Tools for Monitoring Activity Peptide-based substrates and inhibitors bearing the sequence VDVAD have been widely used as tools to monitor caspase-2 activity. However, because these reagents also target caspases-3 and -7 (Mcstay et al., 2008; Pereira and Song, 2008), they cannot be used in cell-based experiments to specifically pinpoint events that involve caspase-2. A different fluorogenic peptide, Ac-VDTTD-AFC, was significantly more selective than

158

M.A. Miles et al.

Ac-VDVAD-AFC for caspase-2 over caspase-3 (Kitevska et al., 2014), but unfortunately no substrates have yet been created that are truly caspase-2 specific, and this deficiency has confounded investigations into the roles played by caspase-2. Almost a decade ago, Schweizer et al. reported their creation of an ankyrin repeat protein (designated “AR_F8”) that could selectively and potently inhibit caspase-2 but not caspases-3, -7, -8, or -9 in vitro (Schweizer et al., 2007). This specificity was achieved via an allosteric interaction between AR_F8 and regions of caspase-2 that, unlike its active site, are not shared with other caspases (Schweizer et al., 2007). Addition of this protein to cell lysates helped define cytoskeletal substrates of caspase-2 (Vakifahmetoglu-Norberg et al., 2013), but AR_F8 has not been widely adopted for research into processes that involve caspase-2. It is unclear whether AR_F8 expression in cells could inhibit caspase-2 activity, as to our knowledge this inhibitor has only been used to date in vitro. A cell-permeable small-molecule inhibitor with equivalent potency and specificity to AR_F8 would be enormously useful. Chemical modification of the peptide-based inhibitor Ac-VDVAD-CHO yielded a reagent (33h) with enhanced selectivity for caspase-2 over caspase-3, but unfortunately the derivative had a slightly lower affinity for caspase-2 than the parent molecule (Maillard et al., 2011). Compound 33h could reduce VDVADase activity in 293T cells transfected with a caspase-2 expression vector 65 times more effectively than it could diminish the DEVDase activity of caspase-3 transfectants, but to date it has not been evaluated in other cellular contexts. Cleavage of procaspase-2, to remove the amino-terminal prodomain and separate the catalytic subunits, is sometimes interpreted as a sign of activity, although as outlined in Section 3.1, is not entirely clear whether or not caspase-2 must be cleaved to be active. An alternative approach to detecting caspase-2 activity relies on the ability of Biotin-Val-Ala-Asp(OMe)fluoromethylketone (bVAD) to enter cells, then inhibit a wide range of caspases to which it irreversibly binds. If cells are incubated with bVAD prior to application of a stimulus, those caspases that are first activated will bind to the reagent and be inhibited (Tinel and Tschopp, 2004; Tu et al., 2006). This blocks any caspase cascade that would otherwise ensue, and the bound initiator caspases can be specifically isolated using streptavidin-bound beads. Immunoblotting can then be used to identify those apical caspases that are activated in response to the applied stimulus. In vitro kinetic assays suggested that VAD-based reagents may only inefficiently target caspase-2: a closely related inhibitor, zVAD-fmk, inhibited caspase-2 activity 615–960 times

Old and Novel Functions of Caspase-2

159

more slowly than caspases-1, -8, or -9, and 56–62 times more slowly than caspases-3 or -7, when used at 1 μM (Garcia-Calvo et al., 1998). Nevertheless, using high concentrations of bVAD (typically 50 μM), this trapping technique has been used to show that caspase-2 plays an initiating role in cell death following optic nerve injury, incubation with Aβ peptide, nerve growth factor deprivation, and exposure to pore-forming toxins (Imre et al., 2012; Ribe et al., 2012; Tizon et al., 2010; Tu et al., 2006; Vigneswara et al., 2012). This technique was also used to examine caspase-2 activation provoked in vitro by incubating cell extracts at 37°C (Read et al., 2002; Tinel and Tschopp, 2004). A sophisticated imaging technique, bimolecular fluorescence complementation (BiFC), can offer spatial information about caspase activation within cells and reveals the proportion of treated cells in which a particular caspase is activated. In this method, amino and carboxyl halves of the fluorescent protein Venus are each fused to the prodomain of the caspase under investigation, so that Venus fluorescence is only emitted when the procaspase monomers are closely associated, as occurs in complexes that prompt activation of initiator caspases (Bouchier-Hayes et al., 2009). This approach has been used to monitor the subcellular site of caspase-2 activation following various stimuli (Ando et al., 2012; Bouchier-Hayes et al., 2009; Imre et al., 2012; Parsons et al., 2016). Two groups generated caspase-2-knockout mice in which an antibiotic resistance cassette replaced a region of the gene encompassing the exon encoding the enzyme’s active site (Bergeron et al., 1998; O’Reilly et al., 2002; Fig. 1). These animals have been widely used to study the in vivo consequences of caspase-2 deficiency on development, aging, and numerous diseases. These mice have been immensely valuable for researching caspase-2 roles; however, it is possible that the constitutive deficiency of

Fig. 1 Genomic environment surrounding the murine caspase-2 gene (around 70 kb), obtained from https://www.ncbi.nlm.nih.gov/gene/12366. The thick blue line shows the caspase-2 gene. Rectangles depict exons, with noncoding portions colored gray. The exon deleted in the caspase-2-knockout mice is indicated with a red cross. Genes neighboring caspase-2 (NM_007610.2) are denoted by thick lines of other colors: GstK1, glutathione S-transferase kappa 1 (NM_029555.2); Tmem139, transmembrane protein 139 (NM_175408.4); Clcn1, chloride channel voltage-sensitive 1 (NM_0.1491.2).

160

M.A. Miles et al.

caspase-2 throughout development and adult life could lead to compensatory changes in expression of other genes, like caspase-9 (Troy et al., 2001), thus obscuring functions of caspase-2. Strain-dependent variability in disease susceptibility can also complicate experimental design and interpretation, particularly when investigating the impact of caspase-2 status on diseases modeled using transgenic mice created on different genetic backgrounds. The impact of the genomic manipulation performed to delete part of the caspase-2 gene could also theoretically affect the expression of nearby genes such as GstK1, Tmem139, or Clcn1, which lie within 40 kb of the region altered to disable caspase-2 (Fig. 1). “Casp2C320S” mice have recently been created using CRISPR/Cas9 technology to precisely replace the active-site cysteine of caspase-2 with serine, without affecting the expression profile of caspase-2 or surrounding genes (Dawar et al., 2016a). As discussed later (Section 4.8) splenocytes from these animals exhibited enhanced aneuploidy, mirroring that seen in caspase-2-knockout cells (Dawar et al., 2016a), confirming that the elevated rates of aneuploidy detected in cells from caspase-2-null mice results from loss of caspase-2 activity, and excluding possible confounding explanations such as perturbation in expression of a neighboring gene. In the future, the Casp2C320S mice could enable researchers to verify the specificity of other phenotypes attributed to loss of caspase-2 on the basis of experiments using the knockout mice.

2.3 Cellular Substrates Caspase-2 is a protease, so it is almost certain that its biological activity would be due to its cleavage of one or more protein substrates. Its residence within the nucleus, Golgi, and mitochondria (reviewed by Kitevska et al., 2009) would enable access to substrates located in these organelles. Prior to 2012, around a dozen proteins had been reported to be sensitive to cleavage by caspase-2 (Kitevska et al., 2009); however, almost all of these cleavage events could also be mediated by caspase-3, and the efficiency of cleavage by caspase-2 was typically not assessed. Notable exceptions were CUX-1 (Truscott et al., 2007) and Golgin-160 (Mancini et al., 2000), which were both cleaved by caspase-2 with efficiencies greater than 104 M1 s1, consistent with physiological relevance (Timmer et al., 2009; Table 1). Processing of the transcription factor CUX-1 by caspase-2 and other caspases (Table 1) removed two repressive domains at the carboxyl terminus of the protein (Truscott et al., 2007). Four possible caspase cleavage sites were identified in the region of CUX-1 between its homeodomain and

Table 1 Substrates Cleaved Relatively Specifically and/or Efficiently by Caspase-2, Ranked by Cleavage Efficiency Substrate

Cleavage Site #

kcat/Km (M–1 s–1)

Site Cleaved More Efficiently by Other Caspases?

References

Yes, cleaved more efficiently by caspases-3 (30) and -8 (9 ), -6 (3 ), and by caspase-7 with similar efficiency

Truscott et al. (2007)

CUX-1

SEGD S and/or DSCD#G

6.2  10

Prothymosin α (ProTα)

M(init)SD#A

1.5  105

No, caspase-2 cleaved 65 more efficiently than caspase-3

Julien et al. (2016)

Golgin-160

ESPD#G

3.3  104

No, but caspase-3 cleaves elsewhere 3 more efficiently

Mancini et al. (2000)

Nucleosome assembly protein 1-like 4 (NAP1L4)

SFSD#G

1.3  104

No, caspase-2 cleaved 6.5  more efficiently than caspase-3

Julien et al. (2016)

Protein transport protein Sec16A

VHPD#S

8.3  103

No, caspase-2 cleaved 4 more efficiently than caspase-3

Julien et al. (2016)

DRAD#S

5.4  103

No, caspase-2 specific Yes, caspase-8 cleaved 7 more efficiently

Guo et al. (2002) and Kitevska et al. (2014)

#

5

Bid

LQTD G

3.2  10

Rho GDI 2

DTKD#G

2.6  103

Yes, caspase-3 cleaved 2.5  more efficiently

Julien et al. (2016)

3

No, caspase-2 specific

Julien et al. (2016)

#

3

Guanine nucleotide-binding protein-like 1

DIND G

2.0  10

Scaffold attachment factor B2

DSRD#G

2.0  103

No, caspase-2 specific

Julien et al. (2016)

Runx1

DVPD#G

1.8  103

No, caspases-3 and -8 cleaved more efficiently at other sites

Kitevska et al. (2014)

BAG-6

DEQD#G

1.6  103

No, caspase-2 specific

Julien et al. (2016)

1.3  10

No, caspase-2 specific

Julien et al. (2016)

Transcriptional-regulating factor 1

#

DTRD G

3

Continued

Table 1 Substrates Cleaved Relatively Specifically and/or Efficiently by Caspase-2, Ranked by Cleavage Efficiency—cont’d Substrate

eIF-4H

Cleavage Site #

DEPD A #

kcat/Km (M–1 s–1)

Site Cleaved More Efficiently by Other Caspases?

References

1.3  10

3

Yes, caspase-3 cleaved 50 more efficiently

Julien et al. (2016)

3

Yes, caspase-3 cleaved 8.5  more efficiently

Julien et al. (2016)

Histone deacetylase 6

DMAD S

1.3  10

Holliday junction recognition protein

DRTD#G

1.2  103

Yes, caspase-3 cleaved 8.3  more efficiently

Julien et al. (2016)

Deubiquitinating protein VCIP135

ETTD#G

1.2  103

No, caspase-2 cleaved 2 more efficiently than caspase-6

Julien et al. (2016)

Ubinuclein-1

DESD#S

1.1  103

Yes, caspase-3 cleaved 11 more efficiently and caspase-7 8 more efficiently

Julien et al. (2016)

Scaffold attachment factor B2

DGTD#G

1.1  103

No, caspase-2 cleaved 13 more efficiently than caspase-6

Julien et al. (2016)

Protein PRRC2B

DQAD#S

1.1  103

No, caspase-2 cleaved 2.6  more efficiently than caspase-6

Julien et al. (2016)

Serine/arginine-related protein 53

IESD#S

1.1  103

Yes, more efficiently cleaved by caspases-8 (2.2 ), -3 (1.8 )

Julien et al. (2016)

GEM-interacting protein

DTSD#G

1.1  103

No, caspase-2 specific

Julien et al. (2016)

No, caspase-2 specific

Julien et al. (2016)

#

Ral GTPase-activating protein subunit alpha-1

TVKD G

1.1  10

C18orf25

VQKD#G

1.0  103

No, caspase-2 cleaved 2 more efficiently than caspase-6

Julien et al. (2016)

eIF-4B

DRKD#G

Not determined

No, caspase-2 cleaved more efficiently than caspases-7 (1.4 ) and -3 (3)

Wejda et al. (2012)

MDM2

DVPD#C

Not determined

Caspase-2 cleaved somewhat more efficiently than caspase-3

Oliver et al. (2011)

3

Old and Novel Functions of Caspase-2

163

the first repressive domain, but mutational experiments suggested a region spanning two possible sites (1333SEGD#SCD#G1340) contributed strongly to cleavage (Truscott et al., 2007). CUX-1 can act as a transcriptional activator or repressor, and abolition of cleavage eliminated both functions. A CUX-1 mutant lacking the caspase cleavage sites failed to induce reporter gene expression from promoters of DNA polymerase α, cyclin A2, DHFR, CAD, or B-myb and failed to prevent transcription from the p21Waf1/Cip1 promoter (Truscott et al., 2007). Many CUX-1 target genes regulate cell cycle checkpoints, and overexpression of full-length CUX-1 or a truncation mutant mimicking the cleaved form stimulated DNA synthesis, whereas a variant lacking the cleavage sites did not alter cell cycle progression (Truscott et al., 2007). Although the specific role of caspase-2 in CUX-1 cleavage has not been formally demonstrated, circumstantial evidence suggests its involvement: caspase-2 is present in the nucleus where CUX-1 resides, cleavage occurred in cells lacking caspases-3 or -8, and cleaved CUX-1 was detected during S-phase of the cell cycle in nonapoptotic cells (Truscott et al., 2007). Since publication of the ability of caspases to cleave CUX-1, important insights have been published about the biological roles of this substrate (Hulea and Nepveu, 2012; Ramdzan and Nepveu, 2014). Intriguingly, many of the functions attributed to CUX-1 intersect with processes reportedly regulated by caspase-2, although in most cases CUX-1 and caspase-2 have opposing effects, arguing against a model in which caspase-2mediated cleavage derepresses CUX-1. Both proteins have been documented to regulate the cell cycle, although CUX-1 cleavage facilitated S-phase entry (Truscott et al., 2007), whereas caspase-2 hindered exit from G1 or G2 (see Section 4.7). CUX-1 enabled base excision repair of oxidative DNA damage, preventing senescence of cells expressing activated Ras, which promotes this damage (Ramdzan et al., 2014). As summarized in Section 4.7, caspase-2 instead tends to promote senescence. Surprisingly, CUX-1 seemed to directly facilitate recruitment of 8-oxoguanine DNA glycosylase 1 to DNA bearing 8-OHdG oxidative lesions, independent of its transcription factor function (Ramdzan et al., 2014, 2015). CUX-1 was also observed to contribute to aneuploidy, through induction of genes that promote multipolar divisions and the survival of tetraploid, genomically unstable cells (Sansregret et al., 2011). In contrast, loss of caspase-2 was associated with aneuploidy (as discussed in Section 4.8). Although CUX-1 cooperated with activated Ras in oncogenesis, tumor suppressor roles have also been assigned to this protein: loss-of-function CUX-1 mutations were identified in 1%–5% of human cancers, and transposon-mediated

164

M.A. Miles et al.

lymphomagenesis frequently involved insertions into the CUX-1 locus (Wong et al., 2014). CUX-1 inhibited tumorigenesis in that model by transcriptional induction of PIK3IP1, a protein that antagonizes PI3K activity (Wong et al., 2014). The impact of caspase-2 status or caspase-mediated cleavage on these recently described functions of CUX-1 has not been examined to date. Golgin-160 is a Golgi-resident protein which may play a structural role within this organelle. It was highly sensitive to cleavage by caspase-2, but caspases-3 and -7 also cleaved Golgin-160 at two other sites (Mancini et al., 2000). Cleavage at the site preferred by caspase-7 could be blocked by phosphorylation (Turowec et al., 2014), but the impact of phosphorylation on sensitivity to caspase-2-mediated proteolysis has not been explored. Probably because a proportion of caspase-2 (unlike caspases-3 and -7) resides in the Golgi, Golgin-160 cleavage was initially caspase-2 dependent, although processing at other sites was detected later in apoptosis, presumably as the dismantling of the cell enabled caspases-3/-7 to access and cleave this substrate (Mancini et al., 2000). Golgin-160 processing was detected in cells exposed to the Alzheimer’s disease-associated peptide Aβ25–35 (Viana et al., 2010). A variant of Golgin-160 that retained the caspase-3 cleavage site but lacked the caspase-2 site delayed apoptotic Golgi disassembly (Mancini et al., 2000), confirming that cleavage contributes to this process; however, the biological significance of caspase-2-mediated Golgi destruction is currently unknown. A triple mutant that prevented cleavage by all caspases rendered cells resistant to apoptosis triggered by death receptor ligation or endoplasmic reticulum (ER) stress, but did not inhibit cell death triggered by exposure to inhibitors of kinases, protein synthesis, or topoisomerase II (Maag et al., 2005). The BH3-only protein Bid was another caspase-2 substrate identified many years ago (Guo et al., 2002). In comparison to CUX-1 and Golgin-160, caspase-2 cleavage of Bid was fairly inefficient (Kitevska et al., 2014). This still may be a biologically relevant event; however, as cleavage of a minority of Bid molecules within a cell may be sufficient to stimulate Bax/Bak activation and mitochondrial outer membrane permeability, leading to apoptosis (Zha et al., 2000), depending on the concentrations of other pro- and antiapoptotic Bcl-2 family members in the cell. It should be noted that caspase-8 cleaved Bid seven times more efficiently than caspase-2 (Kitevska et al., 2014), so Bid cleavage would be only expected to provoke apoptosis in a caspase-2-dependent manner in cells or contexts where caspase-8 is not expressed or not activated. MDM2, a negative

Old and Novel Functions of Caspase-2

165

regulator of p53, was also reported to be sensitive to caspase-2-mediated proteolysis (Oliver et al., 2011). Caspase-3 could also cleave MDM2 in vitro, but somewhat less efficiently (Oliver et al., 2011). Although the biochemical efficiency of caspase-2-mediated cleavage was not examined, MDM2 processing following doxorubicin treatment of osteosarcoma U2OS cells was reduced when caspase-2 was downregulated (Oliver et al., 2011), implying that the efficiency of cleavage was sufficient to be biologically meaningful and was not attributable to caspase-3 in these cells treated with this stimulus. Two comprehensive proteomics studies subsequently identified additional caspase-2 substrates and cleavage sites, by incubating lysates of A549 cells (Wejda et al., 2012) or Jurkat cells (Julien et al., 2016) in vitro with recombinant caspase-2 or other caspases. Julien et al. tracked the individual processing events over time, which enabled estimation of cleavage efficiencies and, importantly, confirmed that the proteolysis was directly attributable to caspase-2, rather than due to a downstream caspase that was activated by the exogenous caspase-2 (Julien et al., 2016). Nineteen cleavage events were mediated solely by caspase-2, and a further 30 sites were cleaved by caspase-2 plus either caspases-3, -6, -7, and/or -8. Physiologically relevant caspase cleavage events tend to feature kcat/Km values of over 10,000 M1 s1 (Timmer et al., 2009). Only two of the caspase-2 substrates identified by Julien et al. were cleaved this efficiently by caspase-2 (Table 1): Prothymosin α (ProTα) and nucleosome assembly protein 1-like 4 (NAP1L4). The small acidic protein ProTα was cleaved very efficiently by caspase-2, at a rate 60 times greater than by caspase-3. Previous work had established that caspase-2 cleaved pentapeptides much more efficiently than tetrapeptides (Talanian et al., 1997) reportedly due to required interactions of the P5 amino acid with Threonine380 and Tyrosine420 of the caspase (Tang et al., 2011), so it was surprising that caspase-2 was able to so potently remove the first three residues of ProTα (or the first two residues, if the initiating methionine is removed posttranslationally). Research to verify this unexpected proteolytic event is warranted, because research implicates ProTα in many of the biological processes reported to involved caspase-2 (described later). ProTα was reportedly essential for mitosis and its levels correlated with proliferative rate (Segade and Gomez-Marquez, 1999), possibly reflecting a role in mitotic spindle organization (Vareli and Frangou-Lazaridis, 2004). It was overexpressed in cancers and has been considered a useful biomarker (Ioannou et al., 2012), but appeared to lack direct

166

M.A. Miles et al.

oncogenic activity (Naylor et al., 1996). ProTα regulated DNA condensation to modulate gene expression through histone binding (Karetsou et al., 2004), and a phosphorylated form was shown to exhibit antiapoptotic activity (Moreira et al., 2013), possibly through apoptosome inhibition (Qi et al., 2010). ProTα regulated the Nrf2-Keap1-mediated response to oxidative stress (Karapetian et al., 2005), and ProTα transgenic mice developed insulin resistance via a pathway involving TLR4 and NF-κB (Su et al., 2015). It should be noted that some of the many functions attributed to ProTα are somewhat controversial (Pineiro et al., 2000), so any link between this candidate substrate, caspase-2, and these diverse biological processes would have to be carefully interrogated. Interestingly, Wejda et al. reported that caspases-2 and -3 could process this protein at an alternative site (EEED#G) but that caspase-7 cleaved at that position around five times more efficiently. Given that this protein was evidently present in the lysates used by both groups, it is surprising that they identified distinct cleavage sites within it. This may reflect differences between Jurkat and A549 cells regarding ProTα’s conformation or interactions with binding partners that could affect caspase access to particular sites. Another possibility is that some of the apparently caspase-2-mediated cleavage events detected by Wejda et al. may have been caused by caspase-7, if this enzyme was activated in the lysates by recombinant caspase-2. Although initial data suggested caspase-2 were incapable of efficiently processing executioner caspases (Van De Craen et al., 1999), subsequent work revealed that caspase-2 could cleave caspase-7 in vitro and this led to its activation in yeast (Ho et al., 2005). The third most efficiently cleaved caspase-2 substrate, NAP1L4, is a histone-binding protein previously documented to regulate nucleosome assembly and disassembly (Okuwaki et al., 2010). The caspase-2 cleavage site identified by Julien et al. lies between residues 8 and 9. Although the functional significance of removing the first eight residues of this protein is unknown, a 49-amino acid amino-terminal truncation significantly reduced the protein’s activity (Okuwaki et al., 2010), so it is conceivable that caspase-2 cleavage in this vicinity may also impair function. Over a third of the caspase-2 cleavage sites identified by Julien et al. were resistant to proteolysis by caspases-3, -6, -7, or -8 (Julien et al., 2016); however, Wejda et al. failed to find any caspase-2-specific substrates in their proteomic analysis (Wejda et al., 2012). Only one protein, eukaryotic translation initiation factor 4B, was more efficiently cleaved by caspase-2 than caspases-3 and/or -7 in the A549 lysates analyzed by Wejda et al.

Old and Novel Functions of Caspase-2

167

(2012). Although this theoretically may reflect actual biological differences between Jurkat vs A549 cells, it seems more likely that methodological variations are responsible. Julien et al. lysed their cells using Triton X-100. Over a third of the caspase-2 substrates they identified, and many of those preferentially cleaved by caspase-2, were nuclear. In contrast, Wejda et al. cited a freeze–thaw lysis method that involved freeze–thawing cycles of cells in isotonic buffer (Demon et al., 2009). It seems possible that this method may have not extracted nuclear proteins as efficiently as detergent-based methods.

3. ACTIVATION 3.1 Cleavage Executioner apoptotic caspases, like mammalian caspases-3 and -7, bear short prodomains amino-terminal to the subunits that comprise the active enzymes. These zymogens are activated by proteolysis, which separates the subunits and enables them to adopt a heterotetramer conformation that forms the active site and permits substrate cleavage. In contrast, caspases with long prodomains, like caspases-8 (Schleich et al., 2016) and -9 (Hu et al., 2014), are activated by recruitment into molecular complexes that facilitate conformational changes to generate active proteases. Intersubunit cleavage typically occurs and can boost activity (Oberst et al., 2010), but it is not required. Although caspase-2 also has a long prodomain (called a caspase activation and recruitment domain, CARD), yeast and in vitro studies revealed that proteolytic dissociation of the prodomain, large, and small units was important to achieve enzymatic activity: mutation of human caspase-2 to prevent separation of the subunits yielded variants that were much less active than the wild-type protein (Ho et al., 2005). Equivalent mutagenesis of the murine orthologue had a less detrimental impact on activity (Baliga et al., 2004; Butt et al., 1998). Caspases-3 and -8 could efficiently cleave the caspase-2 precursor (Van De Craen et al., 1999) so it is possible that cleavage-mediated activation could occur in vivo, although cells containing high levels of active caspases-3 or -8 would probably, although not necessarily (Feinstein-Rotkopf and Arama, 2009), be doomed to die, so this activation mechanism would be unlikely to account for caspase-2-dependent biological processes. Indeed, caspase-9-dependent processing of caspase-2 processing was observed in irradiated thymocytes, but caspase-2 deficiency did not inhibit their apoptosis (O’Reilly et al.,

168

M.A. Miles et al.

2002), implying that although cleavage of caspase-2 occurred, this did not contribute to cell death in this context. It seems probable that caspase2-dependent outcomes would result from initial activation of this protease in cells devoid of other active caspases.

3.2 Induced Proximity Activates Caspase-2 Within the PIDDosome The most obvious caspase-2 activation mechanism that would not require cleavage as an initiating step involves CARD-mediated assembly of a molecular complex, within which caspase-2 could be forced to adopt an active conformation. Almost two decades ago, a homotypic interaction was documented between the CARD domains of caspase-2 and the adaptor protein RAIDD (Duan and Dixit, 1997), a protein recently implicated in cerebral cortex development (Di Donato et al., 2016). RAIDD could also interact with the kinase RIPK1 (Duan and Dixit, 1997), but RAIDD was not detected in the “ripoptosome” complex through which RIPK1 promotes apoptosis (Tenev et al., 2011). It emerged that RAIDD could bind to a p53-inducible protein, PIDD (Lin et al., 2000; Telliez et al., 2000; Tinel and Tschopp, 2004). A recent structural study revealed that a layer of five PIDD death domains could interact with a layer of five RAIDD proteins, with or without 1–2 additional loosely bound RAIDD monomers on top of the PIDD layer (Nematollahi et al., 2015). This interaction apparently liberates the CARD of RAIDD (Jang and Park, 2013), enabling it to recruit procaspase-2 into a complex termed the “PIDDosome” (Tinel and Tschopp, 2004), within which caspase-2 acquires catalytic activity (Fig. 2). PIDD possesses a potent autocleaving activity that initially removes an amino-terminal inhibitory domain-containing leucine-rich repeats (LRRs) and the first of two ZU-5 domains, producing a protein referred to as “PIDD-C” (Tinel et al., 2007). It is possible, by analogy with other LRR-containing proteins that participate in the inflammasome (Franchi et al., 2012), that this autocleavage event may be regulated through molecular interactions with PIDD’s LRRs, but no such interaction has been reported as yet, and in most cells, PIDD is constitutively autoprocessed. Mutational abolition of this cleavage event prevented the nuclear translocation of PIDD that otherwise occurred after DNA damage (Tinel et al., 2007). The PIDD-C fragment can subsequently autoprocess at a second site to discard the remaining ZU-5 domain, yielding a shorter product

Old and Novel Functions of Caspase-2

169

Fig. 2 PIDDosome-regulated caspase-2 activation. Events controlling the formation of the PIDD–RAIDD–caspase-2 and the PIDD–RIPK1–Nemo complexes are summarized. See Sections 3.2 and 4 of the text for details and references.

designated “PIDD-CC,” to which RAIDD could bind and to which PIDD’s caspase-2-activating function has been ascribed (Tinel et al., 2007). The PIDDosome was identified via experiments in which cell lysates were incubated at 37°C (Tinel and Tschopp, 2004). Caspase-2 was initially detected in low-molecular-weight lysate fractions, yet it appeared in higher-molecular-weight fractions, and became active, after the extracts were heated (Read et al., 2002; Tinel and Tschopp, 2004). PIDD and RAIDD were identified within these fractions (Tinel and Tschopp, 2004), yet caspase-2 activation and its shift to high-molecular-weight fractions could also be detected in warmed extracts from cells lacking PIDD (Manzl et al., 2009). Thus, PIDD is not required for activation of caspase-2 or its participation in a high-molecular-weight complex in this (quite artificial) context. Caspase-2 activation does appear to require RAIDD and PIDD in some biological settings (as outlined later). However, other caspase-2-dependent outcomes, discussed later, have been published to be PIDD independent. Thus, the PIDDosome appears unlikely to be the only platform that can provoke the activation of caspase-2.

170

M.A. Miles et al.

4. CASPASE-2-MEDIATED RESPONSES TO DNA DAMAGE AND MITOTIC STRESS 4.1 DNA Damage-Induced Caspase-2 PIDDosome Formation PIDD-dependent caspase-2 activation following DNA damage required phosphorylation of Thr788 within the RAIDD-binding region of PIDD-CC (Ando et al., 2012). Fluorescence microscopy with a phospho-specific antibody revealed that this phosphorylated form of PIDD colocated with chromosomes within cells bearing DNA damage (Thompson et al., 2015). This phosphorylation event could be performed by ATM (Ando et al., 2012), a kinase that facilitates cellular responses to DNA damage (Marechal and Zou, 2013; Fig. 2). A stable interaction was detected between ATM and the amino-terminal LRR-containing region of full-length PIDD (Ando et al., 2012). The significance of this interaction has not been determined, and recombinant GST-PIDD-CC (which lacks PIDD’s ATM-binding site) could be phosphorylated by ATM in vitro (Ando et al., 2012). However, if the association between LRR domain of PIDD and ATM enabled the ATM-mediated phosphorylation reaction within the carboxyl portion of PIDD, the interaction presumably occurred in the cytosol, prior to the rapid autoprocessing events that separate these fragments. It is tempting to speculate that a molecular race between PIDD autocleavage vs ATM-mediated phosphorylation may limit PIDDosome formation to cells bearing high levels of active ATM and/or cellular contexts that slow the rate of PIDD autocleavage.

4.2 CHK1 Inhibition of PIDDosome Formation and Its Regulation by p53 In cells lacking p53 function, irradiation-induced ATM-mediated PIDD phosphorylation and the resultant PIDDosome formation could be dramatically enhanced by G€ o6976 (Ando et al., 2012; Manzl et al., 2013), a drug that efficiently inhibits the kinases CHK1, MAPKAP-K1b, MSK1, PKCα, and PHK (Davies et al., 2000). It seems likely that CHK1 was the most important G€ o6976 target in this context, as downregulation of CHK1 expression also boosted caspase-2 activation following treatment with doxorubicin, gemcitabine, or irradiation (Carroll et al., 2016; Del Nagro et al., 2014; Sidi et al., 2008; Thompson et al., 2015). The mechanism by which CHK1 blocks Thr788 phosphorylation of PIDD has not yet been elucidated.

Old and Novel Functions of Caspase-2

171

The PIDDosome-repressing activity of CHK1 could, in at least some circumstances, be antagonized by p53 (Fig. 2). Doxorubicin treatment of wild-type p53 cells, but not p53 mutant or null cells, led to downregulation of CHK1 expression, boosting caspase-2 activation, and processing (Carroll et al., 2016). The mechanism by which p53 reduced CHK1 levels is unknown, but may involve p53-induced expression of CHK1-silencing microRNAs (Lezina et al., 2013). The ability of caspase-2 to proteolytically disable MDM2, which would otherwise promote p53 degradation (Oliver et al., 2011), may amplify this pathway by keeping p53 levels high and CHK1 levels low, hence reinforcing PIDD-CC generation and caspase-2 activity in cells where DNA damage stimulates ATM activation (Terry et al., 2015). Caspase-2 has also been reported to enhance p53 phosphorylation and activity in a distinct, MDM2-independent manner, which has not yet been fully defined (Dorstyn et al., 2012).

4.3 Inconsistencies Regarding Outcomes of PIDDosome Signaling and Its Inhibition by CHK1 PIDDosome activation, with or without CHK1 inhibition, can trigger apoptosis, but the frequency of this outcome seems to vary between cell types. Only around a 10th of zebrafish cells lacking p53 and CHK1 function were apoptotic after irradiation (Sidi et al., 2008). In MCF7, HCT116, SAOS2, MDA-MB-435, and LN428 cells, doxorubicin or irradiation coupled with CHK1 inhibition only triggered apoptosis in around a fifth of the cells (Ando et al., 2012; Carroll et al., 2016; Sidi et al., 2008). Primary p53-null murine embryonic fibroblasts (MEFs) were moderately sensitive to irradiation-induced apoptosis, regardless of caspase-2 status or G€ o6976 cotreatment (Manzl et al., 2013). HeLa cells were somewhat more efficiently killed, with between a quarter and two-thirds dying after irradiation plus CHK1 inhibition (Ando et al., 2012; Manzl et al., 2013; Sidi et al., 2008). Irradiation-induced killing of p53-deficient murine thymic lymphoma cells was ameliorated by genetic deletion of caspase-2 and augmented by cotreatment with G€ o6976 (Manzl et al., 2013); however, the caspase-2 status of primary thymocytes and T and B cell blasts had very little impact on their sensitivity to irradiation with or without G€ o6976, although irradiation did stimulate processing of caspase-2 (and -3) (Manzl et al., 2013). Irradiation killed p53-deficient HCT116 cells more efficiently than those expressing p53, whether or not caspase-2 was downregulated (Manzl et al., 2013). G€ o6976 did not substantially influence radiation sensitivity in p53-proficient or -deficient HCT116 cells, but did cooperate with

172

M.A. Miles et al.

irradiation to promote caspase-2 processing in cells bearing and lacking p53 (Manzl et al., 2013). G€ o6976 killed around a third of unirradiated p53deficient cells in which caspase-2 was downregulated (Manzl et al., 2013). There appears to be general consensus that irradiation coupled with CHK1 inhibition or downregulation can promote ATM-mediated PIDDosome formation, which results in caspase-2 activation. Whether or not p53 proficiency obviates the necessity for CHK1 inhibition is less clear and may be cell type dependent. The biochemical and cellular consequences of the caspase-2 activity provoked by this pathway are currently unclear, and the propensity for this pathway to induce apoptosis is highly variable.

4.4 PIDDosome Inhibition by BubR1 During mitosis, PIDDosome formation in response to DNA damage was also subject to inhibition by BubR1 (Thompson et al., 2015), a component of the mitotic checkpoint complex. ATM-phosphorylated PIDD was sequestered at kinetochores during prophase by BubR1, preventing recruitment of RAIDD and impeding caspase-2 activation (Thompson et al., 2015). Silencing of BubR1 sensitized HeLa cells to irradiation-induced apoptosis about half as efficiently as treatment with G€ o6976 (which inhibits CHK1, among other kinases) via a mechanism that required caspase-2, RAIDD, and PIDD (Thompson et al., 2015). Thompson et al. proposed that BubR1 inhibited PIDDosome formation by competing with RAIDD for binding to PIDD-CC (Fig. 2). Intriguingly, caspase-2 had previously been detected at the centrosome of mitotic cells (Narine et al., 2010), but its activation status at this site was not explored.

4.5 An Alternative PIDD Complex Induces NF-κB PIDD can also participate in at least one other molecular complex, apart from the caspase-2-activating PIDDosome. Via an interaction with RIPK1, PIDD can activate Nemo, leading to NF-κB activation and induction of many genes including some involved in DNA repair, survival, and inflammation (Janssens et al., 2005). In contrast to the caspase-2-activating platform, which uses the RAIDD-binding PIDD-CC fragment, this NF-κ B-activating pathway required the PIDD-C fragment (Tinel et al., 2007; Fig. 2). An unphosphorylatable mutant of PIDD (PIDDT788A) was unable to bind to RAIDD but interacted more strongly with RIPK1, suggesting that the extent of PIDD phosphorylation at Thr788 determined whether DNA damage elicited caspase-2 activation or NF-κB activation (Ando

Old and Novel Functions of Caspase-2

173

et al., 2012). CHK1 activity was reported to suppress PIDD phosphorylation at Thr788, thereby favoring NF-κB activation (Ando et al., 2012; Carroll et al., 2016). The participation of PIDD in distinct molecular complexes that exert dramatically different biological effects can confound researchers’ attempts to discern its involvement in caspase-2-dependent processes, because the absence of PIDD (through genetic deletion or knockdown) could alter caspase-2-mediated and/or caspase-2-independent pathways. In addition, alternative splicing could produce transcripts encoding PIDD proteins that would be predicted to activate NF-κB but not induce apoptosis (Cuenin et al., 2007). If these alternative isoforms are translated in certain cell types, they may skew the balance of PIDD-directed pathways in favor of Nemo-regulated NF-κB activation rather than RAIDD/caspase-2-mediated outcomes.

4.6 Phosphorylation in Prodomain or Linker Prevents Caspase-2 Activation Direct phosphorylation of caspase-2 at three sites has been reported to suppress its activation (Fig. 3). In xenopus oocytes, CaMKII-mediated

Fig. 3 Regulation of caspase-2 activation by phosphorylation. Phosphorylation of three residues of caspase-2 has been reported to inhibit its activation. See Section 4.6 of the text for details and references. Residue numbering is for human caspase-2, but note that phosphorylation at residue 164 has not been demonstrated in mammalian cells.

174

M.A. Miles et al.

phosphorylation within the caspase-2 prodomain discouraged RAIDD binding when NADPH levels were high (Nutt et al., 2005). This phosphorylation was reversed by the phosphatase PP1, but dephosphorylation was suppressed by binding of 14-3-3ζ to the phosphorylated caspase-2 prodomain (Nutt et al., 2009). The prodomain of human caspase-2 could also be phosphorylated at Ser164 (Mccoy et al., 2012) by purified CaMKII and by xenopus oocyte extracts (Nutt et al., 2005), but whether this occurs within human cells has not been established. Likewise, phosphorylation by PKCK2 at a nearby site within the prodomain, Ser157, also impaired activation (Shin et al., 2005). Cdk1–cyclin B1 was responsible for phosphorylating caspase-2 during mitosis on a separate residue, Ser340, which is located within the linker that separates the large and small catalytic subunits within the procaspase zymogen (Andersen et al., 2009). PP1 could also dephosphorylate this residue (Andersen et al., 2009). Caspase-2 downregulation by siRNA suppressed the apoptotic effects of the microtubule poison nocodazole but enforced expression of an unphosphorylatable S340A mutant exacerbated the apoptotic effect of this treatment (Andersen et al., 2009). Hence phosphorylation-mediated oscillation in caspase-2 activity through the cell cycle seems to afford some apoptotic protection to healthy mitotic cells, but defects in architecture of the microtubule spindle may trigger caspase-2-mediated apoptosis. The precise mechanism connecting spindle defects to caspase-2 activation is still somewhat obscure. The simplest explanation—that mitotic poisons like nocodazole stimulate dephosphorylation of caspase-2 at Ser340—was not experimentally supported, as nocodazole exposure promoted rather than reduced phosphorylation at this site (Andersen et al., 2009).

4.7 Caspase-2-Mediated Cell Cycle Arrest vs Apoptosis, Following DNA Damage or Mitotic Stress Fibroblasts from caspase-2-deficient mice proliferated at a faster rate than those from wild-type animals (Dorstyn et al., 2012; Ho et al., 2009) and a greater proportion of tumor cells from knockout mice than wild-type animals were in S and G2/M phases of the cell cycle (Manzl et al., 2012; Parsons et al., 2013). Consistent with the notion that caspase-2-deficiency alters cell cycle control, a greater proportion of knockout than wild-type cells recommenced cycling 24 h after reintroduction of serum following a period of serum deprivation (Parsons et al., 2013). Loss of caspase-2 enabled cells to

Old and Novel Functions of Caspase-2

175

avoid senescence triggered by in vitro passaging of untransformed MEFs (Dorstyn et al., 2012) or enforced coexpression of MEK and TERT (Gitenay et al., 2014). In contrast, more senescent cells were detected in liver sections obtained from aged (24–26 months old) caspase-2-null mice than wild-type animals of equivalent age (Shalini et al., 2012), a difference postulated to result from lifelong exposure to excessive intracellular oxidative stress (see Section 6). The antiproliferative activity of caspase-2 was more pronounced following DNA damage: unlike wild-type cells, caspase-2-deficient cells continued to incorporate BrdU after irradiation (Ho et al., 2009). When cells were untreated, or serum deprived then refed, more caspase-2-deficient cells than wild-type cells were in S or G2/M cell cycle phases (Parsons et al., 2013). However, following 5-fluorouracil or nocodazole treatment, more caspase-2-deficient cells were tetraploid but fewer were synthesizing DNA, compared to wild-type cells (Parsons et al., 2013). These data suggest that caspase-2 retards exit from G1 or G2 in healthy cells but predominantly regulates a mitotic checkpoint in cells treated with nucleotide analogs or mitotic poisons. Caspase-2 activation can lead to apoptosis, possibly through proteolytic activation of Bid (Guo et al., 2002) and/or transcriptional upregulation of another BH3-only protein, Bim (Jean et al., 2013; Ribe et al., 2012). In p53-proficient cells, caspase-2-mediated MDM2 cleavage (Oliver et al., 2011) could prevent p53 degradation, permitting its induction of other proapoptotic Bcl-2 relatives, Puma and Noxa (Villunger et al., 2003). Accumulation of these proapoptotic BH3-only proteins could trigger intrinsic apoptosis, if insufficient antiapoptotic Bcl-2 relatives were present to prevent this (Fig. 4). The mechanism by which caspase-2 causes cell cycle arrest is less obvious. Caspase-2-mediated MDM2 inactivation could cause mitotic arrest due to p53-mediated induction of cyclin-dependent kinase inhibitor 21 (p21Waf1/Cip1) (Fig. 4). Indeed, caspase-2-deficient E1A/ Ras-transformed MEFs expressed less p21 than wild-type cells (Parsons et al., 2013) and caspase-2 silencing negated the ability of irradiation to boost p21Waf1/Cip1 levels in p53 wild-type HCT116 cells (Sohn et al., 2011). However, surprisingly, this was due to an effect on p21Waf1/Cip1 translation, not transcription (Sohn et al., 2011). p21Waf1/Cip1 transcripts were only slightly more highly induced by cisplatin in caspase-2-proficient than caspase-2-null lung tumor cells (Terry et al., 2015).

176

M.A. Miles et al.

Fig. 4 Cell death and cell cycle arrest downstream of caspase-2. Pathways leading to cell death (red lines) and/or cycle arrest (green lines) are shown. Black lines indicate steps that can lead to both outcomes, depending on the molecular context. See Sections 4.7 and 10.4 of the text for details and references.

4.8 Genomic Instability and Aneuploidy Due to Caspase-2 Inhibition As discussed earlier, BubR1 prevented ATM-mediated caspase-2 activation during prophase, when the mitotic spindle forms (Thompson et al., 2015), but caspase-2 has been strongly implicated in cell death resulting from defective mitotic spindle formation and activity. Caspase-2-knockout MEFs were resistant to death induced by the microtubule poisons vincristine, paclitaxel, or colchicine (Ho et al., 2008; Tiwari et al., 2014), which prevent normal segregation of chromosomes during mitosis, emphasizing that cell death due to cytoskeletal disruption requires caspase-2. Caspase-2 deficiency led to impaired DNA repair, genomic instability, and aneuploidy following irradiation in MEFs and lymphoma cells (Dorstyn et al., 2012). Increased rates of aneuploidy were also recently observed in primary splenocytes from caspase-2 null or Casp2C320S mice exposed to mitotic poisons (Dawar et al., 2016a) and in bone marrow cells from aged knockout mice (Dawar et al., 2016b). Caspase-2-dependent cell death was detected following treatment of HeLa syncytia with a CHK2 inhibitor (Castedo et al., 2004). When this “mitotic catastrophe” was blocked by suppressing caspase-2 expression or activity, aneuploidy resulted (Castedo et al., 2004). Upon mitotic arrest,

Old and Novel Functions of Caspase-2

177

around twice as many SV-40 immortalized MEFs derived from caspase-2null mice accumulated micronuclei compared with cells from wild-type mice (Peintner et al., 2015). Cells from RAIDD-knockout mice were as likely as those from wild-type mice to contain micronuclei (Peintner et al., 2015), however, implying that the mechanism by which caspase-2 prevents micronuclei formation (or eliminates cells containing them) is PIDDosome independent. Caspase-2 has been implicated in the degradation of cytoskeletal components: treatment with 5-fluorouracil, doxorubicin, or staurosporin led to caspase-2 processing and subsequent reduction in the cytoskeletal proteins tropomyosin, myotrophinin, profiling, and stathmin-1 in HCT116 cells, as the cells succumbed to caspase-2-mediated apoptosis (VakifahmetogluNorberg et al., 2013). Interestingly, this decrease was not due to direct cleavage of these proteins by caspase-2, but through targeting them for proteosomal degradation (Vakifahmetoglu-Norberg et al., 2013).

5. RESPONSES TO ER STRESS Caspase-2 downregulation or inhibition blocked Bid cleavage and protected cells from apoptosis triggered by thapsigargin or brefeldin A, agents that inhibit the ER calcium pump and ER–Golgi transport, respectively (Upton et al., 2008). Activation of caspase-2 by these agents was proposed to result from derepression of caspase-2 translation by ER transmembrane receptor (IRE1α) (Upton et al., 2012). Perplexingly, a separate group of researchers failed to reproduce these results, finding that caspase-2 deficiency did not modify the sensitivity of numerous cell types to ER stress triggered by thapsigargin or brefeldin A (Sandow et al., 2013). The basis for this seemingly direct discrepancy is unclear.

5.1 Brucella Infection-Mediated ER Stress Caspase-2 was required for macrophages and dendritic cells to die following infection with some strains of the bacterial species Brucella abortus (Chen and He, 2009; Li and He, 2012). Researchers recently dissected the mechanism by which caspase-2 modulates responses to ER stress provoked by Brucella infection. Bronner et al. triggered ER stress in bone marrow-derived macrophages via infection with an attenuated strain (RB51) of B. abortus (Bronner et al., 2015). This microbe replicates within vacuoles that commandeer membranes from the ER, provoking the reorganization of this

178

M.A. Miles et al.

organelle (Celli et al., 2003). The resulting ER stress promoted activity of IRE1α which led, via TXNIP upregulation, to NLRP3 translocation from the ER to mitochondria in infected cells (Bronner et al., 2015). NLRP3 is a major component of an inflammasome, a molecular complex that forms following recognition of molecules associated with pathogen infection or cellular damage, to promote activation of inflammatory caspases and production of cytokines such as IL-1β and IL-18 (Lamkanfi and Dixit, 2014). By an undefined mechanism, NLRP3 stimulated the mitochondrial translocation of caspase-2 and its cleavage. In cultured macrophages, caspase-2 was required for efficient cleavage of Bid following Brucella infection, release of mitochondrial DNA and cytochrome c via the mitochondrial permeability transition pore, caspase-1 activation and the resultant proteolytic maturation and release of IL-1β (Bronner et al., 2015). Intriguingly, in contrast to previous studies (Zhou et al., 2011), Bronner et al. demonstrated that movement of NLRP3 to mitochondria in the context of Brucella infection was a cause, rather than solely a consequence, of mitochondrial damage. Confirming the relevance of this pathway in vivo, levels of IL-1β in the serum of Brucella-infected mice were dramatically reduced in animals lacking caspase-2 (Bronner et al., 2015). To date, the impact of caspase-2 status on the natural history of unattenuated brucellosis has not been reported. Previous work established that infected macrophages ultimately succumb to caspase-2-dependent cell death (Bronner et al., 2013). If caspase-2 activity in initially infected cells leads to their destruction, caspase-2 may prevent propagation and hence “nip-in-the-bud” Brucella infections. On the other hand, inflammation is associated with brucellosis symptoms (Baldi and Giambartolomei, 2013), so caspase-2-mediated IL-1β production from infected cells may contribute to the pathogenicity of Brucella infections.

5.2 Rhabdovirus Infection-Mediated ER Stress Viruses that replicate in the cytoplasm, like rhabdoviruses, also dramatically perturb ER homeostasis (Romero-Brey and Bartenschlager, 2016). Infection with the Maraba rhabdovirus coupled with downregulation of IRE1α stimulated caspase-2 activation, as revealed by BiFC and caspase-2 processing, followed by cleavage of caspases-8, -9, -3, and PARP and death of the infected cells (Mahoney et al., 2011). The cells were protected by downregulation of caspase-2 or RAIDD, but PIDD silencing had no effect (Mahoney et al., 2011).

Old and Novel Functions of Caspase-2

179

6. RELIEVING OXIDATIVE STRESS 6.1 Aging Two independently generated strains of caspase-2-deficient mice achieved similar median life spans to wild-type animals (Shalini et al., 2012; Zhang et al., 2007), but the maximal life span in both caspase-2-knockout strains was substantially shorter than that of caspase-2-proficient mice, with around an eighth to a quarter of wild-type mice surviving after all caspase-2deficient animals had died (Shalini et al., 2012; Zhang et al., 2007). The knockout mice exhibited a suite of symptoms consistent with accelerated aging, including graying fur and loss of adipose tissue and bone density. Slight skewing of hemopoiesis during aging of caspase-2-null mice was also recently reported, with the bone marrow of these animals containing slightly more stem cells and myeloid progenitors than caspase-2-proficient elderly mice (Dawar et al., 2016b). Interestingly, this skewing during hemopoiesis did not alter the peripheral blood composition of aged caspase-2-knockout mice (Shalini et al., 2012). Aging is associated with proteomic and metabolomic alterations, including elevation in levels of enzymes that metabolize carbohydrates and amino acids but less abundant mitochondrial and ribosomal proteins. The proteome of young vs old wild-type mice differed more substantially than that of caspase-2-knockout mice of different ages (Wilson et al., 2015). Interestingly, the abundance of a number of proteins and metabolites detected in the liver and serum of young caspase-2-deficient mice resembled their profiles in older wild-type mice (Wilson et al., 2015), suggesting that the loss of caspase-2 leads to biochemical as well as physiological features of aging.

6.2 Amelioration of Oxidative Stress Although some of the age-related differences itemized above may relate to metabolic functions of caspase-2 (as discussed in Section 7), caspase-2deficient young mice, like older wild-type animals, had decreased expression of citrate synthase and OXPHOS complex III (Wilson et al., 2015), consistent with the notion that defective mitochondrial function and increased reactive oxygen species (ROS) may lead to some aging-related characteristics (Lopez-Cruzan and Herman, 2013). Caspase-2-knockout MEFs also exhibited higher levels of superoxide, peroxide, and carbonylated proteins than wild-type cells (Tiwari et al., 2014). The ability of caspase-2 to reduce

180

M.A. Miles et al.

oxidative stress appears to be PIDDosome dependent, as neuronal cell death induced by rotenone—an inhibitor of the electron transport chain that stimulates production of ROS—could be inhibited by cell-permeable peptides derived from RAIDD or PIDD (Jang et al., 2016). Other data support a general role for caspase-2 in orchestrating cellular responses to oxidative stress. The oxidizing herbicide paraquat provoked caspase-2 cleavage, and presumably activation, in the lungs of mice administered a single low dose (Shalini et al., 2015b). In wild-type mice, this treatment triggered upregulation of superoxide dismutase (SOD), which presumably limited the oxidation-induced damage. Mice lacking caspase-2 experienced slightly more severe pulmonary oedema following paraquat treatment, seemingly due to increased necrosis of the microvascular endothelium and epithelial cells lining the alveolar walls, and this necrosis was associated with elevated levels of inflammatory cytokines IL-6 and IL-1β in the lungs of the treated caspase-2-null mice (Shalini et al., 2015b). The authors postulated that caspase-2 enabled the cells to either sense and/or respond to oxidative stress, so its loss rendered cells and animals more sensitive to the effects of oxidative damage and secondary consequences such as inflammation (Shalini et al., 2015b). Suppression of caspase-2 expression led to downregulation of the mRNA encoding forkhead transcription factors FoxO1 and FoxO3a, and the consequent reduction in expression of target genes including the antioxidant enzymes SOD and glutathione peroxidase (GSH-Px) (Shalini et al., 2012). It seems likely that this antioxidant deficiency contributed to the elevated levels of ROS detected in cells lacking caspase-2. The mechanism by which caspase-2 regulates levels of FoxO1/3a mRNA has not been elucidated, but proteins implicated in controlling (particularly suppressing) expression of these transcription factors in response to ROS (Klotz et al., 2015) would be attractive candidate substrates whose cleavage by caspase-2 may account for its ability to help cells respond to oxidative stress.

6.3 Suppression of Autophagy The complex I inhibitor rotenone perturbs mitochondrial oxidative phosphorylation and provokes oxidative stress. Treatment of cortical neurons with rotenone provoked superoxide-dependent processing and activation of caspase-2 (Tiwari et al., 2011). Activation of caspase-2 was confirmed to be an initiating molecular event using bVAD trapping (see Section 2.2) (Tiwari et al., 2011). Treatment of wild-type neurons with

Old and Novel Functions of Caspase-2

181

rotenone provoked intrinsic apoptosis featuring Bid cleavage, cytochrome c release, and activation of Bax, caspases-9 and -3. In contrast, caspase-2deficient cells were totally protected during rotenone incubations up to 24 h. By 72 h, around half as many caspase-2-null cells had died as wild-type cells, and these dying caspase-2-deficient cells appeared necrotic (Tiwari et al., 2011). Autophagy was induced in rotenone-treated cells lacking caspase-2: LC3-II and Beclin-1 levels were elevated in these cells, and this effect could be reversed by prior exposure to the antioxidant N-acetylcysteine. The authors speculated that the ultimate necrotic death of caspase-2-deficient cells following prolonged rotenone treatment reflected autophagic processes being overwhelmed by oxidatively damaged cellular material. Evidence has also been published for an antiautophagic function of caspase-2 in vivo: an autophagy marker (GFP-LC3) was detected at higher levels in brain tissue from caspase-2-null mice than wild-type animals (Tiwari et al., 2014). Caspase-2 could also suppress autophagy in nonneuronal cells. Autophagic markers like LC3-II were somewhat elevated in fibroblasts, astrocytes, and osteoclasts lacking caspase-2, even under normal growth conditions (Tiwari et al., 2014). Rotenone, hydrogen peroxide, or heat shock were all substantially more toxic to wild type than caspase-2-null MEFs, but the caspase-2-knockout cells were sensitized to these stimuli by the autophagy inhibitor LY294002 (Tiwari et al., 2014). The ability of caspase-2 deficiency to promote autophagy was impeded by downregulation of the proautophagy kinases PRKAA1/2 or gene deletion of ATG5 and 7. Rapamycin-mediated inhibition of mammalian target of rapamycin (mTOR) promoted LC3-II expression in caspase-2-proficient cells (Tiwari et al., 2014). Consistent with the notion that ROS promote caspase-2-inhibitable autophagy, treatment with a superoxide scavenger suppressed autophagy markers in caspase-2-null cells to around levels seen in wild-type cells (Tiwari et al., 2014). A related phenomenon was recently reported in vivo, with autophagy occurring more prominently in the liver and muscle tissue of starved caspase-2-deficient mice than starved wild-type animals (Wilson et al., 2016).

6.4 Suppression of ROS-Driven Osteoclastogenesis Differentiation of monocyte/macrophage progenitors into osteoclasts, cells that reabsorb bone and counter the bone-forming function of osteoblasts, is largely driven by TNF-family cytokines such as RANKL (Katagiri and

182

M.A. Miles et al.

Takahashi, 2002) via a ROS-dependent mechanism (Ha et al., 2004). The ability of RANKL to promote ROS-dependent differentiation of osteoclasts from macrophages was substantially augmented by deletion or downregulation of caspase-2, with the later steps in osteoclastogenesis being more strongly influenced by caspase-2 levels than earlier stages (Callaway et al., 2015). Presumably this ability of caspase-2 to suppress ROS-driven osteoclastogenesis accounts for the observation that age-related loss of bone density was more pronounced in animals lacking caspase-2 (Shalini et al., 2012; Zhang et al., 2007).

7. METABOLISM The impact of nutritional conditions on caspase-2 activity was first observed in xenopus oocyte extracts. Spontaneous or RAIDD-induced processing of procaspase-2 in egg extracts was abolished by supplementation with metabolites of the pentose phosphate but not glycolytic pathway (Nutt et al., 2005). This inhibition was attributed to CaMKII-mediated phosphorylation of a residue within the CARD of procaspase-2 (Nutt et al., 2005; Fig. 3). A connection between caspase-2 and lipid levels was also documented around the same time: incubating HCT116 cells with lipid-lowering drugs activated sterol regulatory element-binding proteins, which induced caspase-2 transcription (Logette et al., 2005). The influence of caspase-2 status on metabolic pathways, particularly during aging, has received substantial additional research attention in the last few years. Livers from young caspase-2-null mice had lower NADPH levels, and their mitochondria had less oxidative phosphorylation complex III and more citrate synthase, compared to samples from wild-type animals of similar age (Wilson et al., 2015). As the mice aged, more marked differences in proteomic and metabolomic patterns became apparent between caspase-2proficient and -deficient animals. Some changes that normally occur with age were evident even in young caspase-2-null mice, reinforcing the notion that caspase-2 loss phenocopies accelerated aging. Interestingly, although older caspase-2-deficient mice were generally slightly less healthy than aged wild-type mice, they had lower fasting blood glucose levels than wild-type animals and better glucose tolerance (Wilson et al., 2015). The metabolic benefits of caspase-2 loss were exaggerated when mice were fed a diet high in fats and sugars: caspase-2-deficient mice were less likely than wild-type animals to develop conditions frequently suffered by humans who consume high fat/sugar (Western) diets, including diabetes

Old and Novel Functions of Caspase-2

183

mellitus, dyslipidemia, and hepatic steatosis (Machado et al., 2016). Wild-type mice that ate a “Western” diet had about twice the body fat of wild-type mice that ate regular chow, but caspase-2-null mice fed the high fat/sugar diet only had slightly more body fat than knockout mice that were fed chow (Machado et al., 2016). This effect was especially pronounced when epididymal fat was measured. Wild-type mice that ate chow exhibited similar glucose tolerance to caspase-2-null mice fed either chow or high-fat/high-sugar diet, but the wild-type animals that ate the “Western” diet were markedly insulin resistant (Machado et al., 2016). Adipocyte size increased markedly in normal mice fed the high-fat/high-sugar diet, but the adipocytes of caspase-2-null mice fed either diet remained small (Wilson et al., 2015). Caspase-2 deficiency also ameliorated the impact of a diet high in fat but not sugar. Consumption of high-fat diets boosted caspase-2 levels in adipose tissue of rats (Jobgen et al., 2009). Liver sections from patients suffering from nonalcoholic fatty liver disease contained unusually high levels of caspase-2, and the levels were higher still in individuals with more severe nonalcoholic steatohepatitis (NASH), and this mirrored the higher caspase-2 levels seen in multiple animal models of NASH (Machado et al., 2015). Elevated caspase-2 appeared to be a cause, rather than a consequence, of this condition: wild-type mice eating high-fat methionine choline-deficient diets developed characteristic NASH symptoms including severe liver damage, but the livers of caspase-2-null animals fed this diet were less severely affected (Machado et al., 2015). The liver injury in wild-type animals was found to stimulate a regenerative process which led to fibrosis through a mechanism involving Shh ligands (Machado et al., 2015). Because the livers of caspase-2-null mice were much less affected by high dietary fat, less Shh ligands were produced in their livers, and less secondary fibrosis was observed (Machado et al., 2015). The aforementioned studies examined the impact of caspase-2 deficiency on metabolism in male mice. Differences in metabolism of caspase-2-proficient males and females are well established, so researchers compared the impact of caspase-2 loss on starvation responses and other metabolic readouts in male and female mice. In general, caspase-2 status impacted more strongly on liver metabolism in males than females. Caspase-2 status did not modify the glucose tolerance of female mice but, as was reported previously (Wilson et al., 2015), male mice lacking caspase-2 had improved glucose tolerance relative to wild-type males (Wilson et al., 2016). Starvation forces animals to sequentially obtain energy from glycogen

184

M.A. Miles et al.

stored in the liver, then lipids from white adipose tissue and finally muscle protein. Caspase-2 deficiency enhanced lipid breakdown in fasting males, but no such difference was observed between fasting females of the different genotypes (Wilson et al., 2016). On a cellular level, autophagy can also be exploited to recycle macromolecules under conditions of nutrient deprivation, and caspase-2 deficiency promoted autophagic responses to fasting in the livers and muscles of both sexes (Wilson et al., 2016). The papers cited earlier addressed the role of caspase-2 in the breakdown of macromolecules such as lipids for energy, but caspase-2 has also been implicated in cellular responses to perturbations in the synthesis of lipids. Caspase-2 was required for inhibition of fatty acid biosynthesis to efficiently kill ovarian cancer cells: siRNA-mediated downregulation of caspase-2 protected about half of the cells from death induced by Orlistat, a fatty acid synthase inhibitor (Yang et al., 2015). Orlistat treatment led to ATF4dependent transcriptional upregulation of REDD1, which inhibited mTOR, leading to dimerization-induced activation of caspase-2 (Yang et al., 2015). The mechanism by which mTOR suppression triggers caspase-2 activation is presently unknown. The process by which caspase-2 kills cells when activated in this way also awaits full definition, although Bax activation and cleavage of caspases-3 and -8 following Orlistat treatment were less prominent when caspase-2 was downregulated, and PARP cleavage was abolished (Yang et al., 2015), suggesting that caspase-2 acts upstream of Bax, caspases-8 and -3 in cell death caused by inhibition of fatty acid biosynthesis.

8. CANCER 8.1 Mixed Messages From Human and Mouse Research The caspase-2 locus was mapped to the large arm of human chromosome 7 (Kumar et al., 1995). This region is relatively frequently deleted in myeloid cancers, implying that it harbors one or more tumor suppressor genes (Honda et al., 2015). High-resolution single-nucleotide polymorphism (SNP) analyses recently enabled researchers to tightly define three “commonly deleted regions” (CDRs) within 7q in acute myeloid leukemias (Da Silva et al., 2016), which were predicted to contain the putative tumor suppressor genes whose inactivation may contribute to development of myeloid malignancies. A CDR within band 7q22 (Da Silva et al., 2016) encompassed the gene encoding the caspase-2 substrate CUX-1 (Ramdzan and Nepveu, 2014; Wong et al., 2014). Another, harboring

Old and Novel Functions of Caspase-2

185

the putative tumor suppressor EZH2 (Sashida and Iwama, 2016) mapped to 7q35–36 (Da Silva et al., 2016). A third CDR was located within 7q34 (Da Silva et al., 2016). Occupation of this chromosomal band by the caspase-2 gene prompted early speculation that deletion-mediated caspase-2 deficiency encourages the development of myeloid cancers. However, the site of the caspase-2 gene (designated to be 143288215–143307696 by the NCBI Homo sapiens annotation release 108) lies outside the 7q34 CDR identified by the SNP analyses (137841484–139319208) (Da Silva et al., 2016). Other genes located within this CDR, such as LUC7L2, have been postulated to account for the tumor suppressor function associated with 7q34 (Singh et al., 2013). It should be noted, however, that genes like caspase-2 that reside outside of the CDRs identified by Da Silva et al. may still play important tumor suppressor roles. MLL3, a myeloid tumor suppressor (Chen et al., 2014), provides a precedent for this: its locus is slightly telomeric to the CDR within 7q35–36. Very few mutations in the coding region of caspase-2 have been documented in human cancers. Of a total of 460 leukemias, gastric, colorectal, breast, liver, and lung cancers analyzed by Kim et al., none featured homozygous caspase-2 mutations and only four harbored heterozygous missense mutations (Kim et al., 2011a,b). It is not clear whether this frequency exceeded that expected by chance within genomically unstable tumors. Caspase-2 expression within leukemic samples has been correlated both positively (Holleman et al., 2005) and negatively (Dorstyn et al., 2014; Estrov et al., 1998; Faderl et al., 1999) with survival outcomes and/or therapeutic responses. A detailed analysis of caspase-2 status within human cancers is outside of the scope of this review, but recent cancer genome and expression studies have produced a huge volume of data that could help clarify the involvement of caspase-2 alterations in human cancers. Most published evidence pertaining to a link between caspase-2 and cancer has emerged from studies using knockout mice. Caspase-2-null mice were very slightly less cancer prone than wild-type or heterozygous mice over their lifetimes (Shalini et al., 2012; Zhang et al., 2007), demonstrating that caspase-2 does not act as a tumor suppressor under normal (laboratory) circumstances. Nevertheless, much research effort has been invested in exploring the potential for caspase-2 deficiency to enhance the ability of known oncogenic stimuli to promote cancer development, with mixed results. Caspase-2 deletion made no difference to cancer rates in p53 deficient and heterozygous mice, with or without γ-irradiation (Manzl et al., 2013). Similarly, the DNA-damaging compound 3-methylcholanthrene

186

M.A. Miles et al.

triggered fibrosarcoma development equivalently in wild-type and caspase2-deleted mice (but was potentiated by heterozygous loss of p53) (Manzl et al., 2012). As discussed later, caspase-2 deficiency suppressed neuroblastoma formation in N-myc transgenic mice, yet enhanced tumor development in a range of other mouse cancer models.

