SUMOylation and Ubiquitination: Current and Emerging Concepts [1 ed.] 9781912530137, 9781912530120

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SUMOylation and Ubiquitination Current and Emerging Concepts

Edited by

Van G. Wilson Caister Academic Press

SUMOylation and Ubiquitination Current and Emerging Concepts

https://doi.org/10.21775/9781912530120

Edited by Van G. Wilson Department of Microbial Pathogenesis and Immunology College of Medicine Texas A&M University Bryan, TX USA

Caister Academic Press

Copyright © 2019 Caister Academic Press Norfolk, UK www.caister.com British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-1-912530-12-0 (paperback) ISBN: 978-1-912530-13-7 (ebook) Description or mention of instrumentation, software, or other products in this book does not imply endorsement by the author or publisher. The author and publisher do not assume responsibility for the validity of any products or procedures mentioned or described in this book or for the consequences of their use. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior permission of the publisher. No claim to original U.S. Government works. Cover design adapted from Figure 4.3 Ebooks Ebooks supplied to individuals are single-user only and must not be reproduced, copied, stored in a retrieval system, or distributed by any means, electronic, mechanical, photocopying, email, internet or otherwise. Ebooks supplied to academic libraries, corporations, government organizations, public libraries, and school libraries are subject to the terms and conditions specified by the supplier.

Contents

Part I  General Principles

1

1

The Rise of the Ubiquitin Super Family

2

Cracking the Ubiquitin Code: The Ubiquitin Toolbox

15

3

Recent Highlights: Onco Viral Exploitation of the SUMO System

35

4

Progress in the Discovery of Small Molecule Modulators of DeSUMOylation

51

Van G. Wilson

Monique P.C. Mulder, Katharina F. Witting and Huib Ovaa

Domenico Mattoscio, Alessandro Medda and Susanna Chiocca Shiyao Chen, Duoling Dong, Weixiang Xin and Huchen Zhou

Part II  Novel and Advancing Technologies 5

3

69

Identification of SUMOylated and Ubiquitinated Substrates by Mass Spectrometry71

Francis P. McManus and Pierre Thibault

6

Global Proteomic Profiling of SUMO and Ubiquitin

7

Biotin-based Approaches for the Study of Ubiquitin and Ubiquitinlike Protein Modifications

101

Screening Mammalian SUMOylated Proteins by Fluorescence Protein Reconstitution

123

Dissecting Complex SUMOylation Networks in Humans

135

TULIP: Targets of Ubiquitin Ligases Identified by Proteomics

147

Alla Ahmad, Ryan Lumpkin and Elizabeth A. Komives

James D. Sutherland, Orhi Barroso-Gomila and Rosa Barrio

8

Maki Komiya, Mizuki Endo and Takeaki Ozawa

9 10

Ijeoma Uzoma and Heng Zhu

Román González-Prieto and Alfred C.O. Vertegaal

95

iv  | Contents

Part III  Cellular Processes 11

161

Regulation of p53 Family Members by the Ubiquitin and SUMO Modification Systems

163

Interplay between the Ubiquitin Proteasome System and Mitochondria for Protein Homeostasis

193

13

Interplay of Ubiquitination and SUMOylation with miRNAs

217

14

The Role of Ubiquitination and SUMOylation in DNA Replication

231

15

Roles of Ubiquitination and SUMOylation in DNA Damage Response

263

16

The Role of Ubiquitination and SUMOylation in Telomere Biology

289

17

Role of Ubiquitin and SUMO in Intracellular Trafficking

303

18

Roles of Ubiquitination and SUMOylation in the Regulation of Angiogenesis

313

19

The Role of SUMOylation and Ubiquitination in Brain Ischaemia: Critical Concepts and Clinical Implications

331

20

The Role of Ubiquitination and SUMOylation in Autophagy

349

21

Ubiquitin and SUMO Modifications in Caenorhabditis elegans Stress Response363

Viola Calabrò and Maria Vivo

12

Mafalda Escobar-Henriques, Selver Altin and Fabian den Brave Yashika Agrawal and Manas Kumar Santra Tarek Abbas

Siyuan Su, Yanqiong Zhang and Pengda Liu

Michal Zalzman, W. Alex Meltzer, Benjamin A. Portney, Robert A. Brown and Aditi Gupta Maria Sundvall

Andrea Rabellino, Cristina Andreani and Pier Paolo Scaglioni

Joshua D. Bernstock, Daniel G. Ye, Dagoberto Estevez, Gustavo Chagoya, Ya-Chao Wang, Florian Gessler, John M. Hallenbeck and Wei Yang Sushil Devkota

Krzysztof Drabikowski

Part IV  Infection, Immunity and Disease 22

377

Beyond Degradation: Ubiquitination of the Inflammasome Regulates Assembly and Activity

379

23

Ubiquitin and SUMO in Antiviral Defence

389

24

Ubiquitination and SUMOylation in HIV Infection: Friends and Foes

417

Joseph S. Bednash and Rama K. Mallampalli Van G. Wilson

Marta Colomer-Lluch, Sergio Castro-Gonzalez and Ruth Serra-Moreno

Contents |  v

25

Ubiquitination and SUMOylation of Amyloid and Amyloid-like Proteins in Health and Disease

453

Keeping Up with the Pathogens: The Role of SUMOylation in Plant Immunity

489

Lenzie Ford, Luana Fioriti and Eric R. Kandel

26

Rebecca Morrell and Ari Sadanandom

Index501

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Full details at www.caister.com

Preface

The discovery of ubiquitin and the ubiquitin-proteasome system in the late 1970s provided elegant insight into protein degradation as the biochemical process for removing damaged or unwanted proteins. This seminal discovery that polyubiquitination of substrate proteins directed them to the proteasome for subsequent degradation led to the Nobel Prize in 2004 for Drs Aaron Ciechanover, Avram Hershko and Irwin Rose. During the roughly 25 years between the discovery of this process and the award of the Nobel Prize there was an explosion of research demonstrating the breadth and importance of this post-translational modification system. One of the perhaps less expected and more slowly recognized features was the role of polyubiquitin-mediated degradation as a regulatory mechanism for controlling the functional levels of individual proteins and of multi–protein complexes. Proteasomal degradation became not simply a device to remove ageing or defective proteins, but also a powerful system to control levels of functional proteins in complex pathways and thus rapidly modulate the activity of these pathways. In addition, like phosphorylation, ubiquitination became appreciated as a versatile modification that could affect substrate functions in non-degradative ways. Combined with the discovery of mono- and multi-ubiquitination, along with multiple types of linkages for branched forms of polyubiquitin, it became clear that the ubiquitin addition was highly complex with enormous combinatorial capacity. Some of this complexity also stems from the large number of distinct enzymes and co-factors involved in ubiquitin processing and transfer to substrates, with several hundred proteins known to function in this process. Because of this biochemical complexity and diversity of components it is not surprising

that, nearly 40 years later, we are still uncovering the novel features of the ubiquitin system, identifying more and more substrates, and elucidating key cellular regulatory steps controlled by this small, yet profoundly important, protein. A second exciting chapter in the ubiquitin story was the discovery in the late 1980s that there were a number of other ubiquitin-related proteins that together comprise the ubiquitin superfamily. Like ubiquitin, the other Ubiquitin-like proteins (Ubls) are covalently attached via their C-terminus to lysine residues in substrate proteins (although for a few family members ligation to substrates has not yet been established). Each member of the superfamily has its own specific set of enzymes that mediate the addition of the modifier to the substrate, although biochemically all members of the family undergo the same scheme of processing, activation, conjugation and eventual ligation to the substrate. Several members of the superfamily are still poorly characterized and several have fairly limited realms of substrates. However, one member of this super family, the Small Ubiquitin-like Modifier (SUMO) proteins, has been prominently investigated. In humans, there are five related SUMO proteins, SUMOs 1–5. SUMOs 1–3 are widely expressed and well characterized, whereas SUMOs 4 and 5 are more restricted and less is known about their functional roles. In contrast to the ubiquitin system, the SUMO system has far fewer components involved in processing and substrate modification. Nonetheless, sumoylation collectively has been shown to have a large and broad range of substrates (well over 3000 identified) and to be a critical modification during development, as well as for many normal cellular processes. Importantly, there is now wellestablished crosstalk between the ubiquitin and

viii  | Preface

SUMO systems through multiple mechanisms, including competition for the same target lysine in substrates, modification of substrates with both modifiers at different lysines, formation of mixed SUMO–ubiquitin polymers on some substrates, and degradation of substrates through targeting of ubiquitin to sumoylated proteins through SUMOtargeted ubiquitin ligases (STUbLs). The ability of these two modification pathways to function both independently or cooperatively on thousands of substrates is a remarkable observation that underscores just how widespread and entrenched this type of post-translational modification is in cell biology. In late 2017 I was approached about putting together a book on current and emerging concepts in the fields of sumoylation and ubiquitination. The previous 5 years had seen an explosion of new technologies for identifying substrates and mapping modification sites, so there was a wealth of novel biochemical, molecular and biological data about these two systems. All of these new data were perfect fodder for a book project, especially one

focused on the interplay between these systems, so the timing was ideal. Although certainly not allinclusive, I tried to identify topic areas for the book for which there were significant recent advances and/or strong evidence for a functional role of both SUMOs and ubiquitin. I would like to thank all of the authors who so graciously agreed to address these topics and provide chapters for this book. Your individual contributions to the book were uniformly excellent and provided wonderful reviews of your research areas. I hope that this final compilation will prove useful to both novice and seasoned investigators in these fields, and that future readers will learn as much about sumoylation and ubiquitination as I did. Van G. Wilson Department of Microbial Pathogenesis and Immunology College of Medicine Texas A&M University Bryan, TX USA