8.2 Promoting Neuroblastoma Only one study has assigned a cancer-promoting role to caspase-2: N-myc-driven neuroblastomas developed in half as many caspase-2deficient (or heterozygous) mice as wild-type animals, suggesting that caspase-2 could cooperate with N-myc to promote neuroblastoma development (Dorstyn et al., 2014). The authors noted that genetic background heavily modifies neuroblastoma incidence in this model. The N-myc transgene was initially created in SV129J mice, and careful breeding strategies incorporated this transgene into C57BL6 mice of different caspase-2 genotypes. Confirming previous observations (Weiss et al., 1997), inclusion of some C57BL6 genomic content dramatically impeded neuroblastoma formation due to enforced N-myc expression, which was further delayed by the absence of one or both alleles of caspase-2 (Dorstyn et al., 2014). Backcrossing was performed to minimize the impact of strain-dependent factors, but it is theoretically possible that loci which are closely linked to the caspase-2 gene, and divergent in SV129J and C57BL6 mouse strains, may affect the incidence of neuroblastoma induced by N-myc overexpression. The interpretation that caspase-2 promotes neuroblastoma formation was supported by analyses of human neuroblastoma samples: around half of patients whose tumors expressed abundant caspase-2 died within 2 years of diagnosis, whereas the vast majority of the patients whose tumors had low caspase-2 expression were long-term survivors (Dorstyn et al., 2014). Interestingly, this correlation was limited to tumors lacking N-myc amplification.

8.3 Tumor Suppressor Apart from the aforementioned studies, all published investigations into the role of caspase-2 in cancer development have concluded that caspase-2 loss exacerbates the cancer-promoting impact of previously established oncogenic stimuli. The magnitude of the oncogenic cooperation afforded by caspase-2 deficiency varied from relatively modest to substantial. In general, tumors arising in caspase-2-null animals tended to be more mitotically active

Old and Novel Functions of Caspase-2

187

and contain more cells displaying karyotypic and morphological features, consistent with aberrant cell cycle control. Caspase-2-deficient E1A/Ras-transformed MEFs proliferated faster in vitro and exhibited greater clonogenic survival than transformed wild-type MEFs (Ho et al., 2009). The caspase-2-deficient cells also formed rapidly growing tumors upon implantation into athymic mice, in contrast to the transformed wild-type cells that either produced slow-growing tumors or none (Ho et al., 2009). Homozygous (Ho et al., 2009; Manzl et al., 2012; Peintner et al., 2015) or heterozygous (Ho et al., 2009) caspase-2 deficiency accelerated development of lymphomagenesis induced by enforced overexpression of c-Myc in B cells. The lymphomas lacking caspase-2 tended to be more infiltrative (Manzl et al., 2012). Caspase-2 deficiency also enhanced lymphomagenesis due to loss of ATM (Puccini et al., 2013). Mice lacking both caspase-2 and ATM had unusually high postnatal lethality, and those that survived grew more slowly than mice lacking only ATM or caspase-2 (Puccini et al., 2013). Twice as many double-knockout mice developed lymphomas by a year of age than caspase-2-proficient ATM-null mice (Puccini et al., 2013). The lymphomas from mice with the different genotypes contained similar numbers of apoptotic cells, but tumor cells lacking both ATM and caspase-2 proliferated faster than cells from tumors isolated from mice only lacking ATM (Puccini et al., 2013). These data suggest that the power of caspase-2 to reduce lymphomagenesis caused by ATM deficiency reflected the protease’s ability to regulate mitotic checkpoints following DNA damage, rather than a proapoptotic function. Consistent with this notion, most of the cells from lymphomas that arose in animals lacking both caspase-2 and ATM were aneuploid, whereas the majority of ATM-deficient tumor cells were diploid (Puccini et al., 2013). Mammary expression of an activating c-neu allele (Erbb2V664A) in mice promotes formation of tumors, particularly following pregnancy, that resemble luminal human breast cancers. Parsons et al. evaluated the impact of caspase-2 status on the incidence of mammary tumors in “MMTV/ c-neu” mice bearing this transgene (Parsons et al., 2013). Caspase-2 genotype did not significantly affect tumor formation in virgin mice; however, a modest difference was observed in the mice that had produced two or more litters. MMTV/c-neu;caspase-2/ mice all developed tumors between 5 and 9 months of age. The majority of the caspase-2-proficient mice bearing the MMTV/c-neu transgene also developed tumors, at a similar rate but

188

M.A. Miles et al.

with a slightly greater latency than the caspase-2-null animals. Significantly, around a quarter of the MMTV/c-neu;caspase-2+/+ multiparous animals remained tumor free during for over a year, whereas all heterozygous or null mice died by around 9 months of age. Thus, caspase-2 seemed to slightly retard c-neu-mediated tumorigenesis in most mice but also prevented a small proportion of animals from developing cancer (at least within the first year of life). As in the ATM-null lymphomas described earlier, the mammary tumors that arose in MMTV/c-neu;caspase-2/ mice featured a higher proportion of mitotic and multinuclear cells and cells with abnormally large nuclei or aberrant mitotic spindles than those that developed in caspase-2-proficient MMTV/c-neu animals. Inhalation of a Cre-encoding virus was used to drive expression of the oncogenic Ras variant KRasG12D in the lungs of caspase-2-deficient and wild-type mice. After this treatment, the lungs of the caspase-2-null mice contained more tumors than those of caspase-2-proficient mice (Terry et al., 2015). The majority of the cancers that formed in the caspase-2knockout animals were high grade, whereas those that arose in wild-type mice were typically smaller, low-grade tumors containing fewer mitotic cells. Amazingly, despite harboring larger numbers of bigger, higher-grade tumors, the caspase-2-deficient mice succumbed to KRasG12D-driven lung cancers at similar rates to the wild-type animals (Terry et al., 2015). Histochemical staining with an antibody recognizing cleaved caspase-3 implied that cisplatin therapy led to more tumor cell death in the caspase-2-deficient animals than their wild-type counterparts (Terry et al., 2015). However, although the tumors of both caspase-2 and wild-type mice shrank following cisplatin treatment (Terry et al., 2015), those in caspase-2-null mice regrew more rapidly after cessation of therapy, implying that loss of caspase-2 may facilitate reentry into mitosis by tumor cells that survive exposure to chemotherapy. Treatment at 2 weeks of age (but not at 6 weeks) with diethylnitrosamine, a DNA alkylating and ROS-inducing carcinogen, promoted formation of about twice as many hepatocellular carcinomas in the livers of caspase-2knockout mice than wild-type animals (Shalini et al., 2016). Histological examination revealed that the tumors from caspase-2-null animals tended to be more advanced than those in wild-type animals. All of the caspase2-null mice developed tumors, but 2 of 11 wild-type mice remained cancer free at 10 months of age (Shalini et al., 2016). Biochemical markers of oxidative damage such as 8-OHdG and inflammatory cytokines were slightly elevated in untreated caspase-2-deficient mice, relative to wild-type animals,

Old and Novel Functions of Caspase-2

189

and this difference was dramatically exacerbated following exposure to diethylnitrosamine (Shalini et al., 2016). Although inhibition of apoptosis can facilitate cancer development, caspase-2 status did not alter cell death observed in the livers of diethylnitrosamine-treated (or untreated) mice (Shalini et al., 2016), arguing that the tumor suppressor role of caspase-2 does not primarily emanate from a proapoptotic function in this cancer model. Although caspase-2 deletion accelerated lymphoma development trigged by c-Myc overexpression in B cells (Ho et al., 2009; Manzl et al., 2012; Peintner et al., 2015), RAIDD status failed to alter tumor development in this model, in mice proficient or deficient for caspase-2 (Peintner et al., 2015). Absence of the other PIDDosome component, PIDD, dramatically postponed c-Myc-induced lymphomagenesis (Manzl et al., 2012). This result may reflect a dominant procancerous role of the PIDD– RIPK1–Nemo–NF-κB pathway (described in Section 4.5), but the inability of RAIDD or PIDD deficiency to phenocopy that of caspase-2 argues that the tumor suppressor role of caspase-2 in this context is PIDDosome independent.

9. VISION 9.1 Retinal Cell Death Mice lacking a functional Pde6b gene (encoding cGMP phosphodiesterase type 6) experience progressive retinal degeneration due to persistence of cGMP, which impedes closure of cGMP-gated channels (Ionita and Pittler, 2007). These “rd1” mutant mice, like human retinosa pigmentosa patients, suffer primary loss of rods (causing night blindness) followed by secondary cone cell degradation (abolishing vision entirely) after the majority of the rods have died (Han et al., 2013). Oxidative damage is a feature of the cell-autonomous cone cell destruction in this model (Koenekoop, 2009) and may result from exposure of cone cells to higher levels of oxygen after destruction of rod cells (which normally greatly outnumber cone cells and therefore absorb the majority of oxygen supplied to retina). Subretinal introduction of adenoviral constructs directing high-level expression of the antioxidants SOD plus catalase significantly slowed the drop in cone density in rd1 mice (Xiong et al., 2015). Enforced expression of NRF2, a master transcription factor that induces a number of antioxidant genes, had a stronger protective effect (Xiong et al., 2015).

190

M.A. Miles et al.

Nutrient deprivation following rod cell death has also been linked to the loss of cone cells. Cone cell survival was enhanced when insulin levels were boosted (Punzo et al., 2009). During the early stages of multiple models of retinosa pigmentosa, mTOR activation was reduced (Punzo et al., 2012). Constitutive activation of mTOR complex 1, leading to enhanced NADPH production, significantly reduced cone cell loss in mice with retinosa pigmentosa (Venkatesh et al., 2015). It is possible that the apparent protective effect of NADPH in this disease model reflects its antioxidative role, perhaps reconciling the alternative models (Punzo et al., 2012). Intriguingly, cone cell loss in 22-week-old rd1 mice was significantly less pronounced in the absence of caspase-2, although analysis of cone cell density in 10-week-old mice revealed that the initial phase of destruction proceeded similarly in mice with and without caspase-2 (Venkatesh et al., 2015). The authors postulated that, during the initial stages of the disease, when rod cell death disrupts the structure of the retina, cone cells may initially die via caspase-2-independent necrosis, but later the remaining cone cells may subsequently die via caspase-2-dependent apoptosis, which could be triggered by low NADPH levels and/or ROS.

9.2 Loss of Retinal Ganglion Cells Following Optic Nerve Injury Caspase-2 may also play an important role in maintaining the survival of another crucially important cell type in eye biology and disease: retinal ganglion cells (RGCs). These cells transmit visual data collected by photoreceptor cells to the brain via their axons, which comprise the optic nerve. Following optic nerve damage, RGCs undergo apoptosis. Clamping of the axons of rat RGCs led to a 60% decrease in RGC number within 7 days, but this loss was nearly completely prevented by intravitreal administration with a chemically stabilized siRNA-targeting caspase-2 at the time of injury (Ahmed et al., 2011). This protective effect persisted for at least 12 weeks (Vigneswara and Ahmed, 2016). Repeated administration of zVDVAD-fmk, a reagent that inhibits caspase-2 (and also caspases-3 and -7, see Section 2.2), prevented death of about half of the RGCs that were otherwise killed within a fortnight of optic nerve crushing (Vigneswara et al., 2012). To model severe optic nerve damage, researchers severed RGC axons and neural sheaths in rat eyes, which eliminated 90% of RGCs within 14 days. In contrast, only 75% of the RGCs were lost from eyes treated with a caspase-2-downregulating siRNA (Ahmed et al., 2011). The siRNA reagent used in this work (QPI-1007) is presently being

Old and Novel Functions of Caspase-2

191

evaluated for clinical efficacy in treating optic neuropathies (Vigneswara and Ahmed, 2016). Other caspases have also been linked to RGC death. Around half as many RGCs died following a crushing injury to the optic nerve in caspase-7-knockout mice as wild-type animals (Choudhury et al., 2015). Electroretinography measurements confirmed that mice lacking caspase-7 retained significantly greater retinal function after injury than wild-type mice (Choudhury et al., 2015). It is possible that the involvement of caspases-7 and -2 in RGC loss following optic nerve lesions may reflect a linear pathway in which caspase-2 proteolytically activates caspase-7 (Ho et al., 2005), leading to RGC apoptosis. Inhibitors that were designed to disable caspases-3, -6, and -8 rescued up to 60% of RGCs following optic nerve damage (Liu et al., 2015; Monnier et al., 2011). Because these inhibitors lack specificity, additional approaches will be needed to confirm the direct involvement of these proteases in RGC apoptosis. Although caspase-6 (like caspase-3) could not be detected in RGC cells, it was expressed in other retinal cells, and its retinal levels increased after crush injuries to the optic nerve (Vigneswara et al., 2012). Introduction of a dominantly interfering variant of caspase-6 had minimal impact on RGC loss following optic nerve damage and did not augment the protective effect of caspase-2 downregulation (Vigneswara et al., 2014). However, caspase-6 inhibition did cooperate with caspase-2 downregulation to promote modest but significant axonal regeneration after a crush injury, through stimulation of CTNF production by neighboring glial cells (Vigneswara et al., 2014). Hopefully further work will confirm that protection of RGC death achieved through caspase-2 inhibition, with or without caspase-6 inhibition, has the desired functional effect of improving eyesight following optic nerve damage. Presumably ongoing work will also determine the optimal timing and frequency of caspase-2 siRNA administration following such injuries, which will be crucial for assessing clinical utility. Optic nerve damage boosted both the expression level and activity of caspase-2 (Ahmed et al., 2011), but the molecular basis for caspase-2 activation in this context, including any involvement of RAIDD or PIDD, has not yet been elucidated.

10. NEUROLOGICAL CONDITIONS 10.1 Exacerbating Ischemic/Reperfusion Injury Dramatically divergent findings have been reported in relation to the role played by caspase-2 in brain ischemic/reperfusion injury. Occlusion of

192

M.A. Miles et al.

the middle cerebral artery of adult mice for 24 h, or occlusion for 2 h followed by reperfusion for 18 h, led to equivalently sized infarcts in caspase-2-deficient and wild-type mice (Bergeron et al., 1998). A separate group of researchers instead ligated the left carotid artery of 9-day-old mice and subjected them to hypoxia (10% oxygen) for 50 min. Seven days later, infarcts in those caspase-2-null mice were 32% smaller than in their wild-type littermates (Carlsson et al., 2011). Carlsson et al. speculated that they detected an impact of caspase-2 status on brain damage because they used younger mice than Bergeron et al. They presented data showing that caspase-2 expression in the murine brain decreased from robust 3 days after birth to undetectable by 3 weeks of age and hypothesized that caspase-2 plays a crucial role in neuronal cell death during early postnatal life, but other pathways take over later in adult life. Unfortunately Carlsson et al. did not experimentally test this theory by repeating their experiment on older mice. Bergeron et al. did not specify the age of the mice they used for their ischemia experiment. The paper cited for their method only specified that “adult” mice were used (Hara et al., 1997). However, Bergeron et al. included a caspase-2 immunoblot in their paper that appears to show strong expression in the brains of the animals they used to evaluate responses to ischemia, arguing against the notion that their mice were too old to express enough caspase-2 to enable caspase-2-mediated neuronal cell death following ischemia. It is possible that the two research groups observed a differential requirement for caspase-2 because they analyzed the brain damage at different times following the insult. Carlsson et al. measured infarct volumes a week after blood flow was occluded, whereas Bergeron et al. monitored lesions 24 h afterward. Perhaps the initial damage following ischemia occurs via caspase-2-independent mechanisms, but subsequent secondary apoptosis due to aberrant neurological or cytokine signals may require caspase-2. Such a model is reminiscent of the retinal cone cell death described in Section 9.1, in which the initial cell death occurs similarly in caspase-2-proficient and -deficient animals, but the later cone cell loss is less pronounced in animals lacking caspase-2 (Venkatesh et al., 2015).

10.2 Excitotoxicity Carlsson et al. also examined the involvement of caspase-2 in excitotoxicity-induced brain damage by administering intracerebral ibotenate injections to 5-day-old wild-type or caspase-2-deficient mice. This treatment produced mean lesion volumes in cortical and white matter

Old and Novel Functions of Caspase-2

193

regions that were 41% and 59% smaller, respectively, in caspase-2-null mice relative to control animals (Carlsson et al., 2011). Silencing caspase-2 expression by intraparenchymal siRNA administration conferred similar protection against focal lesions provoked by ibotenate (Carlsson et al., 2011).

10.3 Withdrawal of Nerve Growth Factor or Serum Downregulation of caspase-2 (but not caspase-3) was reported to protect axotomized adult dorsal root ganglion neurons and glia from apoptosis following serum withdrawal in vitro (Vigneswara et al., 2013). Although death of neuronal cells from the PC12 cell line triggered by NGF withdrawal was suppressed by antisense downregulation of caspase-2 (Troy et al., 1997), most studies have failed to demonstrate a role for caspase-2 in NGF withdrawal-induced death, and indeed there is some evidence that caspase-2 plays an antiapoptotic role during neuronal development. Caspase-2-deficient dorsal root ganglion cells from 2-day-old mice were as sensitive to killing by NGF withdrawal in vitro as those from wild-type mice (O’Reilly et al., 2002). Sympathetic neurons isolated from the superior cervical ganglia of caspase-2-deficient mice were more sensitive to death from NGF withdrawal than those from wild-type animals (Bergeron et al., 1998), and mice lacking caspase-2 had around 27% fewer facial neurons than their wild-type littermates (Bergeron et al., 1998). Thus, caspase-2 seems to be required for maximal neuronal cell death in some contexts, but is either dispensable or protective in other situations. An explanation for the apparently differential requirements for caspase-2 in these contexts remains elusive.

10.4 Alzheimer’s Disease Incubation of hippocampal or sympathetic neurons with Aβ1–42, a peptide associated with neuronal loss in Alzheimer’s disease, triggered caspase-2 activation (Troy et al., 2000) as revealed by bVAD trapping (Tizon et al., 2010) and cell death (Tizon et al., 2010; Troy et al., 2000). Caspase-2 downregulation or gene deletion protected neurons from this lethality (Troy et al., 2000). This finding was confirmed in vivo: caspase-2-null mice escaped the neurodegeneration induced in wild-type animals by convection-enhanced hippocampal delivery of Aβ1–42 (Jean et al., 2013). Consistent with a role for caspase-2 in Alzheimer’s disease, its levels were dramatically elevated in the brains of patients relative to age-matched controls (Jean et al., 2013).

194

M.A. Miles et al.

Interestingly, Aβ1–42-mediated activation of caspase-2 required RAIDD but not PIDD, and the resulting cell death involved caspase-2-mediated transcription upregulation of Bim (Ribe et al., 2012) which correlated with c-Jun phosphorylation (Jean et al., 2013). Although the mechanistic links between Aβ1–42, caspase-2 activation, c-Jun phosphorylation, and Bim induction have not been defined, a model for the downstream portion of this pathway could be envisaged from data produced in other studies (Fig. 5). JNK, the prototypical c-Jun kinase, was recently shown to indirectly activate FoxO3a (Li et al., 2015), a transcription factor that could directly induce Bim expression (Gilley et al., 2003; Li et al., 2015). It is currently unclear how Aβ1–42 would activate JNK in this context; however, a separate study suggested that caspase-2 activation occurred downstream of JNK in neurons treated with a different Aβ peptide (Aβ25–35) (Viana et al., 2010).

10.5 Huntington’s and Motor Neuron Diseases Huntington’s disease patients express Htt proteins bearing dramatically expanded polyglutamine repeats, and expression of these pathogenic Htt variants (like Htt138) can provoke neuronal cell death in vitro. Caspase2, like caspase-3, could proteolyze Htt after residue Asp522, whereas caspase-6 cleaved after Asp586 (Wellington et al., 2002). Enforced expression of catalytically inactive mutants of caspases-2, -6, or -7 afforded some protection to Htt138-mediated neuronal cell death (Hermel et al., 2004). Abolition of the caspase-6 cleavage site prevented Huntington’s-like pathology in an animal model, but mutating the caspases-2/-3 site was ineffective (Graham et al., 2006), implying that caspase-6-mediated Htt proteolysis had greater pathogenic significance. Nevertheless, caspase-2-deficient mice escaped behavioral symptoms (Carroll et al., 2011) provoked by transgenic expression of human Htt bearing 128 repeats (Lee et al., 2013). The caspase2-null animals expressing Htt128 developed similar reductions in stratum and thalamus volume as caspase-2-proficient mice, but retained better motor and cognitive functions than animals with intact caspase-2 genes (Carroll et al., 2011). Not all neurological diseases require caspase-2: the onset and progression of motor neuron degeneration in SODG93A mutant mice commenced and progressed similarly in wild-type and caspase-2-null mice (Bergeron et al., 1998).

Old and Novel Functions of Caspase-2

195

Fig. 5 Caspase-2-dependent neuronal cell death. Aβ peptide may stimulate caspase-2dependent cell death by the pathways shown. See Section 10.4 of the text for details and references.

11. CONCLUSIONS 11.1 Why Has Caspase-2 Been Evolutionarily Conserved? The retention of caspase-2 throughout mammalian evolution suggests it confers a selective advantage. The subtle phenotypes of caspase-2-deficient

196

M.A. Miles et al.

mice argue that caspase-2 is not required for development or survival under laboratory conditions. Although caspase-2 can act as a tumor suppressor, this role only manifested in animals engineered to express potent oncogenes or lack crucial tumor suppressor genes, so it seems unlikely that caspase-2 was kept throughout mammalian evolution for its cancer-ameliorating function. Likewise, the ability of caspase-2 to rebalance responses to oxidative stress in favor of apoptosis rather than autophagy and/or aneuploidy can be beneficial in experimental contexts, but is unlikely to be highly selective in vivo. Slight extension of maximal life span, without significantly affecting median longevity, would also presumably not confer a significant evolutionary benefit. It has been postulated that apoptosis originally evolved to defend primitive multicellular organisms against intracellular pathogens (Ameisen, 2002). If caspase-2 could protect animals from infection, this may account for its evolutionary conservation. Surprisingly little research has been published regarding the impact of caspase-2 status on responses to infection, but two studies have revealed a role for this enzyme in killing cells experiencing ER stress due to rhabdoviral or bacterial infection (Bronner et al., 2015; Mahoney et al., 2011). Many types of infectious pathogens modify ER structure and function (Escoll et al., 2016; Romero-Brey and Bartenschlager, 2016), leading to strong evolutionary pressure for host cells to detect ER stress that may signify infection, and respond by undergoing apoptosis. Viral genome replication can also trigger DNA damage responses (Escoll et al., 2016; Luftig, 2014). It is therefore possible that the ability of caspase-2 to trigger cell cycle arrest or apoptosis following signals associated with DNA damage reflects an antiviral function. Presumably the hypothesis that caspase-2 may play an important role in host defense against intracellular pathogens could be readily tested by comparing responses of wild-type and caspase-2-null or mutant mice to infection with viruses or microbes.

11.2 Manipulating Caspase-2 for Therapeutic Benefit Genes that were selected for during evolution can sometimes be maladaptive in the context of modern human lifestyles. Loss of caspase-2 protected mice, especially males, from symptoms resulting from high-fat and high-sugar diets, such as diabetes and liver damage. Clearly, a lot more work will be needed to determine the feasibility and efficacy of targeting caspase-2 to

Old and Novel Functions of Caspase-2

197

prevent or treat these conditions in humans, but this is an exciting possible application of caspase-2-related research. Caspase-2 activity was also shown to exacerbate deterioration of vision due to either secondary cone destruction following loss of the rod photoreceptors or optic nerve damage. Animal experiments suggested that eye-specific administration of caspase-2-inhibiting drugs may preserve vision after optic nerve injury. Recruitment is underway for a phase III clinical trial to evaluate one such reagent in patients with acute nonarteritic anterior ischemic optic neuropathy (Vigneswara and Ahmed, 2016). It is possible that caspase-2 inhibitors may also have utility for other neurological conditions, such as Huntington’s disease (and perhaps Alzheimer’s). Prevention of caspase-2 expression in a mouse model of Huntington’s disease alleviated behavioral symptoms, but in this model all cells constitutively lacked caspase-2. In order for drug-mediated caspase-2 inhibition to be effective in a clinical setting, it may be necessary to administer the drug before disease onset and uptake may need to be highly efficient. Because Huntington’s disease can be genetically predicted, it is conceivable that symptom onset could be prevented by early administration of an effective drug. However, unlike in the eye conditions mentioned earlier, achieving uptake by sufficient relevant neurons may be challenging.

11.3 Could a Common Mechanism Underlie the Apparently Disparate Cellular Roles of Caspase-2? Perhaps some of the seemingly diverse cellular phenotypes observed in caspase-2-deficient cells and animals reflect divergent cellular responses to a primary defect in apoptotic signaling. It seems possible, for example, that autophagy occurs in caspase-2-deficient cells as a secondary response to stimuli that would normally lead to apoptosis in cells expressing caspase-2. By analogy, the enhanced rates of aneuploidy seen in caspase2-deficient cells could reflect an ability of caspase-2 to trigger apoptosis in cells undergoing aberrant mitoses, rather than (for example) a role in regulating mitotic spindle formation. However, as discussed in Section 4.7, cells lacking caspase-2 proliferated more rapidly than wild-type cells, arguing that caspase-2 directly regulates a mitotic checkpoint rather than merely facilitating the elimination of cells that fail to successfully complete mitosis. Likewise, caspase-2-proficient cells expressed antioxidant proteins like SOD at higher levels than null cells and contained

198

M.A. Miles et al.

lower concentrations of ROS, arguing that caspase-2 may play a direct antioxidant role rather than, or in addition to, helping to dispose of cells damaged through oxidative stress.

11.4 Molecular Pathways Upstream and Downstream of Caspase-2 The scientific literature relating to caspase-2 is considerably more contentious than that covering other caspases. Previously, the dearth of caspase-2-specific reagents contributed substantially to this problem, but the confounding issue of nonspecific reagents is less significant nowadays: fewer articles have been published in the last few years attributing cellular VDVADase activity to caspase-2, the bVAD trapping method enables researchers to identify contexts in which caspase-2 activation is an apical event, and BiFC helps determine the subcellular site of activation. These advances have helped to drag the field toward consensus in recent years, but it is not there yet! Further improvements in our understanding of this elusive caspase will hopefully emerge from increased adoption of these techniques, the caspase-2-selective reagents described in Section 2.2, and the Casp2C320S mouse and its cells. There is strong evidence that the PIDD–RAIDD–caspase-2 complex can promote caspase-2 activation in vitro, but less evidence that it is required to activate caspase-2 in vivo. Many examples have been published in which caspase-2-dependent events do not require RAIDD and/or PIDD (Table 2). CHK1 inhibitors cooperate strikingly with irradiation to activate caspase-2, but no natural context involving this pathway has yet been described. Caspase-2 can clearly play important roles in vivo (such as in RGCs after optic nerve damage, and macrophages infected with Brucella), so it will be important to define the molecular basis of its activation in these situations. The steps downstream of caspase-2 activation also remain incompletely defined. Initially, caspase-2 was regarded as having very similar biochemical activity to executioner caspases. A number of caspase-2-specific substrates have now been identified (Table 1), but the relevance of many of these to caspase-2-dependent processes awaits investigation. Even caspase-2 substrates that can also be cleaved by other caspases may be exclusively processed by caspase-2 in cells or contexts where other caspases are absent or not active. Recent progress has enhanced our understanding of circumstances under which particular stimuli activate caspase-2 to provoke specific biological effects. Hopefully future searches for the substrate(s) responsible for mediating caspase-2-dependent outcomes will exploit this knowledge to focus on biologically relevant cell types and stimuli.

Table 2 Stimuli-Specific Requirements for PIDD and/or RAIDD for Caspase-2-Dependent Processes PIDD RAIDD Stimulus Cells Outcome Required? Required? References

PIDD overexpression

E1A/Ras-transformed MEFs

Cell death and colony suppression Enforced Yes

Berube et al. (2005)

G€ o6976 + γ-irradiation

HeLa

Caspase-2 processing, cell death

Yes

Yes

Ando et al. (2012)

γ-Irradiation, siBubR1

HeLa

Caspase-2 processing

Yes

Yes

Thompson et al. (2015)

γ-Irradiation

HCT116 (p53 wild type) DEVDase, caspase-2 processing

Not tested

No

Sohn et al. (2011)

LBH589 (deacetylase inhibitor)

ST1 and HuT102

Processing of caspase-9, caspase-3, Yes and PARP, cell death

Yes

Hasegawa et al. (2011)

Heat shock

Activated splenocytes

Caspase-2 binding to bVAD cell death

Not tested

Yes

Tu et al. (2006)

Heat shock

MEFs (transformation status not specified)

Caspase-2 dimerization

Not tested

Yes

Bouchier-Hayes et al. (2009)

In vivo transient global cerebral ischemia

Rat cells in hippocampal Caspase-2, Bid processing, cell CA1 region death

Yes

Not tested Niizuma et al. (2008)

Aβ1–42

Rat hippocampal neurons Caspase-2 processing, cell death

No

Yes

Ribe et al. (2012)

Aβ1–42

Rat hippocampal neurons Bim induction, nuclear phospho-c- Not Jun tested

Yes

Jean et al. (2013) Continued

Table 2 Stimuli-Specific Requirements for PIDD and/or RAIDD for Caspase-2-Dependent Processes—cont’d PIDD RAIDD Stimulus Cells Outcome Required? Required? References

NGF deprivation

Murine sympathetic neurons

Cell death

No

Yes

Ribe et al. (2012)

Paclitaxel and SB-T-1216

SK-BR-3, MCF-7

Cell death

Not tested

No

Jelinek et al. (2013)

5-Fluorouracil

HCT116 (p53 wild type) Caspase-2 processing, cytochrome No c release into cytosol

No

Vakifahmetoglu et al. (2006)

Eμ-Myc expression

Murine B cells

Lymphomagenesis

No

Not tested Manzl et al. (2012)

Eμ-Myc expression

Murine B cells

Lymphomagenesis

Not tested

No

Peintner et al. (2015)

Old and Novel Functions of Caspase-2

201

REFERENCES Ahmed, Z., Kalinski, H., Berry, M., Almasieh, M., Ashush, H., Slager, N., Brafman, A., Spivak, I., Prasad, N., Mett, I., Shalom, E., Alpert, E., Di Polo, A., Feinstein, E., Logan, A., 2011. Ocular neuroprotection by siRNA targeting caspase-2. Cell Death Dis. 2, e173. Ameisen, J.C., 2002. On the origin, evolution, and nature of programmed cell death: a timeline of four billion years. Cell Death Differ. 9, 367–393. Andersen, J.L., Johnson, C.E., Freel, C.D., Parrish, A.B., Day, J.L., Buchakjian, M.R., Nutt, L.K., Thompson, J.W., Moseley, M.A., Kornbluth, S., 2009. Restraint of apoptosis during mitosis through interdomain phosphorylation of caspase-2. EMBO J. 28, 3216–3227. Ando, K., Kernan, J.L., Liu, P.H., Sanda, T., Logette, E., Tschopp, J., Look, A.T., Wang, J., Bouchier-Hayes, L., Sidi, S., 2012. PIDD death-domain phosphorylation by ATM controls prodeath versus prosurvival PIDDosome signaling. Mol. Cell 47, 681–693. Baldi, P.C., Giambartolomei, G.H., 2013. Immunopathology of Brucella infection. Recent Pat. Antiinfect. Drug Discov. 8, 18–26. Baliga, B.C., Read, S.H., Kumar, S., 2004. The biochemical mechanism of caspase-2 activation. Cell Death Differ. 11, 1234–1241. Bergeron, L., Perez, G.I., Macdonald, G., Shi, L., Sun, Y., Jurisicova, A., Varmuza, S., Latham, K.E., Flaws, J.A., Salter, J.C., Hara, H., Moskowitz, M.A., Li, E., Greenberg, A., Tilly, J.L., Yuan, J., 1998. Defects in regulation of apoptosis in caspase-2-deficient mice. Genes Dev. 12, 1304–1314. Berube, C., Boucher, L.M., Ma, W., Wakeham, A., Salmena, L., Hakem, R., Yeh, W.C., Mak, T.W., Benchimol, S., 2005. Apoptosis caused by p53-induced protein with death domain (PIDD) depends on the death adapter protein RAIDD. Proc. Natl. Acad. Sci. U.S.A. 102, 14314–14320. Bouchier-Hayes, L., Oberst, A., Mcstay, G.P., Connell, S., Tait, S.W., Dillon, C.P., Flanagan, J.M., Beere, H.M., Green, D.R., 2009. Characterization of cytoplasmic caspase-2 activation by induced proximity. Mol. Cell 35, 830–840. Bronner, D.N., O’riordan, M.X., He, Y., 2013. Caspase-2 mediates a Brucella abortus RB51-induced hybrid cell death having features of apoptosis and pyroptosis. Front. Cell. Infect. Microbiol. 3, 83. Bronner, D.N., Abuaita, B.H., Chen, X., Fitzgerald, K.A., Nunez, G., He, Y., Yin, X.M., O’riordan, M.X., 2015. Endoplasmic reticulum stress activates the inflammasome via NLRP3- and caspase-2-driven mitochondrial damage. Immunity 43, 451–462. Butt, A.J., Harvey, N.L., Parasivam, G., Kumar, S., 1998. Dimerization and autoprocessing of the Nedd2 (caspase-2) precursor requires both the prodomain and the carboxyl-terminal regions. J. Biol. Chem. 273, 6763–6768. Callaway, D.A., Riquelme, M.A., Sharma, R., Lopez-Cruzan, M., Herman, B.A., Jiang, J.X., 2015. Caspase-2 modulates osteoclastogenesis through down-regulating oxidative stress. Bone 76, 40–48. Carlsson, Y., Schwendimann, L., Vontell, R., Rousset, C.I., Wang, X., Lebon, S., Charriaut-Marlangue, C., Supramaniam, V., Hagberg, H., Gressens, P., Jacotot, E., 2011. Genetic inhibition of caspase-2 reduces hypoxic-ischemic and excitotoxic neonatal brain injury. Ann. Neurol. 70, 781–789. Carroll, J.B., Southwell, A.L., Graham, R.K., Lerch, J.P., Ehrnhoefer, D.E., Cao, L.P., Zhang, W.N., Deng, Y., Bissada, N., Henkelman, R.M., Hayden, M.R., 2011. Mice lacking caspase-2 are protected from behavioral changes, but not pathology, in the YAC128 model of Huntington disease. Mol. Neurodegener. 6, 59.

202

M.A. Miles et al.

Carroll, B.L., Pulkoski-Gross, M.J., Hannun, Y.A., Obeid, L.M., 2016. CHK1 regulates NF-kappaB signaling upon DNA damage in p53-deficient cells and associated tumor-derived microvesicles. Oncotarget 7, 18159–18170. Castedo, M., Perfettini, J.L., Roumier, T., Valent, A., Raslova, H., Yakushijin, K., Horne, D., Feunteun, J., Lenoir, G., Medema, R., Vainchenker, W., Kroemer, G., 2004. Mitotic catastrophe constitutes a special case of apoptosis whose suppression entails aneuploidy. Oncogene 23, 4362–4370. Celli, J., De Chastellier, C., Franchini, D.M., Pizarro-Cerda, J., Moreno, E., Gorvel, J.P., 2003. Brucella evades macrophage killing via VirB-dependent sustained interactions with the endoplasmic reticulum. J. Exp. Med. 198, 545–556. Chen, F., He, Y., 2009. Caspase-2 mediated apoptotic and necrotic murine macrophage cell death induced by rough Brucella abortus. PLoS One 4, e6830. Chen, C., Liu, Y., Rappaport, A.R., Kitzing, T., Schultz, N., Zhao, Z., Shroff, A.S., Dickins, R.A., Vakoc, C.R., Bradner, J.E., Stock, W., Lebeau, M.M., Shannon, K.M., Kogan, S., Zuber, J., Lowe, S.W., 2014. MLL3 is a haploinsufficient 7q tumor suppressor in acute myeloid leukemia. Cancer Cell 25, 652–665. Choudhury, S., Liu, Y., Clark, A.F., Pang, I.H., 2015. Caspase-7: a critical mediator of optic nerve injury-induced retinal ganglion cell death. Mol. Neurodegener. 10, 40. Conradt, B., Wu, Y.C., Xue, D., 2016. Programmed cell death during Caenorhabditis elegans development. Genetics 203, 1533–1562. Cooper, D.M., Granville, D.J., Lowenberger, C., 2009. The insect caspases. Apoptosis 14, 247–256. Cuenin, S., Tinel, A., Janssens, S., Tschopp, J., 2007. p53-induced protein with a death domain (PIDD) isoforms differentially activate nuclear factor-kappaB and caspase-2 in response to genotoxic stress. Oncogene 27, 387–396. Da Silva, F.B., Machado-Neto, J.A., Bertini, V.H., Velloso, E.D., Ratis, C.A., Calado, R.T., Simoes, B.P., Rego, E.M., Traina, F., 2016. Single-nucleotide polymorphism array (SNP-A) improves the identification of chromosomal abnormalities by metaphase cytogenetics in myelodysplastic syndrome. J. Clin. Pathol. 11. http://dx.doi.org/10.1136/ jclinpath-2016-204023. Epub ahead of print. Davies, S.P., Reddy, H., Caivano, M., Cohen, P., 2000. Specificity and mechanism of action of some commonly used protein kinase inhibitors. Biochem. J. 351, 95–105. Dawar, S., Lim, Y., Puccini, J., White, M., Thomas, P., Bouchier-Hayes, L., Green, D., Dorstyn, L., Kumar, S., 2016a. Caspase-2-mediated cell death is required for deleting aneuploid cells. Oncogene. http://dx.doi.org/10.1038/onc.2016.423. Dawar, S., Shahrin, N.H., Sladojevic, N., D’andrea, R.J., Dorstyn, L., Hiwase, D.K., Kumar, S., 2016b. Impaired haematopoietic stem cell differentiation and enhanced skewing towards myeloid progenitors in aged caspase-2-deficient mice. Cell Death Dis. 7, e2509. Del Nagro, C.J., Choi, J., Xiao, Y., Rangell, L., Mohan, S., Pandita, A., Zha, J., Jackson, P.K., O’brien, T., 2014. Chk1 inhibition in p53-deficient cell lines drives rapid chromosome fragmentation followed by caspase-independent cell death. Cell Cycle 13, 303–314. Demon, D., Van Damme, P., Vanden Berghe, T., Deceuninck, A., Van Durme, J., Verspurten, J., Helsens, K., Impens, F., Wejda, M., Schymkowitz, J., Rousseau, F., Madder, A., Vandekerckhove, J., Declercq, W., Gevaert, K., Vandenabeele, P., 2009. Proteome-wide substrate analysis indicates substrate exclusion as a mechanism to generate caspase-7 versus caspase-3 specificity. Mol. Cell. Proteomics 8, 2700–2714. Di Donato, N., Jean, Y.Y., Maga, A.M., Krewson, B.D., Shupp, A.B., Avrutsky, M.I., Roy, A., Collins, S., Olds, C., Willert, R.A., Czaja, A.M., Johnson, R., Stover, J.A., Gottlieb, S., Bartholdi, D., Rauch, A., Goldstein, A., Boyd-Kyle, V., Aldinger, K.A.,

Old and Novel Functions of Caspase-2

203

Mirzaa, G.M., Nissen, A., Brigatti, K.W., Puffenberger, E.G., Millen, K.J., Strauss, K.A., Dobyns, W.B., Troy, C.M., Jinks, R.N., 2016. Mutations in CRADD result in reduced caspase-2-mediated neuronal apoptosis and cause megalencephaly with a rare lissencephaly variant. Am. J. Hum. Genet. 99, 1117–1129. Dorstyn, L., Puccini, J., Wilson, C.H., Shalini, S., Nicola, M., Moore, S., Kumar, S., 2012. Caspase-2 deficiency promotes aberrant DNA-damage response and genetic instability. Cell Death Differ. 19, 1288–1298. Dorstyn, L., Puccini, J., Nikolic, A., Shalini, S., Wilson, C.H., Norris, M.D., Haber, M., Kumar, S., 2014. An unexpected role for caspase-2 in neuroblastoma. Cell Death Dis. 5, e1383. Duan, H., Dixit, V.M., 1997. RAIDD is a new death adaptor molecule. Nature 385, 86–89. Eckhart, L., Ballaun, C., Hermann, M., Vandeberg, J.L., Sipos, W., Uthman, A., Fischer, H., Tschachler, E., 2008. Identification of novel mammalian caspases reveals an important role of gene loss in shaping the human caspase repertoire. Mol. Biol. Evol. 25, 831–841. Escoll, P., Mondino, S., Rolando, M., Buchrieser, C., 2016. Targeting of host organelles by pathogenic bacteria: a sophisticated subversion strategy. Nat. Rev. Microbiol. 14, 5–19. Estrov, Z., Thall, P.F., Talpaz, M., Estey, E.H., Kantarjian, H.M., Andreeff, M., Harris, D., Van, Q., Walterscheid, M., Kornblau, S.M., 1998. Caspase 2 and caspase 3 protein levels as predictors of survival in acute myelogenous leukemia. Blood 92, 3090–3097. Faderl, S., Thall, P.F., Kantarjian, H.M., Talpaz, M., Harris, D., Van, Q., Beran, M., Kornblau, S.M., Pierce, S., Estrov, Z., 1999. Caspase 2 and caspase 3 as predictors of complete remission and survival in adults with acute lymphoblastic leukemia. Clin. Cancer Res. 5, 4041–4047. Feinstein-Rotkopf, Y., Arama, E., 2009. Can’t live without them, can live with them: roles of caspases during vital cellular processes. Apoptosis 14, 980–995. Franchi, L., Munoz-Planillo, R., Nunez, G., 2012. Sensing and reacting to microbes through the inflammasomes. Nat. Immunol. 13, 325–332. Garcia-Calvo, M., Peterson, E.P., Leiting, B., Ruel, R., Nicholson, D.W., Thornberry, N.A., 1998. Inhibition of human caspases by peptide-based and macromolecular inhibitors. J. Biol. Chem. 273, 32608–32613. Gilley, J., Coffer, P.J., Ham, J., 2003. FOXO transcription factors directly activate bim gene expression and promote apoptosis in sympathetic neurons. J. Cell Biol. 162, 613–622. Gitenay, D., Lallet-Daher, H., Bernard, D., 2014. Caspase-2 regulates oncogene-induced senescence. Oncotarget 5, 5845–5847. Graham, R.K., Deng, Y., Slow, E.J., Haigh, B., Bissada, N., Lu, G., Pearson, J., Shehadeh, J., Bertram, L., Murphy, Z., Warby, S.C., Doty, C.N., Roy, S., Wellington, C.L., Leavitt, B.R., Raymond, L.A., Nicholson, D.W., Hayden, M.R., 2006. Cleavage at the caspase-6 site is required for neuronal dysfunction and degeneration due to mutant huntingtin. Cell 125, 1179–1191. Guo, Y., Srinivasula, S.M., Druilhe, A., Fernandes-Alnemri, T., Alnemri, E.S., 2002. Caspase-2 induces apoptosis by releasing proapoptotic proteins from mitochondria. J. Biol. Chem. 277, 13430–13437. Ha, H., Kwak, H.B., Lee, S.W., Jin, H.M., Kim, H.M., Kim, H.H., Lee, Z.H., 2004. Reactive oxygen species mediate RANK signaling in osteoclasts. Exp. Cell Res. 301, 119–127. Han, J., Dinculescu, A., Dai, X., Du, W., Smith, W.C., Pang, J., 2013. Review: the history and role of naturally occurring mouse models with Pde6b mutations. Mol. Vis. 19, 2579–2589. Hara, H., Friedlander, R.M., Gagliardini, V., Ayata, C., Fink, K., Huang, Z., Shimizu-Sasamata, M., Yuan, J., Moskowitz, M.A., 1997. Inhibition of interleukin 1beta converting enzyme family proteases reduces ischemic and excitotoxic neuronal damage. Proc. Natl. Acad. Sci. U.S.A. 94, 2007–2012.

204

M.A. Miles et al.

Hasegawa, H., Yamada, Y., Tsukasaki, K., Mori, N., Tsuruda, K., Sasaki, D., Usui, T., Osaka, A., Atogami, S., Ishikawa, C., Machijima, Y., Sawada, S., Hayashi, T., Miyazaki, Y., Kamihira, S., 2011. LBH589, a deacetylase inhibitor, induces apoptosis in adult T-cell leukemia/lymphoma cells via activation of a novel RAIDD-caspase-2 pathway. Leukemia 25, 575–587. Hermel, E., Gafni, J., Propp, S.S., Leavitt, B.R., Wellington, C.L., Young, J.E., Hackam, A.S., Logvinova, A.V., Peel, A.L., Chen, S.F., Hook, V., Singaraja, R., Krajewski, S., Goldsmith, P.C., Ellerby, H.M., Hayden, M.R., Bredesen, D.E., Ellerby, L.M., 2004. Specific caspase interactions and amplification are involved in selective neuronal vulnerability in Huntington’s disease. Cell Death Differ. 11, 424–438. Ho, P.K., Jabbour, A.M., Ekert, P.G., Hawkins, C.J., 2005. Caspase-2 is resistant to inhibition by inhibitor of apoptosis proteins (IAPs) and can activate caspase-7. FEBS J. 272, 1401–1414. Ho, L.H., Read, S.H., Dorstyn, L., Lambrusco, L., Kumar, S., 2008. Caspase-2 is required for cell death induced by cytoskeletal disruption. Oncogene 27, 3393–3404. Ho, L.H., Taylor, R., Dorstyn, L., Cakouros, D., Bouillet, P., Kumar, S., 2009. A tumor suppressor function for caspase-2. Proc. Natl. Acad. Sci. U.S.A. 106, 5336–5341. Holleman, A., Den Boer, M.L., Kazemier, K.M., Beverloo, H.B., Von Bergh, A.R., Janka-Schaub, G.E., Pieters, R., 2005. Decreased PARP and procaspase-2 protein levels are associated with cellular drug resistance in childhood acute lymphoblastic leukemia. Blood 106, 1817–1823. Honda, H., Nagamachi, A., Inaba, T., 2015. -7/7q- syndrome in myeloid-lineage hematopoietic malignancies: attempts to understand this complex disease entity. Oncogene 34, 2413–2425. http://dx.doi.org/10.1038/onc.2014.196. Epub 2014 Jul 7. Hu, Q., Wu, D., Chen, W., Yan, Z., Yan, C., He, T., Liang, Q., Shi, Y., 2014. Molecular determinants of caspase-9 activation by the Apaf-1 apoptosome. Proc. Natl. Acad. Sci. U.S.A. 111, 16254–16261. Hulea, L., Nepveu, A., 2012. CUX1 transcription factors: from biochemical activities and cell-based assays to mouse models and human diseases. Gene 497, 18–26. Imre, G., Heering, J., Takeda, A.N., Husmann, M., Thiede, B., Zu Heringdorf, D.M., Green, D.R., Van Der Goot, F.G., Sinha, B., Dotsch, V., Rajalingam, K., 2012. Caspase-2 is an initiator caspase responsible for pore-forming toxin-mediated apoptosis. EMBO J. 31, 2615–2628. Ioannou, K., Samara, P., Livaniou, E., Derhovanessian, E., Tsitsilonis, O.E., 2012. Prothymosin alpha: a ubiquitous polypeptide with potential use in cancer diagnosis and therapy. Cancer Immunol. Immunother. 61, 599–614. Ionita, M.A., Pittler, S.J., 2007. Focus on molecules: rod cGMP phosphodiesterase type 6. Exp. Eye Res. 84, 1–2. Epub 2006 Mar 23. Jang, T.H., Park, H.H., 2013. PIDD mediates and stabilizes the interaction between RAIDD and caspase-2 for the PIDDosome assembly. BMB Rep. 46, 471–476. Jang, T.H., Lim, I.H., Kim, C.M., Choi, J.Y., Kim, E.A., Lee, T.J., Park, H.H., 2016. Rescuing neuronal cell death by RAIDD- and PIDD-derived peptides and its implications for therapeutic intervention in neurodegenerative diseases. Sci. Rep. 6, 31198. Janssens, S., Tinel, A., Lippens, S., Tschopp, J., 2005. PIDD mediates NF-kappaB activation in response to DNA damage. Cell 123, 1079–1092. Jean, Y.Y., Ribe, E.M., Pero, M.E., Moskalenko, M., Iqbal, Z., Marks, L.J., Greene, L.A., Troy, C.M., 2013. Caspase-2 is essential for c-Jun transcriptional activation and Bim induction in neuron death. Biochem. J. 455, 15–25. Jelinek, M., Balusikova, K., Kopperova, D., Nemcova-Furstova, V., Sramek, J., Fidlerova, J., Zanardi, I., Ojima, I., Kovar, J., 2013. Caspase-2 is involved in cell death induction by taxanes in breast cancer cells. Cancer Cell Int. 13, 42.

Old and Novel Functions of Caspase-2

205

Jobgen, W., Fu, W.J., Gao, H., Li, P., Meininger, C.J., Smith, S.B., Spencer, T.E., Wu, G., 2009. High fat feeding and dietary L-arginine supplementation differentially regulate gene expression in rat white adipose tissue. Amino Acids 37, 187–198. Julien, O., Zhuang, M., Wiita, A.P., O’donoghue, A.J., Knudsen, G.M., Craik, C.S., Wells, J.A., 2016. Quantitative MS-based enzymology of caspases reveals distinct protein substrate specificities, hierarchies, and cellular roles. Proc. Natl. Acad. Sci. U.S.A. 113, E2001–E2010. Karapetian, R.N., Evstafieva, A.G., Abaeva, I.S., Chichkova, N.V., Filonov, G.S., Rubtsov, Y.P., Sukhacheva, E.A., Melnikov, S.V., Schneider, U., Wanker, E.E., Vartapetian, A.B., 2005. Nuclear oncoprotein prothymosin alpha is a partner of Keap1: implications for expression of oxidative stress-protecting genes. Mol. Cell. Biol. 25, 1089–1099. Karetsou, Z., Martic, G., Tavoulari, S., Christoforidis, S., Wilm, M., Gruss, C., Papamarcaki, T., 2004. Prothymosin alpha associates with the oncoprotein SET and is involved in chromatin decondensation. FEBS Lett. 577, 496–500. Katagiri, T., Takahashi, N., 2002. Regulatory mechanisms of osteoblast and osteoclast differentiation. Oral Dis. 8, 147–159. Kim, M.S., Chung, N.G., Yoo, N.J., Lee, S.H., 2011a. Somatic mutation of proapoptotic caspase-2 gene is rare in acute leukemias and common solid cancers. Eur. J. Haematol. 86, 449–450. Kim, M.S., Kim, H.S., Jeong, E.G., Soung, Y.H., Yoo, N.J., Lee, S.H., 2011b. Somatic mutations of caspase-2 gene in gastric and colorectal cancers. Pathol. Res. Pract. 207, 640–644. Kitevska, T., Spencer, D.M., Hawkins, C.J., 2009. Caspase-2: controversial killer or checkpoint controller? Apoptosis 14, 829–848. Kitevska, T., Roberts, S.J., Pantaki-Eimany, D., Boyd, S.E., Scott, F.L., Hawkins, C.J., 2014. Analysis of the minimal specificity of caspase-2 and identification of Ac-VDTTD-AFC as a caspase-2-selective peptide substrate. Biosci. Rep. 34, e00100. Klotz, L.O., Sanchez-Ramos, C., Prieto-Arroyo, I., Urbanek, P., Steinbrenner, H., Monsalve, M., 2015. Redox regulation of FoxO transcription factors. Redox Biol. 6, 51–72. Koenekoop, R.K., 2009. Why do cone photoreceptors die in rod-specific forms of retinal degenerations? Ophthalmic Genet. 30, 152–154. Kumar, S., White, D.L., Takai, S., Turczynowicz, S., Juttner, C.A., Hughes, T.P., 1995. Apoptosis regulatory gene NEDD2 maps to human chromosome segment 7q34-35, a region frequently affected in haematological neoplasms. Hum. Genet. 95, 641–644. Lamkanfi, M., Dixit, V.M., 2014. Mechanisms and functions of inflammasomes. Cell 157, 1013–1022. Lee, C.Y., Cantle, J.P., Yang, X.W., 2013. Genetic manipulations of mutant huntingtin in mice: new insights into Huntington’s disease pathogenesis. FEBS J. 280, 4382–4394. Lezina, L., Purmessur, N., Antonov, A.V., Ivanova, T., Karpova, E., Krishan, K., Ivan, M., Aksenova, V., Tentler, D., Garabadgiu, A.V., Melino, G., Barlev, N.A., 2013. miR-16 and miR-26a target checkpoint kinases Wee1 and Chk1 in response to p53 activation by genotoxic stress. Cell Death Dis. 4, e953. Li, X., He, Y., 2012. Caspase-2-dependent dendritic cell death, maturation, and priming of T cells in response to Brucella abortus infection. PLoS One 7, e43512. Li, D., Li, X., Wu, J., Li, J., Zhang, L., Xiong, T., Tang, J., Qu, Y., Mu, D., 2015. Involvement of the JNK/FOXO3a/Bim pathway in neuronal apoptosis after hypoxic-ischemic brain damage in neonatal rats. PLoS One 10, e0132998. Lin, Y., Ma, W., Benchimol, S., 2000. PIDD, a new death-domain-containing protein, is induced by p53 and promotes apoptosis. Nat. Genet. 26, 122–127.

206

M.A. Miles et al.

Liu, Y., Yan, H., Chen, S., Sabel, B.A., 2015. Caspase-3 inhibitor Z-DEVD-FMK enhances retinal ganglion cell survival and vision restoration after rabbit traumatic optic nerve injury. Restor. Neurol. Neurosci. 33, 205–220. Logette, E., Le Jossic-Corcos, C., Masson, D., Solier, S., Sequeira-Legrand, A., Dugail, I., Lemaire-Ewing, S., Desoche, L., Solary, E., Corcos, L., 2005. Caspase-2, a novel lipid sensor under the control of sterol regulatory element binding protein 2. Mol. Cell. Biol. 25, 9621–9631. Lopez-Cruzan, M., Herman, B., 2013. Loss of caspase-2 accelerates age-dependent alterations in mitochondrial production of reactive oxygen species. Biogerontology 14, 121–130. Luftig, M.A., 2014. Viruses and the DNA damage response: activation and antagonism. Annu. Rev. Virol. 1, 605–625. Maag, R.S., Mancini, M., Rosen, A., Machamer, C.E., 2005. Caspase-resistant Golgin-160 disrupts apoptosis induced by secretory pathway stress and ligation of death receptors. Mol. Biol. Cell 16, 3019–3027. Machado, M.V., Michelotti, G.A., Pereira Tde, A., Boursier, J., Kruger, L., Swiderska-Syn, M., Karaca, G., Xie, G., Guy, C.D., Bohinc, B., Lindblom, K.R., Johnson, E., Kornbluth, S., Diehl, A.M., 2015. Reduced lipoapoptosis, hedgehog pathway activation and fibrosis in caspase-2 deficient mice with non-alcoholic steatohepatitis. Gut 64, 1148–1157. Machado, M.V., Michelotti, G.A., Jewell, M.L., Pereira, T.A., Xie, G., Premont, R.T., Diehl, A.M., 2016. Caspase-2 promotes obesity, the metabolic syndrome and nonalcoholic fatty liver disease. Cell Death Dis. 7, e2096. Mahoney, D.J., Lefebvre, C., Allan, K., Brun, J., Sanaei, C.A., Baird, S., Pearce, N., Gronberg, S., Wilson, B., Prakesh, M., Aman, A., Isaac, M., Mamai, A., Uehling, D., Al-Awar, R., Falls, T., Alain, T., Stojdl, D.F., 2011. Virus-tumor interactome screen reveals ER stress response can reprogram resistant cancers for oncolytic virus-triggered caspase-2 cell death. Cancer Cell 20, 443–456. Maillard, M.C., Brookfield, F.A., Courtney, S.M., Eustache, F.M., Gemkow, M.J., Handel, R.K., Johnson, L.C., Johnson, P.D., Kerry, M.A., Krieger, F., Meniconi, M., Munoz-Sanjuan, I., Palfrey, J.J., Park, H., Schaertl, S., Taylor, M.G., Weddell, D., Dominguez, C., 2011. Exploiting differences in caspase-2 and -3S(2) subsites for selectivity: structure-based design, solid-phase synthesis and in vitro activity of novel substrate-based caspase-2 inhibitors. Bioorg. Med. Chem. 19, 5833–5851. Mancini, M., Machamer, C.E., Roy, S., Nicholson, D.W., Thornberry, N.A., Casciola-Rosen, L.A., Rosen, A., 2000. Caspase-2 is localized at the Golgi complex and cleaves golgin-160 during apoptosis. J. Cell Biol. 149, 603–612. Manzl, C., Krumschnabel, G., Bock, F., Sohm, B., Labi, V., Baumgartner, F., Logette, E., Tschopp, J., Villunger, A., 2009. Caspase-2 activation in the absence of PIDDosome formation. J. Cell Biol. 185, 291–303. Manzl, C., Peintner, L., Krumschnabel, G., Bock, F., Labi, V., Drach, M., Newbold, A., Johnstone, R., Villunger, A., 2012. PIDDosome-independent tumor suppression by caspase-2. Cell Death Differ. 19, 1722–1732. Manzl, C., Fava, L.L., Krumschnabel, G., Peintner, L., Tanzer, M.C., Soratroi, C., Bock, F.J., Schuler, F., Luef, B., Geley, S., Villunger, A., 2013. Death of p53-defective cells triggered by forced mitotic entry in the presence of DNA damage is not uniquely dependent on caspase-2 or the PIDDosome. Cell Death Dis. 4, e942. Marechal, A., Zou, L., 2013. DNA damage sensing by the ATM and ATR kinases. Cold Spring Harb. Perspect. Biol. 5, a012716. Mccoy, F., Eckard, L., Nutt, L.K., 2012. Janus-faced PIDD: a sensor for DNA damage-induced cell death or survival? Mol. Cell 47, 667–668.

Old and Novel Functions of Caspase-2

207

Mcstay, G.P., Salvesen, G.S., Green, D.R., 2008. Overlapping cleavage motif selectivity of caspases: implications for analysis of apoptotic pathways. Cell Death Differ. 15, 322–331. Monnier, P.P., D’onofrio, P.M., Magharious, M., Hollander, A.C., Tassew, N., Szydlowska, K., Tymianski, M., Koeberle, P.D., 2011. Involvement of caspase-6 and caspase-8 in neuronal apoptosis and the regenerative failure of injured retinal ganglion cells. J. Neurosci. 31, 10494–10505. Moreira, D., Diaz-Jullien, C., Sarandeses, C.S., Covelo, G., Barbeito, P., Freire, M., 2013. The influence of phosphorylation of prothymosin alpha on its nuclear import and antiapoptotic activity. Biochem. Cell Biol. 91, 265–269. Narine, K.A., Keuling, A.M., Gombos, R., Tron, V.A., Andrew, S.E., Young, L.C., 2010. Defining the DNA mismatch repair-dependent apoptotic pathway in primary cells: evidence for p53-independence and involvement of centrosomal caspase 2. DNA Repair (Amst) 9, 161–168. Naylor, P.H., Smith, M.R., Mutchnick, M.G., Naylor, C.W., Dosescu, J., Skunca, M., Moshier, J.A., 1996. Thymosin alpha 1 does not promote growth or oncogenic transformation. Int. J. Immunopharmacol. 18, 321–327. Nematollahi, L.A., Garza-Garcia, A., Bechara, C., Esposito, D., Morgner, N., Robinson, C.V., Driscoll, P.C., 2015. Flexible stoichiometry and asymmetry of the PIDDosome core complex by heteronuclear NMR spectroscopy and mass spectrometry. J. Mol. Biol. 427, 737–752. Niizuma, K., Endo, H., Nito, C., Myer, D.J., Kim, G.S., Chan, P.H., 2008. The PIDDosome mediates delayed death of hippocampal CA1 neurons after transient global cerebral ischemia in rats. Proc. Natl. Acad. Sci. U.S.A. 105, 16368–16373. Nutt, L.K., Margolis, S.S., Jensen, M., Herman, C.E., Dunphy, W.G., Rathmell, J.C., Kornbluth, S., 2005. Metabolic regulation of oocyte cell death through the CaMKII-mediated phosphorylation of caspase-2. Cell 123, 89–103. Nutt, L.K., Buchakjian, M.R., Gan, E., Darbandi, R., Yoon, S.Y., Wu, J.Q., Miyamoto, Y.J., Gibbons, J.A., Andersen, J.L., Freel, C.D., Tang, W., He, C., Kurokawa, M., Wang, Y., Margolis, S.S., Fissore, R.A., Kornbluth, S., 2009. Metabolic control of oocyte apoptosis mediated by 14-3-3zeta-regulated dephosphorylation of caspase-2. Dev. Cell 16, 856–866. Oberst, A., Pop, C., Tremblay, A.G., Blais, V., Denault, J.B., Salvesen, G.S., Green, D.R., 2010. Inducible dimerization and inducible cleavage reveal a requirement for both processes in caspase-8 activation. J. Biol. Chem. 285, 16632–16642. Okuwaki, M., Kato, K., Nagata, K., 2010. Functional characterization of human nucleosome assembly protein 1-like proteins as histone chaperones. Genes Cells 15, 13–27. Oliver, T.G., Meylan, E., Chang, G.P., Xue, W., Burke, J.R., Humpton, T.J., Hubbard, D., Bhutkar, A., Jacks, T., 2011. Caspase-2-mediated cleavage of Mdm2 creates a p53induced positive feedback loop. Mol. Cell 43, 57–71. O’Reilly, L.A., Ekert, P., Harvey, N., Marsden, V., Cullen, L., Vaux, D.L., Hacker, G., Magnusson, C., Pakusch, M., Cecconi, F., Kuida, K., Strasser, A., Huang, D.C., Kumar, S., 2002. Caspase-2 is not required for thymocyte or neuronal apoptosis even though cleavage of caspase-2 is dependent on both Apaf-1 and caspase-9. Cell Death Differ. 9, 832–841. Parsons, M.J., Mccormick, L., Janke, L., Howard, A., Bouchier-Hayes, L., Green, D.R., 2013. Genetic deletion of caspase-2 accelerates MMTV/c-neu-driven mammary carcinogenesis in mice. Cell Death Differ. 20, 1174–1182. Parsons, M.J., Fassio, S.R., Bouchier-Hayes, L., 2016. Detection of initiator caspase induced proximity in single cells by caspase bimolecular fluorescence complementation. Methods Mol. Biol. 1419, 41–56.

208

M.A. Miles et al.

Peintner, L., Dorstyn, L., Kumar, S., Aneichyk, T., Villunger, A., Manzl, C., 2015. The tumor-modulatory effects of caspase-2 and Pidd1 do not require the scaffold protein Raidd. Cell Death Differ. 22, 1803–1811. Pereira, N.A., Song, Z., 2008. Some commonly used caspase substrates and inhibitors lack the specificity required to monitor individual caspase activity. Biochem. Biophys. Res. Commun. 377, 873–877. Pineiro, A., Cordero, O.J., Nogueira, M., 2000. Fifteen years of prothymosin alpha: contradictory past and new horizons. Peptides 21, 1433–1446. Puccini, J., Shalini, S., Voss, A.K., Gatei, M., Wilson, C.H., Hiwase, D.K., Lavin, M.F., Dorstyn, L., Kumar, S., 2013. Loss of caspase-2 augments lymphomagenesis and enhances genomic instability in Atm-deficient mice. Proc. Natl. Acad. Sci. U.S.A. 110, 19920–19925. Punzo, C., Kornacker, K., Cepko, C.L., 2009. Stimulation of the insulin/mTOR pathway delays cone death in a mouse model of retinitis pigmentosa. Nat. Neurosci. 12, 44–52. Punzo, C., Xiong, W., Cepko, C.L., 2012. Loss of daylight vision in retinal degeneration: are oxidative stress and metabolic dysregulation to blame? J. Biol. Chem. 287, 1642–1648. Qi, X., Wang, L., Du, F., 2010. Novel small molecules relieve prothymosin alpha-mediated inhibition of apoptosome formation by blocking its interaction with Apaf-1. Biochemistry 49, 1923–1930. Ramdzan, Z.M., Nepveu, A., 2014. CUX1, a haploinsufficient tumour suppressor gene overexpressed in advanced cancers. Nat. Rev. Cancer 14, 673–682. Ramdzan, Z.M., Vadnais, C., Pal, R., Vandal, G., Cadieux, C., Leduy, L., Davoudi, S., Hulea, L., Yao, L., Karnezis, A.N., Paquet, M., Dankort, D., Nepveu, A., 2014. RAS transformation requires CUX1-dependent repair of oxidative DNA damage. PLoS Biol. 12, e1001807. Ramdzan, Z.M., Pal, R., Kaur, S., Leduy, L., Berube, G., Davoudi, S., Vadnais, C., Nepveu, A., 2015. The function of CUX1 in oxidative DNA damage repair is needed to prevent premature senescence of mouse embryo fibroblasts. Oncotarget 6, 3613–3626. Read, S.H., Baliga, B.C., Ekert, P.G., Vaux, D.L., Kumar, S., 2002. A novel Apaf-1-independent putative caspase-2 activation complex. J. Cell Biol. 159, 739–745. Ribe, E.M., Jean, Y.Y., Goldstein, R.L., Manzl, C., Stefanis, L., Villunger, A., Troy, C.M., 2012. Neuronal caspase 2 activity and function requires RAIDD, but not PIDD. Biochem. J. 444, 591–599. Romero-Brey, I., Bartenschlager, R., 2016. Endoplasmic reticulum: the favorite intracellular niche for viral replication and assembly. Viruses 8 (6), E160. Sandow, J.J., Dorstyn, L., O’reilly, L.A., Tailler, M., Kumar, S., Strasser, A., Ekert, P.G., 2013. ER stress does not cause upregulation and activation of caspase-2 to initiate apoptosis. Cell Death Differ. 21, 475–480. Sansregret, L., Vadnais, C., Livingstone, J., Kwiatkowski, N., Awan, A., Cadieux, C., Leduy, L., Hallett, M.T., Nepveu, A., 2011. Cut homeobox 1 causes chromosomal instability by promoting bipolar division after cytokinesis failure. Proc. Natl. Acad. Sci. U.S.A. 108, 1949–1954. Sashida, G., Iwama, A., 2016. Multifaceted role of the polycomb-group gene EZH2 in hematological malignancies. Int. J. Hematol. 9, 9. Schleich, K., Buchbinder, J.H., Pietkiewicz, S., Kahne, T., Warnken, U., Ozturk, S., Schnolzer, M., Naumann, M., Krammer, P.H., Lavrik, I.N., 2016. Molecular architecture of the DED chains at the DISC: regulation of procaspase-8 activation by short DED proteins c-FLIP and procaspase-8 prodomain. Cell Death Differ. 23, 681–694. Schweizer, A., Roschitzki-Voser, H., Amstutz, P., Briand, C., Gulotti-Georgieva, M., Prenosil, E., Binz, H.K., Capitani, G., Baici, A., Pluckthun, A., Grutter, M.G., 2007.

Old and Novel Functions of Caspase-2

209

Inhibition of caspase-2 by a designed ankyrin repeat protein: specificity, structure, and inhibition mechanism. Structure 15, 625–636. Segade, F., Gomez-Marquez, J., 1999. Prothymosin alpha. Int. J. Biochem. Cell Biol. 31, 1243–1248. Shalini, S., Dorstyn, L., Wilson, C., Puccini, J., Ho, L., Kumar, S., 2012. Impaired antioxidant defence and accumulation of oxidative stress in caspase-2-deficient mice. Cell Death Differ. 19, 1370–1380. Shalini, S., Dorstyn, L., Dawar, S., Kumar, S., 2015a. Old, new and emerging functions of caspases. Cell Death Differ. 22, 526–539. Shalini, S., Puccini, J., Wilson, C.H., Finnie, J., Dorstyn, L., Kumar, S., 2015b. Caspase-2 protects against oxidative stress in vivo. Oncogene 34, 4995–5002. Shalini, S., Nikolic, A., Wilson, C.H., Puccini, J., Sladojevic, N., Finnie, J., Dorstyn, L., Kumar, S., 2016. Caspase-2 deficiency accelerates chemically induced liver cancer in mice. Cell Death Differ. 12, 81. Shin, S., Lee, Y., Kim, W., Ko, H., Choi, H., Kim, K., 2005. Caspase-2 primes cancer cells for TRAIL-mediated apoptosis by processing procaspase-8. EMBO J. 24, 3532–3542. Sidi, S., Sanda, T., Kennedy, R.D., Hagen, A.T., Jette, C.A., Hoffmans, R., Pascual, J., Imamura, S., Kishi, S., Amatruda, J.F., Kanki, J.P., Green, D.R., D’andrea, A.A., Look, A.T., 2008. Chk1 suppresses a caspase-2 apoptotic response to DNA damage that bypasses p53, Bcl-2, and caspase-3. Cell 133, 864–877. Singh, H., Lane, A.A., Correll, M., Przychodzen, B., Sykes, D.B., Stone, R.M., Ballen, K.K., Amrein, P.C., Maciejewski, J., Attar, E.C., 2013. Putative RNA-splicing gene LUC7L2 on 7q34 represents a candidate gene in pathogenesis of myeloid malignancies. Blood Cancer J. 3e117. Sohn, D., Budach, W., Janicke, R.U., 2011. Caspase-2 is required for DNA damage-induced expression of the CDK inhibitor p21(WAF1/CIP1). Cell Death Differ. 18, 1664–1674. Strasser, B., Mlitz, V., Fischer, H., Tschachler, E., Eckhart, L., 2015. Comparative genomics reveals conservation of filaggrin and loss of caspase-14 in dolphins. Exp. Dermatol. 24, 365–369. Su, Y.C., Ou, H.Y., Wu, H.T., Wu, P., Chen, Y.C., Su, B.H., Shiau, A.L., Chang, C.J., Wu, C.L., 2015. Prothymosin-alpha overexpression contributes to the development of insulin resistance. J. Clin. Endocrinol. Metab. 100, 4114–4123. Talanian, R.V., Quinlan, C., Trautz, S., Hackett, M.C., Mankovich, J.A., Banach, D., Ghayur, T., Brady, K.D., Wong, W.W., 1997. Substrate specificities of caspase family proteases. J. Biol. Chem. 272, 9677–9682. Tang, Y., Wells, J.A., Arkin, M.R., 2011. Structural and enzymatic insights into caspase-2 protein substrate recognition and catalysis. J. Biol. Chem. 286, 34147–34154. Telliez, J.B., Bean, K.M., Lin, L.L., 2000. LRDD, a novel leucine rich repeat and death domain containing protein. Biochim. Biophys. Acta 1478, 280–288. Tenev, T., Bianchi, K., Darding, M., Broemer, M., Langlais, C., Wallberg, F., Zachariou, A., Lopez, J., Macfarlane, M., Cain, K., Meier, P., 2011. The Ripoptosome, a signaling platform that assembles in response to genotoxic stress and loss of IAPs. Mol. Cell 43, 432–448. Terry, M.R., Arya, R., Mukhopadhyay, A., Berrett, K.C., Clair, P.M., Witt, B., Salama, M.E., Bhutkar, A., Oliver, T.G., 2015. Caspase-2 impacts lung tumorigenesis and chemotherapy response in vivo. Cell Death Differ. 22, 719–730. Thompson, R., Shah, R.B., Liu, P.H., Gupta, Y.K., Ando, K., Aggarwal, A.K., Sidi, S., 2015. An inhibitor of PIDDosome formation. Mol. Cell 58, 767–779. Thornberry, N., Rano, T., Peterson, E., Rasper, D., Timkey, T., Garcia-Calvo, M., Houtzager, V., Nordstrom, P., Roy, S., Vaillancourt, J., Chapman, K., Nicholson, D., 1997. A combinatorial approach defines specificities of members of

210

M.A. Miles et al.

the caspase family and granzyme B. Functional relationships established for key mediators of apoptosis. J. Biol. Chem. 272, 17907–17911. Timmer, J.C., Zhu, W., Pop, C., Regan, T., Snipas, S.J., Eroshkin, A.M., Riedl, S.J., Salvesen, G.S., 2009. Structural and kinetic determinants of protease substrates. Nat. Struct. Mol. Biol. 16, 1101–1108. Tinel, A., Tschopp, J., 2004. The PIDDosome, a protein complex implicated in activation of caspase-2 in response to genotoxic stress. Science 304, 843–846. Tinel, A., Janssens, S., Lippens, S., Cuenin, S., Logette, E., Jaccard, B., Quadroni, M., Tschopp, J., 2007. Autoproteolysis of PIDD marks the bifurcation between pro-death caspase-2 and pro-survival NF-kappaB pathway. EMBO J. 26, 197–208. Tiwari, M., Lopez-Cruzan, M., Morgan, W.W., Herman, B., 2011. Loss of caspase-2dependent apoptosis induces autophagy after mitochondrial oxidative stress in primary cultures of young adult cortical neurons. J. Biol. Chem. 286, 8493–8506. Tiwari, M., Sharma, L.K., Vanegas, D., Callaway, D.A., Bai, Y., Lechleiter, J.D., Herman, B., 2014. A nonapoptotic role for CASP2/caspase 2: modulation of autophagy. Autophagy 10, 1054–1070. Tizon, B., Ribe, E.M., Mi, W., Troy, C.M., Levy, E., 2010. Cystatin C protects neuronal cells from amyloid-beta-induced toxicity. J. Alzheimers Dis. 19, 885–894. Troy, C.M., Stefanis, L., Greene, L.A., Shelanski, M.L., 1997. Nedd2 is required for apoptosis after trophic factor withdrawal, but not superoxide dismutase (SOD1) downregulation, in sympathetic neurons and pc12 cells. J. Neurosci. 17, 1911–1918. Troy, C.M., Rabacchi, S.A., Friedman, W.J., Frappier, T.F., Brown, K., Shelanski, M.L., 2000. Caspase-2 mediates neuronal cell death induced by beta-amyloid. J. Neurosci. 20, 1386–1392. Troy, C.M., Rabacchi, S.A., Hohl, J.B., Angelastro, J.M., Greene, L.A., Shelanski, M.L., 2001. Death in the balance: alternative participation of the caspase-2 and -9 pathways in neuronal death induced by nerve growth factor deprivation. J. Neurosci. 21, 5007–5016. Truscott, M., Denault, J.B., Goulet, B., Leduy, L., Salvesen, G.S., Nepveu, A., 2007. Carboxyl-terminal proteolytic processing of CUX1 by a caspase enables transcriptional activation in proliferating cells. J. Biol. Chem. 282, 30216–30226. Tu, S., Mcstay, G.P., Boucher, L.M., Mak, T., Beere, H.M., Green, D.R., 2006. In situ trapping of activated initiator caspases reveals a role for caspase-2 in heat shock-induced apoptosis. Nat. Cell Biol. 8, 72–77. Turowec, J.P., Zukowski, S.A., Knight, J.D., Smalley, D.M., Graves, L.M., Johnson, G.L., Li, S.S., Lajoie, G.A., Litchfield, D.W., 2014. An unbiased proteomic screen reveals caspase cleavage is positively and negatively regulated by substrate phosphorylation. Mol. Cell. Proteomics 13, 1184–1197. Upton, J.P., Austgen, K., Nishino, M., Coakley, K.M., Hagen, A., Han, D., Papa, F.R., Oakes, S.A., 2008. Caspase-2 cleavage of BID is a critical apoptotic signal downstream of endoplasmic reticulum stress. Mol. Cell. Biol. 28, 3943–3951. Upton, J.P., Wang, L., Han, D., Wang, E.S., Huskey, N.E., Lim, L., Truitt, M., Mcmanus, M.T., Ruggero, D., Goga, A., Papa, F.R., Oakes, S.A., 2012. IRE1alpha cleaves select microRNAs during ER stress to derepress translation of proapoptotic caspase-2. Science 338, 818–822. Vakifahmetoglu, H., Olsson, M., Orrenius, S., Zhivotovsky, B., 2006. Functional connection between p53 and caspase-2 is essential for apoptosis induced by DNA damage. Oncogene 25, 5683–5692. Vakifahmetoglu-Norberg, H., Norberg, E., Perdomo, A.B., Olsson, M., Ciccosanti, F., Orrenius, S., Fimia, G.M., Piacentini, M., Zhivotovsky, B., 2013. Caspase-2 promotes cytoskeleton protein degradation during apoptotic cell death. Cell Death Dis. 4e940.

Old and Novel Functions of Caspase-2

211

Van De Craen, M., Declercq, W., Van Den Brande, I., Fiers, W., Vandenabeele, P., 1999. The proteolytic procaspase activation network: an in vitro analysis. Cell Death Differ. 6, 1117–1124. Vareli, K., Frangou-Lazaridis, M., 2004. Prothymosin alpha is localized in mitotic spindle during mitosis. Biol. Cell 96, 421–428. Venkatesh, A., Ma, S., Le, Y.Z., Hall, M.N., Ruegg, M.A., Punzo, C., 2015. Activated mTORC1 promotes long-term cone survival in retinitis pigmentosa mice. J. Clin. Invest. 125, 1446–1458. Viana, R.J., Ramalho, R.M., Nunes, A.F., Steer, C.J., Rodrigues, C.M., 2010. Modulation of amyloid-beta peptide-induced toxicity through inhibition of JNK nuclear localization and caspase-2 activation. J. Alzheimers Dis. 22, 557–568. Vigneswara, V., Ahmed, Z., 2016. Long-term neuroprotection of retinal ganglion cells by inhibiting caspase-2. Cell Death Discov. 2, 16044. Vigneswara, V., Berry, M., Logan, A., Ahmed, Z., 2012. Pharmacological inhibition of caspase-2 protects axotomised retinal ganglion cells from apoptosis in adult rats. PLoS One 7e53473. Vigneswara, V., Berry, M., Logan, A., Ahmed, Z., 2013. Caspase-2 is upregulated after sciatic nerve transection and its inhibition protects dorsal root ganglion neurons from apoptosis after serum withdrawal. PLoS One 8e57861. Vigneswara, V., Akpan, N., Berry, M., Logan, A., Troy, C.M., Ahmed, Z., 2014. Combined suppression of CASP2 and CASP6 protects retinal ganglion cells from apoptosis and promotes axon regeneration through CNTF-mediated JAK/STAT signalling. Brain 137, 1656–1675. Villunger, A., Michalak, E.M., Coultas, L., Mullauer, F., Bock, G., Ausserlechner, M.J., Adams, J.M., Strasser, A., 2003. p53- and drug-induced apoptotic responses mediated by BH3-only proteins Puma and Noxa. Science 302, 1036–1038. Weiss, W.A., Aldape, K., Mohapatra, G., Feuerstein, B.G., Bishop, J.M., 1997. Targeted expression of MYCN causes neuroblastoma in transgenic mice. EMBO J. 16, 2985–2995. Wejda, M., Impens, F., Takahashi, N., Van Damme, P., Gevaert, K., Vandenabeele, P., 2012. Degradomics reveals that cleavage specificity profiles of caspase-2 and effector caspases are alike. J. Biol. Chem. 287, 33983–33995. Wellington, C.L., Ellerby, L.M., Gutekunst, C.A., Rogers, D., Warby, S., Graham, R.K., Loubser, O., Van Raamsdonk, J., Singaraja, R., Yang, Y.Z., Gafni, J., Bredesen, D., Hersch, S.M., Leavitt, B.R., Roy, S., Nicholson, D.W., Hayden, M.R., 2002. Caspase cleavage of mutant huntingtin precedes neurodegeneration in Huntington’s disease. J. Neurosci. 22, 7862–7872. Wilson, C.H., Shalini, S., Filipovska, A., Richman, T.R., Davies, S., Martin, S.D., Mcgee, S.L., Puccini, J., Nikolic, A., Dorstyn, L., Kumar, S., 2015. Age-related proteostasis and metabolic alterations in caspase-2-deficient mice. Cell Death Dis. 6e1615. Wilson, C.H., Nikolic, A., Kentish, S.J., Shalini, S., Hatzinikolas, G., Page, A.J., Dorstyn, L., Kumar, S., 2016. Sex-specific alterations in glucose homeostasis and metabolic parameters during ageing of caspase-2-deficient mice. Cell Death Discov. 2, 16009. Wong, C.C., Martincorena, I., Rust, A.G., Rashid, M., Alifrangis, C., Alexandrov, L.B., Tiffen, J.C., Kober, C., Green, A.R., Massie, C.E., Nangalia, J., Lempidaki, S., Dohner, H., Dohner, K., Bray, S.J., Mcdermott, U., Papaemmanuil, E., Campbell, P.J., Adams, D.J., 2014. Inactivating CUX1 mutations promote tumorigenesis. Nat. Genet. 46, 33–38. Xiong, W., Maccoll Garfinkel, A.E., Li, Y., Benowitz, L.I., Cepko, C.L., 2015. NRF2 promotes neuronal survival in neurodegeneration and acute nerve damage. J. Clin. Invest. 125, 1433–1445.

212

M.A. Miles et al.

Yang, C.S., Matsuura, K., Huang, N.J., Robeson, A.C., Huang, B., Zhang, L., Kornbluth, S., 2015. Fatty acid synthase inhibition engages a novel caspase-2 regulatory mechanism to induce ovarian cancer cell death. Oncogene 34, 3264–3272. Zha, J., Weiler, S., Oh, K.J., Wei, M.C., Korsmeyer, S.J., 2000. Posttranslational N-myristoylation of BID as a molecular switch for targeting mitochondria and apoptosis. Science 290, 1761–1765. Zhang, Y., Padalecki, S.S., Chaudhuri, A.R., De Waal, E., Goins, B.A., Grubbs, B., Ikeno, Y., Richardson, A., Mundy, G.R., Herman, B., 2007. Caspase-2 deficiency enhances aging-related traits in mice. Mech. Ageing Dev. 128, 213–221. Zhou, R., Yazdi, A.S., Menu, P., Tschopp, J., 2011. A role for mitochondria in NLRP3 inflammasome activation. Nature 469, 221–225.

CHAPTER FOUR

Metabolic Reprogramming and Oncogenesis: One Hallmark, Many Organelles A.S.H. Costa, C. Frezza1 Medical Research Council Cancer Unit, University of Cambridge, Hutchison/MRC Research Centre, Cambridge, ENG, United Kingdom 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Hallmarks of Metabolic Transformation in Cancer Cells 3. Compartmentalization of Metabolism 3.1 Mitochondria 3.2 Endoplasmic Reticulum 3.3 Peroxisomes 3.4 Lysosomes 4. Metabolic Cooperation Among Organelles 5. Challenges and Future Directions 6. Conclusions Acknowledgments References

214 214 217 218 219 220 222 223 225 226 227 227

Abstract The process of tumorigenesis can be described by a series of molecular features, among which alteration of cellular metabolism has recently emerged. This metabolic rewiring fulfills the energy and biosynthetic demands of fast proliferating cancer cells and amplifies their metabolic repertoire to survive and proliferate in the poorly oxygenated and nutrient-deprived tumor microenvironment. During the last decade, the complex reprogramming of cancer cell metabolism has been widely investigated, revealing cancer-specific metabolic alterations. These include dysregulation of glucose and glutamine metabolism, alterations of lipid synthesis and oxidation, and a complex rewiring of mitochondrial function. However, mitochondria are not the only metabolically active organelles within the cell, and other organelles, including lysosomes, peroxisomes, and endoplasmic reticulum, harbor components of the metabolic network. Of note, dysregulation of the function of these organelles is increasingly recognized in cancer cells. However, to what extent these organelles contribute to the metabolic reprogramming of cancer is not fully understood. In this review, we describe the main metabolic

International Review of Cell and Molecular Biology, Volume 332 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2017.01.001

#

2017 Elsevier Inc. All rights reserved.

213

214

A.S.H. Costa and C. Frezza

functions of these organelles and provide insights into how they communicate to orchestrate a coordinated metabolic reprogramming during transformation.

1. INTRODUCTION Tumorigenesis is a complex and multifaceted process that ultimately leads to unrestrained growth and proliferation of transformed cells. Over the past decades, researchers have attempted to unravel the complexity of the cellular events that drive tumor formation and its evolution. While the mechanisms that lead to transformation are now established (Hanahan et al., 2011), it is emerging that the core metabolic machinery of cancer cells undergoes complex rewiring to fulfill the energy and biosynthetic needs of cancer cells. Fuelled by the development and improvement of novel biochemical and analytical tools, the field of cancer cell metabolism has expanded together with our understanding of the tumor-associated metabolic transformation. The metabolism of glucose and glutamine has undoubtedly taken the spotlight in the study of cancer metabolism. Nevertheless, it is emerging that cancer cells are able to use many other nutrients, from the amino acids alanine (Sousa et al., 2016), cysteine, and methionine (Huang, 2002; Luo et al., 2010; Mo´dis et al., 2014), to fatty acids (Nath et al., 2016; Nieman et al., 2011), acetate (Schug et al., 2015), choline (Glunde et al., 2011; Zeng et al., 2014), as well as trace metals (Basu et al., 2013; Gupte et al., 2009) and vitamins (Feldman et al., 2014; Wu et al., 2013). What has been partially overlooked in this remarkable scientific endeavor is the notion that the metabolic network is compartmentalized and distributed across various intracellular organelles, including mitochondria, peroxisomes, lysosomes, and endoplasmic reticulum (ER). Importantly, the communication among these organelles is vital for the maintenance of a fully coordinated metabolic activity. Consistently, dysfunction of these organelles can contribute to cell transformation. In this review, we will describe the metabolic reprogramming of cancer focusing on the contribution of each of the different cell organelles and the small molecule metabolites that derive from their activity.

2. HALLMARKS OF METABOLIC TRANSFORMATION IN CANCER CELLS All cells require constant supply of nutrients from the environment. Nutrients are then broken down (catabolism) for generation of ATP,

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

215

required to support endoergonic reactions, and metabolic intermediates to maintain cell homeostasis, including DNA synthesis and protein turnover (anabolism). Malignant transformation of cells not only leads to increased nutrient uptake but also profoundly alters nutrient utilization, shifting the balance from catabolic to anabolic reactions. Systematic investigations of the metabolic need of cancer cells started in the first decades of the 20th century. Otto Warburg, the pioneer of the field, showed that cancer cells substantially increase the demand of glucose even when oxygen is plentiful (Warburg and Dickens, 1927). This process, also known as aerobic glycolysis, gives cancer cells several advantages, beyond generation of ATP. Indeed, a series of pathways branch out from glycolysis: (i) the pentose phosphate pathway (PPP), which generates building blocks for the synthesis of nucleotides and maintains cytosolic NADPH pool; (ii) the biosynthesis of hexosamine from fructose-6-phosphate for protein glycosylation; (iii) the biosynthesis of phospholipids for cell membrane; and (iv) the biosynthesis of serine and glycine for protein and nucleotide biosynthesis (Fig. 1A). Not surprisingly, all of these pathways have been reported to be dysregulated during tumor development and progression (DeBerardinis and Chandel, 2016). In cancer cells, reduced entry of glucose-derived carbons in the mitochondria is frequently mirrored by an increase in glutamine oxidation. The requirement of glutamine for cancer cells was originally demonstrated by Eagle, who showed that HeLa cells require glutamine in up to 100-fold molar excess, relative to other amino acids, in culture medium (Eagle, 1955). Through glutamine catabolism, cells generate carbon and nitrogen for the biosynthesis of lipids and various nitrogen-containing small molecule metabolites, including purine and pyrimidine nucleotides, glucosamine-6-phosphate, and nonessential amino acids (Pavlova et al., 2016). Cancer cells exhibit also substantial alterations of lipid metabolism. Cancer cells rely on the uptake of fatty acids and β-oxidation to sustain their survival and metastasis, even in tissues where the lipogenic activity is already high (Kamphorst et al., 2013). In fact, fatty acid β-oxidation is considered as the major bioenergetic pathway in nonglycolytic tumors (Ahmad et al., 2016; Deep and Schlaepfer, 2016). Moreover, the dependence of cancer cells on the oxidation of fatty acids is further heightened in nutrient- and oxygen-depleted environmental conditions (Eales et al., 2016). Fatty acid synthase expression is also known to be increased in certain cancers. The activity of this enzyme is linked to ether lipid production, which is reportedly increased in particularly aggressive tumors (Benjamin et al., 2013; Lodhi et al., 2014). Although it seems

216

A.S.H. Costa and C. Frezza

Fig. 1 The enhanced map of cancer metabolism. Schematic representation of the major hallmarks of the metabolic transformation in cancer (A) and the compartmentalization of major metabolic pathways across the cell (B) depicting also the metabolic cooperation among different organelles. 2-OG: 2-oxo-glutarate; 6PG, 6-phosphogluconate; ACOX: acyl-coenzyme A oxidase; Ac-CoA: acetyl coenzyme A; AMACR: alpha-methylacylCoA racemase; BCFA: branched chain fatty acids; FA: fatty acids; G6P: glucose 6-phosphate; GSH: glutathione; H6PDH: hexose-6-phosphate dehydrogenase; Idp2p:

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

217

counterintuitive that cancer cells activate both fatty acid oxidation (FAO) and synthesis, it has recently been proposed that these two pathways might coexist in cancer cells (Carracedo et al., 2013). Equally correlated with cancer aggressiveness is the increased lipid droplets and stored-cholesteryl ester content in tumors (Beloribi-Djefaflia et al., 2016). Another emerging source of nutrients for cancer cells is autophagy, a process whereby intracellular macromolecules are digested in regulated fashion to release metabolites that can be used for biosynthetic reactions. For instance, during metabolic stress conditions, and as the production of stress proteins increases, autophagy is the main source of amino acids. These amino acids thus obtained can be converted into intermediates of the tricarboxylic acid (TCA) cycle, thus contributing to generate ATP. Similarly, autophagy can be used to release lipids from intracellular lipid stores, lipid droplets, for subsequent oxidation (Singh et al., 2009). Interestingly, it has been shown that autophagy in stromal cell can release nutrients that are then used by cancer cells for bioenergetics purposes (Sousa et al., 2016). These results suggest that the role of autophagy as energy source might be broader that initially anticipated. As indicated in this brief overview on cancer metabolism, the substantial amount of work carried out in the last decade enabled scientists to draw a map of the major metabolic diversions that promote malignant transformation and/or elicit deregulated cell proliferation (Fig. 1A). However, this map does account for the distribution of these reactions across the many intracellular organelles in the cells. This aspect of cellular metabolism is the subject of the next paragraphs.

3. COMPARTMENTALIZATION OF METABOLISM Compartmentalization of metabolic reactions in distinct subcellular locations is a defining feature of eukaryotic cells and translates into a complex metabolic network connecting the different organelles. However, the fact that metabolic enzymes are often localized in multiple organelles and the lack of appropriate experimental tools to investigate metabolic trafficking are major barriers to a full understanding of cooperation between these cytosolic isocitrate dehydrogenase; Idp3p: peroxisomal isocitrate dehydrogenase; LDH: lactate dehydrogenase; MCFA/LCFA: medium/long-chain fatty acid; MCT: monocarboxylate transporter; OxPhos: oxidative phosphorylation; PPP: pentose phosphate pathway; ROS: reactive oxygen species; TCA: tricarboxylic acids; VLCFA/LCFA: very-long/long-chain fatty acids. Dashed lines indicate a series of reaction in a complex pathway, whereas solid lines indicate a single-step reaction.

218

A.S.H. Costa and C. Frezza

metabolic compartments. Here, we will briefly describe how metabolic reactions are compartmentalized within the cells, and how each organelle could contribute to the metabolic reprogramming of cancer (Fig. 1B).

3.1 Mitochondria Mitochondria are a defining feature of eukaryotic cells. These organelles are limited by a double membrane structure—the outer and the inner mitochondrial membrane. The outer mitochondrial membrane is permeable to small lipophilic molecules and presents channels that allow the import/export of small molecules. In contrast, the inner membrane is composed of a densely packed and hydrophobic combination of phospholipids, of which cardiolipin is unique to mitochondria, rendering it highly impermeable to solutes (Szeto and Schiller, 2011). This feature of the inner mitochondrial membrane allows the generation of a proton gradient, which is then utilized for generation of ATP via oxidative phosphorylation. From a metabolic point of view, mitochondria are key hubs for the cells. They are involved in FAO, glucose, and amino acid metabolism through the TCA cycle, and they generate ATP via oxidative phosphorylation. Furthermore, they are main site of generation of reactive oxygen species (ROS) and play key roles in regulation of cell signaling and cell death (Vyas et al., 2016; Fig. 1B). Due to their multiple roles in normal physiology, mitochondria are implicated in a plethora of human inherited disorders and in common diseases, including cancer. Mutations in nuclear and mitochondrial genes encoding for mitochondrial enzymes have been observed in a variety of tumors (Gaude and Frezza, 2014). As a consequence, for years, it was thought that mitochondrial function was lost in cancer cells. It is now clear that this view was imprecise and that mitochondria can coordinate many of the biosynthesis reactions required for the proliferation of cancer cells, even when partially dysfunctional. For instance, via reductive glutamine metabolism in the TCA cycle, mitochondria supply the cells with citrate that can be used for lipid biosynthesis and protein acetylation when oxygen is limiting (Metallo et al., 2012). Furthermore, mitochondrial function is required for the synthesis of aspartate (Birsoy et al., 2015; Sullivan et al., 2015), a key amino acid for nucleotide and protein biosynthesis. The description of the multiple ways mitochondria react to environmental and genetic cues is beyond the scope of the manuscript and we address the readers to comprehensive reviews on this topic (Weinberg and Chandel, 2014).

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

219

3.2 Endoplasmic Reticulum The ER consists of a continuous system of membrane sheets and tubules that contacts and participates in the communication with other cellular compartments, including lysosomes, peroxisomes, and mitochondria. The ER is a metabolically versatile organelle, with an almost ubiquitous participation in the cell’s anabolic and catabolic processes, including protein synthesis and degradation, gluconeogenesis, glycogen synthesis and breakdown, membrane lipid synthesis and recycling, lipid storage, and hormone and drug metabolism (Mandl et al., 2009). ER is also a main site of oxidative protein folding, which requires reduced glutathione (GSH) and NADPH, the two major redox buffers in the cell. However, these molecules are not freely available to ER and mechanisms are in place to guarantee their exchange with the cytosol. For instance, GSH is imported in the ER via yet-unidentified transporter and GSH/GSSH ratio is maintained via a complex redox relay that involves protein disulfide isomerase PDI and ER oxidoreductin 1-α (Ero1-α) (Oka and Bulleid, 2013). On the other hand, NADPH/NADP+ ratio in the ER is maintained by the activity of NADPH-generating hexose-6-phosphate dehydrogenase (H6PDH), which in turn, requires the transport of glucose-6-phosphate, its major substrate. ER is also an important site of lipid metabolism and hosts the machinery for desaturation, elongation, and esterification of phospholipids (Lagace and Ridgway, 2013; Fig. 1B). As an important site for protein folding, disulfide bond formation, longchain fatty acid extension, and sterol reduction, ER dysfunction could contribute to many pathologies, including cancer. At the same time, the process of transformation is thought to disrupt ER function. Indeed, due to elevated biomass generation, cancer cells increase the rate of protein folding and assembly in the ER, leading to ER stress. Furthermore, the accumulation of aberrantly folded protein in cancer cells is thought to alter protein homeostasis, and, as consequence, to activate the unfolded protein response (UPR). Other features of cancer, such as nutrient starvation, hypoxia, and changes in the redox status, can also contribute to the UPR (Ozcan and Tabas, 2012). Notably, UPR can act as an important cancer cell survival pathway (Bohnert et al., 2016; Cohen et al., 2015; Ryan et al., 2016). Beside the role of ER stress in orchestrating UPR, very little is known about the contribution of altered ER metabolism in cancer. Recently, Marini et al. (2016) showed that H6PDH participate in glucose oxidation in cancer cells, suggesting that ER might be an important site of glucose

220

A.S.H. Costa and C. Frezza

metabolism and that H6PDH might play roles that go beyond maintenance of redox homeostasis. It will be interesting to investigate the metabolic role of ER in cancer cells.

3.3 Peroxisomes Peroxisomes are single membrane-bound organelles present in all eukaryotic cells and are involved in many biological functions. Due to their high plasticity, the peroxisomes have the ability to respond to cellular or environmental perturbations by modifying their size, number, morphology, and function (Goodman, 2012). Our understanding of the metabolic reactions occurring in the peroxisomes and their communication networks is just beginning to emerge (Thoms et al., 2009; Tripathi and Walker, 2016), partially due to the fact that this organelle was not identified until the second half of the 20th century (De Duve and Baudhuin, 1966). Peroxisomes fulfill important metabolic functions in mammalian cells, including branched and very-long-chain fatty acid beta-oxidation; etherphospholipid synthesis; bile acid synthesis; fatty acid alpha-oxidation, glyoxylate metabolism; amino acid catabolism; polyamine oxidation; and the oxidative arm of the PPP (Fransen et al., 2012). The observation that several metabolic pathways occur within the peroxisome suggests that substrates, products, and cofactors of peroxisomal enzymes are transported across peroxisomal membranes. Indeed, it has been shown that the peroxisomes membranes express pore-forming proteins that enable the free exchange of metabolites (reviewed in Antonenkov and Hiltunen, 2012). Interestingly, the peroxisomal membrane can discriminate between the sizes of the transported molecules, preventing diffusion of molecules larger than 300–400 Da, such as NADH and ATP (Antonenkov and Hiltunen, 2012). Given that some biochemical reactions in the peroxisomes require these metabolites, the presence of shuttling mechanisms analogous to the malate-aspartate shuttle for the mitochondria has been postulated (Visser et al., 2007). For instance, the shuttling of NADPH, a cofactor required for peroxisomal redox reactions, is guaranteed by the presence of glucose-6-phosphate dehydrogenase and shuttling of hexoses and the presence of NADP-linked isocitrate dehydrogenases (Fig. 1B). Among the most investigated function of peroxisomes is FAO. Peroxisomal FAO shares some similarities with mitochondrial FAO, albeit with subtle, but biologically relevant, differences. While the first step of mitochondrial FAO is catalyzed by acyl-CoA dehydrogenase, peroxisomal FAO involves FAD-containing acyl-CoA oxidase (ACOX), which donates

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

221

electrons directly to molecular oxygen, yielding hydrogen peroxide that is decomposed by catalase, another peroxisomal matrix enzyme (Camo˜es et al., 2015). Furthermore, while both peroxisomes and mitochondria can oxidize medium- and long-chain fatty acids, very-long-chain (more than 20 carbons) fatty acids can be oxidized solely in the peroxisomes, thanks to the presence of very-long-chain acyl-CoA synthetase in these organelles (Watkins and Ellis, 2012). However, the current view is that the main function of peroxisomes is not to fully oxidize these fatty acids, but to shorten them in preparation for further oxidation in the mitochondria. Peroxisomes are also unique sites for the synthesis of etherphospholipids, molecules involved in the formation of plasmalogens (Schrader and Yoon, 2007), a group of phospholipids present in various tissues, including heart, liver, kidneys, and brain; and of alpha-oxidation, i.e., oxidation of 3-methyl fatty acids, such as phytanic acid. Finally, peroxisomes play a key role in ROS homeostasis: besides being a main source of hydrogen peroxide, they also contain a set of antioxidant proteins, including catalases—whose deficiency has been linked to increased risk of cancer—and antioxidant metabolites, including plasmalogens (Fransen et al., 2012; Wallner, 2011). Defects in one or more functions of the peroxisome, or in peroxisomes biogenesis, have dramatic effects in metabolic functions such as biosynthesis of cholesterol and plasmalogens, degradation of unsaturated and aromatic fatty acids, and beta-oxidation of fatty acids (Ferdinandusse et al., 2016; Frederiks et al., 2010), with important implications for human health. The importance of peroxisomes in pathophysiology is underscored by a broad spectrum of peroxisome biogenesis disorders (Braverman et al., 2013; Steinberg et al., 2006; Van Veldhoven, 2010). Defects in peroxisomal biogenesis seem also implicated in cancer. For instance, absence of peroxisomes has been observed in hepatoma (Frederiks et al., 2010; Litwin et al., 1999), colon carcinoma (Baur and Wendel, 1980; Lauer et al., 1999), breast cancer (Keller et al., 1993), and renal cell carcinoma (Frederiks et al., 2010). Furthermore, oncogenic signaling cascades, such as those regulated by the hypoxia-inducible factors, can affect peroxisome stability leading to peroxisome depletion in clear cell renal cell carcinoma (Walter et al., 2014). To what extent, peroxisomal dysfunction contributes to the metabolic reprogramming of cancer is still unclear. Intriguingly, mice deficient for the peroxisomal enzyme ACOX develop liver cancer (Meyer et al., 2003) and it has been proposed that the accumulation of toxic metabolites due to impairment of peroxisomal FAO contribute to liver tumorigenesis in this model. Furthermore, high expression of the enzyme alpha-methylacyl-

222

A.S.H. Costa and C. Frezza

CoA racemase (AMACR), a peroxisomal enzyme involved in the oxidation of 2-methyl-branched chain fatty acid, was observed in various cancer types, including prostate (Box et al., 2016). Finally, the ether lipid enzyme alkylgyceronephosphate synthase (AGPS) was found overexpressed in cancers and its silencing impairs cancer cell growth, migration, and invasion (Benjamin et al., 2013). However, how ether lipids affect tumorigenesis is still under investigation, even though recent evidence suggests they might act as self-ligands to activate the immune system (Facciotti et al., 2012). It has also been proposed that peroxisomal function is linked to aerobic glycolysis in cancer cells. In prostate cells, the lactate transporter MCT2 translocates to the peroxisomal membrane where it facilitates pyruvate import to sustain NAD+ regeneration through peroxisomal lactate dehydrogenase and lactate shuttling back in the cytosol. This unexpected activity of glucose-derived metabolites contributes to peroxisomal redox balance and enables the oxidation of very-long-chain fatty acids (McClelland et al., 2003; Valenc¸a et al., 2015).

3.4 Lysosomes Lysosomes are acidic catabolic organelles present in all mammalian cells, with mature erythrocytes being the only exception (de Duve, 2005; Kallunki et al., 2013). This dynamic organelle is situated at the crossroad of the most important cellular pathways and takes part in sensing, signaling, and transcriptional mechanisms in response to shifts in nutrient availability (Settembre and Ballabio, 2014). Although no metabolic pathways are known to occur in the lysosomes, these organelles take part in autophagy and micropinocytosis, two major nutrient scavenging pathways which maintain intracellular amino acid pools (Perera et al., 2015). Besides providing amino acids from these processes, lysosomes participate to macropinocytosis, an endocytic process by which extracellular fluid and its contents are internalized into cells through vesicles called macropinosomes (Commisso et al., 2013). Fusion of the macropinosomes with lysosome then releases frees amino acids through proteolytic degradation of the engulfed proteins (Pavlova et al., 2016; Fig. 1B). Malignant transformation of cells impacts on lysosomes and their function. For instance, upregulation of lysosomal hydrolases is common in human tumors, and it often correlates with increased risk of recurrence and poorer prognosis (Kallunki et al., 2013). Furthermore, defects in autophagy are also a major feature of cancer (Perera et al., 2015). Lysosomes

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

223

also play roles in other opportunistic processes of nutrient acquisition—an hallmark of cancer metabolic transformation—such as entosis and phagocytosis (Kroemer and Perfettini, 2014), or macroautophagy (Yang and Klionsky, 2010). Finally, the expression of lysosomal genes and lysosomal biogenesis transcription has been shown to be regulated by factor EB (TFEB) (Sardiello et al., 2009), which together with TFE3, has been implicated in the chromosomal translocations observed renal cell carcinomas (Ellati et al., 2016; Kuroda et al., 2012).

4. METABOLIC COOPERATION AMONG ORGANELLES From this overview, it emerges that besides mitochondria, which thus far have taken the spotlight in cancer metabolism research, other intracellular organelles are involved in the metabolic reprogramming of cancer. The physical proximity of these organelles suggests the presence of a simultaneous bidirectional transfer of molecules between them (Shai et al., 2016). Nevertheless, our understanding of how these organelles communicate to maintain a synchronized metabolic cascade is still poor. Later, we present the current understanding of this metabolic cooperation. Interactions between mitochondria and ER have been widely investigated in the past, mostly due to the well-established exchange of calcium among them (Marchi et al., 2014). The ER and mitochondria join together at multiple contact sites to form specific domains, the mitochondria–ERassociated membranes (Marchi et al., 2014). The function of these contact sites is under intense investigation. We now know that they are involved in trafficking of lipid species and in autophagosome formation (Hamasaki et al., 2013). The communication between ER and mitochondria has important physiological roles. When ER function is disrupted, the subsequent dysregulation of calcium homeostasis leads to mitochondrial dysfunction (Giorgi et al., 2015; Patergnani et al., 2015). Conversely, impaired mitochondrial function leads to ER stress. To what extent, mitochondrial and ER metabolism are interconnected is still unknown, but it is reasonable to anticipate the existence of a regulated exchange of metabolites and reducing equivalents between these organelles. Another important interaction between organelles is that between mitochondria and peroxisomes. The relevance of the peroxisome–mitochondria axis is corroborated by the observations that both organelles share key proteins for their biogenesis and division machinery (Schrader et al., 2013). Given the lack of TCA cycle and respiratory chain, continued metabolism

224

A.S.H. Costa and C. Frezza

of the peroxisome-generated products requires cross talk with mitochondria (Wanders, 2013). Indeed, as described earlier, peroxisomal lipid oxidation is incomplete and shortened acyl-CoA are sent to mitochondria for full oxidation. At the same time, peroxisomes can initiate the catabolism of verylong-fatty acids that could not be oxidized in the mitochondria, therefore preventing the toxic effects which would arise from their accumulation (Schrader and Yoon, 2007). Different classes of small molecules, including fatty acids, bile acids, and ROS that accumulate in dysfunctional peroxisomes can affect mitochondrial function (Peeters et al., 2015; Salpietro et al., 2015). Conversely, mitochondrial dysfunction could result in the disruption of peroxisomes function (Peeters et al., 2015). The communication between mitochondria and lysosome has been less investigated. Data on lysosomal disorders suggest that disruption of lysosomal function leads to mitochondrial fragmentation, mostly mediated by lipofuscin, a metabolite generated in the lumen of damaged lysosomes (reviewed in Brunk and Terman, 2002). This interorganelle relation is mutual and mitochondrial dysfunction could also lead to alterations of lysosomal function. For instance, it was showed that mitochondrial dysfunction caused by the genetic ablation of the transcription factor Tfam, leads to impaired lysosomal function and sphingomyelin accumulation, at least in T cells (Baixauli et al., 2015). It is worth noting that the above-described metabolic pathways are intertwined with other well-established functions of these organelles, suggesting that any disruption in the metabolic network could have repercussions on other signaling cascades that use these organelles as a platform. For instance, mitochondria are key players in the apoptotic cascade (Tait and Green, 2013). It is therefore reasonable to think that perturbations of metabolic fluxes, indirectly affecting mitochondrial function, could also affect sensitivity to apoptosis. Indeed, by virtue of this nutrient sensing mechanism, mitochondria ensure that only cells exposed to adequate access to nutrients can survive (reviewed in Nunnari and Suomalainen, 2012). Conversely, cell death proteins, such as the BH3-only member BAD, can influence the metabolic efficiency of mitochondria, corroborating the mutual link between mitochondrial metabolism and apoptotic machinery (Danial, 2008). Decoupling this mechanism would enable cells survival even when nutrients are scarce, a prerequisite for cancer growth. ER function is also intimately related to metabolism. For instance, changes in availability of glucose-6phosphate could affect H6PDH activity and therefore NAPDH/NADP ratio in the ER and, as a consequence, protein folding efficiency, potentially

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

225

leading to UPR. Intracellular accumulation of saturated fatty acids and cholesterol could also affect ER and lead to UPR (Wei and Wang, 2006). These results underscore the complex cross talk between metabolism and cell signaling and suggest that the implications of altered metabolism in cancer could be broader than anticipated.

5. CHALLENGES AND FUTURE DIRECTIONS Both biological and technological challenges are hindering our understanding of the complex communication among organelles and its role in cancer. From a cell biology point of view, the mechanisms of regulation of the interorganelles communication are far from understood. If on one hand this communication is orchestrated at the metabolic level, as described earlier, metabolic changes can also be integrated by more complex signaling cascades that result in adaptive cellular responses. One of the most investigated nutrient sensing machinery is that orchestrated by the mechanistic target of rapamycin (mTOR) complex, which involves a plethora of upstream and downstream kinases (Laplante et al., 2012). It is likely that these networks play a prominent role in the communication between the various organelles, in health and in disease. In support to this hypothesis, we now know that components of the mTOR machinery are distributed in various organelles, including lysosomes, mitochondria, and peroxisomes (Betz and Hall, 2013). Understanding how the signaling and metabolic network communicate each other in multiple compartments and how this communication is dysregulated in cancer will be a crucial step in the field. Indeed, it will enable us to understand the contribution of each organelle to cancer transformation and will help to target the effects of antimetabolic drugs to specific compartments, without disrupting unwanted parts of the metabolic network. To improve our knowledge of this complex communication among organelles, we need to overcome several technological hurdles. For instance, current technologies to probe cell metabolism, including mass spectrometry, require cell lysis, which disrupts cell architecture, hampering our possibility to distinguish between different pools of metabolites in the cell. Moreover, we currently lack the tools to selectively perturb specific subcellular compartments, thus allowing us to obtain the spatial and temporal resolution necessary to elucidate the flow of small molecules between the various organelles. An additional aspect to be considered is the propagation of the metabolic cascade upon perturbation of the

226

A.S.H. Costa and C. Frezza

metabolic network. Steady-state measurements seldom inform on the changes in flux through particular pathways. Furthermore, current methods still lack the ability to detect small shifts in the directionality or magnitude of flux which, taken individually, might not be biologically relevant but may interact synergistically in a way that produces profound changes in the network. Finally, the community needs to develop tools to integrate different layers of complexity in the cell. In the past decade, we have expanded remarkably our capacity to probe the cell at various levels—gene, protein, and small molecule—and to interrogate those multiomics datasets. However, each layer of information is often analyzed in parallel, rather than truly integrated. The capacity to integrate these layers would help us to understand how metabolism affect signaling and, vice versa, how signaling affects metabolism.

6. CONCLUSIONS Reprogramming of cell metabolism is a recent hallmark of cancer. In the past decades, the major features of this metabolic reprogramming have been elucidated. However, we now need to consider that metabolism is not a simple bidimensional map, as generally depicted (Fig. 1A), but rather a three-dimensional set of reactions separated in multiple compartments within the cell (Fig. 1B). While compartmentalization of metabolic pathways allows regulatory mechanisms to control them, it also implies that any disturbance in one organelle can reverberate across the entire cell. Each organelle plays multiple roles, depending on the organ, developmental and physiological status, and it has become clear that in order to perform their functions they are intertwined in a vast and complex metabolic network that relies on their cooperation. While our knowledge on the biochemistry of each organelle is vast, the mechanisms and dynamics of organelle interplay in health and in disease remain elusive. We now face the challenge of having to define the boundaries of the metabolic network and to understand the communication and cooperation among the various subcellular compartments. In discovering how organelle cross talk is wired and functions to support pathological events, we will also deepen our understanding of the metabolic reprogramming of cancer, thus enabling novel and more effective antineoplastic strategies to be devised.

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

227

ACKNOWLEDGMENTS The authors would like to thank all the members of the Frezza lab for fruitful discussion and insightful comments. Competing interests: The authors declare no competing interests. Authors’ information: A.S.H.C. is a Research Associate in the laboratory of C.F. C.F. is a group leader at the MRC Cancer Unit, University of Cambridge, Cambridge, UK. A.S.H.C. and C.F. are funded by MRC Core Funding to the MRC Cancer Unit. Authors’ contribution: A.S.H.C. and C.F. jointly wrote the manuscript.

REFERENCES Ahmad, I., et al., 2016. Sleeping Beauty screen reveals Pparg activation in metastatic prostate cancer. Proc. Natl. Acad. Sci. 113 (29), 8290–8295. Antonenkov, V.D., Hiltunen, J.K., 2012. Transfer of metabolites across the peroxisomal membrane. Biochim. Biophys. Acta (BBA)—Mol. Basis Dis. 1822 (9), 1374–1386. Baixauli, F., et al., 2015. Mitochondrial respiration controls lysosomal function during inflammatory T cell responses. Cell Metab. 22 (3), 485–498. Basu, S., et al., 2013. Heavy and trace metals in carcinoma of the gallbladder. World J. Surg. 37 (11), 2641–2646. Baur, G., Wendel, A., 1980. The activity of the peroxide-metabolizing system in human colon carcinoma. J. Cancer Res. Clin. Oncol. 97 (3), 267–273. Beloribi-Djefaflia, S., Vasseur, S., Guillaumond, F., 2016. Lipid metabolic reprogramming in cancer cells. Oncogenesis 5 (1). e189. Benjamin, D.I., et al., 2013. Ether lipid generating enzyme AGPS alters the balance of structural and signaling lipids to fuel cancer pathogenicity. Proc. Natl. Acad. Sci. U. S. A. 110 (37), 14912–14917. Betz, C., Hall, M.N., 2013. Where is mTOR and what is it doing there? J. Cell Biol. 203 (4), 563–574. Birsoy, K., et al., 2015. An essential role of the mitochondrial electron transport chain in cell proliferation is to enable aspartate synthesis. Cell 162 (3), 540–551. Bohnert, K.R., et al., 2016. Inhibition of ER stress and unfolding protein response pathways causes skeletal muscle wasting during cancer cachexia. FASEB J. 30 (9), 3053–3068. Box, A., et al., 2016. High alpha-methylacyl-CoA racemase (AMACR) is associated with ERG expression and with adverse clinical outcome in patients with localized prostate cancer. Tumor Biol. 37 (9), 12287–12299. Braverman, N.E., D’Agostino, M.D., MacLean, G.E., 2013. Peroxisome biogenesis disorders: biological, clinical and pathophysiological perspectives. Dev. Disabil. Res. Rev. 17 (3), 187–196. Brunk, U.T., Terman, A., 2002. The mitochondrial-lysosomal axis theory of aging. Eur. J. Biochem. 269 (8), 1996–2002. Camo˜es, F., et al., 2015. New insights into the peroxisomal protein inventory: acyl-CoA oxidases and -dehydrogenases are an ancient feature of peroxisomes. Biochim. Biophys. Acta 1853 (1), 111–125. Carracedo, A., Cantley, L.C., Pandolfi, P.P., 2013. Cancer metabolism: fatty acid oxidation in the limelight. Nat. Rev. Cancer 13 (4), 227–232. Cohen, R., et al., 2015. Targeting cancer cell metabolism in pancreatic adenocarcinoma. Oncotarget 6 (19), 16832–16847. Commisso, C., et al., 2013. Macropinocytosis of protein is an amino acid supply route in Ras-transformed cells. Nature 497 (7451), 633–637.

228

A.S.H. Costa and C. Frezza

Danial, N.N., 2008. BAD: undertaker by night, candyman by day. Oncogene 27, S53–S70. de Duve, C., 2005. The lysosome turns fifty. Nat. Cell Biol. 7 (9), 847–849. De Duve, C., Baudhuin, P., 1966. Peroxisomes (microbodies and related particles). Physiol. Rev. 46 (2), 323–357. DeBerardinis, R.J., Chandel, N.S., 2016. Fundamentals of cancer metabolism. Sci. Adv. 2 (5). e1600200. Deep, G., Schlaepfer, I., 2016. Aberrant lipid metabolism promotes prostate cancer: role in cell survival under hypoxia and extracellular vesicles biogenesis. Int. J. Mol. Sci. 17 (7), 1061. Eagle, H., 1955. The minimum vitamin requirements of the L and HeLa cells in tissue culture, the production of specific vitamin deficiencies, and their cure. J. Exp. Med. 102 (5), 595–600. Eales, K.L., Hollinshead, K.E.R., Tennant, D.A., 2016. Hypoxia and metabolic adaptation of cancer cells. Oncogenesis 5 (1). e190. Ellati, R.T., et al., 2016. Clinicopathologic features of translocation renal cell carcinoma. Clin. Genitourin. Cancer 15, 112–116. Facciotti, F., et al., 2012. Peroxisome-derived lipids are self antigens that stimulate invariant natural killer T cells in the thymus. Nat. Immunol. 13 (5), 474–480. Feldman, D., et al., 2014. The role of vitamin D in reducing cancer risk and progression. Nat. Rev. Cancer 14 (5), 342–357. Ferdinandusse, S., et al., 2016. The important role of biochemical and functional studies in the diagnostics of peroxisomal disorders. J. Inherit. Metab. Dis. 39 (4), 531–543. Fransen, M., et al., 2012. Role of peroxisomes in ROS/RNS-metabolism: implications for human disease. Biochim. Biophys. Acta—Mol. Basis Dis. 1822 (9), 1363–1373. Frederiks, W.M., et al., 2010. Renal cell carcinoma and oxidative stress: the lack of peroxisomes. Acta Histochem. 112 (4), 364–371. Gaude, E., Frezza, C., 2014. Defects in mitochondrial metabolism and cancer. Cancer Metab. 2, 10. Giorgi, C., et al., 2015. Intravital imaging reveals p53-dependent cancer cell death induced by phototherapy via calcium signaling. Oncotarget 6 (3), 1435–1445. Glunde, K., Bhujwalla, Z.M., Ronen, S.M., 2011. Choline metabolism in malignant transformation. Nat. Rev. Cancer 11 (12), 835. Goodman, C., 2012. Cell biology: peroxisomes come together. Nat. Chem. Biol. 8 (6), 503. Gupte, A., et al., 2009. Elevated copper and oxidative stress in cancer cells as a target for cancer treatment. Cancer Treat. Rev. 35 (1), 32–46. Hamasaki, M., et al., 2013. Autophagosomes form at ER–mitochondria contact sites. Nature 495 (7441), 389–393. Hanahan, D., et al., 2011. Hallmarks of cancer: the next generation. Cell 144 (5), 646–674. Huang, S., 2002. Histone methyltransferases, diet nutrients and tumour suppressors. Nat. Rev. Cancer 2 (6), 469–476. Kallunki, T., Olsen, O.D., J€a€attel€a, M., 2013. Cancer-associated lysosomal changes: friends or foes? Oncogene 32 (16), 1995–2004. Kamphorst, J.J., et al., 2013. Hypoxic and Ras-transformed cells support growth by scavenging unsaturated fatty acids from lysophospholipids. Proc. Natl. Acad. Sci. 110 (22), 8882–8887. Keller, J.-M., et al., 1993. Peroxisome through cell differentiation and neoplasia. Biol. Cell 77 (1), 77–88. Kroemer, G., Perfettini, J.-L., 2014. Entosis, a key player in cancer cell competition. Cell Res. 24 (11), 1280–1281. Kuroda, N., et al., 2012. Review of renal carcinoma associated with Xp11.2 translocations/ TFE3 gene fusions with focus on pathobiological aspect. Histol. Histopathol. 27 (2), 133–140.

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

229

Lagace, T.A., Ridgway, N.D., 2013. The role of phospholipids in the biological activity and structure of the endoplasmic reticulum. Biochim. Biophys. Acta 1833 (11), 2499–2510. Laplante, M., et al., 2012. mTOR signaling in growth control and disease. Cell 149 (2), 274–293. Lauer, C., et al., 1999. Impairment of peroxisomal biogenesis in human colon carcinoma. Carcinogenesis 20 (6), 985–989. Litwin, J.A., et al., 1999. Immunocytochemical investigation of catalase and peroxisomal lipid β-oxidation enzymes in human hepatocellular tumors and liver cirrhosis. Virchows Arch. 435 (5), 486–495. Lodhi, I.J.J., et al., 2014. Peroxisomes: a nexus for lipid metabolism and cellular signaling. Cell Metab. 19 (3), 380–392. Luo, J., et al., 2010. S-adenosylmethionine inhibits the growth of cancer cells by reversing the hypomethylation status of c-myc and H-ras in human gastric cancer and colon cancer. Int. J. Biol. Sci. 6 (7), 784–795. Mandl, J., et al., 2009. Endoplasmic reticulum: nutrient sensor in physiology and pathology. Trends Endocrinol. Metab. 20 (4), 194–201. Marchi, S., Patergnani, S., Pinton, P., 2014. The endoplasmic reticulum–mitochondria connection: one touch, multiple functions. Biochim. Biophys. Acta 1837 (4), 461–469. Marini, C., et al., 2016. Discovery of a novel glucose metabolism in cancer: the role of endoplasmic reticulum beyond glycolysis and pentose phosphate shunt. Sci. Rep. 6, 25092. McClelland, G.B., et al., 2003. Peroxisomal membrane monocarboxylate transporters: evidence for a redox shuttle system? Biochem. Biophys. Res. Commun. 304 (1), 130–135. Metallo, C.M., et al., 2012. Reductive glutamine metabolism by IDH1 mediates lipogenesis under hypoxia. Nature 481 (7381), 380–384. Meyer, K., et al., 2003. Molecular profiling of hepatocellular carcinomas developing spontaneously in acyl-CoA oxidase deficient mice: comparison with liver tumors induced in wild-type mice by a peroxisome proliferator and a genotoxic carcinogen. Carcinogenesis 24 (5), 975–984. Mo´dis, K., et al., 2014. Effect of S-adenosyl-l-methionine (SAM), an allosteric activator of cystathionine-β-synthase (CBS) on colorectal cancer cell proliferation and bioenergetics in vitro. Nitric Oxide 41, 146–156. Nath, A., et al., 2016. Genetic alterations in fatty acid transport and metabolism genes are associated with metastatic progression and poor prognosis of human cancers. Sci. Rep. 6, 18669. Nieman, K.M., et al., 2011. Adipocytes promote ovarian cancer metastasis and provide energy for rapid tumor growth. Nat. Med. 17 (11), 1498–1503. Nunnari, J., Suomalainen, A., 2012. Mitochondria: in sickness and in health. Cell 148 (6), 1145–1159. Oka, O.B.V., Bulleid, N.J., 2013. Forming disulfides in the endoplasmic reticulum. Biochim. Biophys. Acta 1833 (11), 2425–2429. Ozcan, L., Tabas, I., 2012. Role of endoplasmic reticulum stress in metabolic disease and other disorders. Annu. Rev. Med. 63, 317–328. Patergnani, S., et al., 2015. Mitochondria-associated endoplasmic reticulum membranes microenvironment: targeting autophagic and apoptotic pathways in cancer therapy. Front. Oncol. 5, 173. Pavlova, N.N., et al., 2016. The emerging hallmarks of cancer metabolism. Cell Metab. 23 (1), 27–47. Peeters, A., et al., 2015. Mitochondria in peroxisome-deficient hepatocytes exhibit impaired respiration, depleted DNA, and PGC-1α independent proliferation. Biochim. Biophys. Acta 1853 (2), 285–298. Perera, R.M., et al., 2015. Transcriptional control of autophagy–lysosome function drives pancreatic cancer metabolism. Nature 524 (7565), 361–365.

230

A.S.H. Costa and C. Frezza

Ryan, D., et al., 2016. Calnexin, an ER-induced protein, is a prognostic marker and potential therapeutic target in colorectal cancer. J. Transl. Med. 14 (1), 196. Salpietro, V., et al., 2015. Zellweger syndrome and secondary mitochondrial myopathy. Eur. J. Pediatr. 174 (4), 557–563. Sardiello, M., et al., 2009. A gene network regulating lysosomal biogenesis and function. Science (New York, N.Y.) 325 (5939), 473–477. Schrader, M., Yoon, Y., 2007. Mitochondria and peroxisomes: are the “big brother” and the “little sister” closer than assumed? Bioessays 29 (11), 1105–1114. Schrader, M., et al., 2013. Peroxisome interactions and cross-talk with other subcellular compartments in animal cells. In: del Rı´o, L.A. (Ed.), Peroxisomes and Their Key Role in Cellular Signaling and Metabolism. Springer, The Netherlands, pp. 1–22. Schug, Z.T., et al., 2015. Acetyl-CoA synthetase 2 promotes acetate utilization and maintains cancer cell growth under metabolic stress. Cancer Cell 27 (1), 57–71. Settembre, C., Ballabio, A., 2014. Lysosomal adaptation: how the lysosome responds to external cues. Cold Spring Harb. Perspect. Biol. 6 (6), a016907. Shai, N., Schuldiner, M., Zalckvar, E., 2016. No peroxisome is an island—peroxisome contact sites. Biochim. Biophys. Acta 1863 (5), 1061–1069. Singh, R., Kaushik, S., Wang, Y., Xiang, Y., Novak, I., Komatsu, M., Tanaka, K., Cuervo, A.M., Czaja, MJ., 2009. Autophagy regulates lipid metabolism. Nature 458, 1131–1135. Sousa, C.M., et al., 2016. Pancreatic stellate cells support tumour metabolism through autophagic alanine secretion. Nat. Publ. Group 536 (7617), 479–483. Steinberg, S.J., et al., 2006. Peroxisome biogenesis disorders. Biochim. Biophys. Acta 1763 (12), 1733–1748. Sullivan, L.B., et al., 2015. Supporting aspartate biosynthesis is an essential function of respiration in proliferating cells. Cell 162 (3), 552–563. Szeto, H.H., Schiller, P.W., 2011. Novel therapies targeting inner mitochondrial membrane—from discovery to clinical development. Pharm. Res. 28 (11), 2669–2679. Tait, S.W.G., Green, D.R., 2013. Mitochondrial regulation of cell death. Cold Spring Harb. Perspect. Biol. 5 (9), a008706. Thoms, S., Grønborg, S., G€artner, J., 2009. Organelle interplay in peroxisomal disorders. Trends Mol. Med. 15 (7), 293–302. Tripathi, D.N., Walker, C.L., 2016. The peroxisome as a cell signaling organelle. Curr. Opin. Cell Biol. 39, 109–112. Valenc¸a, I., et al., 2015. Localization of MCT2 at peroxisomes is associated with malignant transformation in prostate cancer. J. Cell. Mol. Med. 19 (4), 723–733. Van Veldhoven, P.P., 2010. Biochemistry and genetics of inherited disorders of peroxisomal fatty acid metabolism. J. Lipid Res. 51 (10), 2863–2895. Visser, W.F., et al., 2007. Metabolite transport across the peroxisomal membrane. Biochem. J. 401 (2), 365–375. Vyas, S., et al., 2016. Mitochondria and Cancer. Cell 166 (3), 555–566. Wallner, S., 2011. Plasmalogens the neglected regulatory and scavenging lipid species. Chem. Phys. Lipids 164 (6), 573–589. Walter, K.M., et al., 2014. Hif-2α promotes degradation of mammalian peroxisomes by selective autophagy. Cell Metab. 20 (5), 882–897. Wanders, R.J.A., 2013. Peroxisomes in human health and disease: metabolic pathways, metabolite transport, interplay with other organelles and signal transduction. In: del Rı´o, L.A. (Ed.), Peroxisomes and Their Key Role in Cellular Signaling and Metabolism. In: Subcellular Biochemistry, vol. 69. Springer, The Netherlands, pp. 23–44. Warburg, O., Dickens, F., 1927. The metabolism of tumors in the body. Am. J. Med. Sci. 182 (1), 123.