Part I

General Principles

The Rise of the Ubiquitin Super Family Van G. Wilson*

1

Department of Microbial Pathogenesis and Immunology, College of Medicine, Texas A&M University, Bryan, TX, USA. *Correspondence: [email protected] https://doi.org/10.21775/9781912530120.01

Abstract Ubiquitin and SUMOs are related small proteins that are members of the larger ubiquitin superfamily. Members of this family (Ubls) are post-translational modifiers that are covalently attached to target proteins through lysine residues in the target. Biochemically, the processing and conjugation of these modifiers is remarkably similar, although the individual enzymes and components are usually specific for their individual modifier. The modification process is dynamic with exact demodification occurring through specific proteases that remove the Ubls and restore the target proteins to their original state. Functionally, Ubl modification can influence many aspects of protein biology including activity, localization, stability, and interactions with partners. Importantly, examples of functional crosstalk between Ub and SUMO modifications have been observed, which provides exciting opportunities for combinatorial regulation of target proteins. This chapter will introduce basic principles of ubiquitinylation and sumoylation and will also provide a general overview of important terms and concepts that will be explored in more detail in the remaining chapters. Introduction: the history of discovery The discovery of ubiquitin was rooted in the quest to understand protein degradation. In particular, during the 1970s there was a burgeoning realization that cellular protein turnover could not solely be due to non-specific lysosomal degradative

activity. It was observed that proteins differed in their half-lives and degraded differently under various physiological conditions, which was not consistent with a simple, non-specific process. The quest to find alternative and more specific mechanisms spurred the investigation of new approaches. Ultimately, the development of cell-free rabbit reticulocyte extracts that could degrade proteins in an ATP-dependent fashion was a critical step that allowed the identification of ubiquitin and eventually the proteasome machinery; (Etlinger and Goldberg, 1977; Ciehanover et al., 1978, 1980; Hershko et al., 1979). It was determined that two components were necessary for degradation, the ubiquitin protein and the proteasome, which provided insight into the specificity question. It became apparent that the proteasome was relatively non-specific in its proteolytic activity, and that the specificity for degradation was at the level of ubiquitin conjugation to the substrate. Subsequent decades have elucidated a complex and exquisite control of ubiquitin conjugation through a family of enzymes that recognizes substrates and facilitates their modification (Saeki, 2017). This two-stage process was a critical concept that explained how proteins could remain in the same compartment with the degradative machinery, but without damage. Only when proteins are ubiquitin tagged at the appropriate time are they earmarked for proteolytic removal. The concept that ubiquitin could tag another protein by covalent conjugation was a seminal observation that expanded the repertoire of post-translational modifications beyond small modifiers such as phosphate or methyl groups.

4  | Wilson

Cloning of the yeast (Ozkaynak et al., 1984) and human (Baker and Board, 1987) ubiquitin genes quickly led to the realization that there were a number of other genes whose protein products shared significant homology with ubiquitin and which became the ubiquitin superfamily. The first of these other members to be studied was an interferon-stimulated gene product designated ISG15 (Haas et al., 1987). Like ubiquitin, ISG15 could conjugate to target proteins (Loeb and Haas, 1992). Similarly, four groups each independently identified another member of the ubiquitin superfamily in 1996, a protein that became known as SUMO (Boddy et al., 1996; Matunis et al., 1996; Okura et al., 1996; Shen et al., 1996). SUMO also shared the ability of ubiquitin and ISG15 to conjugate to target proteins, confirming that this is a common function of this type of protein. In subsequent years over 10 other ubiquitin related

proteins have been identified, though not all have been shown to have conjugative ability (Hochstrasser, 2009). Collectively these proteins are known as ubiquitin-like proteins or Ubls (Schwartz and Hochstrasser, 2003). Among the Ubls that have been characterized to some extent, it is clear that modification of substrates can have varied and important consequences, and that even ubiquitin has roles beyond protein degradation. This book will focus on two of the most highly characterized members of this superfamily, the original ubiquitin and the ubiquitous SUMO. Ubl proteins Ubiquitin, the prototype of the Ubls, is a 76 amino acid polypeptide in humans and yeast, and it is highly conserved with 96% homology between these species (Fig. 1.1). There are four human

Figure 1.1  Comparison of the sequence and structures of ubiquitin and SUMOs1–3. (A) Primary sequence comparison of ubiquitin and SUMOs1–3. The amino acid sequences of the four proteins are aligned for homology with the amino acid numbers shown on the right side of each line. Yellow highlights are residues identical in all four proteins, Blue highlights are residues conserved among the four proteins. Pale orange highlights are residues identical among the three SUMOs. (B) Three dimensional ribbon diagrams of ubiquitin and SUMOs1–3. The coloured helices and beta-sheets comprise the conserved ubiquitin fold domain.

The Rise of the Ubiquitin Super Family |  5

genes (UBB, UBC, UBA52, and RPS27A/UBA80) encoding ubiquitin, each producing a fusion protein where the functional ubiquitin moiety (Ub) must be proteolytically freed with an ubiquitin-specific protease. UBB and UBC encode polyubiquitin precursors while UBA52 and RPS27A/UBA80 encode single copy ubiquitin fused to different ribosomal proteins. This redundancy likely reflects both the essential nature of the ubiquitin system and the need for many copies of the ubiquitin protein. The released Ub has a diglycine motif at the C-terminus that is characteristic of conjugation functional Ubls. The diglycine is a critical element for the initiation of the enzymatic cascade leading to attachment of Ub to substrates with the covalent bond forming between the terminal glycine and a lysine residue on the substrate. Importantly, Ub itself contains seven lysine residues, and each of these lysines can be conjugated by other Ub molecules to form multimeric Ub chains (Meierhofer et al., 2008). Additionally, ubiquitin contains a three-dimensional core structure known as the β-grasp fold that is also conserved among the Ubls (Burroughs et al., 2007). The β-grasp fold is characterized by five anti-parallel β strands forming a beta sheet with a juxtaposed helical segment. Recognition of this β-grasp fold region by ubiquitin binding domains (UBDs) mediates many functional interactions between the ubiquitin system and cellular targets. Interestingly, this fold is evolutionarily ancient with extensive diversification in prokaryotes followed by even further expansion in eukaryotes (Burroughs et al., 2012). Like the Ub prototype, SUMO proteins (Fig. 1.1) are highly conserved (Chen et al., 1998), are produced in a precursor form ( Johnson et al., 1997), and are encoded by multiple genes (Wilson, 2017). In humans there are five SUMO proteins (1–5) encoded by separate genes with distinct expression patterns: SUMOs 1–3 are ubiquitously expressed in all cell types, while SUMO4 and SUMO5 have restricted expression patterns. SUMO4 is limited to renal, pancreatic, immune, and placental cells (Wang and She, 2008; Chen et al., 2014; Baczyk et al., 2017), while SUMO5 is found in testes and peripheral blood leucocytes (Liang et al., 2016). SUMO1 is expressed as a 101 amino acid precursor that is processed by removal of four C-terminal residues to generate the diglycine terminus. SUMO1 is 18% homologous to Ub at the amino acid sequence

level and 48% homologous at the tertiary structural level (Bayer et al., 1998). However, Ub is only a 76 amino acid polypeptide, and the difference resides in an N-terminal extension in SUMO that is absent in the shorter Ub protein. SUMO2 and SUMO3 are expressed as precursors of 95 and 103 amino acids, respectively. After processing, the final active forms are 93 and 92 amino acids, respectively, and differ by only three amino acids. SUMO 2/3 are so similar that they are often considered as one, but they only share 48% identity with SUMO1 so are a distinct subgroup with overlapping but different biological activities. SUMO4 is expressed as a 95 amino acid precursor with 85% amino acid identity to pre-SUMO2, and SUMO5 is predicted as a 101 amino acid precursor. Because of their very limited tissue expression, the functional targets of SUMO4 and SUMO5 have been less well studied and their biological role are less clear than for the widely expressed SUMO1–3 group (Guo et al., 2004, 2005; Liang et al., 2016). Ubl enzymology Covalent attachment of Ubls to their target substrates is a multiple-step process that is biochemically similar across all Ubls, but which uses distinct components which are Ubl-specific and which generally do not cross function with other Ubls The overall process can be divided into four basic steps (Fig. 1.2): (1) proteolytic processing of the precursor Ubls to their mature form, (2) activation of the Ubl in an ATP-dependent process resulting in covalent attachment of the Ubl to the activating enzyme (E1) via a thioester linkage, (3) covalent transfer of the Ubl to the conjugating enzyme (E2), again via a thioester linkage, and (4) covalent transfer of the Ubl to a lysine reside on the substrate via a isopeptide bond in a reaction utilizing a ligase (E3). Modification of substrates with Ubls is generally reversible by precise proteolytic removal of the Ubl to regenerate the unmodified substrate. The released Ubl can then reenter the modification cycle at step 2 and be reused. The first step for both Ub and SUMOs is proteolytic cleavage of the precursor forms to yield the active forms that all terminate with C-terminal diglycine motif. For Ub, processing is mediated by deubiquitinases (DUBs), while SUMOs are processed by SENPs (Sentrin proteases). There are

6  | Wilson

Figure 1.2  General scheme for substrate modification by ubiquitin and ubiquitin-like proteins (Ubl). E1, E2, and E3 refer respectively to the activating, conjugating, and ligase enzymes in the medication pathway. Ulps are the ubiquitin proteases involved in precursor processing and/or removal of the Ubl from the substrate. See text for details of the individual steps.

roughly 100 DUBs in mammals, though it is likely that only certain ones are involved in precursor processing with the rest functioning to remove Ub from substrates and/or to reduce poly-Ub chains back to Ub monomers. Some DUBs are classed as cysteine proteases while others are metalloproteases (Amerik and Hochstrasser, 2004). Recent work has implicated five DUBs (UCHL3, USP9X, USP7, USP5, and Otulin/Gumby/Fam105b) as predominantly active in precursor processing (Grou et al., 2015). There appears to be important redundancy in that for each of the four Ub precursor types there was at least two DUBs could cleave the precursor. Grou et al. (2015) also found that the poly-Ub precursors were cleaved through a combination of both co- and post-translational events while the Ub-ribosomal proteins fusions were primarily processed post-translationally. Clearly there is much more work needed to decipher the roles and functions of individual DUBs. In contrast to the 100 DUBs there are only six human SENPs, SENPs 1–3 and 5–7 (Nayak and Müller, 2014), and they are all cysteine proteases with a papain-like fold in the catalytic domain (Kunz et al., 2018). SENPs 1, 2, 3, and 5 evolved from the S. cerevisiae Ulp1 protease while SENPs 6 and 7 derive from the Ulp2 lineage (Hickey et al., 2012). The SENPs vary in their subcellular localization, a property which is determined primarily by

differences in their variable N-terminal regions. Splice variants of SENP2 and SENP7 also exist with different N-terminal sequences which adds further complexity to their localization ( Jiang et al., 2011; Bawa-Khalfe et al., 2012). For example, the long form of SENP2 is nuclear with association to PML bodies and nuclear pores, while a short form is cytosolic ( Jiang et al., 2011). Similarly, the long form of SENP7 is nuclear while the short form is found in the cytoplasm (Bawa-Khalfe et al., 2012). SENP1 shares a similar distribution to long form SENP2 (Chow et al., 2012), though both can shuttle to the cytoplasm (Bailey and O’Hare, 2005). In contrast, SENP3 and SENP5 are primarily localized to the nucleolus (Gong and Yeh, 2006; Haindl et al., 2008) while SENP6 and SENP7 reside mostly in the nucleoplasm with at least partial association with chromatin (Lima and Reverter, 2008; Maison et al., 2012). This differential localization likely contributes to differences in the ability of individual SENPs to desumoylate substrates in the cell. Note that the preferential localization of most of the SENPS to the nuclear compartment correlates with sumoylation being predominantly a nuclear process. In addition to differential localization, the individual SENPs differ in their innate ability to process SUMO precursors and to desumoylate SUMO1 versus SUMO2/3 modified substrates. With

The Rise of the Ubiquitin Super Family |  7

regard to precursor processing, most of the SENPs preferentially cleave preSUMO2/3 compared to preSUMO1 (Kolli et al., 2010), though the cleavage rates for SENP6 and SENP7 are very low suggesting that they may not be significantly involved in generation of the mature forms of the SUMOs. Only SENP1 exhibits a preference for cleaving preSUMO1, so it is the likely protease for processing this SUMO type (Xu and Au, 2005). Unfortunately, most of these studies involved in vitro experiments or transient in vivo expression systems, so the actual biological roles of the various SENPs in processing preSUMO1 versus preSUMO2/3 remain unclear. Similar to differences in processing preSUMOs, the SENPs also display differences in their ability to desumoylate substrates modified with SUMO1 as opposed to SUMO2/3. On model substrates, SENP1 and SENP2 desumoylate all 3 SUMO forms well (Reverter and Lima, 2004; Shen et al., 2006), though other evidence suggests that SENP1 primarily desumoylates SUMO1 conjugates in vivo while SENP2 is responsible for SUMO2/3 conjugates (Sharma et al., 2013). SENP3 and SENP5 also exhibit a preference for SUMO2/3 conjugates with little activity against SUMO1 modified substrates (Gong and Yeh, 2006; Kolli et al., 2010). Interestingly, SENP6 and SENP7 also prefer deconjugation of SUMO2 modified substrates, but their activity on mono-sumoylated proteins is weak and they show strong preference for cleavage of poly-SUMO2/3 chains (Drag et al., 2008; Shen et al., 2009; Kolli et al., 2010). The second step in the conjugation process is for the mature Ubl protein to be activated through a process the results in covalent attachment of the C-terminus of the Ubl to a cysteine residue in the activating enzyme (the E1 enzyme). For most organisms each Ubl system is activated by a single E1 enzyme, and these E1s are Ubl specific with little or no ability to activate non-cognate Ubls. For the SUMO system, the sole E1 enzyme is a heterodimer (SAE1/2). SAE1 and SAE2 are homologous to the N-terminus and C-terminus, respectively, of the classical ubiquitin E1 enzyme, UBE1. The larger subunit, SAE2, contains the catalytic domain, and both subunits contain nuclear localization signals to direct this E1 to the nuclear compartment (Moutty et al., 2011). SAE2 itself is sumoylated in the nucleus at five sites, and this sumoylation is important to prevent nucleocytoplasmic shuttling

and to retain the SAE1/2 complex in the nucleus (Truong et al., 2012). Unlike the SUMO E1 enzyme, the canonical E1 for the Ub system, UBE1 (also known as UBA1), is a monomeric protein, though two isoforms of 110 kDa and 117 kDa have been reported (Cook and Chock, 1995). UBE1 has three distinct domains: (1) an adenylation domain for binding ATP and Ub, (2) a catalytic domain with the cysteine where Ub will be covalently attached, and (3) an ubiquitin-fold domain in the C-terminus which functions to bind the E2 enzymes. The catalytic mechanism of UBE1 has been extensively studied, and a detail review can be found in Schulman and Harper (Schulman and Harper, 2009). Estimates suggest that UBE1 is responsible for greater than 99% of the Ub activation ( Jin et al., 2007). The minor fraction of Ub not activated by UBE1 uses a second E1 enzyme known as UBE1L2 or UBA6 (Chiu et al., 2007; Pelzer et al., 2007). UBA6 is only about 40% identical to UBE1, and surprisingly can activate not only Ub but also another Ubl, FAT10 (Chiu et al., 2007). Since UBE1 and UBA6 interact with only partially overlapping pools of E2 enzymes, this adds another level of complexity to regulation of Ub modification (Groettrup et al., 2008). Subsequent to the activation step the Ubl is transferred from the E1 enzyme to a cysteine on the conjugation enzyme (E2). At this step there are dramatic differences in the Ub and SUMO systems. For sumoylation there is a single E2 conjugating enzyme called Ubc9, while for ubiquitination there are more than 40 human E2 enzymes (Wenzel et al., 2011) that are classified into 17 subfamilies (Michelle et al., 2009). The Ub E2 enzymes share a common ubiquitin-conjugating (UBC) domain within which sits the catalytic cysteine. The UBCs not only share sequence homology, but also have a common tertiary structure with a four strand β-sheet and four α-helixes (Lin et al., 2002). Also within the active site is either a serine or an aspartate residue; E2s with the aspartate are constitutively active while those with the serine must be phosphorylated to stimulate E2 activity (Valimberti et al., 2015). The UBC domain further contains determinants for E2–E1 interactions, though resides outside the UBC domain can also contribute to discrimination between the two Ub E1s. Notably, Ub E2s only bind the two Ub E1 enzymes with high affinity when they are charged

8  | Wilson

with Ub, and do not interact with E1s for other Ubl proteins (Huang et al., 2007). Like the multiple Ub E2s, the sole SUMO E2 (Ubc9) contains the common tertiary folding feature containing the active site cysteine at residue 93 (Tong et al., 1997). Again, binding determinants within Ubc9 are specific for interaction with its cognate E1, and Ubc9 does not recognize the Ub E1s. The final step in the modification cycle is the engagement of a charged E2 with an E3 ligase and the transfer of the Ub or SUMO moiety to the final substrate. There are hundreds of human Ub E3s while only a few SUMO E3s have been identified (Cappadocia and Lima, 2018). The Ub E3s fall into three main classes, RING, HECT, and RBR proteins (Buetow and Huang, 2016), with the remaining E3s designated as atypical. Members of the RING family function through direct transfer of Ub from the E2–Ub complex to the substrate without attachment to the RING E3. In contrast, members of the other families have an active site cysteine that receives Ub from the E2 to form a thioester linkage before transferring the Ub to the substrate. It is the E3s themselves that confer substrate specificity via determinants that recognize distinct target proteins or direct the E3 to subcellular locations where it can modify accessible proteins. Choice of the specific lysine to be modified by Ub likely is determined at least in part by sequence contexts in the substrate (Radivojac et al., 2010). The best studied class of SUMO E3s is the PIAS family proteins. There are four human PIAS genes, but some are expressed as splice variants so that there seven PIAS proteins, all of which possess a RING domain similar to the RING family of Ub E3 ligases (Rabellino et al., 2017). Numerous sumoylation substrates for the PIAS proteins have been identified though these proteins appear to have other functions unrelated to E3 ligase activity (Pichler et al., 2017). In addition to the PIAS family, there are two other types of bona fide SUMO E3s, RanBP2 (Pichler et al., 2002) and the ZNF451 family (Cappadocia et al., 2015). RanBP2 is a component of the nuclear pore complex and is responsible for sumoylating RanGAP1 (Swaminathan et al., 2004), Topo II alpha (Dawlaty et al., 2008), and RanGDP (Sakin et al., 2015). RanBP2 is unrelated to other E3 ligases, and whether there are additional substrates for this unusual E3 is unknown. The ZNF451 family is largely uncharacterized, has E3 activity highly