Organelle Cross Talk in Oncogenic Metabolic Reprogramming

231

Watkins, P.A., Ellis, J.M., 2012. Peroxisomal acyl-CoA synthetases. Biochim. Biophys. Acta—Mol. Basis Dis. 1822 (9), 1411–1420. Wei, Y., Wang, D., 2006. Saturated fatty acids induce endoplasmic reticulum stress and apoptosis independently of ceramide in liver cells. Am. J. Physiol. Endocrinol. Metab. 1571, 275–281. Weinberg, S.E., Chandel, N.S., 2014. Targeting mitochondria metabolism for cancer therapy. Nat. Chem. Biol. 11 (1), 9–15. Wu, W., Kang, S., Zhang, D., 2013. Association of vitamin B6, vitamin B12 and methionine with risk of breast cancer: a dose–response meta-analysis. Br. J. Cancer 109 (7), 1926–1944. Yang, Z., Klionsky, D.J., 2010. Eaten alive: a history of macroautophagy. Nat. Cell Biol. 12 (9), 814–822. Zeng, F., et al., 2014. Choline and betaine intakes are associated with reduced risk of nasopharyngeal carcinoma in adults: a case–control study. Br. J. Cancer 110 (3), 808–816.

CHAPTER FIVE

Molecular Biology Digest of Cell Mitophagy I. Matic*, D. Strobbe†, F. Di Guglielmo*, M. Campanella*,†,{,§,1 *University of Rome Tor Vergata, Rome, Italy † Regina Elena-National Cancer Institute, Rome, Italy { RVC, University of London, London, United Kingdom § UCL Consortium for Mitochondrial Research, London, United Kingdom 1 Corresponding author: e-mail address: [email protected]

Contents 1. Mitochondria Cannot Be Mended but Can Be Checked 2. Keeping the Engine Clean: Mitophagy 3. The Parkin-Dependent Way of Mitophagy 4. The Parkin-Independent Way of Mitophagy 5. Concluding Remarks Acknowledgments References

233 236 239 247 250 251 251

Abstract The homeostasis of eukaryotic cells relies on efficient mitochondrial function. The control of mitochondrial quality is framed by the combination of distinct but interdependent mechanisms spanning biogenesis, regulation of dynamic network, and finely tuned degradation either through ubiquitin–proteasome system or autophagy (mitophagy). There is continuous evolution on the pathways orchestrating the mitochondrial response to stress signals and the organelle adaptation to quality control during acute and subtle dysfunctions. Notably, it remains indeed ill-defined whether active mitophagy leads to cell survival or death by defective mitochondrial degradation. Above all, uncharted is whether and how pharmacologically tackle these mechanisms may lead to conceive novel therapeutic strategies for treating conditions associated with the defective mitochondria. Here, we attempt to provide a chronological and comprehensive overview of the determining discoveries, which have led to the current knowledge of mitophagy.

1. MITOCHONDRIA CANNOT BE MENDED BUT CAN BE CHECKED Since the pioneering bioenergetics studies in the early 1950s (Palade, 1953; Sjostrand, 1953), mitochondria have been commonly described as International Review of Cell and Molecular Biology, Volume 332 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2016.12.003

#

2017 Elsevier Inc. All rights reserved.

233

234

I. Matic et al.

the generators of energy-rich ATP molecules so-called the cellular powerhouses. But these remarkable organelles do far more than simply transforming the available nutrients into energy. Found in almost all eukaryotic cells, they serve a wide spectrum of important functions adjusted to cellular needs: regulation of the levels of intracellular signaling molecules (such as Ca2+), growth factor signaling, fatty acid synthesis, biogenesis of haem and iron–sulpfur proteins, cell senescence, control of cell cycle, and differentiation. Equally, mitochondria represent potential danger for the cell, being the major source of reactive oxygen species (ROS), which may oxidize and damage proteins, lipids, and DNA (Holmstr€ om and Finkel, 2014; Filomeni et al., 2015) creating serious jeopardy for the cell life. Mitochondrial dysfunction is therefore a critical determinant for inherited metabolic disorders (Calvo and Mootha, 2010; Koopman et al., 2012), as well as common disorders such as type 2 diabetes (Kacerovsky-Bielesz et al., 2012) and cardiac dysfunction (Andres et al., 2015), neurodegenerative diseases (Burte et al., 2015), and cancer (Wallace, 2012). Mitochondria play an important role in the aging (Balaban et al., 2005; Biala et al., 2015; Bratic and Larsson, 2013), hence contributing to specific aspects of cellular senescence, chronic inflammation, and the age-dependent decline in stem cell activity (Sun et al., 2016). Pertinent removal of dysfunctional mitochondria occurring in response to stresses, such as hypoxia and nutrient starvation (Ashrafi and Schwarz, 2013), prevents generation of ROS and conserves valuable nutrients (such as oxygen) from being consumed inefficiently, thereby promoting cellular survival under conditions of energetic stress. Thus, it seems that when a cell hangs in delicate balance between life and death, the path to be taken is chosen by the mitochondrion acting as a sort of decisive level of intervention. Nonetheless, in the event in which accumulated toxins upstage prolife signals, which keep the mitochondrial membrane intact and encourage the organelle to keep producing energy, the mitochondrion pushes the cell down a carefully orchestrated pathway to death (Kroemer et al., 1995). For long time, since 1950s, mitochondrial biologists were largely dedicated in working out the details of mitochondrial metabolism. In the early 1990s the breaking discovery the mitochondrion acts at the center of the programmed cell death paved the way for seminal advancements in understanding mitochondrial quality control mechanisms of dysfunctional

Molecular Biology Digest of Cell Mitophagy

235

mitochondria removal and the role of this highly complex process in a myriad of human diseases. The ground-breaking work from the laboratories of Kroemer et al. (1995) led to the formulation of the novel hypothesis by which mitochondria could be “decision makers” for the fate of cells. All started in 1994, when Martin Raff (Jacobson et al., 1994) and Peter Krammer (Schulze-Osthoff et al., 1994) showed that, in addition to the nucleus, a cytoplasmic regulator for apoptosis of human cells must exist. The focus promptly shifted toward mitochondria, since the loss of the mitochondrial membrane potential (△ψ m) marked an irreversible early step of the apoptotic process in different cell types, including epithelial cell lines and neurons shortly before the onset of apoptosis (Kroemer et al., 2007). Kroemer et al. (1995) found that, upon the induction of permeability transition, mitochondria released what was at first called an apoptosis-inducing factor (AIF) (that is now retrospectively known to be a mix of several proteins) (Zamzami et al., 1996). The mitochondrial release of these apoptogenic factors was later found to be controlled by members of proapoptotic Bcl-2 protein family (Susin et al., 1996), and although AIF effects were claimed to be caspase independent (Susin et al., 1996), it was later demonstrated that they mostly occurred before caspases were activated (Vaux, 2011). Along with the flurry of papers which followed, many different proteins capable of regulating the permeability of mitochondrial membranes were identified. The members of the Bcl-2 family were the first to be recognized to regulate mitochondrial permeability (Kroemer, 1997), and thus, it is the mitochondrial membrane permeabilization on the whole to be regarded as “the critical event in the death/life decision beyond the molecular details” (Kroemer, 2004). It is important to underline that stress does not necessarily lead to cell death and in reality, things are overwhelmingly complex: competing and conflicting signals from the cell surface, cytoplasm, and the mitochondria itself all congregate on a myriad of proteins to regulate either cell death pathway (self-killing) or a stress adaptation that avoids cell death (self-eating) (Maiuri et al., 2007). When less than 10 years ago, we learnt that the depolarization of the mitochondrial membrane may not necessarily lead to the programmed cell death but instead cause the activation of cell mitophagy (Ashrafi and Schwarz, 2013), the concept based on which cells could selectively remove depolarized mitochondria without necessarily undergoing death was

236

I. Matic et al.

simply paradigm shifting. Mitophagy as a kind of cellular “self-cleaning” strategy gained much attention among mitochondrial biologists, such that in the recent years intensified research provided many new insights into the function, mechanism, and regulation of this process (Campanella and Klionsky, 2013; East and Campanella, 2016). Several key proteins have been identified in the mechanistic dissection, paving the way for understanding the essential role the mitophagy process plays in the life and death of eukaryotic cells.

2. KEEPING THE ENGINE CLEAN: MITOPHAGY Autophagy has long been considered to be a nonselective bulk degradation pathway that indiscriminately eliminates cellular components including mitochondria. However, autophagy can become highly selective when a specific organelle (e.g., mitochondria) is selected and targeted to autophagosomes, which then deliver their load to the lysosomes for degradation (Zhang, 2013). Mitophagy was first defined by Lemasters (2005), although as early as in 1962 it had been seen that lysosomes in the liver that contained mitochondrial fragments (Ashford and Porter, 1962). Back in the 1977, two independent research studies led scientists to postulate that autophagy could be selective toward mitochondria. The first evidence came from studies on metamorphosis in silkworms, where it was theorized that mitochondria develop functional alterations, which would activate autophagy (Beaulaton and Lockshin, 1977); alongside, Reme et al. discovered mitochondria inside autophagosomes in the photoreceptor cells of the ground squirrel during hibernation (Reme and Young, 1977). Interestingly, the authors found that autophagy was selective only toward smaller mitochondria produced by budding or fragmentation, and all mitochondria found inside autophagosomes were of spherical shape (Reme and Young, 1977). Activation of general autophagy is generally accompanied by increased mitophagy, but under certain conditions, this may not always be the case. Mitophagy could be, nonetheless, induced under nutrient-rich conditions to remove redundant or dysfunctional mitochondria when general bulk autophagy is not even activated (Kobayashi and Liang, 2015; Youle and Narendra, 2011). On the contrary, when nutrient sources are poor and cells start to starve, autophagy (e.g., in muscle cells) is induced but is not necessarily followed by mitochondrial degradation (Raben et al.,

Molecular Biology Digest of Cell Mitophagy

237

2012; Wu et al., 2009). In alternative, mitophagy can even be inhibited during starvation, probably to provide cell with as much energy as possible. As proposed by the Andersen’s group (Kristensen et al., 2008) who discovered that starvation-induced autophagy degradation is performed in an ordered fashion, and not completely arbitrarily as was previously assumed. Their proteome-wide analysis showed that cytosolic and proteasomal proteins were degraded rapidly followed by ribosomal ones. On the other hand, mitochondrial, endoplasmatic reticulum, spliceosome, and vacuolar proteins were increased in size. A confirmation arrived from Scorrano’s group (Gomes et al., 2011), which observed mitochondrial elongation during starvation-induced autophagy in various cell models. These data reveal that general autophagy and selective mitophagy do not necessarily accompany one another; probably due to the different signaling pathways they follow (Cotan et al., 2011; Rodrı´guez-Herna´ndez et al., 2009). There is much yet to be determined about how mitophagy achieves its selectivity and specificity; however, in the light of recent discoveries, we will try to summarize the mechanisms elucidated so far. To identify factors that might account for selective mitophagy, screens for yeast mutants defective in mitophagy were obtained (Kanki et al., 2009; Okamoto et al., 2009). Approximately 40 genes have been found to be required for mitophagy but not bulk autophagy (Ashrafi and Schwarz, 2013). Camougrand’s group showed for the first time that Uth1 protein is required for mitochondrial removal induced by rapamycin treatment or nitrogen starvation (Kissova´ et al., 2004)—common inducers of bulk autophagy. ATG11 was also identified as an important for mitochondrial degradation (Kanki and Klionsky, 2008; Kanki et al., 2010). This protein functions as a basic scaffold in assembling the specific phagophore assembly site by interacting directly with the receptor, an adaptor between ATG8 and several other proteins. Among these proteins is ATG32, localized to mitochondria upon induction of mitophagy, thus recruiting selectively autophagy machinery to the mitochondria. Surprisingly, however, an absence of ATG32 does not affect cellular ROS levels or growth in starving conditions (Kanki et al., 2009), suggesting that alternative mechanisms of mitophagy exist. ATG11 and ATG32 have no particular homologs in higher eukaryotes (Kanki et al., 2009), except for mammals, which have functionally similar proteins. Such is, for example, the ubiquitin-binding adaptor p62 (SQSTM1) which as an ATG11-like role. p62 accumulates on damaged mitochondria and has been reported to recruit mitochondria to the

238

I. Matic et al.

autophagosomes. It remains controversial if binding to ATG8/LC3 is involved in this process (Geisler et al., 2010; Narendra et al., 2010a,b). Another functional homolog, Nix, may act as both ATG32 and ATG11. Nix is constitutively expressed on the mitochondria and has a core LC3interacting domain (LIR) motif required for direct binding to LC3 (Novak et al., 2010). Nix is involved in mitochondrial clearance in developing reticulocytes and erythrocytes (Sandoval et al., 2008; Schweers et al., 2007). Another protein—ubiquitin ligase Parkin is identified to be one of the most important players in recruiting autophagosomes to damaged mitochondria (Youle and Narendra, 2011). Parkin is an E3 ubiquitin ligase, which promotes the ubiquitin–proteasome system of protein degradation in mitochondrial quality control. Loss-of-function mutations in Parkin have been known to cause heritable forms of Parkinson’s disease, and emerging evidence also implicates a role for Parkin in other neurodegenerative diseases: Alzheimer’s disease, amyotrophic lateral sclerosis, and Huntington’s disease (Hebron et al., 2014; Rosen et al., 2010; Tsai et al., 2003). Interestingly, Parkin overexpression has been found in long-lived flies, and this discovery could provide a link between aging process and age-onset neurodegeneration (Rana et al., 2013). The fact that mitophagy is commonly classified into one of two categories—Parkin dependent or Parkin independent, underlies its role in the pathways elucidated so far. Several other molecules have been identified to serve as adaptors or receptors that mediate mitophagy such as histone deacetylase 6 (HDAC6) (Lee et al., 2010a,b), BCL2/adenovirus E1B interacting protein 3 (BNIP3) (Zhang and Ney, 2009), and FUN14 domain containing 1 (FUNDC1) (Liu et al., 2012). Mitophagy is also closely associated with mitochondrial fission, a process that segregates large mitochondria into smaller mitochondria to facilitate their removal by mitophagy. This process is controlled by dynamin-related protein 1 (Drp1) (Twig et al., 2008). When Drp1 is inactivated, mitophagy is impaired, implicating that fission is a requisite for mitophagy (Youle and Narendra, 2011). Drp1 is a cytosolic protein that can be recruited to the outer mitochondrial membrane (OMM) causing mitochondria to constrict and divide a mitochondrion. Drp1 is known to interact with four mitochondrial receptor proteins: fission 1 (Fis1), mitochondria fission factor (Mff ), and mitochondrial dynamics proteins MID49 and MID51. In mammalian cells, the interaction between Fis1 and Drp1 does not seem to be as important as the interactions of Drp1 with the other three receptor proteins for the control of mitochondrial fission

Molecular Biology Digest of Cell Mitophagy

239

(Gandre-Babbe and van der Bliek, 2008; Loso´n et al., 2013; Otera et al., 2010; Palmer et al., 2011; Westermann, 2010). Many lines of evidence suggest that Drp1 function is regulated through its posttranslational modifications (Chang and Blackstone, 2010). Detailed studies of proteins involved in targeting mitochondria to the autophagosome have extensively broadened our understanding of how mitophagy is initiated and executed. Later, we will discuss protein mitophagy receptor systems which have been most extensively mechanistically characterized, namely, the PINK1/Parkin system, as well as BNIP3/ NIX and Ambra-1 that play distinct and nonoverlapping activities to promote mitophagy (Kubli and Gustafsson, 2012). Even though the current set of mitophagy modulators seems fairly limited, additional players (such as nuclear dot protein (NDP52) and optineurin (OPTN)) are emerging (Youle and Narendra, 2011).

3. THE PARKIN-DEPENDENT WAY OF MITOPHAGY The common trigger for mitophagy is generally acknowledged to be the dissolution of ΔΨ m (Narendra et al., 2010a,b). The dogma recites that when dysfunctional mitochondria lose their ΔΨ m, the serine/threonine protein kinase phosphatase and tensin homolog-induced putative kinase 1 (PINK1) become activated and get stabilized on the OMM (Lin and Kang, 2008) (Fig. 1). PINK1 is ubiquitously expressed in mitochondria and comprises 581 amino acids. The molecular basis for PINK1 localization on the OMM of depolarized mitochondria rather than release to the cytosol is currently not entirely elucidated, but an interesting molecular mechanism underlying PINK1 activation on depolarized mitochondria has been recently proposed by Matsuda group (Okatsu et al., 2015). PINK1 contains three domains that are important for submitochondrial localization: an N-terminal mitochondrial targeting signal (MTS), a transmembrane sequence (TM) responsible for retention in inner mitochondrial membrane (IMM), and a C-terminal kinase domain (Trempe and Edward, 2013). Okatsu et al. revealed that the C-terminal kinase domain is located between the MTS and TM and represents the outer mitochondrial localization signal (OMS). PINK1 is usually degraded in energized mitochondria in an MTSand TM-dependent manner; however, dysfunction of the MTS targets PINK1 to the OMM and consequently, PINK1 is activated and transduces the signal to Parkin. The OMS does not depend on ΔΨ m, but on the translocase of the outer membrane (TOM) machinery instead. This feature

240

I. Matic et al.

Fig. 1 The Parkin-dependent way mitophagy. Mitochondrial depolarization leads to activation of PINK1 that phosphorylates Mfn2 as a receptor for Parkin recruitment and activation, which is mediated by PINK1 via phosphorylation of Ser65 within its UBL and phosphorylation of ubiquitin at Ser65. Once activated, Parkin ubiquitinates mitochondrial proteins including VDAC1, Mfn1, and Mfn2 at Lys48 and -63 leading to the recruitment of UBD mitophagic receptors such as OPTN, NBR1, p62, and NDP52. These proteins attach autophagosomal membrane via LC3 through their LIR motif. The OPTN phosphorylation, mediated by TBK1, increases PINK1–Parkin-mediated mitophagy by biding both ubiquitinated OMM proteins and LC3. Moreover, OPTN and NDP52 further recruit autophagy initiating factors, such as ULK1, DFCP1, and WIPI1. The OMM USP 30 and 15 are responsible for the ubiquitination of OMM proteins Parkin-mediated. In response to mitochondrial depolarization, PINK1 recruits Parkin together with Beclin-1 and AMBRA1 at the OMM contributing to Parkin-mediated mitophagy due to AMBRA1 capacity to bind LC3 through a LIR motif.

reminds of the N-terminal signal-anchored proteins that localize to the OMM in a ΔΨ m-independent and Tom40-dependent manner, which have the bulk of the polypeptide exposed to the cytosol (Ahting et al., 2005). Accordingly, Okatsu et al. suggest that PINK1 is a unique N-terminal signal-anchored protein containing an extra MTS and an IMM arrival-dependent degradation signal (TM) (Okatsu et al., 2015). In physiological conditions, the level of PINK1 is quite low because of its constitutive degradation. In healthy cells, PINK1 is rapidly cleaved and

Molecular Biology Digest of Cell Mitophagy

241

degraded by presenilins-associated rhomboid-like protein (PARL) at the IMM (Jin et al., 2010), as a way to prevent mitophagy of healthy mitochondria. However, PINK1 is stabilized on the OMM, where it can recruit autophagy-related proteins to impaired mitochondria when ΔΨ m is dissipated (Jin et al., 2010; Lazarou et al., 2015). Therefore, the bioenergetic state of mitochondria can regulate PINK1 levels and this regulation allows for selective and efficient turnover of mitochondria that have become damaged. The activated PINK1 phosphorylates specific proteins including mitofusin 2 (Mfn2) that serves as a receptor to recruit Parkin (Chen and Dorn, 2013), an E3 ubiquitin (Ub) ligase that normally resides in the cytosol (Fig. 1). Parkin was firstly implicated the mammalian mitophagy pathway in 2008 by Richard Youle’s group (Narendra et al., 2008). PINK1 activates Parkin E3 ligase activity both directly via phosphorylation of Parkin serine 65 (Ser65)—which lies within its ubiquitin-like domain (Ubl)—and indirectly through phosphorylation of ubiquitin at Ser65 (Fig. 1). Interestingly, the full activation of Parkin requires PINK1 to phosphorylate ubiquitin at Ser65 (Kazlauskaite et al., 2015). The activated Parkin then performs both Lys48 and Lys63 ubiquitination of mitochondrial proteins including voltage-dependent anion channel (VDAC1), Mfn1, and Mfn2 (Chan et al., 2011) (Fig. 1). Harper’s group found 36 OMM proteins that are Parkin substrates (Sarraf et al., 2013). Moreover, there are four known E2 ligases that interact with Parkin for mitochondrial protein ubiquitination: UBE2L3, UBE2D2, UBE2D3, and UBE2N (Fiesel et al., 2014; Geisler et al., 2014). It has been documented that mutations in Parkin undermine protein degradation via ubiquitination, leading to the accumulation of toxic misfolded or aggregated proteins (Ciechanover and Brundin, 2003; Um et al., 2010), and substantial focus has been directed toward Parkin’s role in mitochondrial quality control, particularly in neurodegenerative diseases (Ashrafi and Schwarz, 2013; Youle and Narendra, 2011). Proteomic studies suggest that the translocation of Parkin to dysfunctional mitochondria is associated mainly in the Lys48-linked polyubiquitylation and degradation of many proteins. In 2013, Rana et al. showed for the first time that ubiquitous adult-onset and/or neuron-specific overexpression of Parkin was sufficient to enhance longevity or neuron-specific upregulation in Drosophila melanogaster without reducing reproductive output, physical activity, or food intake (Rana et al., 2013). In fact, long-lived Parkin-overexpressing flies showed an increase in Lys48-linked polyubiquitin and reduced levels of

242

I. Matic et al.

protein aggregation during aging. Indirect confirmation of this discovery comes from the recent study of Ryu et al. (2016), showing that induction of mitophagy is able to prevent the accumulation of dysfunctional mitochondria during aging and extend lifespan in Caenorhabditis elegans and different mouse and rat models. The ubiquitinated mitochondria bind p62 and HDAC6 through the ubiquitin-binding domain (UBD) and are transported along microtubules to cluster in the perinuclear region (Lee et al., 2010a,b; Okatsu et al., 2010). p62 then recruits autophagosomal membranes via its LIR motif. Serine/threonine residues within the LIR preceding region were shown to be fundamental for the regulation of the autophagy receptor activity through phosphorylation (Liu et al., 2012; Wild et al., 2011; Zhu et al., 2013). Recently, a protein called OPTN was shown to act as the LIR-dependent autophagy receptor downstream of Parkin activation (Wong and Holzbaur, 2014) (Fig. 1). OPTN can phosphorylate p62 at Ser403 positively regulating p62 ubiquitin binding (Matsumoto et al., 2011), thus enhancing in this way the p62 targeting of mitochondria. The role of p62 in mitophagy has become controversial since it has been shown that p62 is dispensable (Narendra et al., 2010a,b; Okatsu et al., 2010). In fact, a recent study has shown that nuclear dot protein 52 kDa (NDP52) and OPTN were necessary for PINK1/Parkin-dependent mitophagy (Lazarou et al., 2015). PINK1 recruits OPTN and DPN52 to mitochondria (Lazarou et al., 2015), where they further recruit autophagy-related proteins, such as ULK1, DFCP1, WIPI1, and LC3, to initiate autophagy (Lazarou et al., 2015) (Fig. 1). An interesting recent study by Dikic group (Richter et al., 2016) identified Tank-binding kinase (TBK1) as sort of a signal amplifier in selective autophagy of damaged mitochondria since TBK1-mediated phosphorylation of OPTN’s UBAN domain at Ser473 expands the binding capacity of OPTN to diverse ubiquitin chains (Fig. 1). In the quantitative proteomics approach, Richter et al. found that TBK1 directly phosphorylates several autophagy-relevant sites also in other mitophagy receptors (NDP52 and p62), further controlling their binding to ubiquitin chains and regulating autophagy of damaged mitochondria (Richter et al., 2016). The finding that a mutant TBK1, unable to bind to OPTN, fails to translocate to damaged mitochondria, highlights an important role for OPTN in the regulation of TBK1, and suggests the latter to be vital in the control of selective autophagy pathways. Among mitochondrial targets of Parkin is the autophagy-promoting protein AMBRA1 (Van Humbeeck et al., 2011). In response to

Molecular Biology Digest of Cell Mitophagy

243

mitochondrial depolarization, AMBRA1 binds Parkin at the OMM (Strappazzon et al., 2011; Van Humbeeck et al., 2011) (Fig. 1). Although Ambra-1 is not required for Parkin translocation to depolarized mitochondria, it is critically important for subsequent mitochondrial clearance. Moreover, AMBRA1 was recently shown to bind LC3 through a LIR motif during Parkin-mediated mitophagy and forced targeting of AMBRA1 to the OMM resulted in efficient clearance of mitochondria, independent of Parkin (Strappazzon et al., 2015), which will be discussed in the next section. Other proteins found to be involved in activation of the Parkin-mediated mitophagy pathway are Nix and Smurf1. Ding et al. (2010) have shown that Nix is critical to two distinct phases of mitophagy: ROS-mediated autophagy induction and Parkin-ubiquitin-p62-mediated mitochondrial priming. Nix promotes the recruitment of Parkin to depolarized mitochondria upon carbonyl cyanide m-chlorophenyl hydrazone (CCCP) treatment in MEFs, and consistently, Nix-deficient MEF cells remained resistant to mitochondrial depolarization and Parkin recruitment (Ding et al., 2010). Additionally, a stimulating set of studies has been recently published by Levine’s group (Orvedahl et al., 2011). Their image-based genome-wide siRNA screening has identified another E3 ligase SMAD-specific E3 ubiquitin protein ligase 1 (SMURF1) downstream of Parkin, able to promote mitophagy independently of its catalytic activity. Interestingly, MEFs lacking the C2 domain-containing protein, Smurf1, showed to be deficient in the autophagosomal targeting and clearance of damaged mitochondria. Moreover, Smurf1-deficient mice displayed an accumulation of damaged mitochondria in heart, brain, and liver (Orvedahl et al., 2011). Smurf1 is an E3 ubiquitin ligase similar to Parkin. Smurf1 is not necessary for mitophagy, but for engulfment of damaged mitochondria by autophagosomes. Another independent screen has identified ATPase inhibitory factor 1 (ATPIF1) as an important factor for Parkin recruitment and mitophagy in cultured cells (Lefebvre et al., 2013). It has been shown that ATPIF1 acts as a bioenergetic adaptor by blocking the ATPase activity thus promoting collapse of ΔΨ m and enabling Parkin recruitment to the mitochondria (Lefebvre et al., 2013). Recently, our group showed that during ischemia, a shift in the IF1:F1Fo-ATPsynthase expression ratio occurs in neurons. In fact, in ATPIF1 overexpressing neurons ATP depletion is reduced and ΔΨ m resilient to reoxygenation as well as resistant to electrogenic, Ca2+dependent depolarization (Matic et al., 2016). This adaptation of

244

I. Matic et al.

mitochondria in mammalian neurons during hypoxic conditions has a protective role by coordinating prosurvival cell mitophagy and bioenergetics resilience. While protective in neurons, ATPIF1 overexpression in colon cancer cells has been linked with metabolic adaptation of cancer cells to enhanced aerobic glycolysis (Formentini et al., 2012). By proposing that ATPIF1 overexpression triggered production of ROS, which in turn promotes the transcriptional activation of the NFκB pathway, resulting in a cellular adaptive response that includes proliferation and Bcl-xL-mediated resistance to drug-induced cell death. These findings clearly evidence an important role of this mitochondrial protein in cell survival, which is likely to be the focus of therapy-oriented future studies. Parkin-induced mitophagy is also negatively regulated by the mitochondrial 18-kDa translocator protein (TSPO) and the ubiquitin-specific peptidases 30 (USP30) and USP15 (Bingol et al., 2014; Cornelissen et al., 2014; Gatliff et al., 2014). TSPO, firstly discovered as a peripheral mitochondrial benzodiazepines receptor (PBR), is situated on the OMM of mammalian cells in interaction with the VDAC1. Its core biochemical function resides in the translocation of cholesterol in the mitochondrial for metabolism and steroids synthesis (Gatliff and Campanella, 2016). In brain, TSPO is expressed in low levels at physiological conditions but these markedly increase at sites of brain injury and inflammation. In fact, TSPO is considered a biomarker/molecular sensor of active brain disease in experimental animals and in human studies as well as an emerging therapeutic target (Chen and Guilarte, 2008; Papadopoulos and Lecanu, 2009). For over 20 years, (11)C-PK11195 PET, which aims to image expression of the TSPO on activated microglia in the brain, has been used in preclinical and clinical research to investigate neuroinflammation in vivo in patients with brain diseases (Vivash and O’Brien, 2016). In light of a significant clinical potential of TSPO, diverse ligands have demonstrated their biological efficacy in experimental models of neurodegenerative diseases, and some of them are now in clinical trials for the treatment and/or diagnosis of neurodegenerative processes. Among these, one potent cholesterol-like TSPO ligand has been described as a neuroprotective compound (Kim and Pae, 2016). TSPO has been also involved with the deregulation of cell apoptosis (Gatliff and Campanella, 2012) and associated with the limitation of cell mitophagy via a redox regulation of mitochondrial ubiquitination, which represents a breakthrough of its contribution to mitochondrial and cellular biology (Gatliff and Campanella, 2012) (Fig. 2). In fact, Gatliff et al. demonstrated that overexpression of TSPO is associated with changes in mitochondrial morphology, ATP synthesis, Ca2+ signaling, and elevated ROS production that inhibits mitochondrial autophagy via impeding

Molecular Biology Digest of Cell Mitophagy

245

Fig. 2 Role of TSPO in pathology and mitochondrial quality control. TSPO which undermines mitochondrial quality control via inhibition of mitochondrial proteins ubiquitination is involved in neurological disorders, inflammation, and carcinogenesis. Even though the TSPO-mediated mechanisms underlying these physiological alterations are not well understood, it has been hypothesized that factors such as ROS may be responsible for the upregulation of TSPO through the activation of transcription factors such as AP and NFκB involved in cellular proliferation. Importantly, ROS is likely to be directly involved in the regulation of TSPO expression.

Parkin-dependent ubiquitination of proteins and consequently the recruitment of p62 and LC3 (Fig. 2). These data indicate that TSPO as an inducible, prooxidant element that undermines the quality control of mitochondria (Gatliff and Campanella, 2012, 2015, 2016). In addition, the role of TSPO in ischemia–reperfusion injury has been reported in vivo and in vitro porcine models of renal ischemia (Favreau et al., 2009). The overexpression of TSPO was able to rescue the cells from the detrimental effects of hypoxia and reoxygenation due to caspase activation, reduction of ATP content, and H2O2-induced necrosis (Favreau et al., 2009). In humans, TSPO has been implicated in renal ischemia–reperfusion and acute kidney injury (Thuillier and Hauet, 2012). Furthermore, TSPO has been implicated in carcinogenesis

246

I. Matic et al.

probably due to the role of TSPO in regulating transcription factor members involved in cellular proliferation such as AP-1, AP-2, and NF-κB (Batarseh and Papadopoulos, 2010; Batarseh et al., 2010) (Fig. 2). The selective autophagy receptor p62/SQSTM1 brings mitophagy into an intimate relationship with oxidative stress-sensing factors in two ways in mammals: first, p62 is transcriptionally induced upon oxidative stress by the NF-E2-related factor 2 (NRF2) by direct binding to an antioxidant response element (ARE) in the p62 promoter; second, p62 accumulation, occurring when autophagy is impaired, leads to increased p62 binding to the NRF2 inhibitor KEAP1, resulting in reduced proteasomal turnover of NRF2. The activation of AREs is a fine-tuned antioxidant response in attempt to improve the redox quality of the cell in a response to a variety of different stimuli. However, the relationship mitophagy— AREs is still not well understood, and selective pharmacologic modulators of these pathways are almost nonexistent (Matic et al., 2015). In the recent approach to this setback, our group has developed a new compound, named p62-mediated mitophagy inducer (PMI), able to increase the autophagic adaptor molecule p62/SQSTM1. Its action is based on disrupting the interaction between Keap1 and the nuclear factor E2-related factor 2 (Nrf2)-regulated antioxidant cascade, forcing mitochondria into autophagy without collapsing their ΔΨ m (East and Campanella, 2016; East et al., 2014). When speaking about fine-tuned regulation of mitophagy, it is important to mention USP30, a deubiquitinase localized to mitochondria that antagonizes pathways driven by Parkin and PINK1 (Bingol et al., 2014). The authors have shown that overexpression of USP30 removes ubiquitin attached by Parkin onto damaged mitochondria and blocks Parkin’s ability to drive mitophagy, while knockdown of USP30 rescues the defective mitophagy when Parkin or PINK1 activity is compromised (e.g., by pathogenic mutations). Similarly, Cornelissen et al. have shown that USP15, a deubiquitinating enzyme (DUB) widely expressed in brain and other organs, opposes Parkin-mediated mitophagy, while a panel of other DUBs and a catalytically inactive version of USP15 do not (Cornelissen et al., 2014). USP15 has been shown not to affect the ubiquitination status of Parkin or Parkin translocation to mitochondria but counteract instead the Parkin-mediated mitochondrial ubiquitination (Cornelissen et al., 2014). USP30 and USP15 inhibition could be potentially beneficial in conditions (for example, Parkinson’s disease) in which promoting mitochondrial clearance and quality control can help improve the symptoms.

Molecular Biology Digest of Cell Mitophagy

247

In addition to promoting selective autophagy, Parkin was also shown to inhibit autophagy via monoubiquitination of Bcl-2 (Chen et al., 2010). This ubiquitin signal did not increase the degradation of Bcl-2 but increased its stability. An interaction between Parkin and Bcl-2, but not Bcl-XL, has been documented (Chen et al., 2010). The enhanced stability of Bcl-2 resulted in a stronger affinity between Beclin 1 and Bcl-2. By using mutated Parkin, the E3 ubiquitin ligase activity was found to be required for these effects. The answer how Parkin induces mitophagy while inhibiting autophagy in general, probably lies in its localization. Damage to mitochondria recruits Parkin to the mitochondria to promote mitophagic depletion of these organelles, while upon a more global stress like starvation; Parkin is probably retained in the cytosol (Chen et al., 2010), where it serves an antiautophagic function. While significant progress has been made in elucidating mechanisms of Parkin-dependent mitophagy, results emerging from a number of different studies still remain to be reconciled.

4. THE PARKIN-INDEPENDENT WAY OF MITOPHAGY In addition to the PINK1/Parkin mitophagy system, a group of mitophagy receptors constitutively localized at the OMM, contain LIR domain, which is phosphorylated in response to specific stimuli enabling their commitment to mitophagy (Hamacher-Brady and Brady, 2016) (Fig. 2). A common trigger for their activation is hypoxia. One of the most studied is BNIP3, a mitochondrial Bcl-2 homology 3 (BH3) domain-containing proapoptotic protein whose gene expression is induced during hypoxia by hypoxia-inducing factor-1 alpha (HIF-1α) (Bruick, 2000). HIF-1α binds to the HIF-1-response element (HRE) within the BNIP3 gene promoter (Bruick, 2000) (Fig. 3). Nix (BNIP3L) is a homolog of BNIP3 (Chen et al., 1999), whose expression is also induced by HIF1α during hypoxia (Bruick, 2000). Nix is able to bind to Bcl-2 dissociating it from the complex of Bcl-2 and Beclin-1, necessary for initiation of autophagosome formation (Bellot et al., 2009) (Fig. 3). Nix can also interact with the autophagosome membrane protein LC3 (Novak et al., 2010) contributing to selective autophagy by recruiting autophagosomes to damaged mitochondria (Fig. 3). NIX and BNIP3 seem to complement each other in mitophagy induction during hypoxia since the loss of both proteins inhibits hypoxia-induced mitophagy, while individual loss of either BNIP3 or Nix seems irrelevant for autophagy (Bellot et al., 2009).

248

I. Matic et al.

Fig. 3 Parkin-independent way of mitophagy. Mitophagy receptors also include constitutively OMM-anchored proteins such as BNIP3, Nix, and FUNDC1. They are activated by transcription or phosphorylation during hypoxia. BNIP3 and Nix expression is induced by HIF-1α that binds the HRE within the BNIP3 gene promoter. They activate autophagy by binding to Bcl-2 and release Beclin-1, a protein necessary for initiation of autophagosome formation. Nevertheless, BNIP3 mitophagic activity is also due to its phosphorylation of Ser17 and -24 adjacent to LIR motif that promotes the interaction with LC3. FUNDC1 expression is regulated by phosphorylation. Under normal conditions, FUNDC1 is inactivated by phosphorylation at Tyr16 via the Src kinase. During hypoxia or mitochondrial depolatization, PGAM5 dephosphorylates FUNDC1 on Ser13 activating LC3 binding on the autophagosome membrane through LIR motif. Another mitophagy receptor is Ambra-1, upon mitophagy induction, Ambra-1 together with Beclin-1 translocates from the cytoskeleton to the ER to regulate autophagosome nucleation via its phosphorylation mediated by ULK1. Then Ambra-1 promotes via TRAF6 the ubiquitination of ULK-1 through Lys63-linked ubiquitin chain. This modification is essential for self-association of ULK1 and its autophagic activity. ULK1 ubiquitination at Lys63 is prevented by mTORC1 that is kept in an active state by Ambra-1 phosphorylation.

Nix-induced mitophagy is important for other processes nonrelated to stress, such is maturation of red blood cells, which destroy their mitochondria during the maturation process (Sandoval et al., 2008; Schweers et al., 2007). Furthermore, in cells with high oxidative phosphorylation activity Nix has been found in a physical interaction with the small GTPase Rheb recruited to the OMM and the autophagosomal protein LC3-II (Melser et al., 2013). This interaction can prevent mitochondrial damage and subsequent ROS production, mitochondrial dysfunction, cell death, and injury (Melser et al., 2013).

Molecular Biology Digest of Cell Mitophagy

249

Another mitophagy receptor is FUNDC1, an OMM protein that also contributes to mitophagy induction under hypoxic conditions. FUNDC1 activation during hypoxia-induced mitophagy is regulated by phosphorylation. In normal conditions, FUNDC1 is phosphorylated by the Src kinase on Tyr18 (Liu et al., 2012). During hypoxia, the mitochondrial phosphatase phosphoglycerate mutase family member 5 (PGAM5) dephosphorylates FUNDC1 on Ser13 (Chen et al., 2014) (Fig. 3). When FUNDC1 is dephosphorylated, its LIR’s binding affinity for LC3 increases following ULK1 recruitment, allowing FUNDC1 phosphorylation by ULK1 at Ser17 and FUNDC1 interaction with LC3 on the autophagosome membrane (Liu et al., 2012; Wu et al., 2014) (Fig. 3). As discussed earlier, NIX (but also BNIP3) may directly trigger mitochondrial depolarization, which has been shown to be sufficient to induce mitophagy (Elmore et al., 2001; Twig et al., 2008). Thus, it remains unclear how NIX/BNIP3 interact with Parkin to regulate mitophagy. Wang et al. have demonstrated that, ubiquitinated by Parkin, Nix recruits neighbor of BRCA gene 1 protein (NBR1) to remove damaged mitochondria (Gao et al., 2015). Both NIX and BNIP3 can induce cell death but remains to be elucidated how their mitophagy functions are regulated under physiological or pathological conditions (Chu et al., 2013; Zhang and Ney, 2009). In addition to protein mitophagy receptors, the lipids ceramide and cardiolipin can directly bind LC3 and initiate mitophagy (Sentelle et al., 2012). Cardiolipin is synthesized at the IMM (Schlattner et al., 2014) and, similar to Parkin, translocates to the OMM upon mitochondrial depolarization to induce mitophagy (Ren et al., 2014). However, cardiolipin and Parkin seem to mediate mitophagy under different levels of mitochondrial depolarization. Cardiolipin translocates to the OMM after CCCP treatment, but not to the same extent as Parkin, suggesting that there may be some overlap in Parkin and cardiolipin-induced mitophagy pathways or that they both may respond to CCCP treatment (Chu et al., 2013). AMBRA1 is considered to be another mitophagy receptor. It has been defined as Beclin 1 interactor that positively regulates PtdIns-3KC3 (Fimia et al., 2007) (Fig. 3). Upon autophagy induction AMBRA1 abandons the cytoskeleton location and relocalizes to the endoplasmic reticulum (ER) to regulate autophagosome nucleation (Di Bartolomeo et al., 2010). Recently, Strappazzon et al. (2015) showed that AMBRA1 possesses a LIR domain that allows it to bind directly to LC3 upon mitophagy induction (Fig. 1). Furthermore, targeting AMBRA1 to mitochondria (using

250

I. Matic et al.

AMBRA1-ActA construct) is sufficient to induce mitophagy in both parkin-dependent and -independent systems (in HeLa cells, for instance). This work suggests an alternative pathway for mitophagy in the absence of parkin. When AMBRA-1 was targeted to the mitochondria, they succumbed ubiquitylation, as observed in parkin-deficient cells. Moreover, AMBRA1 mediates ubiquitylation of the kinase ULK1, an important in upstream regulation of autophagy, by the E3-ligase TNF receptor-associated factor 6 (TRAF6) (Nazio et al., 2013) (Fig. 3). These data bring to conclusion that most likely, AMBRA1 serves as an adaptor for E3 ligases. Considering the fast advancement in the study of mitophagy mechanisms and their applications into physiopathological processes, it is likely that other Parkin-independent mitophagy players will be exposed in the future.

5. CONCLUDING REMARKS Mitochondria play many important roles in maintaining cellular homeostasis and decision-making when it comes to cell death/survival situation. They act as a sort of death executors as well, considering their ability to release cell death molecules from their intermembrane space or by generating toxic ROS. Therefore, once they are damaged, their removal via pathways of mitophagy becomes of crucial importance for the well-being of the cells. Pioneering studies that have aimed to clarify mitophagy mechanisms individuated two major types of processes: the Parkin-dependent and the Parkin-independent pathways. Although roles and interactions of many proteins/players in these pathways have been documented extensively in the past years, the physiological conditions of their recruitment—specifically, whether or not their interplay leads to mitophagy—remain elusive. Much evidence demonstrates that mitophagy is particularly important for mitochondrial quality control under pathological conditions, such as tumor growth and microbial invasion, which are of great clinical importance. Even though many therapeutic approaches have been premeditated for treating mitochondrial dysfunction underlying disorders, there is an evident lack of valid pharmacological drugs available. This is partly reflected from the inability to find a consensus on the proper conditions with which to study these pathways in a more disease-relevant cell types. However, some promising advances—pharmacological inducers/inhibitors, are emerging and future work in targeting both Parkin-dependent and

Molecular Biology Digest of Cell Mitophagy

251

-independent players in pathophysiological conditions will surely produce more drugs for increasingly attractive mitophagy therapeutic targets. In addition, recent studies on mitophagy mechanisms during aging in invertebrate and rodent models could highlight mitochondrial quality control as attractive target in slowing down aging/prolonging lifespan and preventing age-related diseases.

ACKNOWLEDGMENTS We would like to thank our funders. The research activities led by M.C. are supported by the following funders, which are gratefully acknowledged: Biotechnology and Biological Sciences Research Council (Grant numbers BB/M010384/1 and BB/N007042/1); the Medical Research Council (Grant number G1100809/2), Bloomsbury Colleges Consortium PhD Studentship Scheme; The Petplan Charitable Trust; Umberto Veronesi Foundation Young Investigator Research Programme; Marie Curie Actions [TSPO & Brain (304165)], LAM-Bighi Grant Initiative. FIRB-Research Grant Consolidator Grant 2 (Grant number: RBFR13P392), Italian Ministry of Health Ministero (IFO14/01/R/52). Conflict of Interest: Authors have no conflict to declare.

REFERENCES Ahting, U., Waizenegger, T., Neupert, W., Rapaport, D., 2005. Signal-anchored proteins follow a unique insertion pathway into the outer membrane of mitochondria. J. Biol. Chem. 280, 48–53. Andres, A.M., Stotland, A., Queliconi, B.B., Gottileb, R.A., 2015. A time to reap, a time to sow: mitophagy and biogenesis in cardiac pathophysiology. J. Mol. Cell. Cardiol. 78, 62–72. Ashford, T.P., Porter, K.R., 1962. Cytoplasmic components in hepatic cell lysosomes. J. Cell Biol. 12, 198–202. Ashrafi, G., Schwarz, T.L., 2013. The pathways of mitophagy for quality control and clearance of mitochondria. Cell Death Differ. 20, 31–42. Balaban, R.S., Nemoto, S., Finkel, T., 2005. Mitochondria, oxidants, and aging. Cell 120, 483–495. Batarseh, A., Papadopoulos, V., 2010. Regulation of translocator protein 18 kDa (TSPO) expression in health and disease states. Mol. Cell. Endocrinol. 327 (1–2), 1–12. Batarseh, A., Li, J., Papadopoulos, V., 2010. Protein kinase C epsilon regulation of translocator protein (18 kDa) Tspo gene expression is mediated through a MAPK pathway targeting STAT3 and c-Jun transcription factors. Biochemistry 49 (23), 4766–4778. Beaulaton, J., Lockshin, K.R., 1977. Ultrastructural study of the normal degeneration of the intersegmental muscles of Antheraea polyphemus and Manduca sexta (Insecta, Lepidoptera) with particular reference of cellular autophagy. J. Morphol. 154, 39–57. Bellot, G., Garcia-Medina, R., Gounon, P., Chiche, J., Roux, D., Pouyssegur, J., Mazure, N.M., 2009. Hypoxia-induced autophagy is mediated through hypoxia-inducible factor induction of BNIP3 and BNIP3l via their BH3 domains. Mol. Cell. Biol. 29, 2570–2581. Biala, A.K., Dhingra, R., Kirshenbaum, L.A., 2015. Mitochondrial dynamics: orchestrating the journey to advanced age. J. Mol. Cell. Cardiol. 83, 37–43.

252

I. Matic et al.

Bingol, B., Tea, J.S., Phu, L., Reichelt, M., Bakalarski, C.E., Song, Q., Foreman, O., Kirkpatrick, D.S., Sheng, M., 2014. The mitochondrial deubiquitinase USP30 opposes parkin-mediated mitophagy. Nature 510, 370–375. Bratic, A., Larsson, N.G., 2013. The role of mitochondria in aging. J. Clin. Invest. 123, 951–957. Bruick, R.K., 2000. Expression of the gene encoding the proapoptotic Nip3 protein is induced by hypoxia. Proc. Natl. Acad. Sci. U.S.A. 97, 9082–9087. Burte, F., Carelli, V., Chinnery, P.F., Yu-Wai-Man, P., 2015. Disturbed mitochondrial dynamics and neurodegenerative disorders. Nat. Rev. Neurol. 11 (1), 11–24. Calvo, S.E., Mootha, V.K., 2010. The mitochondrial proteome and human disease. Annu. Rev. Genomics Hum. Genet. 11, 25–44. Campanella, M., Klionsky, D.J., 2013. Keeping the engine clean: a mitophagy task for cellular physiology. Autophagy 9 (11), 1647. Chan, N.C., Salazar, A.M., Phamm, A.H., Sweredoski, M.J., Kolawa, N.J., Graham, R.L., Hess, S., Chan, D.C., 2011. Broad activation of the ubiquitin-proteasome system by Parkin is critical for mitophagy. Hum. Mol. Genet. 20, 1726–1737. Chang, C.R., Blackstone, C., 2010. Dynamic regulation of mitochondrial fission through modification of the dynamin-related protein Drp1. Ann. N. Y. Acad. Sci. 1201, 34–39. Chen, Y., Dorn 2nd., G.W., 2013. PINK1-phosphorylated mitofusin 2 is a Parkin receptor for culling damaged mitochondria. Science 340, 471–475. Chen, M.K., Guilarte, T.R., 2008. Translocator protein 18kDA (TSPO): molecular sensor of brain injury and repair. Pharmacol. Ther. 118 (1), 1–17. Chen, G., Cizeau, J., Vande Velde, C., Park, J.H., Bozek, G., Bolton, J., Shi, L., Dubik, D., Greenberg, A., 1999. Nix and Nip3 form a subfamily of pro-apoptotic mitochondrial proteins. J. Biol. Chem. 274, 7–10. Chen, D., Gao, F., Li, B., Wang, H., Xu, Y., Zhu, C., Wang, G., 2010. Parkin mono-ubiquitinates Bcl-2 and regulates autophagy. J. Biol. Chem. 285, 38214–38223. Chen, G., Han, Z., Feng, D., Chen, Y., Chen, L., Wu, H., Huang, L., Zhou, C., Cai, X., Fu, C., Duan, L., Wang, X., Liu, L., Liu, X., Shen, Y., Zhu, Y., Chen, Q., 2014. A regulatory signaling loop comprising the PGAM5 phosphatase and CK2 controls receptor-mediated mitophagy. Mol. Cell 54, 362–377. Chu, C.T., Ji, J., Dagda, R.K., Jiang, J.F., Tyurina, Y.Y., Kapralov, A.A., Tyurin, V.A., Yanamala, N., Shrivastava, I.H., Mohammadyani, D., 2013. Cardiolipin externalization to the outer mitochondrial membrane acts as an elimination signal for mitophagy in neuronal cells. Nat. Cell Biol. 15, 1197–2105. Ciechanover, A., Brundin, P., 2003. The ubiquitin proteasome system in neurodegenerative diseases: sometimes the chicken, sometimes the egg. Neuron 40 (2), 427–446. Cornelissen, T., Haddad, D., Wauters, F., Van Humbeeck, C., Mandemakers, W., Koentjoro, B., Sue, C., Gevaert, K., De Strooper, B., Verstreken, P., Vandenberghe, W., 2014. The deubiquitinase USP15 antagonizes Parkin-mediated mitochondrial ubiquitination and mitophagy. Hum. Mol. Genet. 23, 5227–5242. Cotan, D., Cordero, M.D., Garrido-Maraver, J., Oropesa-Avila, M., Rodrı´guez-Herna´ndez, A., Gomez Izquierdo, L., de la Mata, M., De Miguel, M., Lorite, J.B., Infante, E.R., Jackson, S., Navas, P., Sa´nchez-Alca´zar, J.A., 2011. Secondary coenzyme Q10 deficiency triggers mitochondria degradation by mitophagy in MELAS fibroblasts. FASEB J. 25, 2669–2687. Di Bartolomeo, S., Corazzari, M., Nazio, F., Oliverio, S., Lisi, G., Antonioli, M., Pagliarini, V., Matteoni, S., Fuoco, C., Giunta, L., D’Amelio, M., Nardacci, R., Romagnoli, A., Piacentini, M., Cecconi, F., Fimia, G.M., 2010. The dynamic interaction of AMBRA1 with the dynein motor complex regulates mammalian autophagy. J. Cell Biol. 191, 155–168.

Molecular Biology Digest of Cell Mitophagy

253

Ding, W.X., Ni, H.M., Li, M., Liao, Y., Chen, X., Stolz, D.B., Dorn 2nd, G.W., Yin, X.M., 2010. Nix is critical to two distinct phases of mitophagy, reactive oxygen species-mediated autophagy induction and Parkin-ubiquitin-p62-mediated mitochondrial priming. J. Biol. Chem. 285, 27879–27890. East, D.A., Campanella, M., 2016. Mitophagy and the therapeutic clearance of damaged mitochondria for neuroprotection. Int. J. Biochem. Cell Biol. 79, 382–387. East, D.A., Fagiani, F., Crosby, J., Georgakopoulos, N.D., Bertrand, H., Schaap, M., Fowkes, A., Wells, G., Campanella, M., 2014. PMI: a ΔΨm independent pharmacological regulator of mitophagy. Chem. Biol. 21, 1585–1596. Elmore, S.P., Qian, T., Grissom, D.F., Lemasters, J.J., 2001. The mitochondrial permeability transition initiates autophagy in rat hepatocytes. FASEB J. 15, 2286–2287. Favreau, F., Rossard, L., Zhang, K., Desurmont, T., Manguy, E., Belliard, A., Fabre, S., Liu, J., Han, Z., Thuillier, R., Papadopoulos, V., Hauet, T., 2009. Expression and modulation of translocator protein and its partners by hypoxia reoxygenation or ischemia and reperfusion in porcine renal models. Am. J. Physiol. Renal Physiol. 297 (1), 177–190. Fiesel, F.C., Moussaud-Lamodie`re, E.L., Ando, M., Springer, W., 2014. A specific subset of E2 ubiquitin-conjugating enzymes regulate Parkin activation and mitophagy differently. J. Cell Sci. 127, 3488–3504. Filomeni, G., De Zio, D., Cecconi, F., 2015. Oxidative stress and autophagy: the clash between damage and metabolic needs. Cell Death Differ. 22, 377–388. Fimia, G.M., Stoykova, A., Romagnoli, A., Giunta, L., Di Bartolomeo, S., Nardacci, R., Corazzari, M., Fuoco, C., Ucar, A., Schwartz, P., Gruss, P., Piacentini, M., Chowdhury, K., Cecconi, F., 2007. Ambra1 regulates autophagy and development of the nervous system. Nature 447, 1121–1125. Formentini, L., Sa´nchez-Arago´, M., Sa´nchez-Cenizo, L., Cuezva, J.M., 2012. The mitochondrial ATPase inhibitory factor 1 triggers a ROS-mediated retrograde prosurvival and proliferative response. Mol. Cell 45, 731–742. Gandre-Babbe, S., van der Bliek, A.M., 2008. The novel tail-anchored membrane protein Mff controls mitochondrial and peroxisomal fission in mammalian cells. Mol. Biol. Cell 19, 2402–2412. Gao, F., Chen, D., Si, J., Hu, Q., Qin, Z., Fang, M., Wang, G., 2015. The mitochondrial protein BNIP3L is the substrate of PARK2 and mediates mitophagy in PINK1/PARK2 pathway. Hum. Mol. Genet. 24, 2528–2538. Gatliff, J., Campanella, M., 2012. The 18 kDa translocator protein (TSPO): a new perspective in mitochondrial biology. Curr. Mol. Med. 12, 356–368. Gatliff, J., Campanella, M., 2015. TSPO is a REDOX regulator of cell mitophagy. Biochem. Soc. Trans. 43, 543–552. Gatliff, J., Campanella, M., 2016. TSPO: kaleidoscopic 18-kDa amid biochemical pharmacology, control and targeting of mitochondria. Biochem. J. 473, 107–121. Gatliff, J., East, D., Crosby, J., Abeti, R., Harvey, R., Craigen, W., Parker, P., Campanella, M., 2014. TSPO interacts with VDAC1 and triggers a ROS-mediated inhibition of mitochondrial quality control. Autophagy 10 (12), 2279–2296. Geisler, S., Holmstr€ om, K.M., Skujat, D., Fiesel, F.C., Rothfuss, O.C., Kahle, P.J., Springer, W., 2010. PINK1/Parkin-mediated mitophagy is dependent on VDAC1 and p62/SQSTM1. Nat. Cell Biol. 12, 119–131. Geisler, S., Vollmer, S., Golombek, S., Kahle, P.J., 2014. The ubiquitin-conjugating enzymes UBE2N, UBE2L3 and UBE2D2/3 are essential for Parkin-dependent mitophagy. J. Cell Sci. 127, 3280–3293. Gomes, L.C., Di Benedetto, G., Scorrano, L., 2011. During autophagy mitochondria elongate, are spared from degradation and sustain cell viability. Nat. Cell Biol. 13, 589–598.

254

I. Matic et al.

Hamacher-Brady, A., Brady, N.R., 2016. Mitophagy programs: mechanisms and physiological implications of mitochondrial targeting by autophagy. Cell. Mol. Life Sci. 73, 775–795. Hebron, M., Chen, W., Miessau, M.J., Lonskaya, I., Moussa, C.E., 2014. Parkin reverses TDP-43-induced cell death and failure of amino acid homeostasis. J. Neurochem. 129, 350–361. Holmstr€ om, K.M., Finkel, T., 2014. Cellular mechanisms and physiological consequences of redox-dependent signalling. Nat. Rev. Mol. Cell Biol. 15, 411–421. Jacobson, M.D., Burne, J.F., Raff, M.C., 1994. Programmed cell death and Bcl-2 protection in the absence of a nucleus. EMBO J. 13, 1899–1910. Jin, S.M., Lazarou, M., Wang, C., Kane, L.A., Narendra, D.P., Youle, R.J., 2010. Mitochondrial membrane potential regulates PINK1 import and proteolytic destabilization by PARL. J. Cell Biol. 191, 933–942. Kacerovsky-Bielesz, G., Kacerovsky, M., Chmelik, M., Farukuoye, M., Ling, C., Pokan, R., Tschan, H., Szendroedi, J., Schmid, A.I., Gruber, S., Herder, C., Wolzt, M., Moser, E., Pacini, G., Smekal, G., Groop, L., Roden, M., 2012. A single nucleotide polymorphism associates with the response of muscle ATP synthesis to long-term exercise training in relatives of type 2 diabetic humans. Diabetes Care 35, 350–357. Kanki, T., Klionsky, D.J., 2008. Mitophagy in yeast occurs through a selective mechanism. J. Biol. Chem. 283, 32386–32393. Kanki, T., Wang, K., Cao, Y., Baba, M., Klionsky, D.J., 2009. Atg32 is a mitochondrial protein that confers selectivity during mitophagy. Dev. Cell 17, 98–109. Kanki, T., Wang, K., Klionsky, D.J., 2010. A genomic screen for yeast mutants defective in mitophagy. Autophagy 6, 278–280. Kazlauskaite, A., Martı´nez-Torres, R.J., Wilkie, S., Kumar, A., Peltier, J., Gonzalez, A., Johnson, C., Zhang, J., Hope, A.G., Peggie, M., Trost, M., van Aalten, D.M., Alessi, D.R., Prescott, A.R., Knebel, A., Walden, H., Muqit, M.M., 2015. Binding to serine 65-phosphorylated ubiquitin primes Parkin for optimal PINK1-dependent phosphorylation and activation. EMBO Rep. 16, 939–954. Kim, T., Pae, A.N., 2016. Translocator protein (TSPO) ligands for the diagnosis or treatment of neurodegenerative diseases: a patent review. Expert Opin. Ther. Pat. 6, 1–14. Kissova´, I., Deffieu, M., Manon, S., Camougrand, N., 2004. Uth1p is involved in the autophagic degradation of mitochondria. J. Biol. Chem. 279, 39068–39074. Kobayashi, S., Liang, Q., 2015. Autophagy and mitophagy in diabetic cardiomyopathy. Biochim. Biophys. Acta 1852, 252–261. Koopman, W.J., Willems, P.H., Smeitink, J.A., 2012. Monogenic mitochondrial disorders. N. Engl. J. Med. 366, 1132–1141. Kristensen, A.R., Schandorff, S., Høyer-Hansen, M., Nielsen, M.O., J€a€attel€a, M., Dengjel, J., Andersen, J.S., 2008. Ordered organelle degradation during starvation-induced autophagy. Mol. Cell. Proteomics 7 (12), 2419–2428. Kroemer, G., 1997. The proto-oncogene Bcl-2 and its role in regulating apoptosis. Nat. Med. 3, 614–620. Kroemer, G., 2004. Early work on the role of mitochondria in apoptosis, an interview with Guido Kroemer. Cell Death Differ. 11, S33–S36. Kroemer, G., Petit, P., Zamzami, N., Vayssie`re, J.L., Mignotte, B., 1995. The biochemistry of programmed cell death. FASEB J. 9, 1277–1287. Kroemer, G., Galluzzi, L., Brenner, C., 2007. Mitochondrial membrane permeabilization in cell death. Physiol. Rev. 87, 99–163. Kubli, D.A., Gustafsson, A˚.B., 2012. Mitochondria and mitophagy: the yin and yang of cell death control. Circ. Res. 111 (9), 1208–1221. Lazarou, M., Sliter, D.A., Kane, L.A., Sarraf, S.A., Wang, C., Burman, J.L., Sideris, D.P., Fogel, A.I., Youle, R.J., 2015. The ubiquitin kinase PINK1 recruits autophagy receptors to induce mitophagy. Nature 524, 309–314.

Molecular Biology Digest of Cell Mitophagy

255

Lee, J.Y., Koga, H., Kawaguchi, Y., Tang, W., Wong, E., Gao, Y.S., Pandey, U.B., Kaushik, S., Tresse, E., Lu, J., Taylor, J.P., Cuervo, A.M., Yao, T.P., 2010a. HDAC6 controls autophagosome maturation essential for ubiquitin-selective quality-control autophagy. EMBO J. 29, 969–980. Lee, J.Y., Nagano, Y., Taylor, J.P., Lim, K.L., Yao, T.P., 2010b. Disease-causing mutations in parkin impair mitochondrial ubiquitination, aggregation, and HDAC6-dependent mitophagy. J. Cell Biol. 189, 671–679. Lefebvre, V., Du, Q., Baird, S., Ng, A.C., Nascimento, M., Campanella, M., McBride, H.M., Screaton, R.A., 2013. Genome-wide RNAi screen identifies ATPase inhibitory factor 1 (ATPIF1) as essential for PARK2 recruitment and mitophagy. Autophagy 9 (11), 1770–1779. Lemasters, J.J., 2005. Selective mitochondrial autophagy, or mitophagy, as a targeted defense against oxidative stress, mitochondrial dysfunction, and aging. Rejuvenation Res. 8, 3–5. Lin, W., Kang, U.J., 2008. Characterization of PINK1 processing, stability, and subcellular localization. J. Neurochem. 106, 464–474. Liu, L., Feng, D., Chen, G., Chen, M., Zheng, Q., Song, P., Ma, Q., Zhu, C., Wang, R., Qi, W., Huang, L., Xue, P., Li, B., Wang, X., Jin, H., Wang, J., Yang, F., Liu, P., Zhu, Y., Sui, S., Chen, Q., 2012. Mitochondrial outer-membrane protein FUNDC1 mediates hypoxia-induced mitophagy in mammalian cells. Nat. Cell Biol. 14, 177–185. Loso´n, O.C., Song, Z., Chen, H., Chan, D.C., 2013. Fis1, Mff, MiD49, and MiD51 mediate Drp1 recruitment in mitochondrial fission. Mol. Biol. Cell 24, 659–667. Maiuri, M.C., Zalckvar, E., Kimchi, A., Kroemer, G., 2007. Self-eating and self-killing: crosstalk between autophagy and apoptosis. Nat. Rev. Mol. Cell Biol. 8, 741–752. Matic, I., Strobbe, D., Frison, M., Campanella, M., 2015. Controlled and impaired mitochondrial quality in neurons: molecular physiology and prospective pharmacology. Pharmacol. Res. 99, 410–424. Matic, I., Cocco, S., Ferraina, C., Martin-Jimenez, R., Florenzano, F., Crosby, J., Lupi, R., Amadoro, G., Russell, C., Pignataro, G., Annunziato, L., Abramov, A.Y., Campanella, M., 2016. Neuroprotective coordination of cell mitophagy by the ATPase inhibitory factor 1. Pharmacol. Res. 103, 56–68. Matsumoto, G., Wada, K., Okuno, M., Kurosawa, M., Nukina, N., 2011. Serine 403 phosphorylation of p62/SQSTM1 regulates selective autophagic clearance of ubiquitinated proteins. Mol. Cell 44, 279–289. Melser, S., Chatelain, E.H., Lavie, J., Mahfouf, W., Jose, C., Obre, E., Goorden, S., Priault, M., Elgersma, Y., Rezvani, H.R., Rossignol, R., Benard, G., 2013. Rheb regulates mitophagy induced by mitochondrial energetic status. Cell Metab. 17, 719–730. Narendra, D., Tanaka, A., Suen, D.F., Youle, R.J., 2008. Parkin is recruited selectively to impaired mitochondria and promotes their autophagy. J. Cell Biol. 183, 795–803. Narendra, D.P., Jin, S.M., Tanaka, A., Suen, D.F., Gautier, C.A., Shen, J., Cookson, M.R., Youle, R.J., 2010a. PINK1 is selectively stabilized on impaired mitochondria to activate Parkin. PLoS Biol. 8, e1000298–e1000299. Narendra, D., Kane, L.A., Hauser, D.N., Fearnley, I.M., Youle, R.J., 2010b. p62/SQSTM1 is required for Parkin-induced mitochondrial clustering but not mitophagy; VDAC1 is dispensable for both. Autophagy 6, 1090–1106. Nazio, F., Strappazzon, F., Antonioli, M., Bielli, P., Cianfanelli, V., Bordi, M., Gretzmeier, C., Dengjel, J., Piacentini, M., Fimia, G.M., Cecconi, F., 2013. mTOR inhibits autophagy by controlling ULK1 ubiquitylation, self-association and function through AMBRA1 and TRAF6. Nat. Cell Biol. 15 (4), 406–416. Novak, I., Kirkin, V., McEwan, D.G., Zhang, J., Wild, P., Rozenknop, A., Rogov, V., L€ ohr, F., Popovic, D., Occhipinti, A., Reichert, A.S., Terzic, J., D€ otsch, V., Ney, P.A., Dikic, I., 2010. Nix is a selective autophagy receptor for mitochondrial clearance. EMBO Rep. 11, 45–51.

256

I. Matic et al.

Okamoto, K., Kondo-Okamoto, N., Ohsumi, Y., 2009. Mitochondria-anchored receptor Atg32 mediates degradation of mitochondria via selective autophagy. Dev. Cell 17, 87–89. Okatsu, K., Saisho, K., Shimanuki, M., Nakada, K., Shitara, H., Sou, Y.S., Kimura, M., Sato, S., Hattori, N., Komatsu, M., Tanaka, K., Matsuda, N., 2010. p62/SQSTM1 cooperates with Parkin for perinuclear clustering of depolarized mitochondria. Genes Cells 15, 887–900. Okatsu, K., Kimura, M., Oka, T., Tanaka, K., Matsuda, N., 2015. Unconventional PINK1 localization to the outer membrane of depolarized mitochondria drives Parkin recruitment. J. Cell Sci. 128 (5), 964–978. Orvedahl, A., Sumpter Jr., R., Xiao, G., Ng, A., Zou, Z., Tang, Y., Narimatsu, M., Gilpin, C., Sun, Q., Roth, M., Forst, C.V., Wrana, J.L., Zhang, Y.E., Luby-Phelps, K., Xavier, R.J., Xie, Y., Levine, B., 2011. Image-based genome-wide siRNA screen identifies selective autophagy factors. Nature 480, 113–117. Otera, H., Wang, C., Cleland, M.M., Setoguchi, K., Yokota, S., Youle, R.J., Mihara, K., 2010. Mff is an essential factor for mitochondrial recruitment of Drp1 during mitochondrial fission in mammalian cells. J. Cell Biol. 191, 141–1158. Palade, G.E., 1953. An electron microscope study of the mitochondrial structure. J. Histochem. Cytochem. 1, 188–211. Palmer, C.S., Osellame, L.D., Laine, D., Koutsopoulos, O.S., Frazier, A.E., Ryan, M.T., 2011. MiD49 and MiD51, new components of the mitochondrial fission machinery. EMBO Rep. 12, 565–573. Papadopoulos, V., Lecanu, L., 2009. Translocator protein (18 kDa) TSPO: an emerging therapeutic target in neurotrauma. Exp. Neurol. 219 (1), 53–57. Raben, N., Wong, A., Ralston, E., Myerowitz, R., 2012. Autophagy and mitochondria in Pompe disease: nothing is so new as what has long been forgotten. Am. J. Med. Genet. C Semin. Med. Genet. 160C (1), 13–21. Rana, A., Rera, M., Walker, D.W., 2013. Parkin overexpression during aging reduces proteotoxicity, alters mitochondrial dynamics, and extends lifespan. Proc. Natl. Acad. Sci. U.S.A. 110 (21), 8638–8643. Reme, C.E., Young, R.W., 1977. The effects of hibernation on cone visual cells in the ground squirrel. Invest. Ophthalmol. Vis. Sci. 16, 815–840. Ren, M., Phoon, C.K., Schlame, M., 2014. Metabolism and function of mitochondrial cardiolipin. Prog. Lipid Res. 55, 1–16. Richter, B., Sliter, D.A., Herhaus, L., Stolz, A., Wang, C., Beli, P., Zaffagnini, G., Wild, P., Martens, S., Wagner, S.A., Youle, R.J., Dikic, I., 2016. Phosphorylation of OPTN by TBK1 enhances its binding to Ub chains and promotes selective autophagy of damaged mitochondria. Proc. Natl. Acad. Sci. U.S.A. 113 (15), 4039–4044. Rodrı´guez-Herna´ndez, A., Cordero, M.D., Salviati, L., Artuch, R., Pineda, M., Briones, P., Go´mez Izquierdo, L., Cota´n, D., Navas, P., Sa´nchez-Alca´zar, J.A., 2009. Coenzyme Q deficiency triggers mitochondria degradation by mitophagy. Autophagy 5, 19–32. Rosen, K.M., Moussa, C.E., Lee, H.K., Kumar, P., Kitada, T., Qin, G., Fu, Q., Querfurth, H.W., 2010. Parkin reverses intracellular beta-amyloid accumulation and its negative effects on proteasome function. J. Neurosci. Res. 88, 167–178. Ryu, D., Mouchiroud, L., Andreux, P.A., Katsyuba, E., Moullan, N., Nicolet-Dit-Felix, A.A., Williams, E.G., Jha, P., Lo Sasso, G., Huzard, D., Aebischer, P., Sandi, C., Rinsch, C., Auwerx, J., 2016. Urolithin A induces mitophagy and prolongs lifespan in C. elegans and increases muscle function in rodents. Nat. Med. 22 (8), 879–888. Sandoval, H., Thiagarajan, P., Dasgupta, S.K., Schumacher, A., Prchal, J.T., Chen, M., Wang, J., 2008. Essential role for Nix in autophagic maturation of erythroid cells. Nature 454, 232–235. Sarraf, S.A., Raman, M., Guarani-Pereira, V., Sowa, M.E., Huttlin, E.L., Gygi, S.P., Harper, J.W., 2013. Landscape of the PARKIN-dependent ubiquitylome in response to mitochondrial depolarization. Nature 496, 372–376.

Molecular Biology Digest of Cell Mitophagy

257

Schlattner, U., Tokarska-Schlattner, M., Rousseau, D., Boissan, M., Mannella, C., Epand, R., Lacombe, M.L., 2014. Mitochondrial cardiolipin/phospholipid trafficking: the role of membrane contact site complexes and lipid transfer proteins. Chem. Phys. Lipids 179, 32–41. Schulze-Osthoff, K., Walczak, H., Droge, W., Krammer, P.H., 1994. Cell nucleus and DNA fragmentation are not required for apoptosis. J. Cell. Biol. 127, 15–20. Schweers, R.L., Zhang, J., Randall, M.S., Loyd, M.R., Li, W., Dorsey, F.C., Kundu, M., Opferman, J.T., Cleveland, J.L., Miller, J.L., Ney, P.A., 2007. NIX is required for programmed mitochondrial clearance during reticulocyte maturation. Proc. Natl. Acad. Sci. U.S.A. 104, 19500–19505. Sentelle, R.D., Senkal, C.E., Jiang, W., Ponnusamy, S., Gencer, S., Selvam, S.P., Ramshesh, V.K., Peterson, Y.K., Lemasters, J.J., Szulc, Z.M., Bielawski, J., Ogretmen, B., 2012. Ceramide targets autophagosomes to mitochondria and induces lethal mitophagy. Nat. Chem. Biol. 8, 831–838. Sjostrand, F.S., 1953. Electron microscopy of mitochondria and cytoplasmic double membranes. Nature 171, 30–32. Strappazzon, F., Vietri-Rudan, M., Campello, S., Nazio, F., Florenzano, F., Fimia, G.M., Piacentini, M., Levine, B., Cecconi, F., 2011. Mitochondrial BCL-2 inhibits AMBRA1-induced autophagy. EMBO J. 30, 1195–1208. Strappazzon, F., Nazio, F., Corrado, M., Cianfanelli, V., Romagnoli, A., Fimia, G.M., Campello, S., Nardacci, R., Piacentini, M., Campanella, M., Cecconi, F., 2015. AMBRA1 is able to induce mitophagy via LC3 binding, regardless of PARKIN and p62/SQSTM1. Cell Death Differ. 22, 419–432. Sun, N., Youle, R.J., Finkel, T., 2016. The mitochondrial basis of aging. Mol. Cell 61, 654–666. Susin, S.A., Zamzami, N., Castedo, M., Hirsch, T., Marchetti, P., Macho, A., Daugas, E., Geuskens, M., Kroemer, G.A., 1996. Bcl-2 inhibits the mitochondrial release of an apoptogenic protease. J. Exp. Med. 184, 1331–1342. Thuillier, R., Hauet, T.T., 2012. Role of translocator protein in renal ischemia reperfusion, renal preservation and acute kidney injury. Curr. Mol. Med. 12 (4), 413–425. Trempe, J.F., Edward, A.F., 2013. Structure and function of Parkin, PINK1, and DJ-1, the three musketeers of neuroprotection. Front. Neurol. 4, 38. Tsai, Y.C., Fishman, P.S., Thakor, N.V., Oyler, G.A., 2003. Parkin facilitates the elimination of expanded polyglutamine proteins and leads to preservation of proteasome function. J. Biol. Chem. 278, 22044–22055. Twig, G., Elorza, A., Molina, A.J., Mohamed, H., Wikstrom, J.D., Walzer, G., Stiles, L., Haigh, S.E., Katz, S., Las, G., Alroy, J., Wu, M., Py, B.F., Yuan, J., Deeney, J.T., Corkey, B.E., Shirihai, O.S., 2008. Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. EMBO J. 27, 433–446. Um, J.W., Im, E., Lee, H.J., Min, B., Yoo, L., Yoo, J., L€ ubbert, H., Stichel-Gunke, l.C., Cho, H.S., Yoon, J.B., Chung, K.C., 2010. Parkin directly modulates 26S proteasome activity. J. Neurosci. 30 (35), 11805–11814. Van Humbeeck, C., Cornelissen, T., Hofkens, H., Mandemakers, W., Gevaert, K., De Strooper, B., Vandenberghe, W., 2011. Parkin interacts with Ambra1 to induce mitophagy. J. Neurosci. 31, 10249–10261. Vaux, D.L., 2011. Apoptogenic factors released from mitochondria. Biochim. Biophys. Acta 1813, 546–550. Vivash, L., O’Brien, T.J., 2016. Imaging microglial activation with TSPO PET: lighting up neurologic diseases? J. Nucl. Med. 57 (2), 165–168. Wallace, D.C., 2012. Mitochondria and cancer. Nat. Rev. Cancer 12, 685–698. Westermann, B., 2010. Mitochondrial fusion and fission in cell life and death. Nat. Rev. Mol. Cell Biol. 11, 872–884.

258

I. Matic et al.

Wild, P., Farhan, H., McEwan, D.G., Wagner, S., Rogov, V.V., Brady, N.R., Richter, B., Korac, J., Waidmann, O., Choudhary, C., D€ otsch, V., Bumann, D., Dikic, I., 2011. Phosphorylation of the autophagy receptor optineurin restricts Salmonella growth. Science 333, 228–233. Wong, Y.C., Holzbaur, E.L., 2014. Optineurin is an autophagy receptor for damaged mitochondria in Parkin-mediated mitophagy that is disrupted by an ALS-linked mutation. Proc. Natl. Acad. Sci. U.S.A. 111, E4439–E4448. Wu, J.J., Quijano, C., Chen, E., Liu, H., Cao, L., Fergusson, M.M., Rovira, I.I., Gutkind, S., Daniels, M.P., Komatsu, M., Finkel, T., 2009. Mitochondrial dysfunction and oxidative stress mediate the physiological impairment induced by the disruption of autophagy. Aging 1 (4), 425–437. Wu, W., Tian, W., Hu, Z., Chen, G., Huang, L., Li, W., Zhang, X., Xue, P., Zhou, C., Liu, L., Zhu, Y., Zhang, X., Li, L., Zhang, L., Sui, S., Zhao, B., Feng, D., 2014. ULK1 translocates to mitochondria and phosphorylates FUNDC1 to regulate mitophagy. EMBO Rep. 15, 566–575. Youle, R.J., Narendra, D.P., 2011. Mechanisms of mitophagy. Nat. Rev. Mol. Cell Biol. 12, 9–14. Zamzami, N., Susin, S.A., Marchetti, P., Hirsch, T., Go´mez-Monterrey, I., Castedo, M., Kroemer, G., 1996. Mitochondrial control of nuclear apoptosis. J. Exp. Med. 183, 1533–1544. Zhang, J., 2013. Autophagy and mitophagy in cellular damage control. Redox Biol. 1 (1), 19–23. Zhang, J., Ney, P.A., 2009. Role of BNIP3 and NIX in cell death, autophagy, and mitophagy. Cell Death Differ. 16, 939–946. Zhu, Y., Massen, S., Terenzio, M., Lang, V., Chen-Lindner, S., Eils, R., Novak, I., Dikic, I., Hamacher-Brady, A., Brady, N.R., 2013. Modulation of serines 17 and 24 in the LC3interacting region of Bnip3 determines pro-survival mitophagy versus apoptosis. J. Biol. Chem. 288, 1099–1113.

CHAPTER SIX

Regulation of Cell Calcium and Role of Plasma Membrane Calcium ATPases T. Calì*, M. Brini*, E. Carafoli†,1 *University of Padova, Padova, Italy † Venetian Institute of Molecular Medicine, Padova, Italy 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. PMCA Pump 3. Isoforms of PMCA Pump 4. Regulation of PMCA Pump 5. Role of PMCA Pump in Regulating Cell Ca2+ 6. PMCA Pumps and Pathology 7. Concluding Remarks Acknowledgments References

260 261 264 271 279 282 286 287 287

Abstract The plasma membrane Ca2+ ATPase (PMCA pump) is a member of the superfamily of P-type pumps. It has 10 transmembrane helices and 2 cytosolic loops, one of which contains the catalytic center. Its most distinctive feature is a C-terminal tail that contains most of the regulatory sites including that for calmodulin. The pump is also regulated by acidic phospholipids, kinases, a dimerization process, and numerous protein interactors. In mammals, four genes code for the four basic isoforms. Isoform complexity is increased by alternative splicing of primary transcripts. Pumps 2 and 3 are expressed preferentially in the nervous system. The pumps coexist with more powerful systems that clear Ca2+ from the bulk cytosol: their role is thus the regulation of Ca2+ in selected subplasma membrane microdomains, where a number of important Ca2+-dependent enzymes interact with them. Malfunctions of the pump lead to disease phenotypes that affect the nervous system preferentially.

International Review of Cell and Molecular Biology, Volume 332 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2017.01.002

#

2017 Elsevier Inc. All rights reserved.

259

260

T. Calì et al.

1. INTRODUCTION Ca2+ conveys messages to virtually all cell functions. The complexity of its signaling role has increased with the transition from unicellular to multicellular life but is still based on a single general mechanism: the reversible binding to specific proteins. The basic principle of regulation is reversibility, and thus, messengers that transmit regulatory signals to cell functions must be able to do so reversibly. In choosing Ca2+ as a transmitter of regulatory messages evolution has exploited its peculiar coordination chemistry, which has permitted the modulation of its concentration, and thus of the information it carried, in the environment of targets. Basically (Brini et al., 2013a,c; Carafoli and Krebs, 2016; Carafoli et al., 2001), the chemistry of Ca2+ allows it to optimally accept binding sites of very irregular geometry, at sharp variance, for instance, with that of Mg2+, which would demand perfectly octahedral-binding sites. Perfectly octahedral sites do not come about in the macromolecules available in cells, e.g., proteins. Which instead offer a large cadre if sites able to bind Ca2+ with high affinity in the presence of large excesses of cations like Mg2+, thus permitting the lowering of its concentration to the very low levels demanded by the signaling function. As is self-evident, a carrier of signals must be maintained within cells at extremely low levels to prevent unsustainable energy expenditures to vary its concentration as the signaling function would demand. All Ca2+-binding proteins that have been developed during evolution contribute to the buffering of cell Ca2+ to these very low levels, but do so in different ways. They could be soluble in the cytosol or inside organelles and simply bind it reversibly. Or they could be intrinsic to membranes and buffer Ca2+ by transporting it across them. Importantly, some of the soluble proteins (the Ca2+ sensors), in addition to buffering Ca2+, also decode its message for the benefit of the targets. The Ca2+-binding proteins intrinsic to membranes belong to different mechanistic classes: ATPases (colloquially called pumps), located in the endo(sarco)plasmic reticulum, the Golgi membranes, and the plasma membrane; exchangers (mostly Na+/Ca2+-exchangers), channels and “electrophoretic uniporters” (in the inner membrane of mitochondria). The properties of all these transporters have been extensively covered in numerous reviews (Axelsen and Palmgren, 1998; Brini and Carafoli, 2009). This chapter will deal with the Ca2+ pump of the plasma membrane: it will focus on a number of recent findings, also coming from the area of pathology, that have led to novel concepts on some aspects of its activity and on the significance of its role in the regulation of cellular Ca2+ signaling.

Plasma Membrane Ca2 + Pumps

261

2. PMCA PUMP The PMCA pump belongs to the superfamily of ion and lipid transporting ATPases. The superfamily is phylogenetically partitioned into five subfamilies (P1–P5), each divided into subgroups (A, B, C, D, etc.). The pumps of the superfamily all share a single catalytic subunit, the formation of a transient phosphorylated intermediate at a conserved Asp residue during each catalytic cycle, and the ability to undergo large structural changes during the transport process. They differ in the transported species and in the mechanism of regulation. In general, P1-ATPases are K+ (A) and heavy metal (B) pumps; P2-ATPases are Ca2+ (A and B), Na+/K+ and H+/K+ (C), and Na+ (D) pumps; P3A-ATPases are H+ pumps; P4-ATPases are putative lipid flippases; and P5-ATPase have still no ion transport specificity (Axelsen and Palmgren, 1998). In animal cells, the P2A Ca2+-ATPases are the sarco(endo)plasmic reticulum Ca2+ (SERCA) pumps and the secretory pathway Ca2+-ATPase (SPCA) primarily found in the membranes of the Golgi apparatus, the P2B are the plasma membrane Ca2+-ATPases (PMCA). They transport Ca2+ against steep transmembrane gradients: for each ATP hydrolyzed, the SERCA pumps transfer two Ca2+ ions within the sarco(endo)plasmic reticulum and counter transport H+ to the cytosol, the PMCA pumps, export one Ca2+ ion against a steady-state Ca2+ gradient of 10,000-fold across the plasma membrane. One feature makes the PMCAs unique among the P-type ATPases, i.e., they are activated by calmodulin (CaM), which binds at a C-terminal extension that functions as a pump autoinhibitory domain (Brini and Carafoli, 2009). The solution of the crystal structures of the SERCA pump in different conformations (Toyoshima and Inesi, 2004; Toyoshima et al., 2000) has established its three-dimensional arrangement which, despite the low degree of sequence conservation, is similar to that of the other P-type ATPases whose structures have been solved and has defined the molecular steps of the catalytic cycle (Brini and Carafoli, 2009; Palmgren and Nissen, 2011). First discovered in erythrocytes (Schatzmann, 1966), the PMCA was then described in numerous other cell types. It was purified in 1979 using a calmodulin affinity column (Niggli et al., 1979) and cloned about 10 years later (Shull and Greeb, 1988; Verma et al., 1988). It shows the same membrane topology properties of the SERCA pump, using the structure of the latter as a template (Toyoshima et al., 2000), the same general structural fea˚ long and 40 A ˚ wide tures can be predicted (Fig. 1): the pump is 120 A with one end inserted in the membrane, a minimal fraction exposed to the

262

T. Calì et al.

Fig. 1 The Ca2+-ATPases transport cycle. Two main conformational states of the PMCA modeled on that of the SERCA pump are shown: E1 binds Ca2+ with high affinity at the cytosolic site (PDB files 1SU4), E2 has a low affinity for Ca2+, releasing it to the opposite site of the membrane (PDB files 1IWO). The conserved catalytic Asp residue is shown in blue, the A-, P-, and the N-cytosolic domains are shown in green, magenta, and red, respectively. The transmembrane domain and the extracellular domain are shown in light brown and black, respectively. Ca2+-bound calmodulin activates the pump by inducing conformational changes that involve both the cytosolic domains and the intermembrane helices. In the active state the structure of the cytosolic portion of the pump is more open (see text). The calmodulin-binding domain of the PMCA and calmodulin is shown in yellow and light blue (PDB files 2KNE and 1CFC) and has been manually superimposed on the main core of the pump.

extracellular side, and a large cytoplasmic portion with the 3 canonical cytoplasmic domains (A, actuator; N, nucleotide binding; P, phosphorylation) and the 10 transmembrane domains (M1–10). N- and C-terminal regulatory (R) domains containing inhibitory and activatory sites (see below) can also

Plasma Membrane Ca2 + Pumps

263

be defined. The A-domain is predicted as a globular protein phosphatase connected to the transmembrane region by three long and flexible linker sequences attached to transmembrane helices M1, M2, and M3 (Takahashi et al., 2007). It contains the conserved Thr-Gly-Glu (TGE) motif which is important during the catalytic cycle, one of the two sites that mediate the activation of the pump by acidic phospholipids, and one of the two sites for the autoinhibitory interaction of the C-terminal calmodulin-binding domain with the main core of the pump. In addition, the A-loop also contains site A, which is one of the two main sites of alternative splicing that generate pump variants (see below). The phosphorylated invariant Asp (D)-residue is contained in a highly conserved SDKTGT[L/I/ V/M][T/I/S] sequence in the central surface pointing away from the membrane of the pump (the P-domain). Two conserved sequences in this domain, Thr-Gly-Asp-Asn (TGDN) and Gly-Asp-Glyx-Asn-Asp (GDGXND), are involved in the coordination of an Mg2+ ion for ATP binding at the N-domain, which is responsible for the phosphorylation of the catalytic Asp in the P-domain, to which it is connected by a narrow hinge. Thus, the A-domain is an extension of the first cytoplasmic loop, and the N-domain is an insertion into the P-domain, both of which are part of the second large cytoplasmic loop, which also contains the second autoinhibitory-binding site for the C-terminal calmodulin-binding domain. The M2, M4, and M6 helices of the transmembrane domain are involved in the formation of half-channels that harbor the Ca2+-binding site and are alternatively exposed to the cytoplasm and the extracellular side (Toyoshima and Nomura, 2002). The Ca2+-binding site is half-way in the transmembrane region of the pump and alternates between high affinity (exposed to the cytoplasmic side) and low affinity (exposed to the extracellular side), corresponding to the E1 and the E2 form of the enzyme, respectively. Ca2+ binding to the high-affinity cytoplasmic site induces conformational changes that generate a Mg2+-binding site in proximity of the invariant Asp thus allowing the binding of ATP in the N-domain. The conserved Asp residue can now be phosphorylated to produce the E1-P form and, as a result, the A-domain undergoes large conformational changes that obscure the high-affinity Ca2+-binding site and create an exit channel on the other side of the membrane. The TGE motif of the A-domain lies close to the phosphorylated Asp. This is the rate-limiting step of the cycle and corresponds to the transition to the E2-P form. Ca2+ is now free to diffuse away from the extracellular side of the pump. A conformational change that shifts the Glu residue of the TGE motif

264

T. Calì et al.

toward the phosphorylated Asp stimulates its dephosphorylation and the release of phosphate (E2 state). The return to the relaxed E1 state is preceded by a movement of the A-domain with concomitant rearrangements of the transmembrane domain that reestablish the high-affinity Ca2+ site. The C-terminal R-domain of the PMCA pumps makes them peculiar among all P-type ATPases. This long (120 aa) tail contains the calmodulin-binding domain (James et al., 1988) (which, however, also binds activatory acidic phospholipids (Brodin et al., 1992)) that autoinhibits the pump by binding to conserved “acceptor” sites (residues 206–271 and 537–544 in the hPMCA4) in the first and second cytoplasmic loops of the pump, respectively (Falchetto et al., 1991, 1992). Additional studies on the autoinhibition process have suggested that additional regions downstream of the “canonical” CaM-BD could also play a role in the autoinhibition, with the Asp 170 residue potentially involved in the stabilization of the inactive state (Bredeston and Adamo, 2004). Interestingly, in the plant Ca2+ pumps the calmodulin-binding domain is placed at the N-terminus of the protein (Ekberg et al., 2010) and is functionally interchangeable with that of the animal Ca2+ ATPases (Bonza and Luoni, 2010). In all pumps, calmodulin displaces the autoinhibitory C-terminal domain from the conserved “acceptor” sites in the main body of the pump molecule relieving the autoinhibition, i.e., decreasing the Kd for Ca2+ from 10 to 20 μM to less than 1 μM. An additional calmodulin-binding domain downstream of the canonical one has recently been identified in some variants of the pump. It has been suggested that it would permit the regulation of the pump both in the nM and μM ranges of Ca2+ concentration (Tidow et al., 2012).

3. ISOFORMS OF PMCA PUMP The PMCA pump is the product of a multigene family. In mammals four basic genes (ATP2B1–4) code for four main gene products, classified as isoforms 1–4. But, starting from four different transcripts, a complex alternative splicing process increases the total number of isoforms to about 30, which differ in organ and tissue distribution, and in the abundance of their expression changes during development. The existence of such a large group of PMCA isoforms and splice variants immediately raises the question of the functional significance of this diversity and of the rationale of this apparent redundancy. According to a simplified vision of the problem, pumps 1 and 4 are ubiquitous and have lower calmodulin affinity (Kd  30–50 nM), pumps

Plasma Membrane Ca2 + Pumps

265

2 and 3 have higher calmodulin sensitivity (Kd  2–8 nM) and tissue-restricted expression: PMCA2 is expressed prominently in the nervous system and in the mammary gland, PMCA3 in the nervous tissue and the muscle system (Strehler and Zacharias, 2001). As mentioned, this is a simplified version: the distribution of the pumps in the plasma membrane is not homogeneous but is regulated by their colocalization with partners proteins and, in polarized cells, by isoform-specific targeting to different plasma membrane domains, i.e., basolateral or apical plasma membrane (Guerini and Carafoli, 1999; Ortega et al., 2007; Strehler and Zacharias, 2001). All the four PMCA transcripts undergo alternative splicing at two sites (site A and site C): site A is located upstream of the phospholipid-binding domain in the first cytosolic loop of the pump, and site C is in the C-terminal calmodulin-binding domain. The splicing process at site A leads to the insertion of one exon (leading to the x variant in the case of PMCA1, 3, and 4) or up to three exons in PMCA2, thus generating variants w (three exons included), y (two exons included), and x (only one exon included). The A site inserts are always in frame; thus, they do not substantially alter the structure of the pumps but change their properties or their plasma membrane distribution. The z variants have no inserts. Variant z is not found in PMCA1, as all mature transcripts of this isoform invariably contain one inserted exon, i.e., they correspond to the x variant. PMCA3 and 4 are only present as x or z variants. The splicing process at site C is characterized by the inclusion of one (in PMCA1 and 4) or two (in PMCA2 and 3) extra exons that change the reading frame and introduce a premature stop codon leading to truncated proteins designated as variant a. Pumps in which no insertions occur at site C are designated instead as b or full-length variants. The insertion of a single exon, however, can occur piecemeal, leading to variants c, d, e, and f according to the isoform when the insertion of portions of the single exon has occurred (in the case of PMCA3, in addition to the exon, the following intron and a polyA tail can also be introduced). A detailed description of the splicing variants is found in Brini and Carafoli (2009). Table 1 summarizes the tissue distribution and the abundance of the main PMCA splicing variants (Strehler, 2015). A number of studies on isolated PMCAs, or on pumps expressed as recombinant proteins, have determined the enzymatic, regulatory, and structural differences among the splice variants. They have established that the different PMCA variants differ in enzyme kinetics (Vmax, Km for Ca2+)

Table 1 Isoforms of PMCA Pumps Isoforms of the PMCA Pump Temporal Expression of the Splicing Variant (in Brain)

Size in aa (in Tissue or Cell Distribution in Human) Humans

PMCA1 x/a

1a embryonic 18 day

1176

Brain

x/b

1b embryonic 10 day

1220

Ubiquitous; lung; small intestine; kidney

x/c

1249

Skeletal muscle; heart

x/d

1258

Skeletal muscle

x/e

1171

Brain

1199

Brain; cochlear outer hair cells

1168

Brain; hippocampal presynaptic terminals

1154

Brain; excitable issue

1243

Brain; lactating mammary epithelial cells; pancreatic beta cells

x/b

1212

Brain; cerebellar Purkinje cells; spinal cord

z/b

1198

Brain; excitable tissue

Temporal Expression of the Main Transcript in Mouse (First Detection at mRNA Level)

Isoform

Embryonic 9, 5 day

Embryonic 12, 5 day

Human Splicing Variant

PMCA2 w/a x/a

2a embryonic 18 day, but 2a greatly increases at postnatal 2–30 days in cerebellum

z/a w/b

2b embryonic 18 day

Embryonic 12, 5 day

PMCA3 x/a

3a embryonic 18 day

1173

Brain; spinal cord

1159

Brain; pancreatic beta cells

1220

Brain; adrenal gland; skeletal muscle

z/b

1206

Brain

PMCA4 x/a

1170

Smooth muscle; bladder; uterus; heart

z/a

1158

Smooth muscle; heart

x/b

1205

Ubiquitous; heart; kidney

z/b

1193

Heart

x/d

1241

Heart

z/d

1229

Heart

x/e

1164

Brain, bladder

1152

Brain, bladder

z/a x/b

Embryonic 12, 5 day (but the level is low during the development, except than in liver)

3b embryonic 10 day

z/e 2+

Modified from Strehler, E.E., 2015. Plasma membrane calcium ATPases: from generic Ca Commun. 460, 26–33.

pumps to versatile systems for fine-tuning cellular Ca2+. Biochem. Biophys. Res.

268

T. Calì et al.

and other functional aspects, but mostly in the interplay with calmodulin. The C-terminal truncation generated by the C site inserts in the a variants lowers the affinity of the pumps for calmodulin (Brini et al., 2003; Preiano et al., 1996). Most of these studies have been performed in vitro; thus, they have provided only limited information on the true physiological properties of the splice isoforms. Unfortunately, one major problem in studying PMCA pump activity in intact cells is that multiple isoforms and splice variants are generally present simultaneously, and isoform-specific inhibitors are available. Nevertheless, Ca2+ measurements in cells overexpressing different pump isoforms have revealed that the a and b variants of the neuron-specific PMCA2 and 3 isoforms were much more effective in counteracting cytosolic transients generated by cell stimulation than the ubiquitously expressed PMCA1 and 4 isoforms (Brini et al., 2003). They have failed to show major differences in Ca2+ extrusion ability between the truncated and full-length variants of PMCA3 and 4 isoforms, suggesting that either in intact cells calmodulin was not a limiting factor or that their differences in calmodulin affinity were overcome under condition of maximal activation (Brini et al., 2003). The analysis of the joint contribution of site A and site C splicing to the Ca2+ handling ability of the PMCA2 pump has instead revealed that the z/a, w/b, and z/b splicing variants are all very active. However, the doubly inserted w/a variant had about the same high nonstimulated activity of the full-length z/b variant in resting conditions, but only limited ability to rapidly increase activity when challenged with a Ca2+ pulse (Ficarella et al., 2007). Measurements of ATPase activity in microsomal membranes of transfected CHO cells have shown that the PMCA2w/a variant, in addition to being less sensitive to calmodulin than the z/b and w/b isoforms, was also less sensitive to phosphatidylserine (PS); thus underlining the role of acidic phospholipids in the regulation of pump activity by interacting with the calmodulin-binding domain and opening the question of whether the site A inserts, which are close to the binding site for acidic phospholipids in the first cytosolic loop, could also affect the modulation of pump activity by phospholipids. The finding that the z/b and w/b isoforms had the same response to PS stimulation suggests that the A site splicing insertion has no role in the phospholipid sensitivity of the pump (Brini et al., 2010). An interesting indication on the possible physiological meaning of the splicing at site A has come from the finding that the insertion of the three exons at site A of the PMCA2 isoform targets it to the apical domain of polarized cells, whereas smaller inserts sort the protein to the basolateral

Plasma Membrane Ca2 + Pumps

269

domain of the plasma membrane (Chicka and Strehler, 2003). Interestingly, the splicing differentially affects the lipid interactions of PMCA pump with the membrane and the apical localization of this PMCA pump variant is lipid raft-dependent and sensitive to cholesterol depletion (Xiong et al., 2009). An elegant analysis in intact cells has revealed a functional interplay between PMCA isoforms (PMCA4a, PMCA4b, and PMCA2b) and the store-operated Ca2+ entry (SOCE), i.e., the influx of Ca2+ from the extracellular ambient promoted be the depletion of Ca2+ stores. The slow PMCA4b isoform produced long-lasting Ca2+ oscillations in response to the activation of SOCE, whereas the rapid PMCA2b isoform resulted in the rapid clearance of SOCE-mediated Ca2+ transients. The activation of the PMCA4a isoform reduced the cytosolic Ca2+ transient induced by Ca2+ entry but resulted in the establishment of a higher basal cytosolic Ca2+ concentration indicating that this isoform has the ability to respond to repeated stimuli. The study included a mathematical modeling that described how the distinct properties of the three isoforms differentially affected the shape and the kinetics of the Ca2+ transients generated by SOCE activation and how and thus their relative abundance in different cell types could lead to the specific activation of downstream signaling pathways (Paszty et al., 2015). Overall, Ca2+ signaling assays in cultured cells have clearly shown that the PMCA isoforms and splice variants may play different roles. They may be differentially involved in the housekeeping role of controlling bulk Ca2+ and in the fine tuning of local Ca2+ signaling events in localized plasma membrane microdomains, where specific variants may be differentially confined by targeted signals generated by the splicing mechanism (see below). The complexity of PMCA isoform production and expression in the control of Ca2+ signals is made even more evident by the simultaneous expression, with distinct subcellular localization, of multiple PMCA isoforms within the same tissue or cell type. Highly polarized cells are perhaps the most striking example of the situation: in retinal photoreceptor cells, PMCA1b is present in the inner segment and synaptic terminal membrane. Whereas the PMCA4b variant is restricted to the synaptic terminals (Krizaj et al., 2002). In cochlear hair cells, PMCA2w/a is exclusively present in the stereocilia of the apical membrane and PMCA1x/b in the basolateral membrane (Dumont et al., 2001). A particularly striking example of the role of PMCA pump variants in specifying specialized function has emerged from the identification of pathologies associated with the malfunctioning of specific isoform. This

270

T. Calì et al.

aspect of the isoforms’ role will be discussed in more detail in a later section of the contribution, but the point should be mentioned here that the loss of function or the reduced activity of one specific PMCA isoform, even in the presence of other normally functioning PMCA variants, is sufficient to affect Ca2+ handling to a point that renders cells more susceptible to additional stress agents. One last comment on the matter of PMCA isoforms and splicing variants concerns the changes of their expression during the embryonic development and the differentiation of cells. In muscle, the alternative splicing events occur during myogenic differentiation: the application of the muscle differentiation factor myogenin to L6 myoblasts cell lines induced them (Hammes et al., 1994). Interestingly, the induction of the splice form 1c of PMCA1 occurs upon myotube formation (Brandt et al., 1992; De Jaegere et al., 1993). Nerve growth factor treatment of PC12 pheochromocytoma cells leads to the appearance of the “differentiation-specific” splice variants of PMCAs 1, 2, and 4 (i.e., 1c, 2a, 4a) (Hammes et al., 1994). Similarly, a marked upregulation of PMCA1a, 2, and 3 at the mRNA and protein level occurs in rat cerebellar granule cells kept under depolarizing conditions for several days (leading to increased Ca2+ influx) (Guerini et al., 1999). In contrast, elevation of intracellular Ca2+ resulted in a rapid (within hours) and specific downregulation of the PMCA4a splice variant by a process mediated by the Ca2+/calmodulin-sensitive phosphatase calcineurin (Guerini et al., 2000). In the human neuroblastoma cell line IMR32, differentiation is accompanied by a marked upregulation of PMCA isoforms 2 and 4 (and to a lesser extent, of PMCA1), which in turn leads to an improved Ca2+ extrusion efficiency (Usachev et al., 2001). Another interesting example is the upregulation of PMCA4b expression occurring during colon and gastric cancer cells differentiation that closely correlates with the induction of established differentiation markers, suggesting that the increase in the PMCA-dependent Ca2+ transport activity characterizes the differentiation of these cancer cells (Ribiczey et al., 2007). As will be discussed later, PMCA pump isoforms, including their splice variants are selectively recruited to plasma membrane compartments/ domains by the interaction with specific proteins. The functional meaning of the interaction is double: specific interactors recruit PMCA pump isoforms to subplasma membrane domains, but in turn the Ca2+-ejection properties of PMCA maintain intracellular Ca2+ in the specific subplasma membrane microdomains at the set point demanded by the interacting

Plasma Membrane Ca2 + Pumps

271

proteins which are important components of Ca2+-sensitive transduction pathways.

4. REGULATION OF PMCA PUMP As mentioned earlier, a distinguishing feature of the PMCA pumps is the wealth of their regulatory processes. This had been recognized very early in the studies on the pump, and the activation by calmodulin (Gopinath and Vincenzi, 1979; Jarrett and Penniston, 1977) was actually instrumental in devising the protocol that led to the purification of the enzyme. Basically, the regulatory mechanisms that have become known so far can either act rapidly and reversibly (short-term regulation), or more slowly and with longer duration (long-term regulation). Table 2 lists them, and also adds the activation of the pump by partial proteolysis (James et al., 1989a; Schwab et al., 2002) and the effects on the activity of the pump by the alternative splicing processes: which, being irreversible, cannot be called sensu stricto regulatory mechanisms. Calmodulin is the classical short-term regulator. As mentioned earlier, it interacts with high affinity with a binding site in the C-terminal cytosolic tail of the pump, removing it from its anchoring sites in the main body of the enzyme, relieving the state of autoinhibition. The interaction of calmodulin with its binding site is Ca2+ dependent, and this confers to its regulatory mechanism a property that deserves a comment. The arrival of a Ca2+ burst in the cytosolic ambient of the pump will promote the binding of calmodulin and initiate the pump activation process. This will necessarily decrease local Ca2+ in the cytosol and lead to the detachment of calmodulin from the pump and to the cessation of pump activation. At this point, Ca2+ in the ambient will again increase, beginning a Table 2 Regulatory Mechanisms of the PMCA Pumps Regulation of the PMCA Pump Short-Term

Long-Term

Irreversible

Calmodulin

Acidic phospholipids

Proteasesa

PIP2

Protein kinasesb

Alternative splicing

Protein interactors

Oligomerization Protein interactors

a

Calpain, caspases. PKA, PKC. The effects of the protein interactors could be either short- or long-term.

b

272

T. Calì et al.

next round of calmodulin binding/pump activation. Quite simply, then, the regulation of the pump by calmodulin is not only short-term but also basically oscillatory (Carafoli and Krebs, 2016; Lopreiato et al., 2014). The activation of the pump by acidic phospholipids was recognized very early, in experiments in which the treatment of erythrocyte ghosts with phospholipases reduced the activity of their Ca2+ ATPase (Ronner et al., 1977): the activity was then restored to normal by PS. The effect of PS was confirmed, and extended to other acidic phospholipids, on the enzyme that had just been purified on calmodulin columns (Niggli et al., 1981). Zwitterionic phospholipids had no effect. The work on the purified pump showed that acidic phospholipids increased very significantly the affinity of the pump for Ca2+, reducing its apparent Km (Ca2+) to even lower values than calmodulin. It also showed that, at least in the erythrocyte membrane, the amount of acidic phospholipids (essentially PS) presumably surrounding the pump in the leaflet of the plasma membrane bilayer facing the cytosol was in principle sufficient for about 50% of maximal pump activity (Niggli et al., 1981). Since the activation of the pump by PS and other acidic phospholipids is clearly long-term, it must be seen as a means that provides a basic, essentially constant level of pump activity, as the amount of PS and similar acidic phospholipids in the inner leaflet of the membrane cannot be modulated by agonist-mediated mechanisms. Prima facie, this seems a reasonable and attractive possibility. However, in considering the activation of the pump by acidic phospholipids (in fact, also by the other regulatory mechanisms to be discussed later) it must be kept in mind that the activity of the pump requires by definition the presence of Ca2+, which will thus also necessarily trigger the activation by calmodulin. It follows that in the in vivo environment it becomes impossible to attribute activatory effects on the pump to regulators without simultaneously considering those of calmodulin. Somehow, these rather obvious points have not been considered in the discussions of the regulatory mechanisms of the pump. A reasonable assumption would be that even at very low Ca2+ concentrations a low basic level of pump activity is provided by calmodulin in collaboration with the acidic phospholipids of the membrane environment. Concerning the phospholipids, the interesting possibility has recently been proposed (Lopreiato et al., 2014) that the externalization of PS to the cell surface by the action of Ca2+-dependent scramblases (Bevers and Williamson, 2010) which occurs early in apoptosis (Fadok et al., 1992; Verhoven et al., 1995) could decrease pump activity, and thus exacerbate the cytosolic Ca2+ overload of the apoptotic condition. Inhibition of Ca2+ ejection has indeed been

Plasma Membrane Ca2 + Pumps

273

demonstrated in endothelial cells of the umbilical vein by treatments that induce the externalization of PS (Zhang et al., 2009). There is an exception to the rule that phospholipids in the membrane do not change in the short range time scale and that is the doubly phosphorylated product of phosphatidylinositol (PIP2), which is a particularly effective activator of the pump (Choquette et al., 1984). The level of PIP2 in the membrane changes rapidly in response to the cell stimulation by agonists which induce its hydrolysis to the products inositol 1,4,5-trisphosphate (InsP3) and diacylglycerol (DAG) which are inactive on pump activity (but see below). A reversible rapid activation/deactivation cycle of the pump by PIP2 has thus been proposed (Choquette et al., 1984). An additional interesting point on PIP2 has recently been made in a recent study (Penniston et al., 2014) which has shown that PMCAs protect PIP2 in the plasma membrane from phospholipase C. Two mechanisms have been proposed for the protection: the extrusion of Ca2+ by the PMCA pump could limit its availability to PLC, and/or the pump could bind PIP2 reducing its accessibility for PLC. In both cases less InsP₃ would be produced, and less Ca2+ would be released from the intracellular store. As will be discussed in detail later, these observations support the view that PMCA pumps play a role in the regulation of Ca2+ signaling that is not immediately related to the control of bulk cytosolic Ca2+ homeostasis. Acidic phospholipids interact with the pump at two sites (see above), one in the first cytosolic loop and the other in the C-terminal calmodulin-binding domain (Brodin et al., 1992; Zvaritch et al., 1990): the respective contribution of the two sites to the activation is not clear (Brini et al., 2010). One problem in the matter of phospholipids as activators is their spatial separation from the catalytic core of the enzyme: the physical interplay with its protruding cytosolic portion would only be possible if the latter would somehow come in contact with their polar groups on the surface of the membrane. Possibly, acidic phospholipids could act structurally, for instance by modulating the access of substrates to the catalytic center of the pump. Effects of acidic phospholipids on steps of the catalytic cycle have been described that could perhaps be explained by such structural role (Filomatori and Rega, 2003). Phospholipds, however, could also influence the activity of the pump indirectly: the pump could for instance be reversibly sequestered in membrane domains of particular phospholipid composition and/or fluidity. The fluidity of the lipid membrane ambient has indeed been shown to influence the activity of the pump (Jiang et al., 2007; Pang et al.,

274

T. Calì et al.

2005; Tang et al., 2006). It is also known that PMCA pumps associate preferentially with plasma membrane domains like caveolae and rafts that have special lipid composition (Fujimoto, 1993; Schnitzer et al., 1995; Sepulveda et al., 2006), and which also concentrate pump interactors. Two protein kinases (PKA and PKC) have been shown to modulate the activity of the PMCA pumps. PKA phosphorylates a Ser residue downstream of the C-terminal calmodulin-binding domain of isoform 1, increasing its affinity for Ca2+ (James et al., 1989b). The matter of PKC is more complex: the original finding (Smallwood et al., 1988), which was made on inside out erythrocyte ghosts which only express isoforms 1 and 4 of the pump, had shown a five- to sevenfold increase of the Vmax of the Ca2+ uptake by the ghosts, without changes of their affinity for Ca2+ (the activity of the purified pump was also stimulated by PKC, although the effect was smaller). Intriguingly, the effect of PKC was additive with that of calmodulin. Later work (Wang et al., 1991) confirmed the stimulation of the ATPase purified from erythrocytes by PKC but found that calmodulin actually antagonized the effect of the kinase. It also found that PKC phosphorylated a Thr residue within the calmodulin-binding domain, and a Ser that was C-terminal to it. The effect of PKC on the PMCA pump(s) was then explored in a number of other studies (Balasubramanyam and Gardner, 1995; Enyedi et al., 1996, 1997; Kosk-Kosicka and Bzdega, 1988; Usachev et al., 2002; Verma et al., 1999). They all confirmed the stimulation of the PMCA pump by PKC but greatly extended the analysis of its differential effects on its isoforms: even if neurons, in which isoforms 2 and 3 are supposed to predominate, were considered, most studies have concentrated on the most extensively studied isoform 4 and its splice variants. Summarizing, the most specific PKC phosphorylation site has been identified in a domain about 20 residues downstream of the calmodulin-binding domain, which has been claimed to be also involved in the pump autoinhibition process in addition to the two sites in the calmodulin domain (Falchetto et al., 1991, 1992). PKC activated the pump only partially, as it only removed the autoinhibition due to this domain, whereas calmodulin activated the pump more fully, as it removed the entire autoinhibitory domain (Enyedi et al., 1996). It was also found the PKC phosphorylated a Ser (not a Thr) within the calmodulin-binding domain of isoform 4. Whereas phosphorylation of this residue did not interfere with the binding of calmodulin, the previous binding of calmodulin to its domain prevented the phosphorylation of the Ser residue (Verma et al., 1999). Finally, another study considered isoforms 2 and 3 overexpressed in model cells, finding that the truncated “a” variants of

Plasma Membrane Ca2 + Pumps

275

both isoforms were easily phosphorylated by PKC, whereas only the full-length “2b” variant was phosphorylated by PKC: the “3b” full-length variant was not. The phosphorylation of the “a” variants inhibited the binding of calmodulin to the truncated domain and the stimulation of the activity of the pump (Enyedi et al., 1997). One problem that complicates the matter of the activation of the PMCA pump by the kinases is the Ca2+ requirement for the activation of PKA and the DAG requirement for the activation of PKC. Both the increase of Ca2+ in the environment by InsP3 and the generation of DAG demand the hydrolysis of PIP2, which (see above) is a strong activator of the pump. The activatory action of the kinases should thus in principle by counteracted by the disappearance of PIP2: which seems to be a paradox. Possibly, temporal aspects in the hydrolysis/activation processes could rationalize these opposite effects. Perhaps the most intriguing mechanism for the regulation of the PMCA pump is that based on its dimerization. Dimerization has often been suggested for a number of P-type pumps: that the PMCA pumps could exist in the membrane as dimers was indicated by radiation inactivation experiments (Cavieres, 1984; Minocherhomjee et al., 1983) on erythrocyte ghosts. These early studies showed that the estimated molecular mass of the pump was between 251 and 290 kDa, approximately corresponding to twice that of the monomeric erythrocyte enzyme. Following these early findings, a number of other experimental approaches demonstrated reversible self-association of the pump in the purified state and in phospholipid–detergent micelles (Bredeston and Rega, 1999; Coelho-Sampaio et al., 1991; Kosk-Kosicka et al., 1995). Importantly, nearly all studies showed that the self-association activated the enzyme: one study, in particular, showed that the activity of the purified pump increased as a function of its concentration (Kosk-Kosicka and Bzdega, 1988). As it increased, its low, calmodulin-dependent, activity changed to fully active, calmodulin independent. At higher pump concentrations the interaction between pump monomers replaced the pump–calmodulin interaction that was necessary for full activity at low enzyme concentrations. The mechanism of the activation process was studied in more detail on the purified enzyme by Vorherr et al. (1991): it was found that the calmodulin-binding domain mediated the dimerization (oligomerization) of the pump. Synthetic peptides corresponding to the calmodulin-binding domain stimulated the activity of the pump, suggesting the formation of heterodimers, whereas antibodies against the calmodulin-binding domain

276

T. Calì et al.

depressed the activity of the pump and inhibited its self-association. Calmodulin inhibited the dimerization, although it bound to the dimerized enzyme: the dimer bound one calmodulin molecule, which implied that one of the calmodulin-binding domains in the dimer remained free for further interactions. A later study (Levi et al., 2000) suggested that the free calmodulin domain in the dimer would bind to at least one of the two autoinhibitory domains in one of the two pump molecules (Falchetto et al., 1991, 1992). The matter of the self-association of the PMCAs in the membrane has a number of complexities, and its significance as a mechanism for the regulation of the activity of the enzyme in intact cells is questionable. Major perplexities arise from the demonstration that other P-type pumps have been conclusively demonstrated to be capable of full activity in the monomeric state, and the fact that the PMCA pump is a trace enzyme, present in the membrane in amounts far lower than, for instance, those other P-type pumps like the SERCA pump or the Na+/K+-ATPase. The likelihood of random in vivo association of monomers, which could be imagined in the case of abundant pumps, is much less easily to occur in the case of PMCA pumps. As mentioned, the partial proteolysis of the PMCA pumps, being by definition irreversible, cannot be considered a true regulatory mechanism. However, since it occurs in vivo, and may have significance to cell physiology and pathology, it could still be discusses in the context of all other regulatory mechanisms. Two proteases have been shown to attack the PMCA pumps, irreversibly influencing their activity. One is calpain, which has particularly interest because of its Ca2+ dependence, which could depict some sort of autoregulation of the Ca2+ signal. The other are caspases that have obvious significance in the important process of apoptosis. Calpain prefers substrates that contain calmodulin-binding domains and was first shown to activate the PMCA pump of erythrocyte membranes in 1987 (Au, 1987). The mechanisms of the activation process were then studied in detail on the enzyme in the membrane and in the purified state, and shown to be due to the removal of the C-terminal calmodulin-binding domain in two steps. The first cut occurred in the middle of the calmodulin-binding domain and a second closer to the N-terminus of the domain, eventually resulting in the production of a 124-kDa fragment that was maximally active, and had naturally lost the sensitivity to calmodulin (James et al., 1988). The 124-kDa truncated fragment tended to persist for a relatively long time, but in vivo, at least in erythrocytes, the degradation proceeded further, until the pump disappeared completely

Plasma Membrane Ca2 + Pumps

277

(Salamino et al., 1994): possibly, other proteases completed the degradation of the pump in vivo after the first two calpain cuts. One problem with calpain is the μM concentration of Ca2+ necessary to its action, which implies that it could only cleave massively the pump under conditions of cellular Ca2+ overload, i.e., in pathology or, locally, when activated by restricted high Ca2+ microdomains that can transitory be generated by the opening of plasma membrane Ca2+ channels, thus involving in the cleavage only a limited pool of pumps. In the former case it would possibly as a last ditch attempt of the cell to get rid of the excess of Ca2+ in the cytosol. However, the attempt would only be temporary, as in the in vivo condition the degradation of the pump would proceed past the constitutively active 124 kDa fragment, exacerbating the Ca2+ overload condition. In the latter, a very sophisticated mechanism could account for locally reducing Ca2+ concentration in confined subdomains, thus contributing to the pump signaling function. The other proteases that have been shown to act on the PMCA pumps are the caspases. They have been shown to cleave the neuron-specific PMCA2 isoform in vivo following brain ischemia and in cultured neurons undergoing apoptosis (Schwab et al., 2002), and also cleaved the ubiquitous PMCA4 isoform in nonneuronal cells induced to apoptosis (Paszty et al., 2002). The cleavage of the pumps induced Ca2+ overload and secondary necrosis: this would suggest inactivation of the cleaved pumps; however, the measurements of the activity of the pump after exposure to caspases (mostly caspase 3) have yielded conflicting results. The caspase cleavage site was identified in the C-terminal portion of the pump (Paszty et al., 2002; Schwab et al., 2002) at a DEID consensus sequence which is less than 10 residues upstream of the beginning of the calmodulin-binding domain. At variance with the 124-kDa fragment produced by calpain, the 123-kDa fragment produced by the caspase cut was found to be less active in one report that used the formation of the phosphoenzyme as the measuring method (Schwab et al., 2002), and constitutively active by another that used Ca2+ transport by microsomal vesicles (Paszty et al., 2002). The mechanism of the production of the Ca2+ overload in the cell systems induced to apoptosis, and the role of the caspase-induced cleavage of the PMCA pump are thus unclear. A recent report (Pottorf et al., 2006) on the internalization of PMCA pumps in hippocampal neurons treated with excitotoxic concentrations of glutamate may be relevant to the problem. The last entry in the discussion of the regulation mechanisms of PMCA pumps deals with their interactions with molecules involved in signaling

278

T. Calì et al.

pathways. This topic will be discussed in more detail in the next chapter, as it deals with a function of the PMCA pumps that is not related to the housekeeping role of PMCA to restore basal Ca2+ levels by exporting it in the outside medium, but to the recently recognized additional role of modulating signal transduction by locally generating Ca2+ microenvironments adequate for the functioning of other enzymes/proteins. As a rule, the interplay with the pump with partners influences the activity of the latter; however, the interactors may also affect the activity of the pump: thus, they may be mentioned here. The interactors recognize binding sites that are located along the entire length of the molecule and may be isoform specific. Most of them interact with the C-terminal region of the pump (Fig. 2), which ends with four conserved residues (ET/SXV) that match the minimal consensus site of protein ligands for Type I PDZ domains. PDZ domains are 80–90 amino acid protein–protein interaction motifs present in hundreds

Fig. 2 Topology model of the PMCA pump with its interacting protein partners. PMCA-binding proteins are shown in proximity of their interaction sites on the PMCA. The two main C-terminal splice variants “a” and “b” are shown. The PDZ domain-containing proteins only bind to the full-length “b” variants of the PMCA.

Plasma Membrane Ca2 + Pumps

279

of signaling proteins that recognize the C-terminus of partner proteins to promote their interplay with signaling pathways. A number of them indeed interact with the C-terminus of the PMCA pumps, among them several members of the MAGUK family of guanylate kinases (DeMarco and Strehler, 2001); a regulatory factor for the Na+/H+-exchanger (NHERF1 and 2, PMCA2 preferring NHERF2 over NHERF1) (DeMarco et al., 2002); the PMCA-interacting single PDZ protein (Goellner et al., 2003), which may play a role in the sorting of PMCAs to and from the plasma membrane; the calmodulin-dependent serine kinase CASK, which is a coactivator of promoters containing T-element-dependent reporter activity (Schuh et al., 2003); NOS 1, which forms a ternary complex with syntrophin and the PMCA pump (Kim et al., 1998); the scaffold protein Homers: Homer1 accelerates the Ca2+ clearance from hippocampal neurons, and thus somehow influences the activity of the PMCA pump (Salm and Thayer, 2012). In parotid gland acinar cells, Homer 2 interacts instead particularly with isoform 4 at a N-terminal PPXXF motif, increasing the Ca2+ ejection activity of the pump (Yang et al., 2014). Other proteins recognize binding sites away from the C-terminus of the pump: calcineurin binds to the main intracellular loop close to the binding site for ATP (Armesilla et al., 2004) as does the Ras-associated factor 1 (RASSF1) (Kim et al., 1998; Williams et al., 2006); the 14-3-3 protein interacts instead with the N-terminal portion of PMCA4, but not of PMCA2, and has inhibitory function (Rimessi et al., 2005). Finally, the actin cytoskeleton has also been shown to interact with the PMCA pump (Vanagas et al., 2007). Interestingly, the interaction modulates the activity of the pump, the state of polymerization of actin determining whether it acts as an activator or as an inhibitor (Vanagas et al., 2013): G-actin and short oligomers activate the pump, and F-actin inhibits it. Although the pump appears to have only one actin-binding site, more than one molecule of G-actin is required for its effect on the activity of the pump. The site of actin binding has not been identified; however, it is not the calmodulin-binding domain (Dalghi et al., 2013).

5. ROLE OF PMCA PUMP IN REGULATING CELL Ca2+ P-type ATPases use ATP to transport ions across membrane boundaries: Ca2+ in the case of the PMCA pump. The pump was discovered in the membrane of erythrocytes, where its role was the regulation of the homeostasis of Ca2+ by ejecting it from the erythrocyte cytosol. As the early

280

T. Calì et al.

erythrocyte studies were extended to other cell types, the export of Ca2+ became accepted as the general role of the PMCA pumps. Naturally the export of Ca2+ from cells is the mechanistic role of the PMCA pumps; however, while in erythrocytes the PMCA pump is the only system that controls the homeostasis of Ca2+, all other eukaryotic cells contain additional membrane transporters that also do that. All cells contain two other Ca2+ pumps that clear Ca2+ from the cytosol, the SERCA pump in the endo(sarco)plasmic reticulum and the SPCA pump in the Golgi system: the Golgi pump is probably of quantitatively minor importance, but the SERCA pump is far more abundant that the PMCA pump in all cells, and is thus quantitatively far more important in the bulk control of cytosolic Ca2+. The Na+/Ca2+ -exchanger of the plasma membrane also exports Ca2+ from the cytosol and does so with high transport capacity. However, it is probably not present in equally significant amounts in all cell types, its role being particularly important in excitable cells, e.g., cardiac myocytes and neurons. In discussing the global regulation of Ca2+ in the cytosol, two aspects of the process must thus be considered: its modulation within the cell without direct exchanges with the external medium, and the modulation linked to the direct transport of Ca2+ across the plasma membrane. There is little doubt that the bulk internal regulation of Ca2+ relies essentially on the systems that move Ca2+ back and forth between the cytosol and the lumen of the endo(sarco)plasmic reticulum (which could be set in motion by trigger Ca2+ admitted into the cytosol by plasma membrane channels): they are the SERCA pump and two receptor channels, the InsP3 channel and the ryanodine receptor. In nonexcitable cells, in which the plasma membrane Na+/Ca2+-exchanger is normally of limited importance, the regulation of cytosolic Ca2+ by its direct export through the plasma membrane relies on the action of the PMCA pumps. In excitable cells, in which the Ca2+ exchanges between the cytosol and the extracellular space are quantitatively very important, the situation is more complex, due to the action of the very powerful plasma membrane Na+/Ca2+-exchanger. The role of the PMCA pump in the bulk regulation of Ca2+ of excitable cells would thus seem minor: the discovery of a PMCA pump in the sarcolemma of heart myocytes (Caroni and Carafoli, 1981) was indeed a surprise and became accepted only with some difficulty. The situation is perhaps less clear-cut in neurons, which are the excitable cells par excellence: the importance of the PMCAs in the export of Ca2+ varies with the neuronal type, with the PMCA pump isoform, and even with the plasma membrane domain in which the pump operates. In some neurons, e.g., those of the dorsal root ganglion (Benham

Plasma Membrane Ca2 + Pumps

281

et al., 1992; Usachev et al., 2002; Werth et al., 1996) or in the stereocilia of the outer hair cells of the Corti organ of the inner ear (Yamoah et al., 1998), the PMCA pumps have a substantial role in the total ejection of Ca2+. Necessarily, however, their action modulates Ca2+ primarily in the subplasma membrane region of the cell. Therefore, if one would only consider the global regulation of Ca2+ in the entire cytosol the PMCA pumps could even appear as physiological paradoxes: indeed, their Ca2+ transport has for instance been shown to be irrelevant to the beat-to-beat regulation of Ca2+ in the contraction–relaxation cycle of heart myocytes (Mohamed et al., 2011). The PMCA pumps are nevertheless essential to the well-being of all cells, as inescapably certified by the disease phenotypes generated by their malfunction. But also, in a more general way, by the great number and complexity of their regulatory mechanisms, including, and especially, the interplay with the numerous important protein partners described earlier. Thus, the concept has gradually emerged that the main role of the PMCA pumps would be the local, as opposed to the global, regulation of cell Ca2+ (Cartwright et al., 2007; Lopreiato et al., 2014). Among the experimental findings that have led to this new way to look at the “real” role of the PMCA pumps, the association of the pump with plasma membrane districts (see above) like the caveolae (Fujimoto, 1993) is particularly significant. These structures of virtually ubiquitous tissue distribution have particular lipid composition (which could per se have a role in the modulation of the activity of the pump): but the most important point is that they recruit through their scaffolding proteins caveolins signaling molecules, facilitating their interplay with the pumps (Cohen et al., 2004). The ability of the C-terminus of the PMCA pumps to recognize the PDZ domains present in many signaling proteins (see above) adds significant weight to the finding. The fact that so many interactors of the PMCA pumps are components of signaling pathways indeed strongly supports the proposal that the principal role of the PMCA pumps is to regulate the concentration of Ca2+ in selected microdomains in the subplasma membrane region of the cell to the benefit of locally recruited Ca2+-dependent enzyme activities of general cell function significance. A striking example of such signaling function of the PMCA pump is the regulation of the contractility of heart myocytes by one of its isoforms (PMCA4) (Mohamed et al., 2011). PMCA4 forms a complex with the Ca2+-dependent nNOS, regulating the production of NO, which is a key regulator of heart contractility (Barouch et al., 2002; Sears et al., 2003). In addition to regulating the activity of nNOS, the pump also tethers it to the microdomain, where the activity of the pump regulates Ca2+. The

282

T. Calì et al.

study has shown that the knocking out of PMCA4 alters both the production of NO by nNOS and the tethering of the enzyme to the selected plasma membrane domain in PMCA4-mice, altering the contractility of heart without effects on the beat-to-beat concentration of diastolic Ca2+.