specific for SUMO2/3 rather than SUMO1, and uses a catalytic mechanism unlike other E3 ligases (Eisenhardt et al., 2015). At least in vitro ZNF451 can catalyse sumoylation of PML protein and PML components, suggesting an important role in regulation of PML body formation (Koidl et al., 2016). Other potential SUMO E3 ligases have been reported, such as Pc2 (Kagey et al., 2003) and TOPORS (Weger et al., 2005), but their status as authentic E3s has not been thoroughly established. For sumoylation, choice of the substrate lysine is usually determined by sequence context. Known consensus motifs for sumoylation include the predominant ψKxD/E motif (Sampson et al., 2001), the inverted motif (E/DxKψ) (Matic et al., 2010), the hydrophobic motif (ψψψKxE) (Matic et al., 2010), a phosphorylation-dependent motif (Hietakangas et al., 2006), and several extended motifs with heavy negative charges (Yang et al., 2006; Picard et al., 2012). However, even though these consensus motifs have been useful for predicting acceptor lysines for sumoylation, many examples exist of sumoylation at non-consensus sites so there is some flexibility in SUMO targeting (Hendriks and Vertegaal, 2016). Ub/SUMO functions The canonical function of the ubiquitin system is to target substrates for proteasomal degradation. The Ub moiety is transferred from the E2–E3 complex onto a lysine residue on the substrate with formation of an isopeptide bond between the carboxyl group of the C-terminal glycine of Ub and the amino group on a lysine side chain of the substrate. Subsequent to the initial addition of an Ub moiety to the substrate, additional Ub molecules are joined onto the first Ub to form a poly-Ub chain (Fig. 1.3). Ub itself has seven lysine residues (K6, K11, K27, K29, K33, K48, and K63) and an N-terminus that can each serve as acceptors for additional Ub attachment (Komander and Rape, 2012). Proteolytic signalling is defined by poly-ubiquitination with linkage through lysine 48 which is the predominant linkage type in cells (Kim et al., 2011). Chains of four or more K48-linked Ub molecules confer recognition by the proteasome leading to subsequent degradation of the substrate (Chau et al., 1989). This mechanism is responsible for much of the normal turnover of ageing or damaged proteins

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Figure 1.3  Examples of modification by single (mono), dual (multiple), and chains (poly) of ubiquitin like proteins (Ulps).

within cells and for regulating cellular processes where degradation of the target protein is used to stop the protein activity. Interestingly, other nonproteolytic functions of K48 chains have also been described (Kuras et al., 2002; Ye, 2006). The second most common form of Ub linkage is poly-K63 which has non-degradative roles in protein kinase activation through the TLR pathways, the TNF receptor, and DNA damage pathways among others (Chen and Sun, 2009). Numerous other examples of linkages through other Ub lysines exist, as well as mixed chains, and functional monoubiquitination (Swatek and Komander, 2016), and much remains to be understood about the biological roles of these diverse modification patterns. One thing that is clear is that the ubiquitin code is complex and critically important for numerous cellular processes (Fig. 1.3). Unlike the predominant activity of Ub in protein degradation, modification by SUMO does not typically direct proteins to the proteasome and instead has diverse effects including altering substrate function, localization, or interaction with partner proteins. Proteomic studies have demonstrated thousands of targets for sumoylation, supporting the critical and widespread nature of this modification (Hendriks and Vertegaal, 2016). However, sumoylation is more a nuclear phenomenon than is ubiquitinylation which occurs throughout the cell. Examples of important nuclear functions of sumoylation include regulation of transcription factor activity (Rosonina et al., 2017), PML body assembly (Sahin et al., 2014), and DNA damage response (Morris and Garvin, 2017). Importantly, there are clear biological and functional distinctions between SUMO1 versus SUMO2/3 conjugation though there often appears to be redundancy with many substrates capable of being modified with SUMO1 or SUMO2/3 (Evdokimov et al., 2008).

SUMO1 is mostly found conjugated to substrates while SUMO2/3 is largely in free pools until stimulated for conjugation by stress conditions (Golebiowski et al., 2009; Castorálová et al., 2012). Additionally, SUMO2/3 proteins themselves contain a sumoylation consensus sequence which allows poly-SUMO2/3 chain formation (Tatham et al., 2001). SUMO2 chains have also been detected with modifications at different lysines, though how this affects biological activity is unresolved (Tammsalu et al., 2014). While SUMO1 lacks a consensus modification site, both SUMO1 chains and mixed SUMO1/SUMO2 chains have been detected (Hendriks et al., 2017), so the complexity of chain formation seen with Ub is at least partially observed for SUMOs. Much work remains to decipher the complete SUMO code and to relate it systematically to functional effects of sumoylation. For both Ub and SUMO modification, many of their functional consequences on substrates require interaction with other proteins that recognize the Ub or SUMO moieties. Defined recognition motifs have been identified which bind Ub such as the ubiquitin-interacting motif (UIM) and the ubiquitin-associated domain (UBA) (Hofmann and Bucher, 1996; Young et al., 1998). Proteins may possess single or multiple copies of UIM or UBA domains, leading to differences in their binding affinity for ubiquitinylated proteins as well as specificity with regard to number of Ub copies required for binding (Hurley et al., 2006). Likewise, for SUMOs there are defined SUMO-interaction motifs (SIMs) that mediate the binding of some proteins to sumoylated partners (Hecker et al., 2006). The SIM motif has a hydrophobic region flanked by acidic and/or serine residues (Song et al., 2004; Hecker et al., 2006). As for Ub, the existence of proteins with multiple SIMS, coupled with multi- and poly-sumoylated substrates, allows

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for complex combinatorial interactions that likely account for much of the specificity of interactions between sumoylated proteins and their potential partners. Many of these important pairings for Ub and SUMOs will be discussed in detail in subsequent chapters. Ubiquitin-SUMO crosstalk In addition to their varied, complex, and independent functions, many elegant studies have revealed that there are significant interactions between the ubiquitin and SUMO systems. Among the earliest observed examples of interplay between Ub and SUMO was competition for the same acceptor lysine residue 164 of PCNA (Hoege et al., 2002). Mono-ubiquitination (Bienko et al., 2005), K63 type poly-ubiquitination (Branzei and Foiani, 2010), and sumoylation (Gali et al., 2012) at lysine 164 each have different effects on PCNA function in DNA damage repair, and collectively these competing modifications regulate activity. Another example of competition is blocking of proteasomal degradation by sumoylation of the lysine used for poly-ubiquitination (Klenk et al., 2006; EscobarRamirez et al., 2015). As up to 25% of human sumoylation sites are also known ubiquitination sites, functional competition may be a very widespread event (Hendriks et al., 2014). Alternatively, sumoylation is known to promote proteasomal degradation in some cases through recruitment of SUMO-targeted ubiquitin ligases (STUbls) (Prudden et al., 2007; Sun et al., 2007; Xie et al., 2007). STUbls are RING domain Ub E3 ligases that contain multiple SIMs and thus are recruited to poly-sumoylated proteins (Mullen and Brill, 2008; Tatham et al., 2008). The primary human STUbl is RNF4 which has roles in degradation of PML proteins (Geoffroy et al., 2010) and other DNA repair proteins (Psakhye and Jentsch, 2012). In addition to known roles for SUMO–Ub crosstalk in regulation of degradation, a large number of proteins contain independent sites for both Ub and SUMO addition (McManus et al., 2017), suggesting significant opportunities for synergistic and antagonistic effects on the activities of dually modified proteins. Interestingly, Ub can be sumoylated and SUMO can be ubiquitinated, and the formation of heterologous SUMO-Ub chains adds further regulatory complexity for control of protein

activity as has been shown for IĸBα (Aillet et al., 2012). In summary, it is clear that there are several known mechanisms for Ub–SUMO crosstalk, but that potential consequences of these interactions on target proteins is still largely undefined. Continually defining this regulatory network will be an exciting focus for future work in this field. Conclusions In the forty years since the discovery of ubiquitin, the elucidation of the larger Ub superfamily has revealed an ancient and highly conserved regulatory network that touches virtually everything in the cell through modification with Ubls. Between them, Ub and SUMOs modify thousands of cellular proteins, often at multiple sites per protein leading to synergistic or antagonistic effects on protein activity and stability. Both modifiers can be added as monomers or polymers, and examples are now known of mixed polymers consisting of mixtures of different SUMO types or of SUMO–Ub hybrid chains. These observations clearly demonstrate an enormous combinatorial complexity to these modification systems which allows them to help fine tune the activity of important proteins throughout the cell. This book seeks to explore the interface between the Ub and SUMO systems to provide a comprehensive picture of our current knowledge of these two branches of the Ub superfamily. References

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Cracking the Ubiquitin Code: The Ubiquitin Toolbox Monique P.C. Mulder*, Katharina F. Witting and Huib Ovaa*

2

Oncode Institute and Department of Cell and Chemical Biology, Chemical Immunology, Leiden University Medical Centre, Leiden, the Netherlands. *Correspondence: [email protected] and [email protected] https://doi.org/10.21775/9781912530120.02

Abstract Ubiquitination, a post-translational modification, regulates a vast array of fundamental biological processes with dysregulation of the dedicated enzymes giving rise to pathologies such as cancer and neurodegenerative diseases. Assembly and its ensuing removal of this post-translational modification, determining a large variety of biological functions, is executed by a number of enzymes sequentially activating, conjugating, ligating, as well as deubiquitinating. Considering the vast impact of ubiquitination on regulating cellular homeostasis, understanding the function of these vast enzyme networks merits the development and innovation of tools. Thus, advances in synthetic strategies for generating ubiquitin, permitted the development of a plethora of ubiquitin assay reagents and numerous activity-based probes (ABPs) enable the study of enzymes involved in the complex system of ubiquitination. With ubiquitination playing such a pivotal role in the pathogenesis of a multitude of diseases, the identification of inhibitors for ubiquitin enzymes as well as the development of ABPs and high-throughput assay reagents is of utmost importance. Accordingly, this chapter will review the current state-of-the-art activity-based probes, reporter substrates, and other relevant tools based on Ub as a recognition element while highlighting the need of innovative technologies and unique concepts to study emerging facets of ubiquitin biology.