6. PMCA PUMPS AND PATHOLOGY The PMCA pumps are essential to the proper functioning of all animal cells, however, the contribution of the SERCA Ca2+ pump (and of the Na+/Ca2+-exchanger of the plasma membrane) clearly overwhelm that of the PMCA pump when large cytosolic Ca2+ transients must be controlled and shaped. As discussed in detail earlier, the main role of the pump is thus not the global regulation of cytosolic Ca2+ but the fine tuning of Ca2+ variations in selected subplasma membrane microdomains at the precise threshold required to control the activity of Ca2+-dependent pump interactors that are located in these domains. This novel way to look at the function of the PMCA pumps is relevant to the last topic to be discussed in this contribution, i.e., the consequences of the malfunction of the PMCA pumps for the life of the cell. As will be discussed in the next paragraphs, the malfunctions of PMCA pumps that will be described later have altogether minor consequences on the global homeostasis of Ca2+ in the cell but may be of great importance in the Ca2+ balance in the selected subplasma membrane microdomains that are under the sole control of the PMCA pumps. The Ca2+ signal has one important distinctive property and that is the ambivalence. To perform its signaling function, which essential to cells, Ca2+ must be kept under strict control. If this control fails, cellular discomfort follows. The topic of Ca2+ signaling and disease, which was essentially a curiosity a few years ago, has now rapidly grown to become a significant part of the Ca2+ signaling area (Carafoli and Brini, 2007; Krebs and Michalak, 2007). Numerous disease phenotypes, the best studied and most interesting being of genetic origin, are now being related to alterations in the cellular homeostasis of Ca2+. Massive deviations of Ca2+ from the sub-μM concentrations such as those generated by plasma membrane lesions rapidly terminate cell life. However, subtler deviations, most of them of genetic origin, while affecting cell function are still compatible with the cell life. They may be caused by defects in the Ca2+ transporting systems: among them, those involving the PMCA are gradually becoming more numerous. One important point on this matter must be mentioned. Ca2+ in cells is maintained at a

Plasma Membrane Ca2 + Pumps

283

given set point: the pathology-linked deviations from this set point could be in either direction. Ca2+ overload and Ca2+ deprivation are equally dangerous to cell life. The point is important, as the defects of the PMCA pumps that will be discussed later should in principle lead to the increase, as opposed to the decrease, of Ca2+ in the subplasma membrane microdomains where important Ca2+-regulated pump interactors reside. The disease phenotypes linked to the malfunctions of the PMCA pumps affect predominantly the nervous system where isoforms 2 and 3 prevail. From this point of view, the disease phenotypes would thus be due to the inability to keep the Ca2+ levels of selected microdomains of the plasma membrane precisely at the threshold required to control the activity of important Ca2+-dependent interactors, subtle changes above or below the threshold would inevitably lead to their malfunctioning. However, the ubiquitous (1 and 4) isoforms have also been directly or indirectly linked to disease phenotypes: a genome-wide association study has connected a defect of the PMCA1 gene to susceptibility to hypertension (Cho et al., 2009; Tabara et al., 2010; Xi et al., 2014) and coronary artery disease (Lu et al., 2012), suggesting a major role of isoform 1 in the control of Ca2+ handling in smooth muscles and in the regulation of blood pressure. Recently direct evidence has emerged on the involvement of a PMCA4 pump defect in a human disease: a missense mutation (R268Q) cosegregated in a family with autosomal dominant familial spastic paraplegia (Li et al., 2014). The mutation significantly decreased the efficiency of the suppression of depolarization-induced calcium overload (Ho et al., 2015). The finding that the PMCA4 pump is enriched in lipid rafts in cerebellum (Sepulveda et al., 2006) may be relevant to the possibility described earlier that the mutation also affects the interaction with specific partners. However, the pathologies involving the tissue-restricted isoforms PMCA2 and 3 are more frequent and have been studied in more detail. The discussion to follow will focus on the deafness phenotype associated with PMCA2 mutations and the ataxia phenotypes related to PMCA3 mutations. As mentioned, the PMCA2 pump is restricted to the central nervous system and its related tissues (cochlear outer hair cells, Purkinje neurons, and cerebellar granules). Small amounts of it are also present in liver, kidney, uterus, retina, and the mammary gland epithelium (Silverstein and Tempel, 2006). At variance with the other isoforms, the PMCA2 pump is peculiarly active also in the absence of calmodulin (Elwess et al., 1997; Hilfiker et al., 1994). The stereocilia of the outer hair of the Corti organ in the inner ear must control the Ca2+ concentration in the endolymph

284

T. Calì et al.

by the doubly spliced w/a variant of the PMCA2 pump which is responsible for the clearance of Ca2+ from their cytoplasm (Dumont et al., 2001). The opening of the mechanoelectric transduction channels in the stereocilia induced by their bending in response to sound waves induces K+ and Ca2+ entry into the stereocilia to initiate sensory transduction: molecularly the opening of the channels is promoted by the binding of Ca2+ to cadherin 23. The PMCA2 pump then exports Ca2+ back to the endolymph. Mutations in cadherin 23 are associated with nonsyndromic sensorineural hearing loss (Bork et al., 2001); they were found in five affected offspring of a consanguineous family with autosomal recessive, nonsyndromic sensorineural hearing loss (Schultz et al., 2005). Interestingly, three of these subjects, all of whom had severe-to-profound hearing loss affecting all frequencies, were also heterozygous for a point mutation in the PMCA2 gene predicted to result in the substitution of methionine for valine at amino acid position 586. This was the first identified human PMCA2 mutation and led to the proposal that PMCA2 mutations could act as a dominant modifier allele that exacerbated sensorineural hearing loss (Schultz et al., 2005). Slowly thereafter, a second human case of deafness phenotype was reported which was caused by the simultaneous presence of a PMCA2 mutation (a glycine to serine substitution in position 293) and a mutation in cadherin 23 (a threonine to serine substitution in position 1999) (Ficarella et al., 2007). The single mutations, respectively, carried by the healthy mother and the healthy father, were not sufficient to induce the hearing loss phenotype by themselves, suggesting that a digenic mechanism was operating in the affected offspring (Ficarella et al., 2007). The molecular rationale proposed for the hearing loss phenotype was that the PMCA2 mutation would significantly compromise the dissipation of the stereociliar Ca2+ transients induced by the opening of the mechanoelectrical transduction channels, leading to the inability to react to subsequent acoustic stimuli (Ficarella et al., 2007). The effect of the pump mutation was analyzed by overexpressing the mutated pump in model cells. A delay of the return of the Ca2+ transients to baseline after cell stimulation was detected, showing a defect in the removal of Ca2+ from the cytosol (Brini et al., 2013b). PMCA2 pump defects have now been linked to deafness phenotypes in several diseased mice, while in human the pump defect was associated to a simultaneous cadherin 23 mutation, the PMCA2 pump defect, when particularly severe, was found to be per se responsible for the deafness phenotype (Bortolozzi et al., 2010; Ficarella et al., 2007).

Plasma Membrane Ca2 + Pumps

285

Evidence for a pivotal role of the PMCA pumps as important regulators of multisignaling complexes in selected subplasma membrane microdomains has also recently emerged for isoform 2 (Jeong et al., 2016). As mentioned earlier, the PMCA2 pump is also expressed in the lactating mammary gland, where it transports calcium to the milk. Interestingly, its expression is activated in breast cancers and correlates with the levels of human epidermal growth factor receptor 2 (HER2) in them. The PMCA2 pump recruits HER2 to specific actin-rich membrane signaling domains in which intracellular calcium must be kept low to regulate its resistance to endocytosis and allowing continued HER2 signaling. The PMCA2 is thus vital for the localization of HER2 and its partners, EGFR and HER3, to active membrane signaling domains (Jeong et al., 2016). The PMCA3 is the least well characterized of the PMCA isoforms. In the brain, high levels of this isoform are found on the presynaptic side of the axon terminals of cerebellar granule cells (Strehler and Zacharias, 2001), the main splicing variants found there being the a and the b (Stauffer et al., 1997). Genetic defects of the PMCA3 pump associated with human neuronal pathology have appeared only recently, when, by using X-exome sequencing, a missense mutation (G1107D) was identified in a family with X-linked congenital cerebellar ataxia. The mutation was located in the calmodulin-binding domain of the pump (Zanni et al., 2012). A second human mutation (an arginine to histidine substitution (R482H)) was then found in a patient with developmental delay, hypotonia, and cerebellar ataxia (Cali et al., 2015) in analogy with what observed for the PMCA2 mutations, the Ca2+ extrusion ability of the mutated pumps was found to be decreased. Interestingly, in the second case, the patient also carried compound heterozygous mutations in the LAMA1 gene encoding laminin subunit 1α (a maternally inherited T2025M substitution and a paternally inherited R2381C substitution) that appeared to be necessary for the development of the disease. It is of interest that homozygous or compound heterozygous mutations, or deletions which induced protein truncation of the LAMA1 gene are associated with a phenotype of cerebellar dysplasia with cysts (with and without retinal dystrophy) (Aldinger et al., 2014). This finding indicates that the isoform 3 mutations, in analogy with PMCA2 pump mutations, could also play a role as a digenic modulator in Ca2+-linked pathologies. Human PMCA3 mutation have also been reported in nonneuronal disease phenotypes, e.g., a missense T/M substitution between the catalytic Asp and the ATP-binding

286

T. Calì et al.

site has been linked to pancreatic cancer cells (Jones et al., 2008) and an in-frame deletion of two amino acids at positions 425 and 426 in the M4 transmembrane helix has been described in aldosterone-producing adenomas (Beuschlein et al., 2013). No experimental evidence for defects in the Ca2+ extrusion activity of the pump has been provided in these cases; however, considering the type of amino acids deletions, it is likely that in both cases the mutations would have affected the function of the pump. One last point in the ataxic phenotype concerns the PMCA2 pump. The most prominent effect of the mutations of this isoform is the deafness phenotype discussed earlier; however, the PMCA2 mutant mice also have strong motility problems, ranging from unsteady gait, to difficulties in standing up, to defects in walking and maintaining balance (Kozel et al., 1998; Sun et al., 2008). This suggests a functional role of isoform 2 in the cerebellum, also considering its abundant expression in the dendrites and dendritic spines of Purkinje cells (Stahl et al., 1992; Stauffer et al., 1997; Tolosa de Talamoni et al., 1993) and its importance for the development of the dendritic tree and the synaptic connections of the parallel excitatory fibers of Purkinje neurons (Inoue et al., 1993).

7. CONCLUDING REMARKS In the 50 years that have elapsed since its discovery (Schatzmann, 1966), a very considerable amount of information has been accumulated on the PMCA pump. Its 3D structure has not yet been solved, but its essential structural details have been safely predicted from those of the SERCA pump (Fig. 3). However, the PMCA pump also has one unique feature that distinguishes it from all other P-type pumps: a long C-terminal tail that is functionally essential to its operation, and which is also the seat of most of its regulatory mechanisms. This contribution has offered an updated description of the general features of the PMCA pump(s) but has dealt in comparatively greater detail with the functional aspects that are directly or indirectly related with the operation of the C-terminal tail. In doing so, novel ways to look at the operation of the pumps, and at their role in the cell have emerged: on the way in which calmodulin affects the activity of the pumps, on their mutual interplay of the different regulators, chiefly calmodulin and acidic phospholipids, and, especially and more generally, on a shift of the function of the pumps from the global homeostasis of Ca2+ in the cell to the selected regulation of cell signaling in specific subplasma membrane microdomains. Finally, the ambivalence of the Ca2+ signal, and the role of the malfunction of the PMCA pumps in generating

Plasma Membrane Ca2 + Pumps

287

Fig. 3 3D ribbon structure of the PMCA pump (pink) in its Ca2+-free and Ca2+-bound state, modeled on that of the SERCA pump (PDB files 1IWO and 1SU4). The long cytosolic C-terminal tail of the PMCA pump (which is absent in the SERCA pump) is not included in the superimposition. The main domains of the pump are colored as in Fig. 1.

phenotypes of cell discomfort has been discussed in some detail. One important point that has emerged from the discussion of the ambivalence of the Ca2+ signal is that the concentration of cell Ca2+ must be maintained at a precisely defined set point: deviations from it in either direction, i.e., toward Ca2+ overload or Ca2+ deprivation, are incompatible with the correct functioning of cell life.

ACKNOWLEDGMENTS The original work by the authors is supported by Grants from the University of Padova (Progetto Giovani 2012 n° GRIC128SP0 and Progetto di Ateneo 2016 n° CALI_SID16_01) and the Ministry of University and Research (Bando SIR 2014 no RBSI14C65Z) to T.C. and from the University of Padova (Progetto di Ateneo 2015 no CPDA153402) to M.B.

REFERENCES Aldinger, K.A., Mosca, S.J., Tetreault, M., Dempsey, J.C., Ishak, G.E., Hartley, T., Phelps, I.G., Lamont, R.E., O’Day, D.R., Basel, D., Gripp, K.W., Baker, L., Stephan, M.J., Bernier, F.P., Boycott, K.M., Majewski, J., University of Washington Center for Mendelian, G., Care4Rare, C., Parboosingh, J.S., Innes, A.M.,

288

T. Calì et al.

Doherty, D., 2014. Mutations in LAMA1 cause cerebellar dysplasia and cysts with and without retinal dystrophy. Am. J. Hum. Genet. 95, 227–234. Armesilla, A.L., Williams, J.C., Buch, M.H., Pickard, A., Emerson, M., Cartwright, E.J., Oceandy, D., Vos, M.D., Gillies, S., Clark, G.J., Neyses, L., 2004. Novel functional interaction between the plasma membrane Ca2+ pump 4b and the proapoptotic tumor suppressor Ras-associated factor 1 (RASSF1). J. Biol. Chem. 279, 31318–31328. Au, K.S., 1987. Activation of erythrocyte membrane Ca2+-ATPase by calpain. Biochim. Biophys. Acta 905, 273–278. Axelsen, K.B., Palmgren, M.G., 1998. Evolution of substrate specificities in the P-type ATPase superfamily. J. Mol. Evol. 46, 84–101. Balasubramanyam, M., Gardner, J.P., 1995. Protein kinase C modulates cytosolic free calcium by stimulating calcium pump activity in Jurkat T cells. Cell Calcium 18, 526–541. Barouch, L.A., Harrison, R.W., Skaf, M.W., Rosas, G.O., Cappola, T.P., Kobeissi, Z.A., Hobai, I.A., Lemmon, C.A., Burnett, A.L., O’Rourke, B., Rodriguez, E.R., Huang, P.L., Lima, J.A., Berkowitz, D.E., Hare, J.M., 2002. Nitric oxide regulates the heart by spatial confinement of nitric oxide synthase isoforms. Nature 416, 337–339. Benham, C.D., Evans, M.L., McBain, C.J., 1992. Ca2+ efflux mechanisms following depolarization evoked calcium transients in cultured rat sensory neurones. J. Physiol. 455, 567–583. Beuschlein, F., Boulkroun, S., Osswald, A., Wieland, T., Nielsen, H.N., Lichtenauer, U.D., Penton, D., Schack, V.R., Amar, L., Fischer, E., Walther, A., Tauber, P., Schwarzmayr, T., Diener, S., Graf, E., Allolio, B., Samson-Couterie, B., Benecke, A., Quinkler, M., Fallo, F., Plouin, P.F., Mantero, F., Meitinger, T., Mulatero, P., Jeunemaitre, X., Warth, R., Vilsen, B., Zennaro, M.C., Strom, T.M., Reincke, M., 2013. Somatic mutations in ATP1A1 and ATP2B3 lead to aldosterone-producing adenomas and secondary hypertension. Nat. Genet. 45, 440–444. 444e441-442. Bevers, E.M., Williamson, P.L., 2010. Phospholipid scramblase: an update. FEBS Lett. 584, 2724–2730. Bonza, M.C., Luoni, L., 2010. Plant and animal type 2B Ca2+-ATPases: evidence for a common auto-inhibitory mechanism. FEBS Lett. 584, 4783–4788. Bork, J.M., Peters, L.M., Riazuddin, S., Bernstein, S.L., Ahmed, Z.M., Ness, S.L., Polomeno, R., Ramesh, A., Schloss, M., Srisailpathy, C.R., Wayne, S., Bellman, S., Desmukh, D., Ahmed, Z., Khan, S.N., Kaloustian, V.M., Li, X.C., Lalwani, A., Riazuddin, S., Bitner-Glindzicz, M., Nance, W.E., Liu, X.Z., Wistow, G., Smith, R.J., Griffith, A.J., Wilcox, E.R., Friedman, T.B., Morell, R.J., 2001. Usher syndrome 1D and nonsyndromic autosomal recessive deafness DFNB12 are caused by allelic mutations of the novel cadherin-like gene CDH23. Am. J. Hum. Genet. 68, 26–37. Bortolozzi, M., Brini, M., Parkinson, N., Crispino, G., Scimemi, P., De Siati, R.D., Di Leva, F., Parker, A., Ortolano, S., Arslan, E., Brown, S.D., Carafoli, E., Mammano, F., 2010. The novel PMCA2 pump mutation Tommy impairs cytosolic calcium clearance in hair cells and links to deafness in mice. J. Biol. Chem. 285, 37693–37703. Brandt, P., Ibrahim, E., Bruns, G.A., Neve, R.L., 1992. Determination of the nucleotide sequence and chromosomal localization of the ATP2B2 gene encoding human Ca(2 +)-pumping ATPase isoform PMCA2. Genomics 14, 484–487. Bredeston, L.M., Adamo, H.P., 2004. Loss of autoinhibition of the plasma membrane Ca(2 +) pump by substitution of aspartic 170 by asparagin. Activation of plasma membrane calcium ATPase 4 without disruption of the interaction between the catalytic core and the C-terminal regulatory domain. J. Biol. Chem. 279, 41619–41625. Bredeston, L.M., Rega, A.F., 1999. Phosphatidylcholine makes specific activity of the purified Ca(2 +)-ATPase from plasma membranes independent of enzyme concentration. Biochim. Biophys. Acta 1420, 57–62.

Plasma Membrane Ca2 + Pumps

289

Brini, M., Carafoli, E., 2009. Calcium pumps in health and disease. Physiol. Rev. 89, 1341–1378. Brini, M., Coletto, L., Pierobon, N., Kraev, N., Guerini, D., Carafoli, E., 2003. A comparative functional analysis of plasma membrane Ca2 + pump isoforms in intact cells. J. Biol. Chem. 278, 24500–24508. Brini, M., Di Leva, F., Ortega, C.K., Domi, T., Ottolini, D., Leonardi, E., Tosatto, S.C., Carafoli, E., 2010. Deletions and mutations in the acidic lipid-binding region of the plasma membrane Ca2 + pump: a study on different splicing variants of isoform 2. J. Biol. Chem. 285, 30779–30791. Brini, M., Calı`, T., Ottolini, D., Carafoli, E., 2013a. Intracellular calcium homeostasis and signaling. Met. Ions Life Sci. 12, 119–168. Brini, M., Calı`, T., Ottolini, D., Carafoli, E., 2013b. The plasma membrane calcium pump in health and disease. FEBS J. 280, 5385–5397. Brini, M., Ottolini, D., Cali, T., Carafoli, E., 2013c. Calcium in health and disease. Met. Ions Life Sci. 13, 81–137. Brodin, P., Falchetto, R., Vorherr, T., Carafoli, E., 1992. Identification of two domains which mediate the binding of activating phospholipids to the plasma-membrane Ca2 + pump. Eur. J. Biochem. 204, 939–946. Cali, T., Lopreiato, R., Shimony, J., Vineyard, M., Frizzarin, M., Zanni, G., Zanotti, G., Brini, M., Shinawi, M., Carafoli, E., 2015. A novel mutation in isoform 3 of the plasma membrane Ca2+ pump impairs cellular Ca2+ homeostasis in a patient with cerebellar ataxia and laminin subunit 1alpha mutations. J. Biol. Chem. 290, 16132–16141. Carafoli, E., Brini, M., 2007. Calcium Signalling and Disease: Molecular Pathology of Calcium. Springer, New York, NY. Carafoli, E., Krebs, J., 2016. Why calcium? How calcium became the best communicator. J. Biol. Chem. 291 (40), 20849–20857. Carafoli, E., Santella, L., Branca, D., Brini, M., 2001. Generation, control, and processing of cellular calcium signals. Crit. Rev. Biochem. Mol. Biol. 36, 107–260. Caroni, P., Carafoli, E., 1981. The Ca2 + -pumping ATPase of heart sarcolemma. Characterization, calmodulin dependence, and partial purification. J. Biol. Chem. 256, 3263–3270. Cartwright, E.J., Oceandy, D., Neyses, L., 2007. Plasma membrane calcium ATPase and its relationship to nitric oxide signaling in the heart. Ann. N. Y. Acad. Sci. 1099, 247–253. Cavieres, J.D., 1984. Calmodulin and the target size of the (Ca2+ + Mg2 +)-ATPase of human red-cell ghosts. Biochim. Biophys. Acta 771, 241–244. Chicka, M.C., Strehler, E.E., 2003. Alternative splicing of the first intracellular loop of plasma membrane Ca2+-ATPase isoform 2 alters its membrane targeting. J. Biol. Chem. 278, 18464–18470. Cho, Y.S., Go, M.J., Kim, Y.J., Heo, J.Y., Oh, J.H., Ban, H.J., Yoon, D., Lee, M.H., Kim, D.J., Park, M., Cha, S.H., Kim, J.W., Han, B.G., Min, H., Ahn, Y., Park, M.S., Han, H.R., Jang, H.Y., Cho, E.Y., Lee, J.E., Cho, N.H., Shin, C., Park, T., Park, J.W., Lee, J.K., Cardon, L., Clarke, G., McCarthy, M.I., Lee, J.Y., Oh, B., Kim, H.L., 2009. A large-scale genome-wide association study of Asian populations uncovers genetic factors influencing eight quantitative traits. Nat. Genet. 41, 527–534. Choquette, D., Hakim, G., Filoteo, A.G., Plishker, G.A., Bostwick, J.R., Penniston, J.T., 1984. Regulation of plasma membrane Ca2+ ATPases by lipids of the phosphatidylinositol cycle. Biochem. Biophys. Res. Commun. 125, 908–915. Coelho-Sampaio, T., Ferreira, S.T., Benaim, G., Vieyra, A., 1991. Dissociation of purified erythrocyte Ca(2 +)-ATPase by hydrostatic pressure. J. Biol. Chem. 266, 22266–22272. Cohen, A.W., Hnasko, R., Schubert, W., Lisanti, M.P., 2004. Role of caveolae and caveolins in health and disease. Physiol. Rev. 84, 1341–1379.

290

T. Calì et al.

Dalghi, M.G., Fernandez, M.M., Ferreira-Gomes, M., Mangialavori, I.C., Malchiodi, E.L., Strehler, E.E., Rossi, J.P., 2013. Plasma membrane calcium ATPase activity is regulated by actin oligomers through direct interaction. J. Biol. Chem. 288, 23380–23393. De Jaegere, S., Wuytack, F., De Smedt, H., Van den Bosch, L., Casteels, R., 1993. Alternative processing of the gene transcripts encoding a plasma-membrane and a sarco/endoplasmic reticulum Ca2+ pump during differentiation of BC3H1 muscle cells. Biochim. Biophys. Acta 1173, 188–194. DeMarco, S.J., Strehler, E.E., 2001. Plasma membrane Ca2 + -atpase isoforms 2b and 4b interact promiscuously and selectively with members of the membrane-associated guanylate kinase family of PDZ (PSD95/Dlg/ZO-1) domain-containing proteins. J. Biol. Chem. 276, 21594–21600. DeMarco, S.J., Chicka, M.C., Strehler, E.E., 2002. Plasma membrane Ca2 + ATPase isoform 2b interacts preferentially with Na +/H + exchanger regulatory factor 2 in apical plasma membranes. J. Biol. Chem. 277, 10506–10511. Dumont, R.A., Lins, U., Filoteo, A.G., Penniston, J.T., Kachar, B., Gillespie, P.G., 2001. Plasma membrane Ca2+-ATPase isoform 2a is the PMCA of hair bundles. J. Neurosci. 21, 5066–5078. Ekberg, K., Palmgren, M.G., Veierskov, B., Buch-Pedersen, M.J., 2010. A novel mechanism of P-type ATPase autoinhibition involving both termini of the protein. J. Biol. Chem. 285, 7344–7350. Elwess, N.L., Filoteo, A.G., Enyedi, A., Penniston, J.T., 1997. Plasma membrane Ca2 + pump isoforms 2a and 2b are unusually responsive to calmodulin and Ca2 +. J. Biol. Chem. 272, 17981–17986. Enyedi, A., Verma, A.K., Filoteo, A.G., Penniston, J.T., 1996. Protein kinase C activates the plasma membrane Ca2+ pump isoform 4b by phosphorylation of an inhibitory region downstream of the calmodulin-binding domain. J. Biol. Chem. 271, 32461–32467. Enyedi, A., Elwess, N.L., Filoteo, A.G., Verma, A.K., Paszty, K., Penniston, J.T., 1997. Protein kinase C phosphorylates the "a" forms of plasma membrane Ca2 + pump isoforms 2 and 3 and prevents binding of calmodulin. J. Biol. Chem. 272, 27525–27528. Fadok, V.A., Voelker, D.R., Campbell, P.A., Cohen, J.J., Bratton, D.L., Henson, P.M., 1992. Exposure of phosphatidylserine on the surface of apoptotic lymphocytes triggers specific recognition and removal by macrophages. J. Immunol. 148, 2207–2216. Falchetto, R., Vorherr, T., Brunner, J., Carafoli, E., 1991. The plasma membrane Ca2 + pump contains a site that interacts with its calmodulin-binding domain. J. Biol. Chem. 266, 2930–2936. Falchetto, R., Vorherr, T., Carafoli, E., 1992. The calmodulin-binding site of the plasma membrane Ca2 + pump interacts with the transduction domain of the enzyme. Protein Sci. 1, 1613–1621. Ficarella, R., Di Leva, F., Bortolozzi, M., Ortolano, S., Donaudy, F., Petrillo, M., Melchionda, S., Lelli, A., Domi, T., Fedrizzi, L., Lim, D., Shull, G.E., Gasparini, P., Brini, M., Mammano, F., Carafoli, E., 2007. A functional study of plasma-membrane calcium-pump isoform 2 mutants causing digenic deafness. Proc. Natl. Acad. Sci. U.S.A. 104, 1516–1521. Filomatori, C.V., Rega, A.F., 2003. On the mechanism of activation of the plasma membrane Ca2+-ATPase by ATP and acidic phospholipids. J. Biol. Chem. 278, 22265–22271. Fujimoto, T., 1993. Calcium pump of the plasma membrane is localized in caveolae. J. Cell Biol. 120, 1147–1157. Goellner, G.M., DeMarco, S.J., Strehler, E.E., 2003. Characterization of PISP, a novel single-PDZ protein that binds to all plasma membrane Ca2+-ATPase b-splice variants. Ann. N. Y. Acad. Sci. 986, 461–471.

Plasma Membrane Ca2 + Pumps

291

Gopinath, R.M., Vincenzi, F.F., 1979. (Ca2++Mg2 +)-ATPase activity of sickle cell membranes: decreased activation by red blood cell cytoplasmic activator. Am. J. Hematol. 7, 303–312. Guerini, D., Carafoli, E., 1999. The calcium pumps. In: Carafoli, E., Klee, C. (Eds.), In: Calcium as a Cellular Regulators, vol. 10. Oxford University Press, New York, pp. 249–278. Guerini, D., Garcia-Martin, E., Gerber, A., Volbracht, C., Leist, M., Merino, C.G., Carafoli, E., 1999. The expression of plasma membrane Ca2 + pump isoforms in cerebellar granule neurons is modulated by Ca2+. J. Biol. Chem. 274, 1667–1676. Guerini, D., Wang, X., Li, L., Genazzani, A., Carafoli, E., 2000. Calcineurin controls the expression of isoform 4CII of the plasma membrane Ca(2 +) pump in neurons. J. Biol. Chem. 275, 3706–3712. Hammes, A., Oberdorf, S., Strehler, E.E., Stauffer, T., Carafoli, E., Vetter, H., Neyses, L., 1994. Differentiation-specific isoform mRNA expression of the calmodulin-dependent plasma membrane Ca(2 +)-ATPase. FASEB J. 8, 428–435. Hilfiker, H., Guerini, D., Carafoli, E., 1994. Cloning and expression of isoform 2 of the human plasma membrane Ca2+ ATPase. Functional properties of the enzyme and its splicing products. J. Biol. Chem. 269, 26178–26183. Ho, P.W., Pang, S.Y., Li, M., Tse, Z.H., Kung, M.H., Sham, P.C., Ho, S.L., 2015. PMCA4 (ATP2B4) mutation in familial spastic paraplegia causes delay in intracellular calcium extrusion. Brain Behav. 5, e00321. Inoue, Y., Matsumura, Y., Inoue, K., Ichikawa, R., Takayama, C., 1993. Abnormal synaptic architecture in the cerebellar cortex of a new dystonic mutant mouse, Wriggle Mouse Sagami. Neurosci. Res. 16, 39–48. James, P., Maeda, M., Fischer, R., Verma, A.K., Krebs, J., Penniston, J.T., Carafoli, E., 1988. Identification and primary structure of a calmodulin binding domain of the Ca2 + pump of human erythrocytes. J. Biol. Chem. 263, 2905–2910. James, P., Vorherr, T., Krebs, J., Morelli, A., Castello, G., McCormick, D.J., Penniston, J.T., De Flora, A., Carafoli, E., 1989a. Modulation of erythrocyte Ca2 + -ATPase by selective calpain cleavage of the calmodulin-binding domain. J. Biol. Chem. 264, 8289–8296. James, P.H., Pruschy, M., Vorherr, T.E., Penniston, J.T., Carafoli, E., 1989b. Primary structure of the cAMP-dependent phosphorylation site of the plasma membrane calcium pump. Biochemistry 28, 4253–4258. Jarrett, H.W., Penniston, J.T., 1977. Partial purification of the Ca2 + -Mg2 + ATPase activator from human erythrocytes: its similarity to the activator of 3’:5’-cyclic nucleotide phosphodiesterase. Biochem. Biophys. Res. Commun. 77, 1210–1216. Jeong, J., VanHouten, J.N., Dann, P., Kim, W., Sullivan, C., Yu, H., Liotta, L., Espina, V., Stern, D.F., Friedman, P.A., Wysolmerski, J.J., 2016. PMCA2 regulates HER2 protein kinase localization and signaling and promotes HER2-mediated breast cancer. Proc. Natl. Acad. Sci. U.S.A. 113, E282–E290. Jiang, L., Fernandes, D., Mehta, N., Bean, J.L., Michaelis, M.L., Zaidi, A., 2007. Partitioning of the plasma membrane Ca2 + -ATPase into lipid rafts in primary neurons: effects of cholesterol depletion. J. Neurochem. 102, 378–388. Jones, S., Zhang, X., Parsons, D.W., Lin, J.C., Leary, R.J., Angenendt, P., Mankoo, P., Carter, H., Kamiyama, H., Jimeno, A., Hong, S.M., Fu, B., Lin, M.T., Calhoun, E.S., Kamiyama, M., Walter, K., Nikolskaya, T., Nikolsky, Y., Hartigan, J., Smith, D.R., Hidalgo, M., Leach, S.D., Klein, A.P., Jaffee, E.M., Goggins, M., Maitra, A., Iacobuzio-Donahue, C., Eshleman, J.R., Kern, S.E., Hruban, R.H., Karchin, R., Papadopoulos, N., Parmigiani, G., Vogelstein, B., Velculescu, V.E., Kinzler, K.W., 2008. Core signaling pathways in human pancreatic cancers revealed by global genomic analyses. Science 321, 1801–1806.

292

T. Calì et al.

Kim, E., DeMarco, S.J., Marfatia, S.M., Chishti, A.H., Sheng, M., Strehler, E.E., 1998. Plasma membrane Ca2+ ATPase isoform 4b binds to membrane-associated guanylate kinase (MAGUK) proteins via their PDZ (PSD-95/Dlg/ZO-1) domains. J. Biol. Chem. 273, 1591–1595. Kosk-Kosicka, D., Bzdega, T., 1988. Activation of the erythrocyte Ca2 + -ATPase by either self-association or interaction with calmodulin. J. Biol. Chem. 263, 18184–18189. Kosk-Kosicka, D., Lopez, M.M., Fomitcheva, I., Lew, V.L., 1995. Self-association of plasma membrane Ca(2 +)-ATPase by volume exclusion. FEBS Lett. 371, 57–60. Kozel, P.J., Friedman, R.A., Erway, L.C., Yamoah, E.N., Liu, L.H., Riddle, T., Duffy, J.J., Doetschman, T., Miller, M.L., Cardell, E.L., Shull, G.E., 1998. Balance and hearing deficits in mice with a null mutation in the gene encoding plasma membrane Ca2+-ATPase isoform 2. J. Biol. Chem. 273 (30), 18693–18696. PubMed PMID: 9668038. Krebs, J., Michalak, M., 2007. Calcium: A Matter of Life or Death. Elsevier, Oxford, UK. Krizaj, D., Demarco, S.J., Johnson, J., Strehler, E.E., Copenhagen, D.R., 2002. Cell-specific expression of plasma membrane calcium ATPase isoforms in retinal neurons. J. Comp. Neurol. 451, 1–21. Levi, V., Rossi, J.P., Castello, P.R., Gonzalez Flecha, F.L., 2000. Oligomerization of the plasma membrane calcium pump involves two regions with different thermal stability. FEBS Lett. 483, 99–103. Li, M., Ho, P.W., Pang, S.Y., Tse, Z.H., Kung, M.H., Sham, P.C., Ho, S.L., 2014. PMCA4 (ATP2B4) mutation in familial spastic paraplegia. PLoS One 9, e104790. Lopreiato, R., Giacomello, M., Carafoli, E., 2014. The plasma membrane calcium pump: new ways to look at an old enzyme. J. Biol. Chem. 289, 10261–10268. Lu, X., Wang, L., Chen, S., He, L., Yang, X., Shi, Y., Cheng, J., Zhang, L., Gu, C.C., Huang, J., Wu, T., Ma, Y., Li, J., Cao, J., Chen, J., Ge, D., Fan, Z., Li, Y., Zhao, L., Li, H., Zhou, X., Chen, L., Liu, D., Duan, X., Hao, Y., Lu, F., Liu, Z., Yao, C., Shen, C., Pu, X., Yu, L., Fang, X., Xu, L., Mu, J., Wu, X., Zheng, R., Wu, N., Zhao, Q., Liu, X., Wang, M., Yu, D., Hu, D., Ji, X., Guo, D., Sun, D., Wang, Q., Yang, Y., Liu, F., Mao, Q., Liang, X., Ji, J., Chen, P., Mo, X., Li, D., Chai, G., Tang, Y., Li, X., Du, Z., Dou, C., Yang, Z., Meng, Q., Wang, D., Wang, R., Yang, J., Schunkert, H., Samani, N.J., Kathiresan, S., Reilly, M.P., Erdmann, J., Peng, X., Chen, R., Qiang, B., Gu, D., 2012. Genome-wide association study in Han Chinese identifies four new susceptibility loci for coronary artery disease. Nat. Genet. 44, 890–894. Minocherhomjee, A.M., Beauregard, G., Potier, M., Roufogalis, B.D., 1983. The molecular weight of the calcium-transport-ATPase of the human red blood cell determined by radiation inactivation. Biochem. Biophys. Res. Commun. 116, 895–900. Mohamed, T.M., Oceandy, D., Zi, M., Prehar, S., Alatwi, N., Wang, Y., Shaheen, M.A., Abou-Leisa, R., Schelcher, C., Hegab, Z., Baudoin, F., Emerson, M., Mamas, M., Di Benedetto, G., Zaccolo, M., Lei, M., Cartwright, E.J., Neyses, L., 2011. Plasma membrane calcium pump (PMCA4)-neuronal nitric-oxide synthase complex regulates cardiac contractility through modulation of a compartmentalized cyclic nucleotide microdomain. J. Biol. Chem. 286, 41520–41529. Niggli, V., Penniston, J.T., Carafoli, E., 1979. Purification of the (Ca2 + -Mg2 +)-ATPase from human erythrocyte membranes using a calmodulin affinity column. J. Biol. Chem. 254, 9955–9958. Niggli, V., Adunyah, E.S., Carafoli, E., 1981. Acidic phospholipids, unsaturated fatty acids, and limited proteolysis mimic the effect of calmodulin on the purified erythrocyte Ca2+-ATPase. J. Biol. Chem. 256, 8588–8592. Ortega, C., Ortolano, S., Carafoli, E., 2007. The plasma membrane calcium pump. In: Krebs, J., Michalak, M. (Eds.), In: Calcium: A Matter of Life or Death, vol. 41. Springer, New York, NY, pp. 179–197.

Plasma Membrane Ca2 + Pumps

293

Palmgren, M.G., Nissen, P., 2011. P-type ATPases. Annu. Rev. Biophys. 40, 243–266. Pang, Y., Zhu, H., Wu, P., Chen, J., 2005. The characterization of plasma membrane Ca2 + -ATPase in rich sphingomyelin-cholesterol domains. FEBS Lett. 579, 2397–2403. Paszty, K., Penheiter, A.R., Verma, A.K., Padanyi, R., Filoteo, A.G., Penniston, J.T., Enyedi, A., 2002. Asp1080 upstream of the calmodulin-binding domain is critical for autoinhibition of hPMCA4b. J. Biol. Chem. 277, 36146–36151. Paszty, K., Caride, A.J., Bajzer, Z., Offord, C.P., Padanyi, R., Hegedus, L., Varga, K., Strehler, E.E., Enyedi, A., 2015. Plasma membrane Ca(2)(+)-ATPases can shape the pattern of Ca(2)(+) transients induced by store-operated Ca(2)(+) entry. Sci. Signal. 8, ra19. Penniston, J.T., Padanyi, R., Paszty, K., Varga, K., Hegedus, L., Enyedi, A., 2014. Apart from its known function, the plasma membrane Ca(2)(+)ATPase can regulate Ca(2) (+) signaling by controlling phosphatidylinositol 4,5-bisphosphate levels. J. Cell Sci. 127, 72–84. Pottorf 2nd, W.J., Johanns, T.M., Derrington, S.M., Strehler, E.E., Enyedi, A., Thayer, S.A., 2006. Glutamate-induced protease-mediated loss of plasma membrane Ca2 + pump activity in rat hippocampal neurons. J. Neurochem. 98, 1646–1656. Preiano, B.S., Guerini, D., Carafoli, E., 1996. Expression and functional characterization of isoforms 4 of the plasma membrane calcium pump. Biochemistry 35, 7946–7953. Ribiczey, P., Tordai, A., Andrikovics, H., Filoteo, A.G., Penniston, J.T., Enouf, J., Enyedi, A., Papp, B., Kovacs, T., 2007. Isoform-specific up-regulation of plasma membrane Ca2 + ATPase expression during colon and gastric cancer cell differentiation. Cell Calcium 42, 590–605. Rimessi, A., Coletto, L., Pinton, P., Rizzuto, R., Brini, M., Carafoli, E., 2005. Inhibitory interaction of the 14-3-3{epsilon} protein with isoform 4 of the plasma membrane Ca(2 +)-ATPase pump. J. Biol. Chem. 280, 37195–37203. Ronner, P., Gazzotti, P., Carafoli, E., 1977. A lipid requirement for the (Ca2 + + Mg2 +)activated ATPase of erythrocyte membranes. Arch. Biochem. Biophys. 179, 578–583. Salamino, F., Sparatore, B., Melloni, E., Michetti, M., Viotti, P.L., Pontremoli, S., Carafoli, E., 1994. The plasma membrane calcium pump is the preferred calpain substrate within the erythrocyte. Cell Calcium 15, 28–35. Salm, E.J., Thayer, S.A., 2012. Homer proteins accelerate Ca2 + clearance mediated by the plasma membrane Ca2+ pump in hippocampal neurons. Biochem. Biophys. Res. Commun. 424, 76–81. Schatzmann, H.J., 1966. ATP-dependent Ca++-extrusion from human red cells. Experientia 22, 364–365. Schnitzer, J.E., Oh, P., Jacobson, B.S., Dvorak, A.M., 1995. Caveolae from luminal plasmalemma of rat lung endothelium: microdomains enriched in caveolin, Ca(2 +)-ATPase, and inositol trisphosphate receptor. Proc. Natl. Acad. Sci. U.S.A. 92, 1759–1763. Schuh, K., Uldrijan, S., Gambaryan, S., Roethlein, N., Neyses, L., 2003. Interaction of the plasma membrane Ca2+ pump 4b/CI with the Ca2 +/calmodulin-dependent membrane-associated kinase CASK. J. Biol. Chem. 278, 9778–9783. Schultz, J.M., Yang, Y., Caride, A.J., Filoteo, A.G., Penheiter, A.R., Lagziel, A., Morell, R.J., Mohiddin, S.A., Fananapazir, L., Madeo, A.C., Penniston, J.T., Griffith, A.J., 2005. Modification of human hearing loss by plasma-membrane calcium pump PMCA2. N. Engl. J. Med. 352, 1557–1564. Schwab, B.L., Guerini, D., Didszun, C., Bano, D., Ferrando-May, E., Fava, E., Tam, J., Xu, D., Xanthoudakis, S., Nicholson, D.W., Carafoli, E., Nicotera, P., 2002. Cleavage of plasma membrane calcium pumps by caspases: a link between apoptosis and necrosis. Cell Death Differ. 9, 818–831. Sears, C.E., Bryant, S.M., Ashley, E.A., Lygate, C.A., Rakovic, S., Wallis, H.L., Neubauer, S., Terrar, D.A., Casadei, B., 2003. Cardiac neuronal nitric oxide synthase isoform regulates myocardial contraction and calcium handling. Circ. Res. 92, e52–e59.

294

T. Calì et al.

Sepulveda, M.R., Berrocal-Carrillo, M., Gasset, M., Mata, A.M., 2006. The plasma membrane Ca2 + -ATPase isoform 4 is localized in lipid rafts of cerebellum synaptic plasma membranes. J. Biol. Chem. 281, 447–453. Shull, G.E., Greeb, J., 1988. Molecular cloning of two isoforms of the plasma membrane Ca2 + -transporting ATPase from rat brain. Structural and functional domains exhibit similarity to Na +, K+ and other cation transport ATPases. J. Biol. Chem. 263, 8646–8657. Silverstein, R.S., Tempel, B.L., 2006. Atp2b2, encoding plasma membrane Ca2+-ATPase type 2, (PMCA2) exhibits tissue-specific first exon usage in hair cells, neurons, and mammary glands of mice. Neuroscience 141, 245–257. Smallwood, J.I., Gugi, B., Rasmussen, H., 1988. Regulation of erythrocyte Ca2 + pump activity by protein kinase C. J. Biol. Chem. 263, 2195–2202. Stahl, W.L., Eakin, T.J., Owens Jr., J.W., Breininger, J.F., Filuk, P.E., Anderson, W.R., 1992. Plasma membrane Ca(2 +)-ATPase isoforms: distribution of mRNAs in rat brain by in situ hybridization. Brain Res. Mol. Brain Res. 16, 223–231. Stauffer, T.P., Guerini, D., Celio, M.R., Carafoli, E., 1997. Immunolocalization of the plasma membrane Ca2+ pump isoforms in the rat brain. Brain Res. 748, 21–29. Strehler, E.E., 2015. Plasma membrane calcium ATPases: from generic Ca(2 +) sump pumps to versatile systems for fine-tuning cellular Ca(2.). Biochem. Biophys. Res. Commun. 460, 26–33. Strehler, E.E., Zacharias, D.A., 2001. Role of alternative splicing in generating isoform diversity among plasma membrane calcium pumps. Physiol. Rev. 81, 21–50. Sun, X.Y., Chen, Z.Y., Hayashi, Y., Kanou, Y., Takagishi, Y., Oda, S., Murata, Y., 2008. Insertion of an intracisternal A particle retrotransposon element in plasma membrane calcium ATPase 2 gene attenuates its expression and produces an ataxic phenotype in joggle mutant mice. Gene 411 (1–2), 94–102. http://dx.doi.org/10.1016/j.gene. 2008.01.013. PubMed PMID: 18280673. Tabara, Y., Kohara, K., Kita, Y., Hirawa, N., Katsuya, T., Ohkubo, T., Hiura, Y., Tajima, A., Morisaki, T., Miyata, T., Nakayama, T., Takashima, N., Nakura, J., Kawamoto, R., Takahashi, N., Hata, A., Soma, M., Imai, Y., Kokubo, Y., Okamura, T., Tomoike, H., Iwai, N., Ogihara, T., Inoue, I., Tokunaga, K., Johnson, T., Caulfield, M., Munroe, P., Global Blood Pressure Genetics, C., Umemura, S., Ueshima, H., Miki, T., 2010. Common variants in the ATP2B1 gene are associated with susceptibility to hypertension: the Japanese Millennium Genome Project. Hypertension 56, 973–980. Takahashi, M., Kondou, Y., Toyoshima, C., 2007. Interdomain communication in calcium pump as revealed in the crystal structures with transmembrane inhibitors. Proc. Natl. Acad. Sci. U.S.A. 104, 5800–5805. Tang, D., Dean, W.L., Borchman, D., Paterson, C.A., 2006. The influence of membrane lipid structure on plasma membrane Ca2+-ATPase activity. Cell Calcium 39, 209–216. Tidow, H., Poulsen, L.R., Andreeva, A., Knudsen, M., Hein, K.L., Wiuf, C., Palmgren, M.G., Nissen, P., 2012. A bimodular mechanism of calcium control in eukaryotes. Nature 491, 468–472. Tolosa de Talamoni, N., Smith, C.A., Wasserman, R.H., Beltramino, C., Fullmer, C.S., Penniston, J.T., 1993. Immunocytochemical localization of the plasma membrane calcium pump, calbindin-D28k, and parvalbumin in Purkinje cells of avian and mammalian cerebellum. Proc. Natl. Acad. Sci. U.S.A. 90, 11949–11953. Toyoshima, C., Inesi, G., 2004. Structural basis of ion pumping by Ca2 + -ATPase of the sarcoplasmic reticulum. Annu. Rev. Biochem. 73, 269–292. Toyoshima, C., Nomura, H., 2002. Structural changes in the calcium pump accompanying the dissociation of calcium. Nature 418, 605–611.

Plasma Membrane Ca2 + Pumps

295

Toyoshima, C., Nakasako, M., Nomura, H., Ogawa, H., 2000. Crystal structure of the calcium pump of sarcoplasmic reticulum at 2.6 A resolution. Nature 405, 647–655. Usachev, Y.M., Toutenhoofd, S.L., Goellner, G.M., Strehler, E.E., Thayer, S.A., 2001. Differentiation induces up-regulation of plasma membrane Ca(2 +)-ATPase and concomitant increase in Ca(2 +) efflux in human neuroblastoma cell line IMR-32. J. Neurochem. 76, 1756–1765. Usachev, Y.M., DeMarco, S.J., Campbell, C., Strehler, E.E., Thayer, S.A., 2002. Bradykinin and ATP accelerate Ca(2 +) efflux from rat sensory neurons via protein kinase C and the plasma membrane Ca(2 +) pump isoform 4. Neuron 33, 113–122. Vanagas, L., Rossi, R.C., Caride, A.J., Filoteo, A.G., Strehler, E.E., Rossi, J.P., 2007. Plasma membrane calcium pump activity is affected by the membrane protein concentration: evidence for the involvement of the actin cytoskeleton. Biochim. Biophys. Acta 1768, 1641–1649. Vanagas, L., de La Fuente, M.C., Dalghi, M., Ferreira-Gomes, M., Rossi, R.C., Strehler, E.E., Mangialavori, I.C., Rossi, J.P., 2013. Differential effects of G- and F-actin on the plasma membrane calcium pump activity. Cell Biochem. Biophys. 66, 187–198. Verhoven, B., Schlegel, R.A., Williamson, P., 1995. Mechanisms of phosphatidylserine exposure, a phagocyte recognition signal, on apoptotic T lymphocytes. J. Exp. Med. 182, 1597–1601. Verma, A.K., Filoteo, A.G., Stanford, D.R., Wieben, E.D., Penniston, J.T., Strehler, E.E., Fischer, R., Heim, R., Vogel, G., Mathews, S., et al., 1988. Complete primary structure of a human plasma membrane Ca2+ pump. J. Biol. Chem. 263, 14152–14159. Verma, A.K., Paszty, K., Filoteo, A.G., Penniston, J.T., Enyedi, A., 1999. Protein kinase C phosphorylates plasma membrane Ca2+ pump isoform 4a at its calmodulin binding domain. J. Biol. Chem. 274, 527–531. Vorherr, T., Kessler, T., Hofmann, F., Carafoli, E., 1991. The calmodulin-binding domain mediates the self-association of the plasma membrane Ca2 + pump. J. Biol. Chem. 266, 22–27. Wang, K.K., Wright, L.C., Machan, C.L., Allen, B.G., Conigrave, A.D., Roufogalis, B.D., 1991. Protein kinase C phosphorylates the carboxyl terminus of the plasma membrane Ca(2 +)-ATPase from human erythrocytes. J. Biol. Chem. 266, 9078–9085. Werth, J.L., Usachev, Y.M., Thayer, S.A., 1996. Modulation of calcium efflux from cultured rat dorsal root ganglion neurons. J. Neurosci. 16, 1008–1015. Williams, J.C., Armesilla, A.L., Mohamed, T.M., Hagarty, C.L., McIntyre, F.H., Schomburg, S., Zaki, A.O., Oceandy, D., Cartwright, E.J., Buch, M.H., Emerson, M., Neyses, L., 2006. The sarcolemmal calcium pump, alpha-1 syntrophin, and neuronal nitric-oxide synthase are parts of a macromolecular protein complex. J. Biol. Chem. 281, 23341–23348. Xi, B., Shen, Y., Zhao, X., Chandak, G.R., Cheng, H., Hou, D., Li, Y., Ott, J., Zhang, Y., Wang, X., Mi, J., 2014. Association of common variants in/near six genes (ATP2B1, CSK, MTHFR, CYP17A1, STK39 and FGF5) with blood pressure/hypertension risk in Chinese children. J. Hum. Hypertens. 28, 32–36. Xiong, Y., Antalffy, G., Enyedi, A., Strehler, E.E., 2009. Apical localization of PMCA2w/b is lipid raft-dependent. Biochem. Biophys. Res. Commun. 384, 32–36. Yamoah, E.N., Lumpkin, E.A., Dumont, R.A., Smith, P.J., Hudspeth, A.J., Gillespie, P.G., 1998. Plasma membrane Ca2 + -ATPase extrudes Ca2 + from hair cell stereocilia. J. Neurosci. 18, 610–624. Yang, Y.M., Lee, J., Jo, H., Park, S., Chang, I., Muallem, S., Shin, D.M., 2014. Homer2 protein regulates plasma membrane Ca(2)(+)-ATPase-mediated Ca(2)(+) signaling in mouse parotid gland acinar cells. J. Biol. Chem. 289, 24971–24979.

296

T. Calì et al.

Zanni, G., Calı`, T., Kalscheuer, V.M., Ottolini, D., Barresi, S., Lebrun, N., Montecchi-Palazzi, L., Hu, H., Chelly, J., Bertini, E., Brini, M., Carafoli, E., 2012. Mutation of plasma membrane Ca2+ ATPase isoform 3 in a family with X-linked congenital cerebellar ataxia impairs Ca2+ homeostasis. Proc. Natl. Acad. Sci. U.S.A. 109, 14514–14519. Zhang, J., Xiao, P., Zhang, X., 2009. Phosphatidylserine externalization in caveolae inhibits Ca2 + efflux through plasma membrane Ca2+-ATPase in ECV304. Cell Calcium 45, 177–184. Zvaritch, E., James, P., Vorherr, T., Falchetto, R., Modyanov, N., Carafoli, E., 1990. Mapping of functional domains in the plasma membrane Ca2 + pump using trypsin proteolysis. Biochemistry 29, 8070–8076.

CHAPTER SEVEN

Emerging Mechanisms and Roles for Asymmetric Cytokinesis C. Thieleke-Matos*,†,1, D.S. Osório*,{,1, A.X. Carvalho*,{, E. Morais-de-Sá*,†,2 *i3S—Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de, Universidade do Porto, Porto, Portugal † Cell Division and Genomic stability, IBMC, Instituto de Biologia Molecular e Celular, and i3S, Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de, Universidade do Porto, Porto, Portugal { Cytoskeletal Dynamics, IBMC, Instituto de Biologia Molecular e Celular, and i3S, Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de, Universidade do Porto, Porto, Portugal 2 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Primer on Animal Cell Cytokinesis 2.1 Organization of Cytokinesis Contractile Ring 2.2 Regulation of Cytokinesis by Small GTPases Rho and Rac 2.3 Midbody Formation and Abscission 2.4 Overview of Epithelial Organization and Cytokinesis 3. Molecular and Cellular Mechanisms Controlling Asymmetric Cytokinesis 3.1 Intrinsic Regulation of Asymmetric Ring Constriction 3.2 Extrinsic Regulation of Epithelial Cytokinesis 3.3 Asymmetric Midbody Inheritance 4. Polarized Cytokinesis: Defining Cellular Identity and Shape 4.1 Importance of Midbody in Cell-Fate Determination 4.2 Axis Specification 4.3 Midbody as Spatial Determinant of Epithelial Architecture 5. Concluding Remarks Acknowledgments References

298 300 300 304 306 307 310 311 313 319 322 322 324 326 330 331 331

Abstract Cytokinesis completes cell division by physically separating the contents of the mother cell between the two daughter cells. This event requires the highly coordinated reorganization of the cytoskeleton within a precise window of time to ensure faithful genomic segregation. In addition, recent progress in the field highlighted the importance of cytokinesis in providing particularly important cues in the context of multicellular tissues. The organization of the cytokinetic machinery and the asymmetric localization 1

Authors contributed equally.

International Review of Cell and Molecular Biology, Volume 332 ISSN 1937-6448 http://dx.doi.org/10.1016/bs.ircmb.2017.01.004

#

2017 Elsevier Inc. All rights reserved.

297

298

C. Thieleke-Matos et al.

or inheritance of the midbody remnants is critical to define the spatial distribution of mechanical and biochemical signals. After a brief overview of the conserved steps of animal cytokinesis, we review the mechanisms controlling polarized cytokinesis focusing on the challenges of epithelial cytokinesis. Finally, we discuss the significance of these asymmetries in defining embryonic body axes, determining cell fate, and ensuring the correct propagation of epithelial organization during proliferation.

1. INTRODUCTION Cell division ensures the proliferation of unicellular organisms as well as the development and renewal of tissues in multicellular organisms. Cytokinesis, the last step of cell division segregates the cytoplasm of the mother cell between the two daughter cells. The timing of daughter cell separation must be tightly coordinated with chromosome segregation to ensure proper partitioning of the genetic material. For many decades, research on the field has largely focused on the molecular characterization of the cytokinesis machinery and on the mechanisms that contribute to timely assemble and position the contractile ring, the transient structure that allows cytokinesis to occur. Studies using unicellular organisms, mammalian cell cultures, and one-cell zygotes remarkably advanced the current understanding of these processes (D’Avino et al., 2015; Fededa and Gerlich, 2012; Green et al., 2012). However, the complex biomechanical interactions within a multicellular environment and the need to synchronize cell division with the spatial organization of tissue architecture and cell fate determination raise a number of new challenges, which are just beginning to be addressed. The event of cytokinesis is also specifically tailored to support very distinct developmental functions, mostly via the generation of asymmetries. In many cases, the contractile ring is asymmetrically positioned along the dividing cell axis, producing daughter cells with different sizes. These events, generally termed asymmetric cell division, are critical to generate cellular diversity in multicellular organisms and occur for instance in Drosophila and Caenorhabditis elegans neuroblasts, C. elegans zygotes, and mouse oocytes. Asymmetric cell division has been extensively discussed elsewhere, for comprehensive reviews of the subject the reader is referred to other publications (Gonczy, 2008; Lu and Johnston, 2013; Roubinet and Cabernard, 2014). A second type of asymmetry can occur within the division plane when the constriction of the contractile ring occurs preferentially from one of the sides. The C. elegans one-cell stage embryo provides a good example to

Asymmetric Cytokinesis in Animal Cells

299

distinguish these two types of asymmetries: (1) the embryo is divided into two daughter cells of different sizes and (2) the contractile ring normally constricts asymmetrically along the division plane (Fig. 1; Audhya et al., 2005; Maddox et al., 2007). Although the exact role of asymmetric cytokinesis in the C. elegans zygote is still a matter of debate, more recent studies in Drosophila and Xenopus highlight its particular importance in the context of an epithelium. It is now clear that the multicellular context of the epithelial tissue demands an elaborated coordination of adhesion between dividing and neighboring cells, which promotes cytokinesis asymmetries, and ensures the correct spatial organization of new cellular interfaces (Firmino et al., 2016; Founounou et al., 2013; Guillot and Lecuit, 2013; Herszterg et al., 2013, 2014; Higashi et al., 2016; Morais-de-Sa and Sunkel, 2013). In this review, we will first briefly describe the cytokinesis machinery and then focus on the physiological relevance and molecular mechanisms that control asymmetric cytokinesis and polarized midbody localization or inheritance in a variety of biological settings, with a particular emphasis on epithelial cytokinesis.

Fig. 1 Asymmetries during the first embryonic division in C. elegans. (A) Scheme of a one-cell C. elegans embryo depicting two types of asymmetry: (1) embryonic asymmetry on the anterior (A)–posterior (P) axis (anterior cell is larger than posterior cell, blue arrows) and (2) the asymmetry of ring constriction as the furrow progresses unilaterally from one of the sides and only later from the opposite (green arrows). (B) Snapshots of a dividing one-cell embryo-expressing GFP-tagged myosin II (NMY-2; green) and an mCherry-tagged pleckstrin homology (PH) domain to label the plasma membrane (red) at 50% ingression and after cytokinesis completion showing the generation of a larger anterior AB cell and a smaller posterior P1 cell (blue arrows). Size bar, 10 μm. (C) Kymograph of the equatorial region (dashed lines in B) of a dividing one-cell embryo showing the asymmetry of constriction as furrow progresses unilaterally. Fluorescent markers as in (B). Time interval between each frame is 10 s. Size bar, 10 μm.

300

C. Thieleke-Matos et al.

2. PRIMER ON ANIMAL CELL CYTOKINESIS In animal cells, cytokinesis depends on a contractile actomyosin ring that assembles after the onset of anaphase at the cell equator and then constricts toward the center of the cell while folding the plasma membrane behind it. Contractile ring constriction progresses until only a narrow bridge of cytoplasm and compacted central spindle connects the two daughter cells. The process of cytokinetic abscission then finalizes daughter cell separation by microtubule and membrane scission. In this section, we will first introduce the key players and general regulation of cytokinesis and abscission and then briefly overview epithelial cell organization and cytokinesis.

2.1 Organization of Cytokinesis Contractile Ring Pioneering electron microscopy studies by Schroeder in the 1970s showed that the cytokinetic contractile ring is a thin (0.1–0.2 μm), filamentous structure that assembles beneath the plasma membrane at the cell equator (Schroeder, 1968, 1972). Its molecular composition has been characterized through a combination of genetic perturbations, small molecule inhibitors, and proteomics approaches (Eggert et al., 2006; Glotzer, 2005; Skop et al., 2004). The contractile ring is mainly composed of filamentous actin (F-actin) and nonmuscle myosin II (hereafter myosin II) together with several cross-linking and regulatory proteins that modulate their behavior (Green et al., 2012). The scaffold of the contractile ring is composed of nonbranched actin filaments. Actin filaments treadmill naturally, monomers incorporate preferentially at the plus-end of the filament (barbed-end) and disassemble at the minus-end (pointed-end). Actin dynamics plays an important role during cytokinesis and results from the interplay of different actin-binding proteins (Blanchoin et al., 2014; Guha et al., 2005; Murthy and Wadsworth, 2005). Nucleation and elongation of nonbranched filaments are promoted by formins of the diaphanous family (CYK-1 in C. elegans, Diaphanous in Drosophila, and Dia2 in mammalians) that localize to the cytokinetic furrow and are required for cytokinesis in several species (Castrillon and Wasserman, 1994; Severson et al., 2002; Swan et al., 1998; Watanabe et al., 2008). The severing and disassembly of filaments is promoted by Cofilin whose activity is modulated by Aip1 (actin interacting protein 1) and coronin (Chen et al., 2015; Ishikawa-Ankerhold et al., 2010; Lin et al., 2010; Ono, 2003). The Arp-2/3 complex promotes the formation

Asymmetric Cytokinesis in Animal Cells

301

of branched actin filament networks and is thought to be inhibited in the cleavage furrow region (Canman et al., 2008; Rotty et al., 2013, see more in Section 2.2). Profilin binds to monomeric actin controlling actin monomer distribution between F-actin nucleated by the formin and the Arp-2/3 complex (Suarez et al., 2015) and also modulating F-actin dynamics by directly binding to barbed ends (Pernier et al., 2016). Myosin II is a motor protein that forms bipolar filaments capable of sliding actin filaments past one another by moving toward their barbed ends at the expense of ATP. It is also able to cross-link actin filaments and allows force transmission throughout an actin filament network. Myosin II is a conventional myosin composed of two heavy chains (NMHC: NMY-1/2 in C. elegans, Zip in Drosophila, MYH9/10/14 in mammalians), two essential light chains (ELC: MLC-5 in C. elegans, Mlc-c in Drosophila, MYL6 in mammalian), and two regulatory light chains (RLC: MLC-4 in C. elegans, Sqh in Drosophila, MRLCs in mammalians). The activation of myosin II requires the phosphorylation of two conserved serines in the N-terminus of the RLC by Rho kinase (ROCK: LET-502 in C. elegans, Rok in Drosophila) and other kinases (see Section 2.2). Phosphorylation of the RLC promotes a conformational shift of the myosin II molecule from a closed (inactive) state to an opened (active) state that is able to form bipolar tail-to-tail filaments (see Vicente-Manzanares et al., 2009 for a review). These filaments are approximately 300 nm long and include a few dozen myosin heads and are believed to be the main functional form of the protein (Niederman and Pollard, 1975; Pollard, 1982). Myosin II was one of the earliest molecules to be implicated in cytokinesis as its depletion or inhibition cause cytokinesis failure and multinucleation (De Lozanne and Spudich, 1987; Mabuchi and Okuno, 1977; Straight et al., 2003). In addition to actomyosin, anillin and septins are the most intensively studied components of the contractile ring. Anillin is a conserved multidomain protein that was first identified in Drosophila embryos through its ability to bind to F-actin (Miller et al., 1989). In mammalian cells and Drosophila, anillin localizes to the nucleus in interphase and then in the cortex from nuclear envelope breakdown onward. After anaphase, anillin localizes to the contractile ring and then to the midbody after full constriction (Field and Alberts, 1995; Oegema et al., 2000). Anillin has been shown to interact with a myriad of contractile ring components thus placing it as a potential hub for cytokinesis organization. In addition to being responsible for recruiting the septins to the contractile ring in some systems (D’Avino et al., 2008; Goldbach et al., 2010; Hickson and

302

C. Thieleke-Matos et al.

O’Farrell, 2008; Maddox et al., 2005), anillin also binds F-actin, myosin II (independently of F-actin), mammalian Dia2, RhoA, and its regulators Ect2 and RacGAP50C among others (Piekny and Maddox, 2010). The myosin II and actin-binding sites were mapped to the N-terminal part of the protein whereas the C-terminal half contains the anillin homology domain (AHD) that is important for RhoA binding (Piekny and Glotzer, 2008). The C-terminal end contains a pleckstrin homology (PH) domain that can associate with membranes and is important for the interaction with septins. The PH domain of anillin interacts with phosphatidylinositol phosphate lipids (PIPs), including phosphatidylinositol 4,5-bisphosphate (PtdIns (4,5)P2 or PIP2), which is enriched in the furrow in mammalian cells (Liu et al., 2012; Logan and Mandato, 2006). Additionally, two cryptic motifs for membrane association have recently been identified in the structure of human anillin (Sun et al., 2015): (1) the C2 domain that precedes the PH domain and (2) the Rho binding domain. Due to their membrane interacting abilities anillin and septins have been proposed to constitute the link between the contractile ring and the plasma membrane. Accordingly, Drosophila and mammalian cells depleted of anillin can fail cytokinesis presenting an ingressing furrow that displaces back and forth along the plasma membrane. These oscillatory movements suggest improper anchorage to the plasma membrane (Hickson and O’Farrell, 2008; Kechad et al., 2012; Piekny and Glotzer, 2008; Straight et al., 2005). Similar oscillations were not observed during the first embryonic division in C. elegans where anillin (ANI-1) depletion caused the loss of ring constriction asymmetry (see more in Section 3.1) but did not affect constriction rate or completion of cytokinesis (Maddox et al., 2007). Nonetheless, anillin was shown to promote the topological transition from an actomyosin disc to a stack of rings that occurs during meiotic cytokinesis (Dorn et al., 2010; Maddox et al., 2005). In the postembryonic context, ANI-1 depletion causes several developmental defects including cuticle, gonad, and vulva malformations (Maddox et al., 2005). Analysis of cytokinesis in vulval precursor cells (epithelial cells that form the vulva) upon ANI-1 depletion showed that although constriction was still asymmetric toward the apical side (typical in epithelial tissues, see more in Sections 2.4, 3.1, and 3.2), the constriction rate was lower indicating that ANI-1 is required for cytokinesis in epithelial cells (Bourdages et al., 2014). Septins were first identified as the main components of the filamentous rings (septin collars) that encircle the bud neck of dividing S. cerevisiae (Byers and Goetsch, 1976). Septins are conserved in animals with several

Asymmetric Cytokinesis in Animal Cells

303

orthologues described: two septins in C. elegans (UNC-59 and UNC-61) (John et al., 2007), five in Drosophila (Pnut, Sep1, Sep2, Sep4, and Sep5), and 13 in mammalian systems (divided in four groups: SEPT2 (SEPT1, 2, 4, 5), SEPT6 (SEPT6, 8, 10, 11, 14), SEPT7, and SEPT9 (SEPT3, 9, 12) (Weirich et al., 2008). Septins are GTP-binding proteins that form heterooligomers that can further assemble into apolar filaments. The N-terminal part contains a proline rich polybasic domain that is important for lipid interaction preceding a central GTPase domain that is followed by a C-terminal coiled-coil-rich septin unique element (Mostowy and Cossart, 2012). Septin filaments are considered to be the fourth element of the cytoskeleton. In contrast to other cytoskeletal components, septins self-assemble into rings in vitro and in the absence of actin form rings in the cytosol of mammalian cells (Kinoshita et al., 2002). Septins have been proposed to perform diverse functions by serving as scaffolds or diffusion barriers between different membrane compartments (Caudron and Barral, 2009). They often accumulate at curved membrane structures like the cytokinetic furrow and have been shown to sense micron-scale membrane curvature (Bridges et al., 2016). Additionally, septins can also promote the formation of F-actin rings by cross-linking filaments into curved bundles (Mavrakis et al., 2014). Finally, septins can induce membrane tubule formation from liposomes, in particular when these are enriched with PIP2 (Tanaka-Takiguchi et al., 2009). The roles of septins during cytokinesis in metazoans are tissue and developmental stage-dependent. In C. elegans, septins localize to the contractile ring and midbody but are mostly not required for embryonic divisions. Instead, septin mutants display anomalies in postembryonic development of the vulva, sensory neurons, and gonads (Finger et al., 2003; Nguyen et al., 2000). Additionally, septins together with anilins play a role in asymmetric furrow closure in the first embryonic division (Dorn et al., 2016; Maddox et al., 2007, see more in Section 3.1). In the Drosophila early embryo, Pnut localizes to the cellularization membranes, cleavage furrows of dividing cells, and epithelial leading edges during dorsal closure. Pnut is not required for embryonic divisions but mutants exhibited defects later during cellularization (Adam et al., 2000; Fares et al., 1995; Neufeld and Rubin, 1994). Septins are also specifically required for cytokinesis during planar division in the Drosophila pupal neuroepithelium, being dispensable in the orthogonal divisions where the contractile ring does not face the constraints of intercellular adhesion (Founounou et al., 2013). In mammalian tissue culture cells inhibition or depletion of different septin isoforms causes failure in cytokinesis (Estey et al., 2010; Kinoshita et al., 1997; Nagata et al., 2003;

304

C. Thieleke-Matos et al.

Spiliotis et al., 2005; Surka et al., 2002). Sept7 knockout mice displayed embryonic lethality, however, Sept7 / bone marrow-derived stem cells were able to proliferate independently of septins (Ageta-Ishihara et al., 2013; Menon et al., 2014). Overall, it is possible that a pattern of septin requirement exists: embryonic divisions, nonadherent cells, and cells in simpler tissues are mostly able to perform cytokinesis in the absence of septins, whereas strongly adherent cells and cells in complex tissues and constrained epithelia require septins for cytokinesis (reviewed in Menon and Gaestel, 2015, see more in Section 3).

2.2 Regulation of Cytokinesis by Small GTPases Rho and Rac Like most actomyosin processes, cytokinesis is regulated by members of the Rho family of small GTPases. These molecular binary switches cycle between an active GTP-bound and an inactive GDP-bound form. This switch is controlled by guanine nucleotide exchange factors (GEFs) that promote activation via GDP to GTP exchange and GTPase-activating proteins (GAPs) that promote the hydrolysis of GTP leading to inactivation. Additionally, posttranslational lipid modifications allow them to associate with the plasma membrane where they can initiate signaling cascades activating their downstream effectors. RhoA (RHO-I in C. elegans, Rho1 in Drosophila) is the most upstream factor of cytokinesis and localizes to the nascent cytokinetic furrow early in cytokinesis (Nishimura et al., 1998; Takaishi et al., 1995). At this stage, a zone of active, plasma membrane-associated RhoA forms at the division plane in a spindle-dependent manner (Bement et al., 2005; Murthy and Wadsworth, 2008; Rappaport, 1985). RhoA promotes cytokinesis via several downstream effectors of which the best characterized are: (1) Rho kinase and citron kinase (Sticky in Drosophila), two kinases that can activate myosin II via myosin phosphorylation (Amano et al., 1996; Riento and Ridley, 2003; Yamashiro et al., 2003); (2) anilin that directly binds activated RhoA (Hickson and O’Farrell, 2008; Piekny and Glotzer, 2008); and (3) diaphanous formins that are released from their autoinhibited state to promote actin filament elongation via direct interaction with RhoA (Otomo et al., 2005; Rose et al., 2005; Watanabe et al., 1997,1997, 2010). The cycling of RhoA between the active and inactive forms (RhoA flux) is important for the maintenance of the active RhoA zone and is thought to be promoted by the sequential action of its GEFs and GAPs (Miller and Bement, 2009; Zhang and Glotzer, 2015).

Asymmetric Cytokinesis in Animal Cells

305

ECT-2 (epithelial cell-transforming sequence 2, LET-21/ECT-2 in C. elegans, Pebble in Drosophila) is the main GEF for RhoA during cytokinesis (Tatsumoto et al., 1999) with its depletion in C. elegans (Schonegg and Hyman, 2006; Sonnichsen et al., 2005), Drosophila (Prokopenko et al., 1999), or mammalian cells (Kim et al., 2005; Yuce et al., 2005) leading to complete inhibition of ring constriction and cytokinesis failure, similar to what is observed in RhoA depletions. Upon chromosome segregation at anaphase, an antiparallel microtubule array termed spindle midzone forms between the two chromosomal masses. The spindle midzone concentrates cytokinesis regulators including the centralspindlin complex comprising kinesin-6 MKLP-1 (ZEN-4 in C. elegans and Pavarotti in Drosophila) and MgcRacGAP (CYK-4 in C. elegans and Tumbleweed/RacGAP50C in Drosophila) ( Jantsch-Plunger et al., 2000; Mishima et al., 2002; Pavicic-Kaltenbrunner et al., 2007). ECT-2 also concentrates in this region by interacting with the centralspindlin component MgcRacGap/CYK-4 in a Plk-1 phosphorylation-dependent manner (Kim et al., 2014; Wolfe et al., 2009; Zou et al., 2014). The interaction between the centralspindlin complex and ECT-2 is essential for RhoA activation (Basant et al., 2015; Burkard et al., 2007, 2009; Lekomtsev et al., 2012; Niiya et al., 2006; Wolfe et al., 2009; Zhang and Glotzer, 2015). In C. elegans, where Rho activation has been characterized in more detail, full RhoA activation also involves the activity of the GAP CYK-4 that functions noncanonically contributing to the activation of ECT-2 (Zhang and Glotzer, 2015). In addition, it involves NOP-1, a serine-rich protein with no recognizable domains and no identified orthologs in other systems (Tse et al., 2012). Astral microtubules touching the cortex were proposed to restrict the accumulation of active RhoA and contractile ring proteins at the poles contributing to their concentration in the equatorial region (Dassow et al., 2009; Lewellyn et al., 2010). Rac1 has mostly been reported as a negative regulator of cytokinesis by controlling the interplay between cell adhesion and division. This interplay is most critical in adherent cells and tissues since cytokinesis involves shape changes that are counteracted by substrate and cell–cell adhesions (see more in Section 3). Rac1 inactivation at the equatorial region is important to locally reduce cell adhesions and allow for contractile ring constriction (Bastos et al., 2012). Additionally, Rac1 activation of Arp-2/3 complex via WASP in the remainder cortex is thought to counteract contractile ring ingression. Consistent with this, depletion of Rac1 or the Arp-2 subunit of the Arp-2/3 complex enables cytokinesis completion in ECT-2 impaired C. elegans embryos (Loria et al., 2012).

306

C. Thieleke-Matos et al.

2.3 Midbody Formation and Abscission The physical isolation of the two daughter cells occurs during abscission, the last step of cytokinesis. The process of abscission is better characterized in human tissue culture cells (for a review, see Mierzwa and Gerlich, 2014). By late telophase, the contractile ring has reached the spindle midzone and encircles the compacted midzone microtubules. At this point, the contractile ring is denominated midbody ring. The midbody ring together with the central microtubule region is designated midbody and the flanking microtubule regions are called midbody arms. The abscission machinery assembles at the midbody and requires the evolutionarily conserved family of ESCRT (endosomal sorting complex required for transport) proteins. ESCRTs form modular complexes involved in membrane remodeling in different cellular processes. Early ESCRTs (ESCRT-0, -I, -II) exist between cytoplasmic and membrane-associated states, whereas ESCRT-III are cytoplasmic monomers shown to polymerize into different types of filaments in vitro (Guizetti et al., 2011). Finally, the AAA-ATPase Vps4 catalyzes membrane scission and the disassembly of ESCRT-III filaments (reviewed in Olmos and Carlton, 2016). The centrosomal-associated protein CEP-55 is recruited from centrosomes to the spindle midzone after phosphorylation by mitotic kinases and is an important regulator of abscission in mammalian cells (Fabbro et al., 2005; Martinez-Garay et al., 2006; Zhao et al., 2006). CEP-55 interacts with the centralspindlin component MKLP-1 and is responsible for recruiting two components of the ESCRT machinery, ALIX and the ESCRT-I complex (Lee et al., 2008; Morita et al., 2007) that can independently promote the assembly of spiral ESCRT-III filaments on one of the midbody arms (Christ et al., 2016; Elia et al., 2011; Guizetti et al., 2011; Lafaurie-Janvore et al., 2013). Microtubules are then progressively severed by the action of spastin and katanin (Agromayor et al., 2009; Matsuo et al., 2013; Reid et al., 2005; Yang et al., 2008) and membrane scission is accomplished by Vps4, leading to the cleavage of the intracellular bridge (Mierzwa and Gerlich, 2014, see further in Section 3). In HeLa cells, cytoplasmic isolation of the two daughter cells occurs around 50–60 min after the end of constriction, coincidently with microtubule bundle severing and ESCRT-III-mediated scission (Guizetti et al., 2011). Our knowledge on abscission at the embryonic and developmental level and how it compares to cells in culture is still limited. Unlike tissue culture cells, recent work in C. elegans early embryos (Green et al., 2013)

Asymmetric Cytokinesis in Animal Cells

307

demonstrated that abscission involves two steps: (1) cytoplasm isolation (early abscission) that takes place immediately when contractile ingression stops and is septin, but not ESCRT, dependent and (2) midbody remnant release (late abscission) that only occurs at the subsequent cell division involving membrane shedding in an ESCRT-III-dependent manner. Additionally, midbody microtubules are not required for abscission as both abscission stages occurred normally in embryos depleted of SPD-1/Prc-1 that do not have a spindle midzone. This work suggested a stronger role for the midbody ring instead of midbody microtubules in coordinating abscission of the first C. elegans division (Green et al., 2013). In Drosophila oogenesis, the female germline stem cell (fGSC) divides asymmetrically to give rise to another fGSC and a daughter cell cystoblast. Recent studies showed that cytokinesis completion in fGSC depends on a complex formed between ALIX and the ESCRT-III component shrub that localizes to the midbody/midbody rings. Depletion of either component delayed fGSC abscission leading to the formation of chains of daughter cells that remained connected to the fGSC via midbody rings (Eikenes et al., 2015; Matias et al., 2015). This study indicates some degree of conservation of the pathways previously described in human cells. Nonetheless, no CEP-55 orthologues are known in C. elegans or Drosophila, indicating that the ESCRT complex is either directly recruited or that there are alternative recruitment pathways.

2.4 Overview of Epithelial Organization and Cytokinesis Complex life emerged from the development of dynamic physiological barriers that protect the internal from the external environment and allow organ compartmentalization. These barriers are efficiently established by the epithelial tissue and formed by polarized sheets of cells that display a biochemical and functional asymmetry along their apicobasal axis (Rodriguez-Boulan and Macara, 2014; Fig. 2A). The cortex of epithelial cells is organized into distinct subcellular domains: an apical domain (normally facing the exterior of the epithelia or the lumen of an organ); a basal domain on the opposite side, in close contact with the underlying extracellular matrix; a lateral domain contacting neighboring cells; and a junctional domain formed at the apicolateral border that contains a number of specialized intercellular junctions, and whose organization presents some differences across evolution (Rodriguez-Boulan and Macara, 2014). The typical Drosophila epithelial cell presents adherens junctions (AJs) connecting the cortex of neighboring cells just below the apical domain. These structures are assembled by the intercellular interactions of the

308

C. Thieleke-Matos et al.

Fig. 2 Overview of epithelial organization and cytokinesis asymmetry. (A) Basic subcellular organization of Drosophila and vertebrate epithelia. Cortical domains polarized along the apicobasal axis are defined by the activity of evolutionarily conserved polarity complexes. Antagonistic (red) and positive interactions (blue) between the main polarity complexes are represented, as well as the position of the set of junctions that control adhesion (adherens junctions) and paracellular diffusion (septate junctions and tight junctions). (B) Epithelial cell cytokinesis is asymmetric along the apicobasal axis (endon views of the contractile ring are shown). The scheme shows how asymmetric furrowing (black arrows) and ring constriction position the midbody close to the apical domain. A time-lapse series of Drosophila follicular epithelial cells expressing GFPtagged anillin (green in the top panel; gray in the bottom panel) to mark the contractile/midbody ring and a plasma membrane marker (red, top panel) is shown on the right. Note that asymmetric ring constriction positions the midbody close to the apical domain (arrow).

extracellular domain of the transmembrane protein E-Cadherin, which is linked to the intracellular actin cytoskeleton by catenins. AJs control adhesion and shape, not only by resisting detachment but also via modulation of the actomyosin cytoskeleton in response to forces transmitted along the epithelial tissue

Asymmetric Cytokinesis in Animal Cells

309

(Lecuit and Yap, 2015). Most types of vertebrate epithelia contain a set of junctions close to the apical side (AJs and tight junctions (TJs)), which are globally termed the apical junctional complex (Fig. 2A). TJs are positioned above the AJ, and function as sealing junctions to segregate the internal from the external environment (Fig. 2A). In Drosophila, the order of sealing junctions (septate junction) and AJs is inverted, whereas in C. elegans, AJs and sealing junctions are combined into a single electron-dense apical junction (McMahon et al., 2001). Setting apicobasal polarity and junction positioning requires the integration of a network of biochemical interactions, based on mutual antagonism and positive-feedback loops between evolutionarily conserved protein complexes with discrete subcellular localizations (Flores-Benitez and Knust, 2016; Rodriguez-Boulan and Macara, 2014; Fig. 2A). The aPKC (consisting of Par-3 or Bazooka in Drosophila, aPKC, Par-6, and CDC42) and Crumbs (consisting of Crumbs, PALS1, or Stardust in Drosophila, and PATJ) complexes define the apical domain, whereas the basolateral domain is determined by members of the Lgl module (Lgl, Scrib, and Dlg), the protein kinase Par-1 and the Yurt module (Yurt/Coracle/Neurexin-IV/Na+,K+ATPase). The presence of Par-3/Baz within the apical complex is tightly controlled as it also interacts with components of the cell adhesion complexes, regulating the stability and localization of AJs and TJs (Ebnet et al., 2001; Takekuni et al., 2003; Wei et al., 2005). Both the apical kinase aPKC and the basolateral kinase Par-1 phosphorylate Par-3/Baz to limit its extension on the apical and basolateral side, refining the position of the junctional domain at the apicolateral border (Benton and St Johnston, 2003; Krahn et al., 2010a; Morais-de-Sa et al., 2010; Suzuki et al., 2004; Walther and Pichaud, 2010). In addition to determining the size of each spatial domain and the positioning of specific junctional complexes, epithelial polarity also ultimately orients vesicle trafficking and cytoskeleton organization, which in turn play critical roles controlling epithelial organization (Apodaca et al., 2012; Eaton and Martin-Belmonte, 2014; RodriguezBoulan and Macara, 2014). For instance, regulated trafficking of the transmembrane proteins E-Cadherin and Crumbs regulates AJ stability and apicobasal polarity, respectively (Langevin et al., 2005; Lu and Bilder, 2005; Roeth et al., 2009; Woichansky et al., 2016). During cell division, epithelial cells orient the mitotic spindle, either planar to the epithelial layer to maintain a monolayer organization or orthogonally to form stratified epithelia and control cell fate. Mitotic

310

C. Thieleke-Matos et al.

spindle orientation and cell rounding require extensive morphological and cytoskeleton changes, which are often coupled with the reorganization of core components of planar and apicobasal polarity (Bell et al., 2015; Bergstralh et al., 2013; Carvalho et al., 2015; Devenport et al., 2011; Morais-de-Sa and Sunkel, 2013; Rosa et al., 2015). In contrast, epithelia maintain the localization of apical adhesion complexes during mitosis to preserve tissue cohesiveness (Baker and Garrod, 1993; Founounou et al., 2013; Guillot and Lecuit, 2013; Herszterg et al., 2013; Jinguji and Ishikawa, 1992; Morais-de-Sa and Sunkel, 2013; Reinsch and Karsenti, 1994). Cytokinesis recently assumed particular relevance in the epithelial environment due to in vivo studies in Drosophila and C. elegans, which described a multicellular coordination of forces between neighboring and dividing cells and common features that include asymmetric ring constriction and apical midbody positioning (Fig. 2B; see more in Section 3; Bourdages et al., 2014; Founounou et al., 2013; Guillot and Lecuit, 2013; Herszterg et al., 2013, 2014; Morais-de-Sa and Sunkel, 2013). Asymmetric furrowing has also been reported in very distinct types of vertebrate epithelia, including the mouse small intestine (Fleming et al., 2007; Jinguji and Ishikawa, 1992), mouse and Zebrafish neuroepithelia (Das et al., 2003; Kosodo et al., 2004, 2008), Xenopus embryos (Hatte et al., 2014; Higashi et al., 2016), cultured hepatocytes, Madin-Darby canine kidney (MDCK), and Caco-2 cells ( Jaffe et al., 2008; Reinsch and Karsenti, 1994; Taneja et al., 2016; Wang et al., 2014).

3. MOLECULAR AND CELLULAR MECHANISMS CONTROLLING ASYMMETRIC CYTOKINESIS Unilateral, asymmetric, polarized, or nonconcentric are common terminologies to describe an asymmetry within the contractile ring plane. This kind of asymmetry is observed throughout animal evolution, having also been documented in unicellular eukaryotes, such as Trypanosome brucei, where the cleavage furrow follows unidirectional ingression during division (Wheeler et al., 2013). Asymmetric cytokinesis may also refer to situations where the midbody is asymmetrically inherited by one of the daughter cells (Kuo et al., 2011; Paolini et al., 2015; Salzmann et al., 2014). In this chapter, we will cover the molecular mechanisms that control these asymmetries. Polarized furrowing and ring constriction entail different physical and

Asymmetric Cytokinesis in Animal Cells

311

biochemical properties between the opposite sides of the equatorial region of the dividing cells. Both asymmetries in protein distribution around the ring circumference and mechanisms extrinsic to the ring contribute to polarized cytokinesis.

3.1 Intrinsic Regulation of Asymmetric Ring Constriction Asymmetric contractile ring constriction can result from the heterogeneous distribution of structural components around the ring circumference. An example of this phenomenon occurs during the first zygotic division in C. elegans (Maddox et al., 2007; Fig. 1). Early in cytokinesis, embryos display a shallow deformation around the division plane but then ingression happens only from one side. Furrowing proceeds unilaterally and only later does the opposite side undertake ingression, resulting in nonconcentric cytokinesis (Fig. 1B and C). Components of the contractile ring, such as myosin II, anillin, and septins, display higher concentration on the side of the furrow that ingresses first. Depletion of anillin or septins leads to uniform myosin II distribution and symmetric furrowing, suggesting that these proteins are regulators of asymmetric ring contraction (Maddox et al., 2007). Nonetheless, anillin-independent mechanisms for generating asymmetric constriction must exist as NOP-1 impaired embryos (see Section 2.2) still constrict asymmetrically in the absence of anillin (Tse et al., 2012). The C. elegans zygote is thought to lack polarized distribution of spatial cues at the cell equator. The symmetry breaking event is therefore random and possibly arises from the amplification of small stochastic differences in the recruitment of contractile ring components to the cell equator. This may lead to localized membrane deformation and result in the formation of a back-to-back plasma membrane configuration at only one side of the cell equator. A recent theoretical model proposed that the localized high curvature of this membrane configuration leads to the stochastic asymmetry of furrowing. The local alignment of membrane-associated actin filaments parallel to the division plane would facilitate local actomyosin contractility and further promote furrowing. Filament alignment and increased contractility would gradually propagate around the ring circumference, potentiating furrowing asymmetry (Dorn et al., 2016). According to this model, symmetric furrowing would be less advantageous to the cell because uniform F-actin alignment would significantly increase the rigidity of the contractile ring and delay

312

C. Thieleke-Matos et al.

constriction. Therefore, asymmetric constriction could serve as an energyefficient way to perform fast cytokinesis. Asymmetric furrowing with basal to apical directional asymmetry positions the midbody apically and seems to be the prevalent mode of cytokinesis in epithelial tissue (for a review, see Herszterg et al., 2014). Although the underlying causes of asymmetric furrowing seem to be related to extrinsic factors (see Section 3.2), intrinsic anisotropy of ring contractility can also play a role in epithelia (Fig. 3A). Live cell imaging in the Drosophila pupal notum revealed that the ring constricts preferentially from the basal side (Founounou et al., 2013; Herszterg et al., 2013). Accordingly, myosin II and septins accumulate on the basolateral cortex during the asymmetric phase of ring constriction. However, the distribution of ring components is variable in other epithelial tissues where it does not correlate with the basal to apical directionality of ring constriction. Moreover, in contrast to what happens in one-cell C. elegans embryos, loss of septin, or anillin function

Fig. 3 Multiple modes controlling asymmetric ring constriction to promote apical midbody positioning in Drosophila epithelia. Basal to apical constriction results from asymmetric distribution of contractile ring components (red) (A) or is promoted by apical anchoring of a homogeneous ring to intercellular adherens junctions (green) (B).

Asymmetric Cytokinesis in Animal Cells

313

does not interfere with apical-directed constriction in several Drosophila epithelial types (Founounou et al., 2013; Guillot and Lecuit, 2013; Herszterg et al., 2013; Morais-de-Sa and Sunkel, 2013). Similarly, the simple epithelium formed by the vulval precursor cells in C. elegans also displays anillin-independent asymmetric cleavage (Bourdages et al., 2014). Thus, an intrinsic asymmetry of septins and anillin distribution in the contractile ring is dispensable to generate asymmetric constriction in an epithelial tissue. Asymmetric ring constriction could alternatively be determined by the uneven distribution of an upstream factor, such as RhoA, along the apicobasal axis of epithelial cells. Curiously, unilateral cleavage furrow ingression is induced in human cells upon Chlamydia infection due to local disruption of RhoA on one side of the cell (Sun et al., 2011). Nevertheless, RhoA distribution is fairly symmetric in the Drosophila follicular epithelium (Morais-de-Sa and Sunkel, 2013), and enrichment of active RhoA on one of the sides of the division plane has yet to be documented in contexts where cytokinesis is normally asymmetric.

3.2 Extrinsic Regulation of Epithelial Cytokinesis The distribution of septins, anillin, and myosin II along the contractile ring circumference in the Drosophila follicular epithelium and embryonic ectoderm does not follow a basal to apical gradient that could explain preferential contractility of the ring on the basal side (Guillot and Lecuit, 2013; Moraisde-Sa and Sunkel, 2013). Moreover, using laser nanodissection to cut the contractile ring in embryonic epithelia, Guillot and Lecuit have shown that the intrinsic ring tension is identical in the apical and basal sides in a context of asymmetric furrowing (Guillot and Lecuit, 2013). Thus, other factors external to the ring must dictate the directionality of constriction. In agreement, recent studies show the involvement of forces exerted on the contractile ring linked to the polarized adhesion to neighboring cells or substrate. For example, despite the generalized idea that isolated single cells display near concentric constriction, polarized ring constriction toward the substrate was observed in cultured HeLa and Drosophila S2 cells (Bourdages et al., 2014; Fishkind and Wang, 1993; Taneja et al., 2016). In these cases, the extent of adhesion to the substrate modulates furrow geometry as higher densities of adhesive substrates induce the formation of larger focal adhesions that resist constriction and cause a reduction of ingression from the contacting side. This resistive force depends on a population of mitotic focal adhesions that bind to the extracellular matrix and are regulated by paxilin,

314

C. Thieleke-Matos et al.

focal adhesion kinase, and vinculin (Taneja et al., 2016). Polarization of cleavage furrow progression during epithelial cytokinesis relies instead on localized intercellular adhesion between epithelial cells. Maintenance of the mechanical coupling of the dividing cell to its neighbors via cell–cell adhesion represents a major challenge to epithelial cells, having critical impact on the specific features of epithelial cytokinesis. In particular, the contractile ring needs to counter forces generated by adhesion to neighboring cells. Recent studies using laser microsurgery and genetic manipulation in Drosophila as well as contributions from C. elegans and vertebrate models have begun to elucidate how epithelial cell division is synchronized with tissue architecture and the importance of cell-cell junctions, in particular AJs, for asymmetric cytokinesis.

3.2.1 Role of AJs in Asymmetric Ring Constriction AJs connect neighboring epithelial cells as they are linked through α-Catenin to an intracellular actomyosin belt that encircles the apicolateral border (Lecuit and Yap, 2015). Laser ablation of a region of the contractile ring in the Drosophila embryonic ectoderm led to the formation of a gap that widened with time. Widening was larger when laser ablation was performed apically, suggesting that the contractile ring is subject to higher extrinsic forces on the side where the AJs are located (Guillot and Lecuit, 2013). Since the apical side of the ring contacts AJs throughout constriction in both Drosophila embryonic ectoderm and follicular epithelium (Guillot and Lecuit, 2013; Morais-de-Sa and Sunkel, 2013), ring anchoring to AJs could result in additional forces resisting constriction (Fig. 3B). In line with this idea, genetic inactivation of AJ core components disrupts the asymmetry of ring constriction (Guillot and Lecuit, 2013; Morais-de-Sa and Sunkel, 2013) and gap widening is reduced when laser ablation is performed on the apical side of the ring in cells depleted of α-Catenin (Guillot and Lecuit, 2013). Furthermore, ectopic polarization of AJ complexes in Drosophila S2 cells directs furrow constriction toward the surface with accumulation of AJ components, showing that AJs determine the orientation of ring constriction in a context lacking other polarized cues (Morais-de-Sa and Sunkel, 2013). Intercellular adhesion also seems to oppose ring constriction leading to nonconcentric closure in the C. elegans epithelial tissue, even though it is unclear whether this asymmetry requires AJs (Bourdages et al., 2014). Furthermore, attachment of the contractile ring apparatus to the apicolateral junctions may also underlie asymmetric cytokinesis in vertebrates as studies

Asymmetric Cytokinesis in Animal Cells

315

in Xenopus, mouse, and human epithelial tissue have shown that AJs and TJs continuously contact the contractile ring during cytokinesis (Baker and Garrod, 1993; Higashi et al., 2016; Jinguji and Ishikawa, 1992). Importantly, AJs may provide an alternative link between the contractile ring and the plasma membrane in both invertebrate and vertebrate cells as overexpression of the Cadherin–Catenin complex rescues the cytokinesis defects produced by loss of anillin and septin function in Drosophila spermatocytes and mouse fibroblasts (Goldbach et al., 2010). We do not yet understand the exact molecular nature of the connection between AJs and the cytokinetic ring, but a number of components of the actomyosin belt associated with AJs during interphase are shared with the protein machinery of the cytokinetic ring (Padmanabhan et al., 2015). The mechanical load produced by a contractile ring and sensed by AJs may stabilize the association of these with the actin filaments in the ring, as application of force increases the lifetime of the interactions between actin and the Cadherin–Catenin complex (Buckley et al., 2014). Furthermore, both E-Cadherin and α-Catenin undergo force-dependent conformational changes that stabilize AJs under tension (Borghi et al., 2012; Buckley et al., 2014; Kim et al., 2015; Rakshit et al., 2012; Yao et al., 2014; Yonemura et al., 2010) and may reinforce the connection with the actomyosin contractile ring. Interestingly, FRAP studies of E-Cadherin and β-Catenin in Xenopus embryos have shown that these proteins are stabilized at the cleavage furrow (Higashi et al., 2016). Moreover, the actin-binding protein vinculin is recruited to AJs at the furrow, as anticipated when AJs are under high tension (Higashi et al., 2016; Yonemura et al., 2010). However, a similar AJ stabilization does not seem to occur in Drosophila as we describe later (Section 3.2.2).

3.2.2 Remodeling of Cell Contacts at Daughter-Neighboring Cell Interface In most epithelia, daughter cells remain neighbors after division, leading to the formation of compact clones after continuous proliferation (Aigouy et al., 2010; Baena-Lopez et al., 2005; Farhadifar et al., 2007; Gibson et al., 2006; Knox and Brown, 2002; Olivier et al., 2010). In Drosophila, efficient abscission and formation of new adhesive contacts between daughter cells involves a transient reduction of adhesion with the neighboring cells (Fig. 4A and B; Founounou et al., 2013; Guillot and Lecuit, 2013; Herszterg et al., 2013; Morais-de-Sa and Sunkel, 2013). Decrease of the E-Cadherin levels at the interface between the cleavage furrow and the

316

C. Thieleke-Matos et al.

Fig. 4 AJ remodeling and regulation of cohesion between dividing and neighboring cell membranes during epithelial cytokinesis. Schematic representation of an apical cross section of dividing Drosophila (A and B) or Xenopus (C) epithelial cells is shown. A balance of forces produced by ring constriction and neighboring cells control tissue cohesion and AJ remodeling. (A) In Drosophila embryonic epithelia, opposing forces of ring constriction (black arrows) and tension transmitted from neighboring cells (blue arrows) induces AJ disengagement and detachment of the dividing from the neighboring cell membrane. Septins are required in the contractile ring to induce AJ detachment and physical separation from neighboring cells. (B) In the Drosophila pupal notum, neighboring cells respond to ring constriction by locally accumulating myosin II in a Rok-dependent manner. This pool of myosin II exerts tension to maintain the membranes of the dividing cells tightly juxtaposed (blue arrows), ensuring maintenance of cohesion between dividing and neighboring cells (C). In the Xenopus gastrula epithelium, the connection to the AJs is maintained during constriction and reinforced by the recruitment of vinculin to the cleavage furrow.

neighboring cells seems to be induced mechanically by a combination of opposing forces, one generated by constriction of the contractile ring and the other resulting from transmission of cortical tension from neighboring cells (Fig. 4A; Founounou et al., 2013; Guillot and Lecuit, 2013). Anillin and septins seem to be necessary to ensure that ring constriction produces sufficient force to overcome adhesion to neighboring cells, as dividing cells depleted of or mutant for either protein constrict slower (Founounou et al., 2013; Guillot and Lecuit, 2013). Septin mutant cells fail cytokinesis but succeed when neighboring cells are laser ablated, further supporting the idea that constricting rings have to overcome resisting forces from neighboring cells (Founounou et al., 2013). As septins are able to cross-link actin filaments into curved bundles in vitro (Mavrakis et al., 2014), they could

Asymmetric Cytokinesis in Animal Cells

317

contribute to the production of force during ring constriction to counteract adhesion to neighboring cells. Alternatively, anillin and septins may ensure that the contractile ring is able to loosen its anchoring to AJs as their depletion or mutation lead to the stable attachment of AJs to the contractile ring (Founounou et al., 2013; Guillot and Lecuit, 2013). Extrinsic tension transmitted by neighboring cells may also challenge the stability of the last stages of ring constriction as septins have been shown to have an important role in anchoring the midbody ring to the plasma membrane (El Amine et al., 2013; Kechad et al., 2012). Accordingly, mutants for the septin pnut tend to fail cytokinesis after the ring has already constricted significantly and only a thin intercellular bridge exists between the two separating cells (Founounou et al., 2013). Although transient loss of adhesion is likely to be induced mechanically, biochemical pathways may also cooperate. Phosphorylation-dependent regulation of the interactions established within the Cadherin–Catenin complex may provide the means for rapid control of E-Cadherin levels at AJs (for a recent review, see Coopman and Djiane, 2016), but has yet to be examined during cytokinesis. Alternatively, localized regulation of vesicle trafficking triggering E-Cadherin endocytosis could also play a role. The E3 ubiquitin ligase Hakai triggers E-Cadherin internalization and lysosomal targeting in mammalian cells (Fujita et al., 2002; Palacios et al., 2005), but does not directly control E-Cadherin degradation in Drosophila (Kaido et al., 2009). Interestingly, proteins that concentrate on the contractile ring, such as Rok and the formin Diaphanous are involved in the activation of E-Cadherin endocytosis during Drosophila epithelial morphogenesis (Levayer et al., 2011). However, their involvement in contractile ring function makes it hard to uncouple the mechanical from the biochemical roles that they may play on the localized loss of E-Cadherin. Equatorial reduction of adhesion at the dividing cell-neighboring cell interface affects the cohesiveness of Drosophila epithelial tissues differently (Fig. 4). Electron microscopy analyses revealed the existence of an extracellular empty space between the ingressing furrow of a dividing cell and neighboring cells in embryonic epithelia (Fig. 4A; Guillot and Lecuit, 2013). In contrast, cohesion of the two plasma membranes is maintained in the pupal notum where the neighboring cell is pulled toward the ingressing furrow of the dividing cell (Fig. 4B; Founounou et al., 2013; Herszterg et al., 2013). In this case, cytokinetic ring constriction induces localized accumulation of active myosin in neighboring cells (Fig. 4B; Founounou et al., 2013; Herszterg et al., 2013). This Rho kinase-dependent myosin II accumulation

318

C. Thieleke-Matos et al.

is thought to generate tension to maintain the juxtaposition of daughter cell membranes during invagination. Thus, localized myosin II accumulation in the neighboring cell facilitates the contact between daughters to define the length of the newly formed AJ interface (Herszterg et al., 2013). What triggers myosin II accumulation in the neighboring cell is unknown. Local changes in tension could trigger it intrinsically but given the mechanotransduction properties of both E-Cadherin and α-Catenin, it is possible that the tension directly exerted by the constricting ring on the AJs formed with the neighboring cell plays an important role. Increased cortical stiffness and accumulation of F-actin is seen on E-Cadherin adhesion sites subjected to force (Barry et al., 2014; le Duc et al., 2010), highlighting the mechanical impact that AJs have on the remodeling of the actomyosin cytoskeleton. However, the simultaneous local decrease of AJ component levels is hard to reconcile with the idea that E-Cadherin-dependent adhesion mediates myosin II accumulation in neighboring cells. Therefore, further research will be necessary to identify how the mechanical signal of ring constriction leads to myosin accumulation. Cell cohesiveness is also maintained during cytokinesis in mammals, as electron microscopy in the mouse small intestine showed the complete juxtaposition between neighboring cells and the furrow of dividing cells (Jinguji and Ishikawa, 1992). Furthermore, maintenance of the occluding TJs throughout cytokinesis in the Xenopus embryonic gastrula ensures that the epithelial tissue functions as an efficient barrier to paracellular diffusion during division (Higashi et al., 2016). Although the aforementioned studies in the Drosophila pupal notum provided an elegant explanation as to how epithelial cohesion is maintained in a context with localized reduction in cell adhesion, recent work in the Xenopus gastrula suggests that vertebrates may display differences regarding AJ remodeling during cytokinesis. In this context, AJs remain continuously connected to the contractile ring during cytokinesis and recruit vinculin to the cleavage furrow (Fig. 4C; Higashi et al., 2016). Inhibition of vinculin is sufficient to induce a transient reduction of AJs at the region that contacts the contractile ring, which results on the acceleration of cleavage furrow ingression. Thus, a specific function for vinculin in reinforcing AJs during vertebrate epithelial cytokinesis provides one possible explanation for the differences observed between AJ behavior in Drosophila, where vinculin is dispensable for viability (Alatortsev et al., 1997). The formation of a new daughter–daughter cell interface is dispensable in some epithelial contexts during mouse, Xenopus, and chicken embryonic

Asymmetric Cytokinesis in Animal Cells

319

development, where tissue rearrangements results from the intercalation of neighboring cells between daughter cells during cell division (Firmino et al., 2016; Hatte et al., 2014; Higashi et al., 2016; Lau et al., 2015; Packard et al., 2013). In chick embryos, the degree of neighboring cell intercalation increases during gastrulation and requires low levels of actomyosin accumulation in neighboring cells to allow junction remodeling (Firmino et al., 2016). Thus, molecular responses in neighboring epithelial cells can also facilitate tissue rearrangement, instead of promoting maintenance of cohesive clonal populations. Epithelial cytokinesis therefore also seems to be tightly coordinated with the control of the geometry of epithelial cell packing, which has particular relevance during developmental stages with extensive tissue remodeling, such as gastrulation (Firmino et al., 2016; Higashi et al., 2016). In conclusion, advances in the field underline the importance of coordinating intrinsic mechanisms controlling asymmetric ring constriction with the extrinsic forces imposed by adhesion to neighboring cells. Recent efforts elucidated important aspects of this multicellular synchronization, highlighting the intricate mechanical interplay between ring constriction and the remodeling of AJs and the connected actomyosin cytoskeleton. Understanding how adhesion is remodeled and how neighboring cells sense ring constriction will now benefit from the large number of in vivo models where epithelial cytokinesis can be studied. Future analysis can also be extended to tubular epithelia, as recent studies in Drosophila tracheal branches and Zebrafish blood vessels have revealed complex AJ remodeling mechanisms coupling morphogenesis with different modes of asymmetric cytokinesis (Aydogan et al., 2015; Denes et al., 2015).

3.3 Asymmetric Midbody Inheritance Although the final destination of the midbody is unclear in most epithelial cell divisions, its biased inheritance has emerged as a critical factor in cell fate decisions and embryonic development (see Section 4 for details). Unilateral severing of intercellular bridge during abscission can lead to the asymmetric inheritance of the midbody by one of the daughter cells. This was first described in vitro in HeLa cell culture, where it may result from the preferential accumulation of abscission-related secretory vesicles in one of the daughter cells (Byers and Abramson, 1968; Gromley et al., 2005; Mishima et al., 2002). Furthermore, asymmetric inheritance may also occur as consequence of release of tension at the intercellular bridge, which

320

C. Thieleke-Matos et al.

controls the assembly of the abscission promoting ESCRT-III complex (Lafaurie-Janvore et al., 2013). The molecular mechanisms underlying asymmetric inheritance of the midbody in vivo are now coming into view, mostly arising from studies using invertebrate models. The mechanical properties of the actomyosin cell cortex also seem to underlie the stereotypical inheritance of the midbody in C. elegans embryos, as they tend to be inherited by the daughter cell that is under lower tension during early embryogenesis (Singh and Pohl, 2014). Anterior–posterior axis establishment in the C. elegans zygote and its asymmetric division result in a two-cell embryo comprising a larger AB cell with high levels of cortical myosin II and higher cortical tension and a smaller P1 cell with low levels of myosin II and lower cortical tension. The midbody remnant of the first cytokinesis localizes at the interface between the AB and P1 cells and is invariably internalized by the EMS cell (Ou et al., 2014; Singh and Pohl, 2014). Before internalization, the midbody is displaced into the future ventral side of the embryo. Midbody displacement along the AB-P1 interface is due to rotational cortical flows of myosin II perpendicular to the anteroposterior axis of the embryo, which occur primarily in the AB cell as it starts to divide. Midbody internalization seems to be due to the interaction between a coat of actin that forms on the midbody site on the P1 side of the cell–cell interface and the P1 mitotic spindle (Singh and Pohl, 2014). Singh and Pohl proposed a model (Fig. 5) where this interaction would mediate: (1) the tethering of the P1 spindle to the midbody and lead to

Fig. 5 Model for the asymmetric midbody localization and inheritance in C. elegans early embryo. (A) During the transition from the two-cell to the four-cell stage of the C. elegans embryo, the midbody of the first embryonic division moves ventrally concomitantly with rotational flows of cortical myosin in the AB cell, which are polarized orthogonally to the anterior-posterior axis. (B) Actin filaments transiently coat the membrane overlaying the midbody remnant and tethers astral microtubules of the mitotic spindle of the P1 cell contributing to its orientation. (C) The midbody remnant is released and inherited by the EMS cell.

Asymmetric Cytokinesis in Animal Cells

321

spindle skewing during cortical flow-mediated midbody displacement (spindle skewing is essential for dorsoventral patterning of the embryo, see more in Section 4.2) and (2) the release of the midbody into the P1 cell. Anterior– posterior polarity limits the formation of the actin coat to the P1 cell as disruption of anterior polarity in par-6 mutants induces actin coat formation in both sides of the midbody remnant, rotation of both AB and P1 spindles and splitting of the midbody between the two cells. Conversely, the disruption of posterior polarity using par-2 mutants blocks actin coat formation altogether and prevents midbody internalization. The nature of the actin accumulation at the midbody requires further characterization as it is unclear how it is promoted and whether the timing of cap formation and disassembly match that of the establishment of the P1 spindle and its orientation. Midbody release also requires the ESCRT-III component VPS-32.1 and the apoptotic cell corpse engulfment genes CED-1 and CED-2 (Fig. 5C; Ou et al., 2014; Singh and Pohl, 2014). Asymmetric midbody inheritance has also been directly connected with the centrosome of the dividing cell. A functional centrosome is necessary for the stereotypical midbody inheritance in Drosophila male and fGSC (Salzmann et al., 2014; Yamashita et al., 2007). Accordingly, if the function of the centrosome is disrupted using dsas-4 mutants that lack centrioles, midbody inheritance is randomized (Salzmann et al., 2014). The centrosomes present at the opposite poles of the mitotic spindle are intrinsically different as each contains a pair of centrioles of different age and molecular composition (Reina and Gonzalez, 2014). Importantly, midbody inheritance is coupled with the daughter (younger) centrosome, regardless of its segregation to the differentiated gonioblast in the male GSC division or to the stem cell in the female GSC division (Salzmann et al., 2014). In contrast, the asymmetric inheritance of the midbody is biased toward cells that retain the mother centrosome in mammalian cells, including pluripotent stem cells and cancer stem cells (Kuo et al., 2011). Although the molecular mechanism that links centrosome age to midbody inheritance is unknown, it is tempting to speculate that the asymmetries in microtubule stability observed between mother and daughter centrosomes (Gasic et al., 2015) may account for the distinct association with the midbody and ultimately to its asymmetric inheritance. However, other mechanisms must also govern this process as the correlation between centrosome age and midbody inheritance seems to be dependent on the cellular context. There is evidence for other mechanisms determining midbody inheritance beside asymmetric organization of the actomyosin cytoskeleton and

322

C. Thieleke-Matos et al.

the connection to centrosomes or astral microtubules. For instance, anillin was shown to be critical for the asymmetric inheritance of the midbody by the retinal ganglion cell during neurogenic divisions in Zebrafish neuroepithelium (Paolini et al., 2015). Current knowledge on asymmetric midbody inheritance is however still very limited and requires further efforts to understand how differences between the two daughter cells signal into the biochemical and mechanical pathways controlling abscission.

4. POLARIZED CYTOKINESIS: DEFINING CELLULAR IDENTITY AND SHAPE Initially viewed as cell division junk, the midbody has emerged as a critical signaling hub. In this sense, polarized or unilateral cytokinesis can ultimately be considered as a mechanism to position the midbody at a specific place or to induce its asymmetric inheritance. The position of the midbody either apically in epithelial cells or ventrally during C. elegans axis formation or its asymmetric retention or release have been proposed to coordinate the architecture of the dividing cells, axis specification, and cell fate determination, respectively. These recent findings will be outlined later.

4.1 Importance of Midbody in Cell-Fate Determination It is well established that the midbody can persist for long periods after cell division. Midbody persistence in cells may relate to its importance as a signaling platform. Moreover, the asymmetric inheritance of the midbody to one of the daughter cells, its release into the extracellular space, degradation, or active retention contribute to determine whether cells maintain their undifferentiated state or initiate a differentiation program after division. Accordingly, the accumulation of midbody remnants seems to correlate with stem cell identity in multiple mouse and human tissues (Kuo et al., 2011). In vitro, the number of midbody remnants decreases as human embryonic stem cells (hESCs) differentiate into fibroblast-like cells, whereas it increases when these cells are reprogrammed into pluripotent stem cells (dH1f-iPS) (Kuo et al., 2011). As mentioned above, the midbody is also asymmetrically inherited by the stem cell during fGSC division in Drosophila (Salzmann et al., 2014). These results could indicate that midbody inheritance at the end of cell division is part of a pathway controlling stemness but this idea is challenged by other findings. For instance, randomization of midbody inheritance via centrosome disruption did not elicit cell fate

Asymmetric Cytokinesis in Animal Cells

323

defects in the Drosophila germline (Stevens et al., 2007; Yamashita et al., 2003). Furthermore, the midbody is inherited by the gonialbasts, the differentiating cells in the Drosophila male germline (Salzmann et al., 2014). Although midbody inheritance does not necessarily determine stem cell fate, midbody remnant removal could be a requisite for differentiation. Midbody remnants are released from the differentiating gonialblasts into the neighboring somatic cells (Salzmann et al., 2014), whereas mammalian stem cells accumulate midbody remnants throughout subsequent divisions by blocking their degradation by autophagy (Kuo et al., 2011). Other examples that couple midbody removal to cell differentiation have been observed in stem cell types (NS-5 and HSCs) and tumor-derived cell lines where increased midbody release occurs in response to differentiating agents (Ettinger et al., 2011). Whether released midbodies simply contribute to the elimination of stem cell fate determinants or if they also serve as mediators of longdistance communication between cells remains unknown. The ability of released midbodies to move along the cell surface and their engulfment by nonsister cells via an actin-dependent phagocytosis-like mechanism may provide an efficient mechanism for long range signaling (Crowell et al., 2014). In addition to studies suggesting that differentiation can be promoted by midbody removal, the midbody can also directly act as a platform to control the distribution of differentiation factors. For instance, asymmetric inheritance of midbody remnants favors differentiation during neurogenic asymmetric cell division in the Zebrafish retina (Paolini et al., 2015). Furthermore, during the early differentiation of Drosophila neurons, the polarization of the first neurite occurs at the site of the midbody during cytokinesis of the precursor cell, through the accumulation of two midbody components, Rho1 and Aurora-A. These initiate a molecular cascade that induces membrane deformation and neurite outgrowth (Pollarolo et al., 2011). Studies using Drosophila sensory organ precursor cells also highlighted that the asymmetric distribution of factors controlling Notch signaling into the two daughter cells (pIIa and pIIb) is only generated at the later stages of cytokinesis, further revealing the importance of synchronizing cytokinesis with cell differentiation (Couturier et al., 2012; Derivery et al., 2015). Proteomic and lipidomic analysis of isolated midbodies revealed a complex mixture of lipids and hundreds of proteins, some of which are involved in multiple signaling pathways and may be important to control the cellular program (Atilla-Gokcumen et al., 2014; Skop et al., 2004). Proteins identified at the midbody whose asymmetric inheritance would be relevant

324

C. Thieleke-Matos et al.

for cell fate determination, include the core binding protein β (CBFβ), which belongs to a family of transcription factors (Lopez-Camacho et al., 2014), components of the Wnt signaling pathway, such as the Frizzled 2, β-Catenin, and Dishevelled (Fumoto et al., 2012; Habib et al., 2013; Kaplan et al., 2004); proteins involved in chemokine receptor signaling as phospholipase C and heterotrimeric Giα (Cho and Kehrl, 2007; Naito et al., 2006); and the components of the MAP kinase signaling pathway, MEK1/2 and ERK1/2 (Kasahara et al., 2007; Willard and Crouch, 2001). Furthermore, the prominin-1 (CD133, prom1) stem cell marker is another promising candidate to link midbody inheritance with cell fate. It is enriched in midbodies that are released into the extracellular neural tube to trigger differentiation of neuroepithelial cells during development of the vertebrate nervous system (Dubreuil et al., 2007; Marzesco et al., 2005). Midbody release involves budding of the apical membrane of neuroepithelial cells leading to loss of prom1 coupled with the reduction in the size of the apical domain. Midbody release therefore can also influence the switch from symmetric self-renewing to asymmetric neurogenic divisions by reducing components of the apical domain of each daughter cell asymmetrically (Dubreuil et al., 2007; Kosodo et al., 2004; Marzesco et al., 2005). Altogether, these studies highlight the importance of the midbody as a source of signaling factors controlling cell fate. However, they also challenge the idea that the midbody generally harbors determinants toward stem cell maintenance or differentiation, as receiving and retaining midbody remnants produce distinct effects depending on cell type, tissue, and species. How the midbody is selectively retained and targeted to autophagy or released in specific cell types remains elusive. Moreover, identifying the molecules regulating proliferation and differentiation that are inherited together with the midbody is an exciting avenue to understand the role of asymmetric midbody inheritance.

4.2 Axis Specification Internal or external asymmetric cues induce the establishment of embryonic axes, setting the development of the overall organism body plan. It has long been documented that asymmetric cytokinesis during the first division of both Cnetophore and Cnidaria embryos plays a critical role in axis specification (Freeman, 1977, 1981). In these organisms, the unilateral furrow ingression

Asymmetric Cytokinesis in Animal Cells

325

is initiated in the vicinity of the polar body and establishes the oral pole of the embryo. The molecular factors linking the cytokinesis machinery and axis determination remain unknown, but classic embryo centrifugation experiments that induced unilateral cleavage away from the polar body directly demonstrated that the site of the first cleavage sets the oral-aboral axis (Freeman, 1977). As mentioned previously, the midbody of the first embryonic division in C. elegans gets displaced along the AB-P1 cell interface due to myosin cortical flows. Concomitantly, a transient increase of actin at the site of the midbody in the P1 cell is thought to tether astral microtubules and skew the P1 mitotic spindle as the midbody is displaced (see Section 3.3). Midbody displacement and consequent P1 spindle skewing define the dorsoventral axis of the embryo (Singh and Pohl, 2014). Indeed, genetic interference with the stereotyped midbody localization or laser ablation of the midbody prevents spindle skewing and perturbs dorsoventral axis formation (Singh and Pohl, 2014). The asymmetrically localized midbody remnant is eventually internalized by the EMS cell at the four-cell stage. This internalization is however dispensable for dorsoventral axis formation as animals mutant for the apoptotic cell corpse engulfment genes ced-1 and ced-2 fail to internalize the midbody and do not present any developmental defects (Ou et al., 2014). These results suggest that in this case, the midbody does not work as an essential signaling platform that controls cell fate but it contributes to dorsoventral axis specification prior to its internalization, by directing the correct orientation of the P1 spindle (Singh and Pohl, 2014; Fig. 5). Interestingly, in the entomopathogenic nematode Romanomermis culicivorax, the midbody from the first embryonic division also maintains its association with the cortex during subsequent divisions and is distributed in a stereotypical manner (Schulze and Schierenberg, 2008). Despite displaying a pattern of early embryonic divisions that is dramatically different from that of C. elegans, R. culicivorax embryos also mobilize the midbody remnants of the first division onto a ventral position. Similar to what has been described in C. elegans, irradiation of the midbody also affects spindle orientation and positioning during the transition from the two-cell to fourcell stage. Thus, mediating the anchoring of astral microtubules could be a conserved feature controlling the development of these two phylogenetically distant nematodes. Whether the localization of midbody remnants from embryonic division’s influences pattern formation during development in more complex organisms remains to be addressed.

326

C. Thieleke-Matos et al.

4.3 Midbody as Spatial Determinant of Epithelial Architecture Epithelial cells must maintain tissue cohesiveness and organize a new polarized interface at the end of cytokinesis. Recent studies in the Drosophila notum and follicular epithelium provided key insights into the role of the midbody as a platform that defines how and where new cell junctions are formed (Herszterg et al., 2013; Morais-de-Sa and Sunkel, 2013). These studies describe a wave of actin polymerization that occurs around the midbody and is dependent on Rac and the Arp-2/3 complex (Fig. 6A). The apical position of the midbody acts as a spatial cue for actin polymerization as midbody mispositioning, via mitotic spindle misorientation (Herszterg et al., 2013) or disruption/mislocalization of AJ components (Morais-deSa and Sunkel, 2013), leads to the corresponding displacement of actin polymerization. Actin accumulation around the midbody may serve multiple purposes in the regulation of epithelial architecture. In the Drosophila notum, it ensures the formation of the interface between the two daughter cells, promoting the withdrawal of the membranes of neighboring cells that are interposed between the new daughter cells (Herszterg et al., 2013). Consistently, the wave of actin polymerization occurs simultaneously with the removal of the neighboring cell membrane. Loss of rac or arp3 activities leads to smaller interfaces and can fully block the formation of new AJs between daughter cells, resulting in a distinct topological arrangement of the tissue (Herszterg et al., 2013). The formation of a new AJ interface also requires the assembly of new Cadherin–Catenin complexes. Cadherin-based adhesion is promoted by the formation of oligomeric cadherin clusters, whose turnover and supra-molecular organization are dependent on the association with actin filaments (Engl et al., 2014; Hong et al., 2013; Truong Quang et al., 2013; Wu et al., 2015). Thus, actin polymerization around the midbody may also promote the assembly of AJs at the interface between new daughter cells. Arp-2/3 is indeed required to induce cell surface extension, fostering the efficient assembly of new adhesions between contacting cells in culture (Verma et al., 2004; Wu et al., 2015). Nevertheless, how nucleation of branched actin filaments is coordinated with the assembly of new E-Cadherin complexes at the daughter cell interface remains unknown. Basal mispositioning of the midbody and its associated branched actin polymerization results in epithelial invaginations in the follicular epithelium, which likely develop by positioning components that normally localize to the apical region and adhesion complexes at the site of the mispositioned

Asymmetric Cytokinesis in Animal Cells

327

Fig. 6 The midbody as a spatial determinant for the formation of new cell–cell contacts and apical polarization. (A) The scheme shows an apical cross section of Drosophila epithelia (enlarged from the region on the right). The apically localized midbody (blue circle) orients a wave of Rac and Arp-2/3-dependent actin polymerization (blue arrows), which promotes the withdrawal of the membranes of neighboring cells (red) and allows the assembly of an adherens junction interface (yellow rectangles) between daughter cells. (B) Midbody-oriented formation of the apical lumen in mammalian 3D cultures. During the first cell division, the midbody serves as a spatial cue for the formation of the apical initiation site (AMIS, blue rectangles). Stable midbody-associated microtubules guide the transport of Rab11-positive endosomes (green circles), powered by the interaction between the molecular motor kinesin-2 and FIP5. These endosomes carry specific cargo, such as the apical proteins Podocalyxin (PODXL) and Crumbs3a, and are then docked at the cleavage site in a mechanism dependent on PODXL binding to GTP-bound Rab35. The delivery of these proteins also relies on Rab8/Rab11-dependent exocytosis, which mediates the initial recruitment of Par3 to the AMIS, where it participates on an amplification loop with aPKC, Par-6 and the Cdc42 GTPase to form the apical domain. Similar mechanisms may couple the apical-directed furrow ingression and midbody positioning with the expansion of the apical lumen during subsequent divisions.

328

C. Thieleke-Matos et al.

midbody (Morais-de-Sa and Sunkel, 2013). However, how the formation of the new apical interface is directed by the midbody and whether there is a connection to actin polymerization remains unknown. The multi-PDZ domain protein Par-3 (Baz in Drosophila), which is required to establish the apical domain and define the position of the apicolateral junction in both vertebrate and invertebrate cells (Section 2.4) may be the central piece of this enigma. During Drosophila embryonic cellularization, an apical actin network is the primary cue for Baz localization at the apical-lateral border, where it mediates the recruitment of Cadherin–Catenin complexes thereby eliciting AJ formation (Harris and Peifer, 2004, 2005; McGill et al., 2009). Likewise, Par-3 specifically localizes to F-actin rich domains, which are essential for cell polarization in the context of endothelial cell adhesion (Galvagni et al., 2012). These studies establish a link between F-actinand Par-3-mediated polarizations and add to multiple studies that identify Par-3/Baz as the connection between cytokinesis and polarization. Firstly, Par-3 accumulation at the cleavage furrow promotes apical lumen formation during neural tube formation in Zebrafish (Buckley et al., 2013; Tawk et al., 2007). Second, clustering of Baz/Par-3 and Cadherin–Catenin complexes immediately after the division of Drosophila sensory neuron precursors marks the place where the first membrane deformation will initiate neuronal polarization (Pollarolo et al., 2011). Finally, in rat hepatocytes, Par-3 relocates from midbody-associated microtubules to the plasma membrane prior to abscission and accumulation of the TJ protein ZO-1, preceding de novo formation of the apical lumen (Wang et al., 2014). Although polarization during mitotic exit seems to rely on recruitment of Par-3/Baz to the plasma membrane in several contexts, how Par-3/Baz is delivered is not fully understood. Interestingly, phosphatidylinositol(4,5)-bisphosphate (PIP2) strongly accumulates at the midbody (Ben El Kadhi et al., 2011; Emoto et al., 2005; Field et al., 2005) and is also a key determinant of the apical domain, recruiting Par-3 and other polarity proteins to the plasma membrane (Bailey and Prehoda, 2015; Claret et al., 2014; Field et al., 2005; Horikoshi et al., 2011; Krahn et al., 2010b; MartinBelmonte et al., 2007). Studies in mammalian epithelial cultured cells have shown that the midbody can also serve as a platform to guide apical determinants and junction components to the plasma membrane by orchestrating the recruitment of specialized trafficking machinery (Fig. 6B). 3D cultured cysts of MDCK and human colorectal cells (Caco-2) form an apical lumen surrounded by a layer of polarized cells. The apical lumen is positioned during cytokinesis

Asymmetric Cytokinesis in Animal Cells

329

of a single epithelial cell when the midbody guides the formation of the apical membrane initiation site (AMIS), by mediating selective transport in Rab11-positive endosomes of apical proteins, such as Podocalyxin (PODXL, also known as glycoprotein 35) and Crumbs3a (Crb3a) from the outer surface to the AMIS (Hobdy-Henderson et al., 2003; Schluter et al., 2009). Stable microtubules associated with the midbody have critical relevance for the directed transport of intracellular vesicles, which is dependent on the interaction between the microtubule-binding motor kinesin-2 and FIP5, a Rab11 interacting protein (Li et al., 2014a). Formation of these apical endocytic carriers is controlled via a phospho-regulated interaction between FIP5 and Sorting Nexin 18, which temporally restricts the transport of FIP5-endosomes along the midbody microtubules to telophase (Li et al., 2014a,b; Willenborg et al., 2011). Fusion of apical proteins with the plasma membrane at the midbody site is then promoted by a combination of mechanisms that tether vesicles transporting apical determinants, recently shown to require binding of GTP-Rab35 to the cytoplasmic tail of PODXL (Klinkert et al., 2016). Furthermore, establishment of the apical domain is dependent on Rab8/Rab11a-directed exocytosis to target Par-3 to the AMIS, where Par-3 directs further apical trafficking, participating on an amplification loop with Cdc42 GTPase and aPKC (Bryant et al., 2010; Fig. 6B). During subsequent epithelial divisions, cleavage furrow ingression always occurs toward the apical surface, positioning the midbody apically, where it may contribute to lumen expansion and apical polarization (Jaffe et al., 2008; Reinsch and Karsenti, 1994; Wang et al., 2014). The precise contribution of the midbody to apical polarization in vivo remains to be shown, but a recent study that addressed TJ biogenesis revealed that two new tricellular TJs are formed in either side of the midbody immediately after cytokinesis in Xenopus embryos (Higashi et al., 2016). In addition, another recent study has shown the importance of the midbody to form a specialized apical structure—the primary cilium (Bernabe-Rubio et al., 2016)—a critical signaling center that projects outward at the surface of the apical membrane. In MDCK cells, the midbody remnant carries components involved in ciliogenesis (intraflagellar transport proteins, Rab8, and exocyst subunits) from a peripheral to a central position along the apical surface until it contacts the centrosome, where primary cilium formation begins (Bernabe-Rubio et al., 2016). Thus, the coordination between the polarity and cytokinesis machinery is relevant to set polarity de novo, and is also important to propagate epithelial organization during proliferation via the spatial localization of the midbody on the apical side of epithelia.

330

C. Thieleke-Matos et al.

5. CONCLUDING REMARKS Asymmetric cleavage furrow ingression and the regulation of the position of the midbody remnants are emerging as widespread events to couple cell division with specific developmental purposes. We are just starting to unravel the molecular mechanisms underlying these events, which depend both on intrinsic mechanisms that polarize the cytokinetic machinery and on the spatial distribution of extrinsic cues, as documented in multicellular tissues. In epithelia, intercellular junctions assume particular importance in cytokinesis both during initial generation of asymmetric ring constriction and also as part of the coordination with neighboring cells to ensure the continued integrity of the tissue. The exact mechanical impact of ring constriction on the molecular organization of cell–cell junctions, and how it is transduced to neighboring cells is not fully understood and seems to be dependent on the developmental context. Although it is now evident that much of what has been reported in specific in vivo models should not be generalized, understanding how differences in the specialized organization of each tissue impacts cytokinesis will enable the identification of key basic principles. To further decipher the interplay between cytokinesis and the organization of tissue architecture and polarity, researchers will benefit from the continuous revolution on microscopy techniques and the power of new optogenetic tools (Sydor et al., 2015; Tischer and Weiner, 2014) to reversibly interfere with protein activity with the necessary temporal and spatial control. Moreover, the development of multiple Foster resonance energy transfer (FRET)-based sensors of force-dependent conformational changes of junctional proteins will help to understand how the force generated by ring constriction could be sensed by cell-cell junctions and transduced to neighboring cells. The complexity of mechanisms underlying the localization, inheritance, and function of the midbody is just starting to be described and appears to be highly dependent on the cell type, microenvironment, and developmental context. Distinct midbody fates have been linked to the control of cell fate and tissue organization in different cell types, but the signaling effectors that associate with the midbody in a cell type-specific manner are mostly unknown. The concentration of several lipid species is dramatically altered during cytokinesis and may impact on the mechanical and signaling properties of the plasma membrane (Arai et al., 2015; Atilla-Gokcumen et al., 2014; Makino et al., 2015). Thus, understanding how particular lipids locally

Asymmetric Cytokinesis in Animal Cells

331

accumulate at the midbody and work at the interface between cytokinesis, polarity, and cell fate machineries will be important future goals. Finally, it is increasingly clear that the midbody acts as an essential cue to spatially couple cell division with apicobasal organization. Recent studies have started to elucidate the molecular basis underlying the interactions between midbodyassociated microtubules, Rab-GTPases, motor proteins, and the exocyst to coordinate the localization of polarity proteins with cytokinesis completion. Although the importance of cell adhesion and polarity protein trafficking during interphase is well established (Apodaca et al., 2012; Eaton and Martin-Belmonte, 2014), direct evidence for the role of midbody-directed vesicle trafficking in epithelial polarization is still lacking in vivo. In conclusion, the midbody can no longer be viewed as a simple by-product of cell division since its role in controlling the organization and fate of daughter cells has now been described in a wide range of organisms and biological settings. Over the next years, it will be important to further understand the mechanisms underlying the asymmetries generated during cytokinesis, and the critical importance of these events in development and disease.