Introduction One of the most versatile post-translational modifications is the attachment of the small protein ubiquitin (Ub) or its polymeric chains to target substrates. The attachment of the 76 amino acid long protein Ub to a nucleophilic functionality in the amino acid side chain of substrate proteins alters the fate of the modified protein, thereby regulating the vast majority of fundamental cellular processes such as DNA damage response (Muratani and Tansey, 2003), cell cycle progression (Kernan et al., 2018), transcription (Hicke, 2001), endocytosis (McCann et al., 2016), as well as apoptosis ( Jackson and Durocher, 2013) and autophagy (Kwon and Ciechanover, 2017). Covalent attachment of Ub to its substrate proteins is orchestrated by the sequential action of three specialized enzyme classes – E1, E2, and E3 enzymes (Fig. 2.1A). However, the combination of E2 and E3 enzymes dictates what type of ubiquitin chain is formed and which substrate protein becomes ubiquitinated. To date, 2 human E1’s, about 40 E2’s and over 600 E3 enzymes are known. Adenylation of the C-terminus of Ub at the expense of ATP yields a high-energy E1-Ub-thioester. Upon activation, Ub is transferred unto the active-site cysteine residue of the E2-enzyme, poising it for transfer unto the lysine residue of its substrates by the cooperation of an E3 enzyme. This final step in Ub-transfer through the E3 enzyme can occur via three main classes of E3 ligases: the homologous to the E6-AP- C terminus (HECT), the really interesting new gene (RING),

16  | Mulder et al. A) O Ub S

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Figure 2.1 The complexity of ubiquitination. (A) The ubiquitination cascade, an orchestrated interplay of enzymes. (B) Self-modification of ubiquitin on one of its seven lysine residues results in a variety of different linkage types. Additionally, Ub can modify itself using the N-terminal methionine residue. (C) Increased complexity can be achieved by linking the Ub-modules in various manners leading to homotypic Ub-chains, in which the same type of Ub linkage is found or as (D) heterotypic linkages, which can either be mixed or branched. (E) Modification by a Ubl yielding hybrid chains.

and the RING-in-between-RING E3 (RBR) E3 enzymes (Vittal et al., 2015). In contrast to HECT E3 ligases, which utilize a direct transfer mechanism to relay activated ubiquitin to its substrate lysines, and RING E3s that employ an indirect scaffolding mechanism, RBR (RING-between-RING) ligases possess a trilateral domain architecture consisting of three zinc-binding domains – a RING1 domain flanked by an in-between-RING (IBR) domain, adjacent to a RING2 domain (Walden and Rittinger, 2018). On E2 Ub thioester recognition by RING1, it is transferred to the catalytic cysteine of RING2, which then facilitates transfer to the lysine of the substrate (Spratt et al., 2014; Walden and Rittinger, 2018).

Importantly, ubiquitination is a reversible process. The ubiquitination status of a protein can be regulated by removal or editing of ubiquitin chains, which is carried out by a group of approximately 100 deubiquitinating enzymes (DUBs) (Fig. 2.1A). Several categories of human DUBs have been identified to date; including the subfamilies of ubiquitin-specific proteases (USPs), ubiquitin C-terminal hydrolases (UCHs), Machado-Joseph disease proteases (MJD), ovarian tumour domain proteases (OTUs), motif interacting with Ubcontaining novel DUB family (MINDYs) and zinc finger with UFM1-specific peptidase domain protein (ZUFSPs) cleaving Ub linkages through a cysteine protease mechanism whereas JAB1/

The Ubiquitin Toolbox |  17

MPN/MOV34 proteases ( JAMMs) are zinc dependent metallo-proteases (Komander et al., 2009; Abdul Rehman et al., 2016; Hermanns et al., 2018). For some of these DUBs, linkage specificity has also been observed, further modulating the cellular response to ubiquitination (Komander and Rape, 2012; Harrigan et al., 2018). Intricate coordination of substrate ubiquitination by E3 ligases and DUBs is integral to maintain cellular homeostasis with deregulation leading to the onset and progression of numerous pathologies including cancer, neurodegenerative diseases, inflammatory, and infectious diseases arising from their deregulation (Scheffner and Kumar, 2014; Harrigan et al., 2018). This complex interplay is perhaps best exemplified by the ubiquitination of the tumour suppressor p53 by the E3 ligase MDM2 which is counterbalanced by USP7 deubiquitination thereby preventing proteasomal degradation but also regulating its expression levels (Nag et al., 2013). To further modulate the biological consequence of Ubiquitination, Ub can undergo self-modification by forming isopeptide bonds between the N-terminal methionine (Met1-linked ubiquitination) or any of the internal seven lysine (Lys6, Lys11, Lys27, Lys29, Lys33, Lys48, Lys63) ε-amines (Lys-linked ubiquitination) of one Ub molecule and the C-terminal carboxylic acid of another Ub molecule (Fig. 2.1B). In this manner, homotypic poly Ub chains of a single linkage type consisting of M1, K6, K11, K27, K29, K33, K48 or K63 can be formed (Fig. 2.1C), each having unique structural features creating distinct signalling events (Komander, 2009). While K48-Ub, one of the most abundant linkage type (Michel et al., 2017), destines substrates for proteasomal degradation, K33-linked Ubiquitin chains mediate protein trafficking (Yuan et al., 2014). All of these linkages have been detected in cells and their abundance changes during specific cellular events, indicative of their various functions (Xu et al., 2009). In addition, heterotypic chains of multiple ubiquitin linkage types adopting mixed or branched topology can be formed (Fig. 2.1D), opening up an even more complex layer of posttranslational modification (Kim et al., 2007). The increased regulation of cellular processes especially by heterotypic ubiquitin chains is underscored by the observation that branched K11/K48 Ubiquitin chains promote proteasomal degradation in vitro

(Meyer and Rape, 2014), while mixed K11/K63 linked Ubiquitin chains regulate the endocytic internalization of the major histocompatibility complex class 1 (MHC1) (Boname et al., 2010). Additionally, Ub itself can be post translationally modified to further modulate the biological fate, most prominently by acetylation, phosphorylation, and more recently ribosylation (Yang et al., 2017). The consequences of such an additional modification is best exemplified by the phosphorylation of Ub by PINK1 resulting in Parkin recruitment and activation (Herhaus and Dikic, 2015). Furthermore, this additional layer of complexity can be expanded to include modification with Ubiquitinlike modifiers (Ubls) – a class of proteins that share high structural similarity and a common β-grasp fold with Ub such as SUMO, NEDD8 and ISG15 (Fig. 2.1E) (Kwon and Ciechanover, 2017). These UBL modifiers are attached to the target protein via their own dedicated E1, E2 and E3 enzymes and deconjugated with dedicated proteases. Discovery of Ub and its role in proteasome mediated protein degradation was awarded with the Nobel Prize in Chemistry in 2004 (Giles, 2004). However, the complexity of the ubiquitination network and its cellular roles are far more diverse than just being a degradation signal. In the past years an enormous biochemical effort has been made in developing reagents and tools to study this complex enzyme cascade. Here, we will discuss the advances made in the chemical toolbox to study a broad range of biochemical and biological aspects of ubiquitin. Chemical approaches to ubiquitination In the past years, an enormous biochemical effort has been made in finding E2–E3 enzyme combinations that can give access to sufficient amounts of di- and polyubiquitin molecules representing all eight different homogenously linked ubiquitin types (Faggiano et al., 2016). In these efforts, people have been hampered by the lack of specific E2 and E3 enzymes to generate the so-called atypical (K6, K11, K27, K29, K33) chains. Only recently enzymatic approaches for making K6-, K11-, K29-, and K33-linked chains (Bremm et al., 2010; Hospenthal et al., 2013; Michel et al., 2015) were reported. Currently, only K27-linked ubiquitin

18  | Mulder et al.

remains enzymatically unattainable. On top of this some of the enzyme combinations reported are not linkage specific and further sample processing using DUBs (with their own specificity issues) is needed. Therefore, much effort has been put into making differentially linked ubiquitin derivatives or ubiquitinated proteins through semi-synthetic and synthetic strategies to circumvent traces of other linkages and assure homogenous preparation. Moreover, for study of the (de)ubiquitination network, modifying Ub derivatives with a specific handle to generate a particular Ub-based probe or enzyme substrate makes it even more challenging to prepare such a modified Ub conjugate enzymatically. Semi-synthetic strategies One of the most powerful semi-synthetic approaches for the production of large peptides and small proteins has been intein-based chemistry. This methodology relies on protein trans-splicing (PTS) which through a series of acyl shifts forms a thioester that can react with thiol or amine nucleophiles (Mootz, 2009). Expansion of the genetic code with unnatural amino acids (UAAs) has further aided the field of protein semi-synthesis and permitted the incorporation of unnatural amino acids facilitating the production of ubiquitin-based reagents (Trang et al., 2012; Wals and Ovaa, 2014; Rösner et al., 2015). While genetic code expansionbased methods are clearly useful, most do require certain expertise that can only be found in specialized labs and often require specific E. coli strains and tRNA pairs that might not be widely accessible. Another semi-synthetic strategy to generate fluorogenic ubiquitin and diubiquitin substrates exploits the E1-enzyme mediated C-terminal amidation reaction to equip the ubiquitin C-terminus with several reactive groups (Wang et al., 2014). Synthetic strategies Although efforts to synthesize ubiquitin have been pioneered by Briand et al. (1989) and Ramage et al. (1994) in the late 1980s, the chemical synthesis of natively linked ubiquitinated peptide conjugates was first established by Muir and co-workers (Chatterjee et al., 2007). Their photo cleavable auxiliary (Aux) mediated ligation approach has paved the way for several chemical strategies for ubiquitination. Recently, two Aux mediated chemical