ACKNOWLEDGMENTS This work was supported by grants from FCT (Fundac¸a˜o para a Ci^encia e a Tecnologia) under projects (PTDC/BIA-BCM/120132/2010) and (PTDC/BEX-BCM/0432/2014) to E.M. and from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation programme to A.X.C. (grant agreement 640553ACTOMYO). A.X.C. and E.M. have FCT Investigator positions funded by FCT and cofunded by ESF through POPH Type 4.2—Promotion of Scientific Employment.

REFERENCES Adam, J.C., Pringle, J.R., Peifer, M., 2000. Evidence for functional differentiation among Drosophila septins in cytokinesis and cellularization. Mol. Biol. Cell 11, 3123–3135. Ageta-Ishihara, N., Miyata, T., Ohshima, C., Watanabe, M., Sato, Y., Hamamura, Y., Higashiyama, T., Mazitschek, R., Bito, H., Kinoshita, M., 2013. Septins promote dendrite and axon development by negatively regulating microtubule stability via HDAC6mediated deacetylation. Nat. Commun. 4, 2532. Agromayor, M., Carlton, J.G., Phelan, J.P., Matthews, D.R., Carlin, L.M., Ameer-Beg, S., Bowers, K., Martin-Serrano, J., 2009. Essential role of hIST1 in cytokinesis. Mol. Biol. Cell 20, 1374–1387. Aigouy, B., Farhadifar, R., Staple, D.B., Sagner, A., Roper, J.C., Julicher, F., Eaton, S., 2010. Cell flow reorients the axis of planar polarity in the wing epithelium of Drosophila. Cell 142, 773–786. Alatortsev, V.E., Kramerova, I.A., Frolov, M.V., Lavrov, S.A., Westphal, E.D., 1997. Vinculin gene is non-essential in Drosophila melanogaster. FEBS Lett. 413, 197–201. Amano, M., Ito, M., Kimura, K., Fukata, Y., Chihara, K., Nakano, T., Matsuura, Y., Kaibuchi, K., 1996. Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase). J. Biol. Chem. 271, 20246–20249.

332

C. Thieleke-Matos et al.

Apodaca, G., Gallo, L.I., Bryant, D.M., 2012. Role of membrane traffic in the generation of epithelial cell asymmetry. Nat. Cell Biol. 14, 1235–1243. Arai, Y., Sampaio, J.L., Wilsch-Brauninger, M., Ettinger, A.W., Haffner, C., Huttner, W.B., 2015. Lipidome of midbody released from neural stem and progenitor cells during mammalian cortical neurogenesis. Front. Cell. Neurosci. 9, 325. Atilla-Gokcumen, G.E., Muro, E., Relat-Goberna, J., Sasse, S., Bedigian, A., Coughlin, M.L., Garcia-Manyes, S., Eggert, U.S., 2014. Dividing cells regulate their lipid composition and localization. Cell 156, 428–439. Audhya, A., Hyndman, F., McLeod, I.X., Maddox, A.S., Yates 3rd, J.R., Desai, A., Oegema, K., 2005. A complex containing the Sm protein CAR-1 and the RNA helicase CGH-1 is required for embryonic cytokinesis in Caenorhabditis elegans. J. Cell Biol. 171, 267–279. Aydogan, V., Lenard, A., Denes, A.S., Sauteur, L., Belting, H.G., Affolter, M., 2015. Endothelial cell division in angiogenic sprouts of differing cellular architecture. Biol. Open 4, 1259–1269. Baena-Lopez, L.A., Baonza, A., Garcia-Bellido, A., 2005. The orientation of cell divisions determines the shape of Drosophila organs. Curr. Biol. 15, 1640–1644. Bailey, M.J., Prehoda, K.E., 2015. Establishment of Par-polarized cortical domains via phosphoregulated membrane motifs. Dev. Cell 35, 199–210. Baker, J., Garrod, D., 1993. Epithelial cells retain junctions during mitosis. J. Cell Sci. 104 (Pt. 2), 415–425. Barry, A.K., Tabdili, H., Muhamed, I., Wu, J., Shashikanth, N., Gomez, G.A., Yap, A.S., Gottardi, C.J., de Rooij, J., Wang, N., Leckband, D.E., 2014. alpha-catenin cytomechanics—role in cadherin-dependent adhesion and mechanotransduction. J. Cell Sci. 127, 1779–1791. Basant, A., Lekomtsev, S., Tse, Y.C., Zhang, D., Longhini, K.M., Petronczki, M., Glotzer, M., 2015. Aurora B kinase promotes cytokinesis by inducing centralspindlin oligomers that associate with the plasma membrane. Dev. Cell 33, 204–215. Bastos, R.N., Penate, X., Bates, M., Hammond, D., Barr, F.A., 2012. CYK4 inhibits Rac1dependent PAK1 and ARHGEF7 effector pathways during cytokinesis. J. Cell Biol. 198, 865–880. Bell, G.P., Fletcher, G.C., Brain, R., Thompson, B.J., 2015. Aurora kinases phosphorylate Lgl to induce mitotic spindle orientation in Drosophila epithelia. Curr. Biol. 25, 61–68. Bement, W.M., Benink, H.A., von Dassow, G., 2005. A microtubule-dependent zone of active RhoA during cleavage plane specification. J. Cell Biol. 170, 91–101. Ben El Kadhi, K., Roubinet, C., Solinet, S., Emery, G., Carreno, S., 2011. The inositol 5-phosphatase dOCRL controls PI(4,5)P2 homeostasis and is necessary for cytokinesis. Curr. Biol. 21, 1074–1079. Benton, R., St Johnston, D., 2003. Drosophila PAR-1 and 14-3-3 inhibit Bazooka/PAR-3 to establish complementary cortical domains in polarized cells. Cell 115, 691–704. Bergstralh, D.T., Lovegrove, H.E., St Johnston, D., 2013. Discs large links spindle orientation to apical-basal polarity in Drosophila epithelia. Curr. Biol. 23, 1707–1712. Bernabe-Rubio, M., Andres, G., Casares-Arias, J., Fernandez-Barrera, J., Rangel, L., Reglero-Real, N., Gershlick, D.C., Fernandez, J.J., Millan, J., Correas, I., Miguez, D.G., Alonso, M.A., 2016. Novel role for the midbody in primary ciliogenesis by polarized epithelial cells. J. Cell Biol. 214, 259–273. Blanchoin, L., Boujemaa-Paterski, R., Sykes, C.C., Plastino, J., 2014. Actin dynamics, architecture, and mechanics in cell motility. Physiol. Rev. 94, 235–263. Borghi, N., Sorokina, M., Shcherbakova, O.G., Weis, W.I., Pruitt, B.L., Nelson, W.J., Dunn, A.R., 2012. E-cadherin is under constitutive actomyosin-generated tension that is increased at cell-cell contacts upon externally applied stretch. Proc. Natl. Acad. Sci. U.S.A. 109, 12568–12573.

Asymmetric Cytokinesis in Animal Cells

333

Bourdages, K.G., Lacroix, B., Dorn, J.F., Descovich, C.P., Maddox, A.S., 2014. Quantitative analysis of cytokinesis In Situ during C. elegans postembryonic development. PLoS One 9, e110689. Bridges, A.A., Jentzsch, M.S., Oakes, P.W., Occhipinti, P., Gladfelter, A.S., 2016. Micronscale plasma membrane curvature is recognized by the septin cytoskeleton. J. Cell Biol. 213, 23–32. Bryant, D.M., Datta, A., Rodriguez-Fraticelli, A.E., Peranen, J., Martin-Belmonte, F., Mostov, K.E., 2010. A molecular network for de novo generation of the apical surface and lumen. Nat. Cell Biol. 12, 1035–1045. Buckley, C.E., Ren, X., Ward, L.C., Girdler, G.C., Araya, C., Green, M.J., Clark, B.S., Link, B.A., Clarke, J.D., 2013. Mirror-symmetric microtubule assembly and cell interactions drive lumen formation in the zebrafish neural rod. Embo J. 32, 30–44. Buckley, C.D., Tan, J., Anderson, K.L., Hanein, D., Volkmann, N., Weis, W.I., Nelson, W.J., Dunn, A.R., 2014. Cell adhesion. The minimal cadherin-catenin complex binds to actin filaments under force. Science 346, 1254211. Burkard, M.E., Randall, C.L., Larochelle, S.P., Zhang, C., Shokat, K.M., Fisher, R.P., Jallepalli, P.V., 2007. Chemical genetics reveals the requirement for Polo-like kinase 1 activity in positioning RhoA and triggering cytokinesis in human cells. Proc. Natl. Acad. Sci. U.S.A. 104, 4383–4388. Burkard, M.E., Maciejowski, J., Rodriguez-Bravo, V.N., Repka, M., Lowery, D.M., Clauser, K.R., Zhang, C., Shokat, K.M., Carr, S.A., Yaffe, M.B., Jallepalli, P.V., 2009. Plk1 self-organization and priming phosphorylation of HsCYK-4 at the spindle midzone regulate the onset of division in human cells. PLoS Biol. 7, e1000111. Byers, B., Abramson, D.H., 1968. Cytokinesis in HeLa: post-telophase delay and microtubule-associated motility. Protoplasma 66, 413–435. Byers, B., Goetsch, L., 1976. A highly ordered ring of membrane-associated filaments in budding yeast. J. Cell Biol. 69, 717–721. Canman, J.C., Lewellyn, L., Laband, K., Smerdon, S.J., Desai, A., Bowerman, B., Oegema, K., 2008. Inhibition of Rac by the GAP activity of centralspindlin is essential for cytokinesis. Science 322, 1543–1546. Carvalho, C.A., Moreira, S., Ventura, G., Sunkel, C.E., Morais-de-Sa, E., 2015. Aurora a triggers Lgl cortical release during symmetric division to control planar spindle orientation. Curr. Biol. 25, 53–60. Castrillon, D.H., Wasserman, S.A., 1994. Diaphanous is required for cytokinesis in Drosophila and shares domains of similarity with the products of the limb deformity gene. Development 120, 3367–3377. Caudron, F., Barral, Y., 2009. Septins and the lateral compartmentalization of eukaryotic membranes. Dev. Cell 16, 493–506. Chen, Q., Courtemanche, N., Pollard, T.D., 2015. Aip1 promotes actin filament severing by cofilin and regulates constriction of the cytokinetic contractile ring. J. Biol. Chem. 290, 2289–2300. Cho, H., Kehrl, J.H., 2007. Localization of Gi alpha proteins in the centrosomes and at the midbody: implication for their role in cell division. J. Cell Biol. 178, 245–255. Christ, L., Wenzel, E.M., Liestol, K., Raiborg, C., Campsteijn, C., Stenmark, H., 2016. ALIX and ESCRT-I/II function as parallel ESCRT-III recruiters in cytokinetic abscission. J. Cell Biol. 212, 499–513. Claret, S., Jouette, J., Benoit, B., Legent, K., Guichet, A., 2014. PI(4,5)P2 produced by the PI4P5K SKTL controls apical size by tethering PAR-3 in Drosophila epithelial cells. Curr. Biol. 24, 1071–1079. Coopman, P., Djiane, A., 2016. Adherens junction and E-Cadherin complex regulation by epithelial polarity. Cell. Mol. Life Sci. 73, 3535–3553.

334

C. Thieleke-Matos et al.

Couturier, L., Vodovar, N., Schweisguth, F., 2012. Endocytosis by Numb breaks Notch symmetry at cytokinesis. Nat. Cell Biol. 14, 131–139. Crowell, E.F., Gaffuri, A.L., Gayraud-Morel, B., Tajbakhsh, S., Echard, A., 2014. Engulfment of the midbody remnant after cytokinesis in mammalian cells. J. Cell Sci. 127, 3840–3851. Das, T., Payer, B., Cayouette, M., Harris, W.A., 2003. In vivo time-lapse imaging of cell divisions during neurogenesis in the developing zebrafish retina. Neuron 37, 597–609. Dassow, G.v., Verbrugghe, K.J.C., Miller, A.L., Sider, J.R., Bement, W.M., 2009. Action at a distance during cytokinesis. J. Cell Biol. 187, 831–845. D’Avino, P.P., Takeda, T., Capalbo, L., Zhang, W., Lilley, K.S., Laue, E.D., Glover, D.M., 2008. Interaction between anillin and RacGAP50C connects the actomyosin contractile ring with spindle microtubules at the cell division site. J. Cell Sci. 121, 1151–1158. D’Avino, P.P., Giansanti, M.G., Petronczki, M., 2015. Cytokinesis in animal cells. Cold Spring Harb. Perspect. Biol. 7, a015834. De Lozanne, A., Spudich, J.A., 1987. Disruption of the Dictyostelium myosin heavy chain gene by homologous recombination. Science (New York, N.Y.) 236, 1086–1091. Denes, A.S., Kanca, O., Affolter, M., 2015. A cellular process that includes asymmetric cytokinesis remodels the dorsal tracheal branches in Drosophila larvae. Development 142, 1794–1805. Derivery, E., Seum, C., Daeden, A., Loubery, S., Holtzer, L., Julicher, F., Gonzalez-Gaitan, M., 2015. Polarized endosome dynamics by spindle asymmetry during asymmetric cell division. Nature 528, 280–285. Devenport, D., Oristian, D., Heller, E., Fuchs, E., 2011. Mitotic internalization of planar cell polarity proteins preserves tissue polarity. Nat. Cell Biol. 13, 893–902. Dorn, J.F., Zhang, L., Paradis, V.r., Edoh-Bedi, D., Jusu, S., Maddox, P.S., Maddox, A.S., 2010. Actomyosin tube formation in polar body cytokinesis requires anillin in C. elegans. Curr. Biol. 20, 2046–2051. Dorn, J.F., Zhang, L., Phi, T.T., Lacroix, B., Maddox, P.S., Liu, J., Maddox, A.S., 2016. A theoretical model of cytokinesis implicates feedback between membrane curvature and cytoskeletal organization in asymmetric cytokinetic furrowing. Mol. Biol. Cell 27, 1286–1299. Dubreuil, V., Marzesco, A.M., Corbeil, D., Huttner, W.B., Wilsch-Brauninger, M., 2007. Midbody and primary cilium of neural progenitors release extracellular membrane particles enriched in the stem cell marker prominin-1. J. Cell Biol. 176, 483–495. Eaton, S., Martin-Belmonte, F., 2014. Cargo sorting in the endocytic pathway: a key regulator of cell polarity and tissue dynamics. Cold Spring Harb. Perspect. Biol. 6, a016899. Ebnet, K., Suzuki, A., Horikoshi, Y., Hirose, T., Meyer Zu Brickwedde, M.K., Ohno, S., Vestweber, D., 2001. The cell polarity protein ASIP/PAR-3 directly associates with junctional adhesion molecule (JAM). Embo J. 20, 3738–3748. Eggert, U.S., Mitchison, T.J., Field, C.M., 2006. Animal cytokinesis: from parts list to mechanisms. Annu. Rev. Biochem. 75, 543–566. Eikenes, A.H., Malerod, L., Lie-Jensen, A., Sem Wegner, C., Brech, A., Liestol, K., Stenmark, H., Haglund, K., 2015. Src64 controls a novel actin network required for proper ring canal formation in the Drosophila male germline. Development 142, 4107–4118. El Amine, N., Kechad, A., Jananji, S., Hickson, G.R., 2013. Opposing actions of septins and Sticky on Anillin promote the transition from contractile to midbody ring. J. Cell Biol. 203, 487–504. Elia, N., Sougrat, R., Spurlin, T.A., Hurley, J.H., Lippincott-Schwartz, J., 2011. Dynamics of endosomal sorting complex required for transport (ESCRT) machinery during cytokinesis and its role in abscission. Proc. Natl. Acad. Sci. U.S.A. 108, 4846–4851.

Asymmetric Cytokinesis in Animal Cells

335

Emoto, K., Inadome, H., Kanaho, Y., Narumiya, S., Umeda, M., 2005. Local change in phospholipid composition at the cleavage furrow is essential for completion of cytokinesis. J. Biol. Chem. 280, 37901–37907. Engl, W., Arasi, B., Yap, L.L., Thiery, J.P., Viasnoff, V., 2014. Actin dynamics modulate mechanosensitive immobilization of E-cadherin at adherens junctions. Nat. Cell Biol. 16, 587–594. Estey, M.P., Ciano-Oliveira, C.D., Froese, C.D., Bejide, M.T., Trimble, W.S., 2010. Distinct roles of septins in cytokinesis: SEPT9 mediates midbody abscission. J. Cell Biol. 191, 741–749. Ettinger, A.W., Wilsch-Brauninger, M., Marzesco, A.M., Bickle, M., Lohmann, A., Maliga, Z., Karbanova, J., Corbeil, D., Hyman, A.A., Huttner, W.B., 2011. Proliferating versus differentiating stem and cancer cells exhibit distinct midbody-release behaviour. Nat. Commun. 2, 503. Fabbro, M., Zhou, B.-B., Takahashi, M., Sarcevic, B., Lal, P., Graham, M.E., Gabrielli, B.G., Robinson, P.J., Nigg, E.A., Ono, Y., Khanna, K.K., 2005. Cdk1/ Erk2- and Plk1-dependent phosphorylation of a centrosome protein, Cep55, is required for its recruitment to midbody and cytokinesis. Dev. Cell 9, 477–488. Fares, H., Peifer, M., Pringle, J.R., 1995. Localization and possible functions of Drosophila septins. Mol. Biol. Cell 6, 1843–1859. Farhadifar, R., Roper, J.C., Aigouy, B., Eaton, S., Julicher, F., 2007. The influence of cell mechanics, cell-cell interactions, and proliferation on epithelial packing. Curr. Biol. 17, 2095–2104. Fededa, J.P., Gerlich, D.W., 2012. Molecular control of animal cell cytokinesis. Nat. Cell Biol. 14, 440–447. Field, C.M., Alberts, B.M., 1995. Anillin, a contractile ring protein that cycles from the nucleus to the cell cortex. J. Cell Biol. 131, 165–178. Field, S.J., Madson, N., Kerr, M.L., Galbraith, K.A., Kennedy, C.E., Tahiliani, M., Wilkins, A., Cantley, L.C., 2005. PtdIns(4,5)P2 functions at the cleavage furrow during cytokinesis. Curr. Biol. 15, 1407–1412. Finger, F.P., Kopish, K.R., White, J.G., 2003. A role for septins in cellular and axonal migration in C. elegans. Dev. Biol. 261, 220–234. Firmino, J., Rocancourt, D., Saadaoui, M., Moreau, C., Gros, J., 2016. Cell division drives epithelial cell rearrangements during gastrulation in chick. Dev. Cell 36, 249–261. Fishkind, D.J., Wang, Y.L., 1993. Orientation and three-dimensional organization of actin filaments in dividing cultured cells. J. Cell Biol. 123, 837–848. Fleming, E.S., Zajac, M., Moschenross, D.M., Montrose, D.C., Rosenberg, D.W., Cowan, A.E., Tirnauer, J.S., 2007. Planar spindle orientation and asymmetric cytokinesis in the mouse small intestine. J. Histochem. Cytochem. 55, 1173–1180. Flores-Benitez, D., Knust, E., 2016. Dynamics of epithelial cell polarity in Drosophila: how to regulate the regulators? Curr. Opin. Cell Biol. 42, 13–21. Founounou, N., Loyer, N., Le Borgne, R., 2013. Septins regulate the contractility of the actomyosin ring to enable adherens junction remodeling during cytokinesis of epithelial cells. Dev. Cell 24, 242–255. Freeman, G., 1977. The establishment of the oral-aboral axis in the ctenophore embryo. J. Embryol. Exp. Morph. 42, 237–260. Freeman, G., 1981. The role of polarity in the development of the hydrozoan planula larva. Rouxs Arch. Dev. Biol. 190, 168–184. Fujita, Y., Krause, G., Scheffner, M., Zechner, D., Leddy, H.E., Behrens, J., Sommer, T., Birchmeier, W., 2002. Hakai, a c-Cbl-like protein, ubiquitinates and induces endocytosis of the E-cadherin complex. Nat. Cell Biol. 4, 222–231. Fumoto, K., Kikuchi, K., Gon, H., Kikuchi, A., 2012. Wnt5a signaling controls cytokinesis by correctly positioning ESCRT-III at the midbody. J. Cell Sci. 125, 4822–4832.

336

C. Thieleke-Matos et al.

Galvagni, F., Baldari, C.T., Oliviero, S., Orlandini, M., 2012. An apical actin-rich domain drives the establishment of cell polarity during cell adhesion. Histochem. Cell Biol. 138, 419–433. Gasic, I., Nerurkar, P., Meraldi, P., 2015. Centrosome age regulates kinetochore-microtubule stability and biases chromosome mis-segregation. Elife 4, e07909. Gibson, M.C., Patel, A.B., Nagpal, R., Perrimon, N., 2006. The emergence of geometric order in proliferating metazoan epithelia. Nature 442, 1038–1041. Glotzer, M., 2005. The molecular requirements for cytokinesis. Science 307, 1735–1739. Goldbach, P., Wong, R., Beise, N., Sarpal, R., Trimble, W.S., Brill, J.A., 2010. Stabilization of the actomyosin ring enables spermatocyte cytokinesis in Drosophila. Mol. Biol. Cell 21, 1482–1493. Gonczy, P., 2008. Mechanisms of asymmetric cell division: flies and worms pave the way. Nat. Rev. Mol. Cell Biol. 9, 355–366. Green, R.A., Paluch, E., Oegema, K., 2012. Cytokinesis in animal cells. Annu. Rev. Cell Dev. Biol. 28, 29–58. Green, R.A., Mayers, J.R., Wang, S., Lewellyn, L., Desai, A., Audhya, A., Oegema, K., 2013. The midbody ring scaffolds the abscission machinery in the absence of midbody microtubules. J. Cell Biol. 203, 505–520. Gromley, A., Yeaman, C., Rosa, J., Redick, S., Chen, C.T., Mirabelle, S., Guha, M., Sillibourne, J., Doxsey, S.J., 2005. Centriolin anchoring of exocyst and SNARE complexes at the midbody is required for secretory-vesicle-mediated abscission. Cell 123, 75–87. Guha, M., Zhou, M., Wang, Y.-l., 2005. Cortical actin turnover during cytokinesis requires myosin II. Curr. Biol. 15, 732–736. Guillot, C., Lecuit, T., 2013. Adhesion disengagement uncouples intrinsic and extrinsic forces to drive cytokinesis in epithelial tissues. Dev. Cell 24, 227–241. Guizetti, J., Schermelleh, L., Mantler, J., Maar, S., Poser, I., Leonhardt, H., Muller-Reichert, T., Gerlich, D.W., 2011. Cortical constriction during abscission involves helices of ESCRT-III-dependent filaments. Science 331, 1616–1620. Habib, S.J., Chen, B.C., Tsai, F.C., Anastassiadis, K., Meyer, T., Betzig, E., Nusse, R., 2013. A localized Wnt signal orients asymmetric stem cell division in vitro. Science 339, 1445–1448. Harris, T.J., Peifer, M., 2004. Adherens junction-dependent and -independent steps in the establishment of epithelial cell polarity in Drosophila. J. Cell Biol. 167, 135–147. Harris, T.J., Peifer, M., 2005. The positioning and segregation of apical cues during epithelial polarity establishment in Drosophila. J. Cell Biol. 170, 813–823. Hatte, G., Tramier, M., Prigent, C., Tassan, J.P., 2014. Epithelial cell division in the Xenopus laevis embryo during gastrulation. Int. J. Dev. Biol. 58, 775–781. Herszterg, S., Leibfried, A., Bosveld, F., Martin, C., Bellaiche, Y., 2013. Interplay between the dividing cell and its neighbors regulates adherens junction formation during cytokinesis in epithelial tissue. Dev. Cell 24, 256–270. Herszterg, S., Pinheiro, D., Bellaiche, Y., 2014. A multicellular view of cytokinesis in epithelial tissue. Trends Cell Biol. 24, 285–293. Hickson, G.R.X., O’Farrell, P.H., 2008. Anillin: a pivotal organizer of the cytokinetic machinery. Biochem. Soc. Trans. 36, 439–441. Higashi, T., Arnold, T.R., Stephenson, R.E., Dinshaw, K.M., Miller, A.L., 2016. Maintenance of the epithelial barrier and remodeling of cell-cell junctions during cytokinesis. Curr. Biol. 26, 1829–1842. Hobdy-Henderson, K.C., Hales, C.M., Lapierre, L.A., Cheney, R.E., Goldenring, J.R., 2003. Dynamics of the apical plasma membrane recycling system during cell division. Traffic 4, 681–693.

Asymmetric Cytokinesis in Animal Cells

337

Hong, S., Troyanovsky, R.B., Troyanovsky, S.M., 2013. Binding to F-actin guides cadherin cluster assembly, stability, and movement. J. Cell Biol. 201, 131–143. Horikoshi, Y., Hamada, S., Ohno, S., Suetsugu, S., 2011. Phosphoinositide binding by par-3 involved in par-3 localization. Cell Struct. Funct. 36, 97–102. Ishikawa-Ankerhold, H.C., Gerisch, G.N., Mueller-Taubenberger, A., 2010. Genetic evidence for concerted control of actin dynamics in cytokinesis, endocytic traffic, and cell motility by coronin and Aip1. Cytoskeleton (Hoboken) 67, 442–455. Jaffe, A.B., Kaji, N., Durgan, J., Hall, A., 2008. Cdc42 controls spindle orientation to position the apical surface during epithelial morphogenesis. J. Cell Biol. 183, 625–633. Jantsch-Plunger, V., Gonczy, P., Romano, A., Schnabel, H., Hamill, D., Schnabel, R., Hyman, A.A., Glotzer, M., 2000. CYK-4: A Rho family gtpase activating protein (GAP) required for central spindle formation and cytokinesis. J. Cell Biol. 149, 1391–1404. Jinguji, Y., Ishikawa, H., 1992. Electron microscopic observations on the maintenance of the tight junction during cell division in the epithelium of the mouse small intestine. Cell Struct. Funct. 17, 27–37. John, C.M., Hite, R.K., Weirich, C.S., Fitzgerald, D.J., Jawhari, H., Faty, M., Schlapfer, D., Kroschewski, R., Winkler, F.K., Walz, T., Barral, Y., Steinmetz, M.O., 2007. The Caenorhabditis elegans septin complex is nonpolar. Embo J. 26, 3296–3307. Kaido, M., Wada, H., Shindo, M., Hayashi, S., 2009. Essential requirement for RING finger E3 ubiquitin ligase Hakai in early embryonic development of Drosophila. Genes Cells 14, 1067–1077. Kaplan, D.D., Meigs, T.E., Kelly, P., Casey, P.J., 2004. Identification of a role for betacatenin in the establishment of a bipolar mitotic spindle. J. Biol. Chem. 279, 10829–10832. Kasahara, K., Nakayama, Y., Nakazato, Y., Ikeda, K., Kuga, T., Yamaguchi, N., 2007. Src signaling regulates completion of abscission in cytokinesis through ERK/MAPK activation at the midbody. J. Biol. Chem. 282, 5327–5339. Kechad, A., Jananji, S., Ruella, Y., Hickson, G.R., 2012. Anillin acts as a bifunctional linker coordinating midbody ring biogenesis during cytokinesis. Curr. Biol. 22, 197–203. Kim, J.-E., Billadeau, D.D., Chen, J., 2005. The tandem BRCT domains of Ect2 are required for both negative and positive regulation of Ect2 in cytokinesis. J. Biol. Chem. 280, 5733–5739. Kim, H., Guo, F., Brahma, S., Xing, Y., Burkard, M.E., 2014. Centralspindlin assembly and 2 phosphorylations on MgcRacGAP by Polo-like kinase 1 initiate Ect2 binding in early cytokinesis. Cell Cycle 13, 2952–2961. Kim, T.J., Zheng, S., Sun, J., Muhamed, I., Wu, J., Lei, L., Kong, X., Leckband, D.E., Wang, Y., 2015. Dynamic visualization of alpha-catenin reveals rapid, reversible conformation switching between tension states. Curr. Biol. 25, 218–224. Kinoshita, M., Kumar, S., Mizoguchi, A., Ide, C., Kinoshita, A., Haraguchi, T., Hiraoka, Y., Noda, M., 1997. Nedd5, a mammalian septin, is a novel cytoskeletal component interacting with actin-based structures. Genes Dev. 11, 1535–1547. Kinoshita, M., Field, C.M., Coughlin, M.L., Straight, A.F., Mitchison, T.J., 2002. Self- and actin-templated assembly of Mammalian septins. Dev. Cell 3, 791–802. Klinkert, K., Rocancourt, M., Houdusse, A., Echard, A., 2016. Rab35 GTPase couples cell division with initiation of epithelial apico-basal polarity and lumen opening. Nat. Commun. 7, 11166. Knox, A.L., Brown, N.H., 2002. Rap1 GTPase regulation of adherens junction positioning and cell adhesion. Science 295, 1285–1288. Kosodo, Y., Roper, K., Haubensak, W., Marzesco, A.M., Corbeil, D., Huttner, W.B., 2004. Asymmetric distribution of the apical plasma membrane during neurogenic divisions of mammalian neuroepithelial cells. Embo J. 23, 2314–2324.

338

C. Thieleke-Matos et al.

Kosodo, Y., Toida, K., Dubreuil, V., Alexandre, P., Schenk, J., Kiyokage, E., Attardo, A., Mora-Bermudez, F., Arii, T., Clarke, J.D., Huttner, W.B., 2008. Cytokinesis of neuroepithelial cells can divide their basal process before anaphase. Embo J. 27, 3151–3163. Krahn, M.P., Buckers, J., Kastrup, L., Wodarz, A., 2010a. Formation of a Bazooka-Stardust complex is essential for plasma membrane polarity in epithelia. J. Cell Biol. 190, 751–760. Krahn, M.P., Klopfenstein, D.R., Fischer, N., Wodarz, A., 2010b. Membrane targeting of Bazooka/PAR-3 is mediated by direct binding to phosphoinositide lipids. Curr. Biol. 20, 636–642. Kuo, T.C., Chen, C.T., Baron, D., Onder, T.T., Loewer, S., Almeida, S., Weismann, C.M., Xu, P., Houghton, J.M., Gao, F.B., Daley, G.Q., Doxsey, S., 2011. Midbody accumulation through evasion of autophagy contributes to cellular reprogramming and tumorigenicity. Nat. Cell Biol. 13, 1214–1223. Lafaurie-Janvore, J., Maiuri, P., Wang, I., Pinot, M., Manneville, J.B., Betz, T., Balland, M., Piel, M., 2013. ESCRT-III assembly and cytokinetic abscission are induced by tension release in the intercellular bridge. Science 339, 1625–1629. Langevin, J., Morgan, M.J., Sibarita, J.B., Aresta, S., Murthy, M., Schwarz, T., Camonis, J., Bellaiche, Y., 2005. Drosophila exocyst components Sec5, Sec6, and Sec15 regulate DE-Cadherin trafficking from recycling endosomes to the plasma membrane. Dev. Cell 9, 365–376. Lau, K., Tao, H., Liu, H., Wen, J., Sturgeon, K., Sorfazlian, N., Lazic, S., Burrows, J.T., Wong, M.D., Li, D., Deimling, S., Ciruna, B., Scott, I., Simmons, C., Henkelman, R.M., Williams, T., Hadjantonakis, A.K., Fernandez-Gonzalez, R., Sun, Y., Hopyan, S., 2015. Anisotropic stress orients remodelling of mammalian limb bud ectoderm. Nat. Cell Biol. 17, 569–579. le Duc, Q., Shi, Q., Blonk, I., Sonnenberg, A., Wang, N., Leckband, D., de Rooij, J., 2010. Vinculin potentiates E-cadherin mechanosensing and is recruited to actin-anchored sites within adherens junctions in a myosin II-dependent manner. J. Cell Biol. 189, 1107–1115. Lecuit, T., Yap, A.S., 2015. E-cadherin junctions as active mechanical integrators in tissue dynamics. Nat. Cell Biol. 17, 533–539. Lee, H.H., Elia, N., Ghirlando, R., Lippincott-Schwartz, J., Hurley, J.H., 2008. Midbody targeting of the ESCRT machinery by a noncanonical coiled coil in CEP55. Science (New York, N.Y.) 322, 576–580. Lekomtsev, S., Su, K.-C., Pye, V.E., Blight, K., Sundaramoorthy, S., Takaki, T., Collinson, L.M., Cherepanov, P., Divecha, N., Petronczki, M., 2012. Centralspindlin links the mitotic spindle to the plasma membrane during cytokinesis. Nature 492, 276–279. Levayer, R., Pelissier-Monier, A., Lecuit, T., 2011. Spatial regulation of Dia and Myosin-II by RhoGEF2 controls initiation of E-cadherin endocytosis during epithelial morphogenesis. Nat. Cell Biol. 13, 529–540. Lewellyn, L., Dumont, J., Desai, A., Oegema, K., 2010. Analyzing the effects of delaying aster separation on furrow formation during cytokinesis in the Caenorhabditis elegans embryo. Mol. Biol. Cell 21, 50–62. Li, D., Kuehn, E.W., Prekeris, R., 2014a. Kinesin-2 mediates apical endosome transport during epithelial lumen formation. Cell Logist. 4, e28928. Li, D., Mangan, A., Cicchini, L., Margolis, B., Prekeris, R., 2014b. FIP5 phosphorylation during mitosis regulates apical trafficking and lumenogenesis. EMBO Rep. 15, 428–437. Lin, M.C., Galletta, B.J., Sept, D., Cooper, J.A., 2010. Overlapping and distinct functions for cofilin, coronin and Aip1 in actin dynamics in vivo. J. Cell Sci. 123, 1329–1342. Liu, J., Fairn, G.D., Ceccarelli, D.F., Sicheri, F., Wilde, A., 2012. Cleavage furrow organization requires PIP2-mediated recruitment of anillin. Curr. Biol. 22, 64–69.

Asymmetric Cytokinesis in Animal Cells

339

Logan, M.R., Mandato, C.A., 2006. Regulation of the actin cytoskeleton by PIP2 in cytokinesis. Biol. Cell 98, 377–388. Lopez-Camacho, C., van Wijnen, A.J., Lian, J.B., Stein, J.L., Stein, G.S., 2014. Core binding factor beta (CBFbeta) is retained in the midbody during cytokinesis. J. Cell. Physiol. 229, 1466–1474. Loria, A., Longhini, K.M., Glotzer, M., 2012. The RhoGAP domain of CYK-4 has an essential role in RhoA activation. Curr. Biol. 22, 213–219. Lu, H., Bilder, D., 2005. Endocytic control of epithelial polarity and proliferation in Drosophila. Nat. Cell Biol. 7, 1232–1239. Lu, M.S., Johnston, C.A., 2013. Molecular pathways regulating mitotic spindle orientation in animal cells. Development 140, 1843–1856. Mabuchi, I., Okuno, M., 1977. The effect of myosin antibody on the division of starfish blastomeres. J. Cell Biol. 74, 251–263. Maddox, A.S., Habermann, B., Desai, A., Oegema, K., 2005. Distinct roles for two C. elegans anillins in the gonad and early embryo. Development 132, 2837–2848. Maddox, A.S., Lewellyn, L., Desai, A., Oegema, K., 2007. Anillin and the septins promote asymmetric ingression of the cytokinetic furrow. Dev. Cell 12, 827–835. Makino, A., Abe, M., Murate, M., Inaba, T., Yilmaz, N., Hullin-Matsuda, F., Kishimoto, T., Schieber, N.L., Taguchi, T., Arai, H., Anderluh, G., Parton, R.G., Kobayashi, T., 2015. Visualization of the heterogeneous membrane distribution of sphingomyelin associated with cytokinesis, cell polarity, and sphingolipidosis. Faseb J. 29, 477–493. Martin-Belmonte, F., Gassama, A., Datta, A., Yu, W., Rescher, U., Gerke, V., Mostov, K., 2007. PTEN-mediated apical segregation of phosphoinositides controls epithelial morphogenesis through Cdc42. Cell 128, 383–397. Martinez-Garay, I., Rustom, A., Gerdes, H.-H., Kutsche, K., 2006. The novel centrosomal associated protein CEP55 is present in the spindle midzone and the midbody. Genomics 87, 243–253. Marzesco, A.M., Janich, P., Wilsch-Brauninger, M., Dubreuil, V., Langenfeld, K., Corbeil, D., Huttner, W.B., 2005. Release of extracellular membrane particles carrying the stem cell marker prominin-1 (CD133) from neural progenitors and other epithelial cells. J. Cell Sci. 118, 2849–2858. Matias, N.R., Mathieu, J., Huynh, J.-R., 2015. Abscission is regulated by the ESCRT-III protein shrub in Drosophila germline stem cells. PLoS Genet. 11, e1004653. Matsuo, M., Shimodaira, T., Kasama, T., Hata, Y., Echigo, A., Okabe, M., Arai, K., Makino, Y., Niwa, S.-I., Saya, H., Kishimoto, T., 2013. Katanin p60 contributes to microtubule instability around the midbody and facilitates cytokinesis in rat cells. PLoS One 8, e80392. Mavrakis, M., Azou-Gros, Y., Tsai, F.C., Alvarado, J., Bertin, A., Iv, F., Kress, A., Brasselet, S., Koenderink, G.H., Lecuit, T., 2014. Septins promote F-actin ring formation by crosslinking actin filaments into curved bundles. Nat. Cell Biol. 16, 322–334. McGill, M.A., McKinley, R.F., Harris, T.J., 2009. Independent cadherin-catenin and Bazooka clusters interact to assemble adherens junctions. J. Cell Biol. 185, 787–796. McMahon, L., Legouis, R., Vonesch, J.L., Labouesse, M., 2001. Assembly of C. elegans apical junctions involves positioning and compaction by LET-413 and protein aggregation by the MAGUK protein DLG-1. J. Cell Sci. 114, 2265–2277. Menon, M.B., Gaestel, M., 2015. Sep(t)arate or not—how some cells take septinindependent routes through cytokinesis. J. Cell Sci. 128, 1877–1886. Menon, M.B., Sawada, A., Chaturvedi, A., Mishra, P., Schuster-Gossler, K., Galla, M., Schambach, A., Gossler, A., Forster, R., Heuser, M., Kotlyarov, A., Kinoshita, M., Gaestel, M., 2014. Genetic deletion of SEPT7 reveals a cell type-specific role of septins in microtubule destabilization for the completion of cytokinesis. PLoS Genet. 10, e1004558.

340

C. Thieleke-Matos et al.

Mierzwa, B., Gerlich, D.W., 2014. Cytokinetic abscission: molecular mechanisms and temporal control. Dev. Cell 31, 525–538. Miller, A.L., Bement, W.M., 2009. Regulation of cytokinesis by Rho GTPase flux. Nat. Cell Biol. 11, 71–77. Miller, K.G., Field, C.M., Alberts, B.M., 1989. Actin-binding proteins from Drosophila embryos: a complex network of interacting proteins detected by F-actin affinity chromatography. J. Cell Biol. 109, 2963–2975. Mishima, M., Kaitna, S., Glotzer, M., 2002. Central spindle assembly and cytokinesis require a kinesin-like protein/RhoGAP complex with microtubule bundling activity. Dev. Cell 2, 41–54. Morais-de-Sa, E., Sunkel, C., 2013. Adherens junctions determine the apical position of the midbody during follicular epithelial cell division. EMBO Rep. 14, 696–703. Morais-de-Sa, E., Mirouse, V., St Johnston, D., 2010. aPKC phosphorylation of Bazooka defines the apical/lateral border in Drosophila epithelial cells. Cell 141, 509–523. Morita, E., Sandrin, V., Chung, H.-Y., Morham, S.G., Gygi, S.P., Rodesch, C.K., Sundquist, W.I., 2007. Human ESCRT and ALIX proteins interact with proteins of the midbody and function in cytokinesis. EMBO J. 26, 4215–4227. Mostowy, S., Cossart, P., 2012. Septins: the fourth component of the cytoskeleton. Nat. Rev. Mol. Cell Biol. 13, 183–194. Murthy, K., Wadsworth, P., 2005. Myosin-II-dependent localization and dynamics of F-actin during cytokinesis. Curr. Biol. 15, 724–731. Murthy, K., Wadsworth, P., 2008. Dual role for microtubules in regulating cortical contractility during cytokinesis. J. Cell Sci. 121, 2350–2359. Nagata, K., Kawajiri, A., Matsui, S., Takagishi, M., Shiromizu, T., Saitoh, N., Izawa, I., Kiyono, T., Itoh, T.J., Hotani, H., Inagaki, M., 2003. Filament formation of MSF-A, a mammalian septin, in human mammary epithelial cells depends on interactions with microtubules. J. Biol. Chem. 278, 18538–18543. Naito, Y., Okada, M., Yagisawa, H., 2006. Phospholipase C isoforms are localized at the cleavage furrow during cytokinesis. J. Biochem. 140, 785–791. Neufeld, T.P., Rubin, G.M., 1994. The Drosophila peanut gene is required for cytokinesis and encodes a protein similar to yeast putative bud neck filament proteins. Cell 77, 371–379. Niederman, R., Pollard, T.D., 1975. Human platelet myosin. II. In vitro assembly and structure of myosin filaments. J. Cell Biol. 67, 72–92. Nguyen, T.Q., Sawa, H., Okano, H., White, J.G., 2000. The C. elegans septin genes, unc59 and unc-61, are required for normal postembryonic cytokineses and morphogenesis but have no essential function in embryogenesis. J. Cell Sci. 113, 3825–3837. Niiya, F., Tatsumoto, T., Lee, K.S., Miki, T., 2006. Phosphorylation of the cytokinesis regulator ECT2 at G2/M phase stimulates association of the mitotic kinase Plk1 and accumulation of GTP-bound RhoA. Oncogene 25, 827–837. Nishimura, Y., Nakano, K., Mabuchi, I., 1998. Localization of Rho GTPase in sea urchin eggs. FEBS Lett. 441, 121–126. Oegema, K., Savoian, M.S., Mitchison, T.J., Field, C.M., 2000. Functional analysis of a human homologue of the Drosophila actin binding protein anillin suggests a role in cytokinesis. J. Cell Biol. 150, 539–552. Olivier, N., Luengo-Oroz, M.A., Duloquin, L., Faure, E., Savy, T., Veilleux, I., Solinas, X., Debarre, D., Bourgine, P., Santos, A., Peyrieras, N., Beaurepaire, E., 2010. Cell lineage reconstruction of early zebrafish embryos using label-free nonlinear microscopy. Science 329, 967–971. Olmos, Y., Carlton, J.G., 2016. The ESCRT machinery: new roles at new holes. Curr. Opin. Cell Biol. 38, 1–11.

Asymmetric Cytokinesis in Animal Cells

341

Ono, S., 2003. Regulation of actin filament dynamics by actin depolymerizing factor/cofilin and actin-interacting protein 1: new blades for twisted filaments. Biochemistry 42, 13363–13370. Otomo, T., Otomo, C., Tomchick, D.R., Machius, M., Rosen, M.K., 2005. Structural basis of Rho GTPase-mediated activation of the formin mDia1. Mol. Cell 18, 273–281. Ou, G., Gentili, C., Gonczy, P., 2014. Stereotyped distribution of midbody remnants in early C. elegans embryos requires cell death genes and is dispensable for development. Cell Res. 24, 251–253. Packard, A., Georgas, K., Michos, O., Riccio, P., Cebrian, C., Combes, A.N., Ju, A., FerrerVaquer, A., Hadjantonakis, A.K., Zong, H., Little, M.H., Costantini, F., 2013. Luminal mitosis drives epithelial cell dispersal within the branching ureteric bud. Dev. Cell 27, 319–330. Padmanabhan, A., Rao, M.V., Wu, Y., Zaidel-Bar, R., 2015. Jack of all trades: functional modularity in the adherens junction. Curr. Opin. Cell Biol. 36, 32–40. Palacios, F., Tushir, J.S., Fujita, Y., D’Souza-Schorey, C., 2005. Lysosomal targeting of E-cadherin: a unique mechanism for the down-regulation of cell-cell adhesion during epithelial to mesenchymal transitions. Mol. Cell. Biol. 25, 389–402. Paolini, A., Duchemin, A.L., Albadri, S., Patzel, E., Bornhorst, D., Gonzalez Avalos, P., Lemke, S., Machate, A., Brand, M., Sel, S., Di Donato, V., Del Bene, F., Zolessi, F.R., Ramialison, M., Poggi, L., 2015. Asymmetric inheritance of the apical domain and self-renewal of retinal ganglion cell progenitors depend on anillin function. Development 142, 832–839. Pavicic-Kaltenbrunner, V., Mishima, M., Glotzer, M., 2007. Cooperative assembly of CYK4/MgcRacGAP and ZEN-4/MKLP1 to form the centralspindlin complex. Mol. Biol. Cell 18, 4992–5003. Pernier, J., Shekhar, S., Jegou, A., Guichard, B., Carlier, M.-F., 2016. Profilin interaction with actin filament barbed end controls dynamic instability, capping, branching, and motility. Dev. Cell 36, 201–214. Piekny, A.J., Glotzer, M., 2008. Anillin is a scaffold protein that links RhoA, actin, and myosin during cytokinesis. Curr. Biol. 18, 30–36. Piekny, A.J., Maddox, A.S., 2010. The myriad roles of anillin during cytokinesis. Semin. Cell Dev. Biol. 21, 881–891. Pollard, T.D., 1982. Structure and polymerization of Acanthamoeba myosin-II filaments. J. Cell Biol. 95, 816–825. Pollarolo, G., Schulz, J.G., Munck, S., Dotti, C.G., 2011. Cytokinesis remnants define first neuronal asymmetry in vivo. Nat. Neurosci. 14, 1525–1533. Prokopenko, S.N., Brumby, A., O’Keefe, L., Prior, L., He, Y., Saint, R., Bellen, H.J., 1999. A putative exchange factor for Rho1 GTPase is required for initiation of cytokinesis in Drosophila. Genes Dev. 13 (17), 2301–2314. Rakshit, S., Zhang, Y., Manibog, K., Shafraz, O., Sivasankar, S., 2012. Ideal, catch, and slip bonds in cadherin adhesion. Proc. Natl. Acad. Sci. U.S.A. 109, 18815–18820. Rappaport, R., 1985. Repeated furrow formation from a single mitotic apparatus in cylindrical sand dollar eggs. J. Exp. Zool. 234, 167–171. Reid, E., Connell, J., Edwards, T.L., Duley, S., Brown, S.E., Sanderson, C.M., 2005. The hereditary spastic paraplegia protein spastin interacts with the ESCRT-III complexassociated endosomal protein CHMP1B. Hum. Mol. Genet. 14, 19–38. Reina, J., Gonzalez, C., 2014. When fate follows age: unequal centrosomes in asymmetric cell division. Philos. Trans. R. Soc. Lond. B Biol. Sci. 369, 20130466. Reinsch, S., Karsenti, E., 1994. Orientation of spindle axis and distribution of plasma membrane proteins during cell division in polarized MDCKII cells. J. Cell Biol. 126, 1509–1526.

342

C. Thieleke-Matos et al.

Riento, K., Ridley, A.J., 2003. ROCKs: multifunctional kinases in cell behaviour. Nat. Rev. Mol. Cell Biol. 4, 446–456. Rodriguez-Boulan, E., Macara, I.G., 2014. Organization and execution of the epithelial polarity programme. Nat. Rev. Mol. Cell Biol. 15, 225–242. Roeth, J.F., Sawyer, J.K., Wilner, D.A., Peifer, M., 2009. Rab11 helps maintain apical crumbs and adherens junctions in the Drosophila embryonic ectoderm. PLoS One 4, e7634. Rosa, A., Vlassaks, E., Pichaud, F., Baum, B., 2015. Ect2/Pbl acts via Rho and polarity proteins to direct the assembly of an isotropic actomyosin cortex upon mitotic entry. Dev. Cell 32, 604–616. Rose, R., Weyand, M., Lammers, M., Ishizaki, T., Ahmadian, M.R., Wittinghofer, A., 2005. Structural and mechanistic insights into the interaction between Rho and mammalian Dia. Nature 435, 513–518. Rotty, J.D., Wu, C., Bear, J.E., 2013. New insights into the regulation and cellular functions of the ARP2/3 complex. Nat. Rev. Mol. Cell Biol. 14, 7–12. Roubinet, C., Cabernard, C., 2014. Control of asymmetric cell division. Curr. Opin. Cell Biol. 31, 84–91. Salzmann, V., Chen, C., Chiang, C.Y., Tiyaboonchai, A., Mayer, M., Yamashita, Y.M., 2014. Centrosome-dependent asymmetric inheritance of the midbody ring in Drosophila germline stem cell division. Mol. Biol. Cell 25, 267–275. Schluter, M.A., Pfarr, C.S., Pieczynski, J., Whiteman, E.L., Hurd, T.W., Fan, S., Liu, C.J., Margolis, B., 2009. Trafficking of Crumbs3 during cytokinesis is crucial for lumen formation. Mol. Biol. Cell 20, 4652–4663. Schonegg, S., Hyman, A.A., 2006. CDC-42 and RHO-1 coordinate acto-myosin contractility and PAR protein localization during polarity establishment in C. elegans embryos. Development 133, 3507–3516. Schroeder, T.E., 1968. Cytokinesis: filaments in the cleavage furrow. Exp. Cell Res. 53, 272–276. Schroeder, T.E., 1972. The contractile ring. II. Determining its brief existence, volumetric changes, and vital role in cleaving Arbacia eggs. J. Cell Biol. 53, 419–434. Schulze, J., Schierenberg, E., 2008. Cellular pattern formation, establishment of polarity and segregation of colored cytoplasm in embryos of the nematode Romanomermis culicivorax. Dev. Biol. 315, 426–436. Severson, A.F., Baillie, D.L., Bowerman, B., 2002. A Formin Homology protein and a profilin are required for cytokinesis and Arp2/3-independent assembly of cortical microfilaments in C. elegans. Curr. Biol. 12, 2066–2075. Singh, D., Pohl, C., 2014. Coupling of rotational cortical flow, asymmetric midbody positioning, and spindle rotation mediates dorsoventral axis formation in C. elegans. Dev. Cell 28, 253–267. Skop, A.R., Liu, H., Yates, J., Meyer, B.J., Heald, R., 2004. Dissection of the mammalian midbody proteome reveals conserved cytokinesis mechanisms. Science (New York, N.Y.) 305, 61–66. Sonnichsen, B., Koski, L.B., Walsh, A., Marschall, P., Neumann, B., Brehm, M., Alleaume, A.M., Artelt, J., Bettencourt, P., Cassin, E., Hewitson, M., Holz, C., Khan, M., Lazik, S., Martin, C., Nitzsche, B., Ruer, M., Stamford, J., Winzi, M., Heinkel, R., Roder, M., Finell, J., Hantsch, H., Jones, S.J., Jones, M., Piano, F., Gunsalus, K.C., Oegema, K., Gonczy, P., Coulson, A., Hyman, A.A., Echeverri, C.J., 2005. Full-genome RNAi profiling of early embryogenesis in Caenorhabditis elegans. Nature 434, 462–469. Spiliotis, E.T., Kinoshita, M., Nelson, W.J., 2005. A mitotic septin scaffold required for Mammalian chromosome congression and segregation. Science 307, 1781–1785. Stevens, N.R., Raposo, A.A., Basto, R., St Johnston, D., Raff, J.W., 2007. From stem cell to embryo without centrioles. Curr. Biol. 17, 1498–1503.

Asymmetric Cytokinesis in Animal Cells

343

Straight, A.F., Cheung, A., Limouze, J., Chen, I., Westwood, N.J., Sellers, J.R., Mitchison, T.J., 2003. Dissecting temporal and spatial control of cytokinesis with a myosin II inhibitor. Science 299, 1743–1747. Straight, A.F., Field, C.M., Mitchison, T.J., 2005. Anillin binds nonmuscle myosin II and regulates the contractile ring. Mol. Biol. Cell 16, 193–201. Suarez, C., Carroll, R.T., Burke, T.A., Christensen, J.R., Bestul, A.J., Sees, J.A., James, M.L., Sirotkin, V., Kovar, D.R., 2015. Profilin regulates F-actin network homeostasis by favoring formin over Arp2/3 complex. Dev. Cell 32, 43–53. Sun, H.S., Wilde, A., Harrison, R.E., 2011. Chlamydia trachomatis inclusions induce asymmetric cleavage furrow formation and ingression failure in host cells. Mol. Cell. Biol. 31, 5011–5022. Sun, L., Guan, R., Lee, I.J., Liu, Y., Chen, M., Wang, J., Wu, J.-Q., Chen, Z., 2015. Mechanistic insights into the anchorage of the contractile ring by anillin and Mid1. Dev. Cell 33, 413–426. Surka, M.C., Tsang, C.W., Trimble, W.S., 2002. The mammalian septin MSF localizes with microtubules and is required for completion of cytokinesis. Mol. Biol. Cell 13, 3532–3545. Suzuki, A., Hirata, M., Kamimura, K., Maniwa, R., Yamanaka, T., Mizuno, K., Kishikawa, M., Hirose, H., Amano, Y., Izumi, N., Miwa, Y., Ohno, S., 2004. aPKC acts upstream of PAR-1b in both the establishment and maintenance of mammalian epithelial polarity. Curr. Biol. 14, 1425–1435. Swan, K.A., Severson, A.F., Carter, J.C., Martin, P.R., Schnabel, H., Schnabel, R., Bowerman, B., 1998. cyk-1: a C. elegans FH gene required for a late step in embryonic cytokinesis. J. Cell Sci. 111, 2017–2027. Sydor, A.M., Czymmek, K.J., Puchner, E.M., Mennella, V., 2015. Super-resolution microscopy: from single molecules to supramolecular assemblies. Trends Cell Biol. 25, 730–748. Takaishi, K., Sasaki, T., Kameyama, T., Tsukita, S., Tsukita, S., Takai, Y., 1995. Translocation of activated Rho from the cytoplasm to membrane ruffling area, cell-cell adhesion sites and cleavage furrows. Oncogene 11, 39–48. Takekuni, K., Ikeda, W., Fujito, T., Morimoto, K., Takeuchi, M., Monden, M., Takai, Y., 2003. Direct binding of cell polarity protein PAR-3 to cell-cell adhesion molecule nectin at neuroepithelial cells of developing mouse. J. Biol. Chem. 278, 5497–5500. Tanaka-Takiguchi, Y., Kinoshita, M., Takiguchi, K., 2009. Septin-mediated uniform bracing of phospholipid membranes. Curr. Biol. 19, 140–145. Taneja, N., Fenix, A.M., Rathbun, L., Millis, B.A., Tyska, M.J., Hehnly, H., Burnette, D.T., 2016. Focal adhesions control cleavage furrow shape and spindle tilt during mitosis. Sci. Rep. 6, 29846. Tatsumoto, T., Xie, X., Blumenthal, R., Okamoto, I., Miki, T., 1999. Human ECT2 is an exchange factor for Rho GTPases, phosphorylated in G2/M phases, and involved in cytokinesis. J. Cell Biol. 147, 921–928. Tawk, M., Araya, C., Lyons, D.A., Reugels, A.M., Girdler, G.C., Bayley, P.R., Hyde, D.R., Tada, M., Clarke, J.D., 2007. A mirror-symmetric cell division that orchestrates neuroepithelial morphogenesis. Nature 446, 797–800. Tischer, D., Weiner, O.D., 2014. Illuminating cell signalling with optogenetic tools. Nat. Rev. Mol. Cell Biol. 15, 551–558. Truong Quang, B.A., Mani, M., Markova, O., Lecuit, T., Lenne, P.F., 2013. Principles of E-cadherin supramolecular organization in vivo. Curr. Biol. 23, 2197–2207. Tse, Y.C., Werner, M., Longhini, K.M., Labbe, J.C., Goldstein, B., Glotzer, M., 2012. RhoA activation during polarization and cytokinesis of the early Caenorhabditis elegans embryo is differentially dependent on NOP-1 and CYK-4. Mol. Biol. Cell 23, 4020–4031. Verma, S., Shewan, A.M., Scott, J.A., Helwani, F.M., den Elzen, N.R., Miki, H., Takenawa, T., Yap, A.S., 2004. Arp2/3 activity is necessary for efficient formation of E-cadherin adhesive contacts. J. Biol. Chem. 279, 34062–34070.

344

C. Thieleke-Matos et al.

Vicente-Manzanares, M., Ma, X., Adelstein, R.S., Horwitz, A.R., 2009. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol. Cell Biol. 10, 778–790. Walther, R.F., Pichaud, F., 2010. Crumbs/DaPKC-dependent apical exclusion of Bazooka promotes photoreceptor polarity remodeling. Curr. Biol. 20, 1065–1074. Wang, T., Yanger, K., Stanger, B.Z., Cassio, D., Bi, E., 2014. Cytokinesis defines a spatial landmark for hepatocyte polarization and apical lumen formation. J. Cell Sci. 127, 2483–2492. Watanabe, N., Madaule, P., Reid, T., Ishizaki, T., Watanabe, G., Kakizuka, A., Saito, Y., Nakao, K., Jockusch, B.M., Narumiya, S., 1997. p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. Embo J. 16, 3044–3056. Watanabe, S., Ando, Y., Yasuda, S., Hosoya, H., Watanabe, N., Ishizaki, T., Narumiya, S., 2008. mDia2 induces the actin scaffold for the contractile ring and stabilizes its position during cytokinesis in NIH 3T3 cells. Mol. Biol. Cell 19, 2328–2338. Watanabe, S., Okawa, K., Miki, T., Sakamoto, S., Morinaga, T., Segawa, K., Arakawa, T., Kinoshita, M., Ishizaki, T., Narumiya, S., 2010. Rho and anillin-dependent control of mDia2 localization and function in cytokinesis. Mol. Biol. Cell 21, 3193–3204. Wei, S.Y., Escudero, L.M., Yu, F., Chang, L.H., Chen, L.Y., Ho, Y.H., Lin, C.M., Chou, C.S., Chia, W., Modolell, J., Hsu, J.C., 2005. Echinoid is a component of adherens junctions that cooperates with DE-Cadherin to mediate cell adhesion. Dev. Cell 8, 493–504. Weirich, C.S., Erzberger, J.P., Barral, Y., 2008. The septin family of GTPases: architecture and dynamics. Nat. Rev. Mol. Cell Biol. 9, 478–489. Wheeler, R.J., Scheumann, N., Wickstead, B., Gull, K., Vaughan, S., 2013. Cytokinesis in Trypanosoma brucei differs between bloodstream and tsetse trypomastigote forms: implications for microtubule-based morphogenesis and mutant analysis. Mol. Microbiol. 90, 1339–1355. Willard, F.S., Crouch, M.F., 2001. MEK, ERK, and p90RSK are present on mitotic tubulin in Swiss 3T3 cells: a role for the MAP kinase pathway in regulating mitotic exit. Cell. Signal. 13, 653–664. Willenborg, C., Jing, J., Wu, C., Matern, H., Schaack, J., Burden, J., Prekeris, R., 2011. Interaction between FIP5 and SNX18 regulates epithelial lumen formation. J. Cell Biol. 195, 71–86. Woichansky, I., Beretta, C.A., Berns, N., Riechmann, V., 2016. Three mechanisms control E-cadherin localization to the zonula adherens. Nat. Commun. 7, 10834. Wolfe, B.A., Takaki, T., Petronczki, M., Glotzer, M., 2009. Polo-like kinase 1 directs assembly of the HsCyk-4 RhoGAP/Ect2 RhoGEF complex to initiate cleavage furrow formation. PLoS Biol. 7, e1000110. Wu, Y., Kanchanawong, P., Zaidel-Bar, R., 2015. Actin-delimited adhesion-independent clustering of E-cadherin forms the nanoscale building blocks of adherens junctions. Dev. Cell 32, 139–154. Yamashiro, S., Totsukawa, G., Yamakita, Y., Sasaki, Y., Madaule, P., Ishizaki, T., Narumiya, S., Matsumura, F., 2003. Citron kinase, a Rho-dependent kinase, induces di-phosphorylation of regulatory light chain of myosin II. Mol. Biol. Cell 14, 1745–1756. Yamashita, Y.M., Jones, D.L., Fuller, M.T., 2003. Orientation of asymmetric stem cell division by the APC tumor suppressor and centrosome. Science 301, 1547–1550. Yamashita, Y.M., Mahowald, A.P., Perlin, J.R., Fuller, M.T., 2007. Asymmetric inheritance of mother versus daughter centrosome in stem cell division. Science 315, 518–521. Yang, D., Rismanchi, N., Renvoise, B., Lippincott-Schwartz, J., Blackstone, C., Hurley, J.H., 2008. Structural basis for midbody targeting of spastin by the ESCRTIII protein CHMP1B. Nat. Struct. Mol. Biol. 15, 1278–1286.

Asymmetric Cytokinesis in Animal Cells

345

Yao, M., Qiu, W., Liu, R., Efremov, A.K., Cong, P., Seddiki, R., Payre, M., Lim, C.T., Ladoux, B., Mege, R.M., Yan, J., 2014. Force-dependent conformational switch of alpha-catenin controls vinculin binding. Nat. Commun. 5, 4525. Yonemura, S., Wada, Y., Watanabe, T., Nagafuchi, A., Shibata, M., 2010. alpha-Catenin as a tension transducer that induces adherens junction development. Nat. Cell Biol. 12, 533–542. Yuce, O., Piekny, A., Glotzer, M., 2005. An ECT2-centralspindlin complex regulates the localization and function of RhoA. J. Cell Biol. 170, 571–582. Zhang, D., Glotzer, M., 2015. The RhoGAP activity of CYK-4/MgcRacGAP functions non-canonically by promoting RhoA activation during cytokinesis. Elife 4, e08898. Zhao, W.-M., Seki, A., Fang, G., 2006. Cep55, a microtubule-bundling protein, associates with centralspindlin to control the midbody integrity and cell abscission during cytokinesis. Mol. Biol. Cell 17, 3881–3896. Zou, Y., Shao, Z., Peng, J., Li, F., Gong, D., Wang, C., Zuo, X., Zhang, Z., Wu, J., Shi, Y., Gong, Q., 2014. Crystal structure of triple-BRCT-domain of ECT2 and insights into the binding characteristics to CYK-4. FEBS Lett. 588, 2911–2920.