ubiquitination methods have been reported. In the first approach Chatterjee and co-workers used a 2-aminooxyethanethiol Aux to mediate chemical ubiquitination (Weller et al., 2014). Their methodology enabled the preparation of the native isopeptide linkage by mild reductive removal of the Aux or alternatively, retention of the ligation Aux yielded protease-resistant non-native analogues of ubiquitinated peptides. Secondly, Liu and coworkers used the trifluoroacetic acid (TFA)-labile 1-(2,4-dimethoxyphenyl)-2-mercaptoethyl Aux to assist the synthesis of K27-linked di- and tri-Ub chains (Pan et al., 2016). The native chemical ligation (NCL) reaction, an important extension of the chemical ligation field, is widely used to construct large poly peptides or proteins by reacting an N-terminal cysteine residue to C-terminal thioester peptide followed by transthiolation and S-to-N-acyl migration giving an amide bond as final product (Dawson et al., 1994). This powerful technique, has been employed by Brik and co-workers and Ovaa and co-workers to synthesize Ub dimers of defined linkage by the incorporation of a δ- or γ-thiolysine moiety at a designated lysine residue to allow NCL with a thioester moiety, which had previously been introduced by Yang et al. (2009) (El Oualid et al., 2010; Kumar et al., 2010). Recently, this methodology was adapted to create Ub mutants containing both a thiolysineand a thioester entity, allowing polymerization under NCL conditions (van der Heden van Noort et al., 2017). The development of γ-thionorleucine (ThioNle) as handle for native chemical ligationdesulfurization has expanded the thiolated amino acid toolbox further and serves as a methionine substitute in NCL, making the N-terminal ubiquitination towards full synthetic linear M1 diubiquitin possible for the first time (Xin et al., 2018). Liu and co-workers describe an alternative NCL strategy that does not require the use of the δ- or γ-thiolysine moieties. Here a premade isopeptidelinked Ub isomer, which has an N-terminal Cys and a C-terminal hydrazide, is the key building block to assemble atypical Ub chains in a modular fashion resulting in the synthesis of several linkage- and length-defined atypical Ub chains, including K27linked tetra-Ub and K11/K48-branched tri-, tetra-, penta-, and hexa-Ubs (Tang et al., 2017). Only the introduction of an efficient linear Fmoc-based solid phase peptide synthesis (SPPS)

The Ubiquitin Toolbox |  19

of Ub unlocked the potential of the above described methodologies. The ubiquitin module can be synthesized with total linear synthesis, or from fragments. During the total linear Fmoc-based SPPS approach, the growing peptide chain is stabilized by the incorporation of special building blocks, that prevent the formation of aggregates as the Ub chain grows (El Oualid et al., 2010). These SPPS strategies have allowed for the site-specific installation of a wide variety of reactive groups, unnatural amino acids, fluorescent labels, or pull-down handles (Hameed et al., 2017). Recently, a microwave assisted SPPS methodology for ubiquitin was reported that avoids the use of aggregation breakers and allows synthesis of isoUb in just one day. Here a four segment three step ligation method is used to synthesize K33/K11 mixed triUb (Qu et al., 2018). Another study, exploits an intermolecular side reaction, observed while synthesizing Ub on a trityl resin, occurring between the N-terminal amine of one Ub molecule and the activated C-terminus of another Ub molecule to obtain natively M1-linked polymeric ubiquitin chains (van der Heden van Noort et al., 2018). The length of these M1-linked poly Ub chains (up to ten Ub-residues) is unprecedented in a single chemical reaction, giving easy access towards bona fide M1 poly Ub chains shown to be fully recognized by the enzymatic ubiquitination cascade, as exemplified by DUB (OTULIN) cleavage and E1 activation (Uba1). This research not only provides a platform for the development of novel tools based on polymeric Ub in the near future, but also highlights new insights important to consider in experimental design for the construction of large peptides (van der Heden van Noort et al., 2018). Despite these technological advances, numerous aspects of Ub signalling are difficult to study with a native isopeptide bond. Since the proteolytic activity of DUBs degrades the poly-Ub chain, crystallization or pulldown experiments are rendered impossible. In order to study stable complexes between poly Ub chains and DUBs, catalytically inactive DUBs are typically used. Yet, this approach yields numerous drawbacks, especially in biological settings necessitating the use of proteolysis-resistant Ub-chains. Utilizing a variety of chemistries, a broad range of poly-Ubiquitin chains of all linkage types can be generated giving access to studying mechanistic aspects of DUB cleavage as well as

elucidating the role of the Ub-chains in a cellular environment. In the field of Ub-chemistry, examples of nonhydrolyzable Ub conjugates generating strategies include the oxime-based ligation (Shanmugham et al., 2010), Huisgen cycloaddition reaction between an alkyne and azide (Flierman et al., 2016) or thiol-ene chemistry leading to a forged thioether bridge (Valkevich et al., 2012). Of note is that the thus generated linkage between two following Ub-modules is not the native isopeptide bond. Some of these unnatural linkages are generally accepted to be adequate amide-bond mimics and several examples show that poly Ub material containing this linkage is tolerated and advantageous in biological settings (Flierman et al., 2016; Zhang et al., 2017). It has however also been shown that slight modifications in this isopeptide linker region can have a dramatic effect on biological function (Haj-Yahya et al., 2012). Although synthetic strategies allow complete control over modifications, the experimental design needs to be carefully evaluated when using these reagents in biological settings to further the understanding of Ubiquitination. Advantage of the chemical approaches described above over biochemical methods is the complete control over regioselectivity in the reaction and thus formation of only the desired (poly-)Ub chain. Another superiority is the potential ease of introducing modifications to the chain such as for instance incorporation of reactive groups on the C-terminal side converting the chains into an activity-based probe. Beyond ubiquitin – crosstalk with other post-translational modifications Ub itself can be post-translationally modified to further modulate the biological fate, and simple PTMs on Ub such as phosphorylation (Huguenin-Dezot et al., 2016) and acetylation (Ohtake et al., 2015) can be incorporated through semisynthetic approaches. However, more complex PTMs such as adenosine diphosphate ribose (ADPr), are more difficult to introduce. Interestingly, ADP-ribosylation of Ub (Arg42) is mediated by a family of effector proteins originating from Legionella pneumophila, the pathogen causing Legionnaires disease in an ATP-independent reaction to hijack the host

20  | Mulder et al.

cells Ub pool, preventing the processing of existing Ub chains by host DUBs, and use it to its own advantage. These SidE effectors are the first reported class of enzymes that are able to ubiquitinate target proteins independent of the normally employed enzymatic cascade of E1, E2, and E3 enzymes (Bhogaraju et al., 2016; Puvar et al., 2017). In a recent study, the design and synthesis of propargylated ADP-ribose building block is presented employing a copper-catalysed cycloaddition reaction in which an Ub azide (Arg42 replaced by azido-homoalanine) an analogue of Ub-ADPr, was prepared. Subsequently, this triazole-containing Ub-ADPr was shown to be recognized in western blot and accepted by SdeA in an auto-ubiquitination assay, instigating a useful platform for the biological interrogation of Ub-ADPr biology (Liu et al., 2018). Additionally, there is a growing evidence implying crosstalk between ubiquitin and ubiquitin-like (UbL) proteins, increasing the complexity and fine-tuning cellular responses further. Best studied is the crosstalk between ubiquitin and SUMO (Nie and Boddy, 2016), but ubiquitinated-NEDD8 chains and crosstalk between Ub and Nedd8 signalling pathways have also been reported (Leidecker et al., 2012; Singh et al., 2014), as well as the existence of ubiquitinated FAT10 (Buchsbaum et al., 2012) and ISGylated ubiquitin (Fan et al., 2015). To address these unmet needs on hybrid chains, (semi-)synthetic strategies for obtaining ubiquitinated Rub1, the yeast NEDD8 homologue (Singh et al., 2014) and SUMO-2–K63diUb hybrid chains (Bondalapati et al., 2017) have already been reported. Despite these advancements, synthetic strategies for obtaining full-length Ubl proteins have long been neglected. Only recently, efforts to devise synthetic strategies for Ubl proteins such as Nedd8 (Ekkebus et al., 2013), SUMO (Dobrotă et al., 2012; Wucherpfennig et al., 2014; Boll et al., 2015; Mulder et al., 2018) and Ufm1 (Ogunkoya et al., 2012; Witting et al., 2018) have been undertaken not only providing access to Ubl reagents allowing research on their respective enzymatic cascades, but also enabling future developments on hybrid chains enabling in depth studies on their crosstalk with ubiquitin.

Visualizing ubiquitin in action – Ub reagents targeting DUBs and ligases Activity-based probes (ABPs) are powerful tools to study enzyme activities in vitro and in vivo and have been helpful for studying the activity of enzymes. They typically consist of three elements – a reactive group, a recognition element and a reporter tag and have been instrumental in not only identifying but also studying DUBs and more recently the conjugating and ligating enzymes of the Ub cascade (Hewings et al., 2017). Additionally, the introduction of a facile linear solid phase peptide synthesis method for ubiquitin, permitted the development of a plethora of ubiquitin assay reagents, such as fluorogenic assays, native and non-hydrolyzable ubiquitin-linkages, and even poly-ubiquitin chains thereby enabling the characterization of these enzymes. Taking a snapshot of DUB activity – ABPs targeting the deconjugation machinery While the first generation of ABPs targeting DUBs utilized Ubiquitin-aldehyde (UbaI) (Pickart and Rose, 1986) and Ub-nitrile (Ub-CN) (Lam et al., 1997), introduction of the vinyl-sulfone (VS) (Borodovsky et al, 2001) as a reactive group led to the development of irreversible DUB ABPs. Since then, a wide variety of electrophilic reactive groups (Borodovsky et al., 2002) have been introduced with the vinyl methyl ester (VME) (Borodovsky et al., 2002; Ovaa et al., 2004) and propargyl amides (PA) (Ekkebus et al., 2013) being the most widespread used ones (Fig. 2.2A). These ABPs furthered the discovery of novel DUBs, as is exemplified not only by the discovery of OTU family of DUBs (Borodovsky et al., 2002; Balakirev et al., 2003), numerous viral (Hewings et al., 2017) and bacterial DUBs (Pruneda et al., 2016), but also by the discovery of a novel bacterial protease class exhibiting both deubiquitinating and deneddylase activity (Grabe et al., 2016). In addition, they have been used in activity profiling, crystallization studies to study the interactions between the protease and Ub in detail as previously reviewed (van Tilburg et al., 2016), as well as inhibitor screening (Reverdy et al., 2012). However, these ABPs bind irreversibly to the active site of the DUB, rendering them

The Ubiquitin Toolbox |  21

A)

B) S2

S1 S1’

C) S1 S1’

DUB

1st Generation

O

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Borodovsky et al. , 2001

O N H

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Triazole-linked diUb - proximal C-terminal PA warhead (all seven linkages) Flierman et al., 2016

S β

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K48C- and K63C- linked VA-diUb Li et al., 2014

O N H

O Ekkebus et al., 2013

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Borodovsky et al., 2002

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N H

(all seven linkages) McGouran et al., 2013

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H N

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catalytic site warhead

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γ β

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O N H

G75 H

N

O

O N H

ε

γ δ

β

K48- and K63- linked Dha-diUb Haj-Yahya et al., 2014

Figure 2.2  Overview of activity-based probes to target DUB activity. (A) First generation DUB probes targeting S1 interactions. (B) Advanced DUB probes allowing S1 and S1’ interactions. (C) Third generation DUB probes, enabling the covalent capture of DUBs preferentially targeting S1–S2 interactions.

inactive. In a recent study, a novel type of ABP containing a methyldisulfide warhead that captures DUBs reversibly, by means of active-site-specific disulphide exchange, allowing the release of an active enzyme was presented (de Jong et al., 2017). The significance of this probe lies in its ability to isolate active DUBs from their cellular environment retaining present cell-specific post-translational modifications that might regulate DUB activity. Although only proof of principal studies have been performed, this novel technology holds great promise for the future capture, release, and follow up investigations of native active cysteine DUBs in cellular contexts.

However, while activity-based probes have greatly increased our understanding of DUB reactivity and have enabled the discovery of new DUBs such as the OTU (Balakirev et al., 2003) and MINDY (MIU-containing novel DUB) classes (Abdul Rehman et al., 2016), these ABPs offer limited information on poly-Ubiquitin chain recognition and processing, since the existing diUbiquitin reagents contained isopeptide-linked Ubiquitin modules. While this characteristic allows the profiling of recombinant deubiquitinating enzymes towards their linkage specificity and kinetics (Mevissen et al., 2013), a major limitation is its incompatibility with the cellular environment

22  | Mulder et al.

which modulates DUB activity, thus necessitating innovative tools specifically addressing these questions. With the advent of synthetic strategies, a 2nd generation of probes has emerged where, between two ubiquitin modules, a reactive group is positioned at the site of proteolytic action of the DUB allowing its covalent capture (Fig. 2.2B). An initial report by Iphofer et al. (2012) show a Michael acceptor linking the C-terminus of a distal Ub and short peptides representing K48 or K63 diUb. Later reports include the entire palette of Ub-chains allowing access to all seven lysine linked diubiquitin probes with a warhead in-between the distal and proximal ubiquitin module. Numerous research groups have independently reported ABPs utilizing a vinyl amide electrophilic trap between non-natively linked Ub moieties linked through a triazole, thiol ether (McGouran et al., 2013; Li et al., 2014) or an amide bond closely resembling the native isopeptide in both length as structure (Mulder et al., 2014). An alternative warhead is described by Haj-Yahya et al., here thiol elimination of Ub(G76C)-Ub results in dehydroalanine (Dha) as an electrophilic trap between two Ub modules (Haj-Yahya et al., 2014). Although these covalent vinyl amide probes have allowed more detailed structural investigation of diubiquitin-specific DUB recognition (Mevissen et al., 2016), they do not allow investigation of additional Ubiquitin-binding sites, referred to as the S1’ (proximal), S1 (middle), and S2 (distal) binding sites (Kulathu, 2016). To investigate the contribution of the Ubiquitin binding sites to polyubiquitin chain processing by DUBs, a third generation of probes (Fig. 2.2C) generated by click chemistry and C-terminally modified with propargyl (PA) were devised (Flierman et al., 2016). Utility of this reagent enabled the structural characterization of the K48 polyubiquitin cleaving mechanism of the SARS DUB PLpro, revealing that the S1-S1’ binding mode of K48linked ubiquitin dictates the enzyme specificity for K48-Ubiquitin over ISG15, which binds only in the S1 site (Békés et al., 2016). Despite the variety of di-ubiquitin-specific ABPs, designing effective tools to study the M1-linked chain type has posed a challenge primarily due to differences in chemistry imposed by the ‘linear’ peptide linkage. In attempts to create an linear diUb ABP, the methionine 1 (M1) of the proximal Ub

was replaced by the electrophilic dehydroalanine (Dha) residue. However, this probe was cleaved by OTULIN and USP2 rather than reacting covalently with the active site cysteine residues. A more recent design addressed this issue by replacing the Gly76 of the distal Ub by Dha (Weber et al., 2017). Although the UbG76Dha-Ub probe showed high selectivity for OTULIN, it did not label other M1-cleaving DUBs, indicating that Gly76 of the distal Ub is essential for recognition and cleavage of linear diUb by other M1 cleaving DUBs. Interestingly, the first report on the fully synthetic preparation of linear diubiquitin reveals that the methionine to norleucine substitution of the proximal Ub affects the hydrolysis rate of DUBs towards the linear diUb chain (Xin et al., 2018). Assessment of DUB-mediated cleavage of the synthetic (NLE1linked) and expressed (M1-linked) linear diUb was assed using OTULIN, USP16 and USP21, known to specifically cleave the linear Ub linkage, demonstrated that synthetic NLE1-linked linear diUb was processed less efficiently than M1-linked linear diUb (Xin et al., 2018). Collectively, these observations indicate a more profound role for methionine and Gly76 in the interaction between M1-linked diubiquitin and DUBs, complicating the way for the design of linear diUb-based activity-based probes and assay reagents. Furthermore, these ABPs together with the insights gained from both structural and biochemical studies underscore that the interaction dynamics of di-Ubiquitin chains are far more complex than previously assumed. The numerous activity-based probes have furthered our mechanistic, kinetic, and biological understanding of DUBs as well as enabled the discovery of new DUB classes, yet these reagents do not target the JAMM/MPN and Machado-JacobDisease protein (MJD) metalloprotease DUBs. Developing such reagents akin to those for the other DUB families is urgently needed in order to dissect the role of these proteases in diseases. While significant advances have been made in the development of a variety of activity-based probes and reagents for DUBs, similar tools are slowly emerging for the proteases specific for ubiquitinlike modifiers, such as for the de-SUMOylating (SENPs) (Mulder et al., 2018), de-NEDDylating (Ekkebus et al., 2013) and de-UFMylating enzymes (Witting et al., 2018).

The Ubiquitin Toolbox |  23

Relaying ubiquitin to its substrate – ABPs targeting the ubiquitin conjugation machinery Whereas DUBs have been extensively profiled using ABPs, the Ub-conjugating and ligating enzymes have only recently become the focus of ABP development. The delay in developing suitable reagents to profile the E1-E2-E3 enzymes is largely due to the challenges attributed with targeting a sequential enzymatic cascade rather than a single enzyme. While ABPs originally designed to specifically target DUBs, such as HA-Ub-VME and Ub-VS, display cross-reactivity with HECT E3 ligases, they are not designed for monitoring Ub-conjugating and ligating enzyme activity concurrently (Borodovsky et al., 2001; Love et al., 2009), necessitating the development of ABPs and reagents specifically devised for the Ub conjugation machinery. At the apex of the ubiquitination cascade, the E1 enzyme activates the C-terminal carboxylate of ubiquitin in an ATP-dependent manner. In this initial step, the Ub-AMP adenylate is formed under the consumption of ATP and magnesium. Subsequently, the intermediate undergoes nucleophilic attack by the adjacent catalytic E1 active site cysteine resulting thioester bond and the simultaneous release of AMP (Olsen and Lima, 2013). Early efforts towards developing Ub-based probes targeting the E1-enzyme were pioneered by Lu et al. (2010), who used a C-terminal 5′-sulfonyladenosine modified Ub or Ubl. This design [Fig. 2.3A(I)] permitted the mechanistic study of the E1-catalysed adenylation and thioesterification by crosslinking it with the Ub/Ubl probe. A major drawback of the semisynthetic approach taken by Lu et al. (2010) is the alteration of the Ub/Ubl sequence. An and Statsyuk (2016) later published a method to efficiently generate the ABPs reported by Lu et al. (2010) while retaining the ‘native’ sequence, utilizing a native chemical ligation strategy followed by the conversion of cysteine to Dha, permitting the trapping of the ‘tetrahedral E1-Ubl-AMP intermediate’. Owing to the mechanism-based approach of these Ub/Ubl-AMP probes, it reacts directly with the E1-Ub/Ubl thioester intermediate resulting in the formation of the covalent Ub/Ubl–ABP1 conjugate structurally mimicking the Ub-AMP intermediate. Other advancements by Statsyuk and co-workers employed a mechanism-based approach [Fig. 2.3A(II)] using an AMP-derived

compound (ABP1), which due to its structural resemblance of the Ub/Ubl-adenylate reacts with the Ub/Ubl substrates rather than the respective E1 enzymes (An and Statsyuk, 2013). However, while this ABP has the advantage of being cellpermeable, cross-reactivity issues limits its utility to monitoring ubiquitination of substrates in vitro. Together, these approaches all mimic the Ub-Ubladenylate intermediate restricting these ABPs to the E1, enabling them to be processed downstream the cascade towards E2 and HECT- and RBR-E3 enzymes. The second step in the cascade involves transfer of the activated Ubiquitin from E1 to E2 via a thioester exchange reaction, a processes that can be trapped and studied using a E2 derived ABP [Fig. 2.3A(III)] (Stanley et al. 2015). Recombinant expression of an E2 and modification with a tosyl-substituted double activated ene-reagent (TDAE) forms an electron poor activated vinylsulfide that on juxta-positioning of the E1’s cysteine is able to form a stable bis-thioether E1– E2 complex (Stanley et al., 2015). To enable the study of enzymes downstream in the cascade a more advanced activity probe was designed (Fig. 2.3B) and generated in an analogues approach, coupling an azide-modified Ub to an alkynemodified tosyl-substituted doubly activated ene (TDAE) using click chemistry (Stanley et al., 2015; Pao et al., 2016). This design enabled the generation of stable E2–Ub conjugates, on reaction with a respective E2 enzyme, and subsequent recruitment of the RBR-E3 ligase Parkin whilst monitoring the transthiolation activity of this ligase (Pao et al., 2016). Of note is that in the TDAE derived probe the C-terminal RGG motif of Ub is replaced by the reactive TDAE element, which might limit the generality of such probes as it is implicated that R74 and the diGly motif can play an important role in recognition of the downstream enzymes (Zhao et al., 2012). In a later stage, Pao et al. (2018) include Arg74 in their TDAE-Ub probe and despite being the improper length, the ABPs described are able to recruit not only HECT/RBR but also RING E3 ligases. Most notably, the authors discover a novel RING E3 ligase – MYCBP2 (or PRH1), which utilizes a unique cysteine relaying mechanism mediating the transfer of activated Ubiquitin onto the threonine and serine residues. This unexpected finding

24  | Mulder et al. A) E1 probes (I) Mimicking the tetrahedral intermediate of the E1–Ub–AMP complex. NH2 O

H N

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An et al., 2016

(II) Mechanism based probe consuming free Ub HN

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An et al., 2013

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Figure 2.3  Current activity-based probes targeting the ubiquitin cascade. (A) Targeting the E1 enzyme in a mechanism-based manner by (I) mimicking the tetrahedral intermediate of the E1–Ub–AMP complex or (II) using an AMP-derived compound (ABP1) or (III) utilizing the E1-transthiolation activity. (B) Capturing Ub–E2–E3 interactions by a modular approach, where Ub-TDAE reacts with an E2 generating an ABP reactive towards HECT- and RBR- E3 ligases. (C) Cascading E1–E2–E3 ABP sequentially reacting with the E1, E2 and E3 enzymes by either forming (i) the thioester yielding the transferable Ub-probe or as (ii) a thioether, which allows irreversibly entrapment of the enzyme.

The Ubiquitin Toolbox |  25

exemplifies the utility and potential of ABPs and foreshadows the extent of future possibilities for these chemical tools (Pao et al., 2018). In order to address the shortcomings of existing Ub-ABPs for studying multiple types of enzymes from the UPS simultaneously, Mulder et al. (2016) developed a mechanistically engaged ABP (Fig. 2.3C). Here the C-terminal Gly76 is replaced by Dha, thereby retaining a native carboxy terminus thus allowing it to be processed by the native Ub conjugation machinery in the same ATP-dependent manner with E1–Ub-based and E2–Ub-based probes transiently formed in situ allowing relay to the E2 and E3 enzymes. Most notably, at each transthiolation step, the probe also has the option of reacting covalently with the active site Cys. However, in contrast to native ubiquitin, this cascading probe is inert towards lysine residues in target proteins, making it applicable to chemo-proteomics approaches. Additionally, its ATP-dependant reactivity is advantageous for proteome‐wide profiling experiments, as ATP-depletion permits facile background subtraction. Beyond its application for chemoproteomics, the utility of this unique cascading ABP has been showcased using living cells, where the effects of E1 enzyme inhibition on ubiquitination were visualized (Mulder et al., 2016). These experiments highlight the power of in-cell enzymology of the entire Ub cascade overcoming the limitation of labelling experiments in lysates, which are devoid of the organization and interaction of cellular structures. The recent emergence of E2–Ub-ABPs and the novel Ub-ABP Ub-Dha greatly expand the Ub toolbox and provide new ways to decipher the cellular functions and structural/biochemical properties of HECT ligases in specific cellular contexts as well as potentially in normal and disease state. However, of the three major classes of E3s, the current probes are only reactive towards HECT/RBR ligases, as these E3 ligases mechanistically rely on an active-site cysteine. RING E3s do not possess such an active site cysteine and merely serve as platforms to bring Ub charged E2’s and substrates together, thereby making them unsuited for direct probing using ABPs.

Assay reagents – real time monitoring of activity Measuring catalytic activity of (de)ubiquitinating enzymes is key not only to understand their biological function but also to inhibitor development efforts. In contrast to the probes described above these reagents lack a Michael acceptor element and thus do not form a covalent complex with their target enzymes, but instead rely on a fluorescent reporter tag allowing correlation of the enzymes native activity and/or specificity. An important class of Ub based assay reagents are the fluorogenic assay reagents where a quenched fluorophore is conjugated via an amide bond at the C-terminal end of Ub. DUB activity and recognition will hydrolyse the amide bond at the C-terminus of Ub, releasing the fluorophore and simultaneously start to fluoresce. Hence the increase in fluorescence is a direct measure of DUB activity. One of the first fluorogenic reagents to measure the catalytic activity of DUBs is Ub aminomethyl coumarin (UbAMC) (Dang et al., 1998). Hassiepen et al. (2007) later report on a substituted rhodamine-110 (Rho110) scaffold with favourable fluorescent properties, making Ub-Rho110 a more preferred reagent in high throughput screening assays due to its non-overlapping spectrum with many small molecule inhibitors). In a similar set up, DUB mediated amino-luciferin release can be assayed in a bioluminescence approach using a luciferase assay, allowing the study of DUBs at lower concentrations (Orcutt et al., 2012). Another striking example illustrating the utility of fluorescent ubiquitin reagents are the non-hydrolyzable di-ubiquitin AMC reagents, which allow the monitoring of chain specific proteolysis mediated by S1–S2 interactions on the DUB. In analogy to the diUb-PRG covalent probes, these substrates allowed mechanistic dissection of DUB specificity and cleavage rate, exemplified by the finding that the S2 ubiquitin binding pocket of OTUD3 confers its preference for K11 Ub-linkages as well as accelerating Ub hydrolysis (Flierman et al., 2016). In all these cases, the reporters did not contain a native isopeptide bond at the side where the DUB would normally perform its proteolytic action, whereas the natural substrates for most

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DUBs would. Given that Ub-linkages govern a plethora of biological processes finetuning the cellular responses to a variety of stimuli, assessing the dynamics of Ubiquitin chain processing by DUBs is critical. Therefore, fluorescent polarization (FP) reagents were developed where Ub is conjugated via a native isopeptide linkage to a fluorophore carrying substrate derived peptide (Tirat et al., 2005; Geurink et al., 2012). Assays with these reagents are based on a change in fluorescence polarization on cleavage of the isopeptide bond between Ub and a fluorophore labelled peptide. While the unprocessed large Ub-FP reagents tumble slowly giving high fluorescence polarization, the processed small fluorophore containing peptide tumbles faster and hence the polarization of light decreases. The synthetic advancements enabled the generation of a palette of FP reagents as well as the generation of FP reagents based on UBLs like the three SUMO isoforms, NEDD8 and ISG15 (Geurink et al., 2012). Another class of reagents are Fluorescent Resonance Energy Transfer (FRET)-based reagents that make use of a fluorophore and quenching moiety in close proximity of each other. On DUB proteolysis, the FRET signal decreases over time, which can be measured in a fluorescence spectrometer enabling the study of enzyme linkage specific kinetics in real time. Geurink et al. (2016) prepared all seven isopeptide-linked diUb FRET assay reagents by native chemical ligation using Rhodamine-Ub as the FRET-donor and TAMRA-Ub as the FRET-acceptor, permitting insights into the catalytic efficiency of vOTU. From the kinetic measurements it became apparent that the preference for K6-linked di-Ubiquitin chains over K48 chains resulted from an increased catalytic turnover rate kcat and not Ubbinding (KM) (Geurink et al., 2016). Using a similar technology, a high-throughput screening (HTS) assay for the E2 enzyme UBC13 was developed by combining a fluorochrome (Fl)conjugated ubiquitin (fluorescence acceptor) with terbium (Tb)-conjugated ubiquitin (fluorescence donor) in a TR-FRET assay, such that the assembly of mixed chains of Fl- and Tb-ubiquitin creates a robust TR-FRET signal. In this particular study, this reagent enabled the identification of E2 inhibitors (Madiraju et al., 2012). While numerous reagents to assay the catalytic activity of DUBs have been reported, the

development of reagents enabling the monitoring of Ubiquitin ligase activity has been lagging behind due to the complexity of these enzymes. An elegant attempt to generate reagents to efficiently monitor the transthiolation activity of HECT- and RBR-E3 ligases is the development of the ‘Bypassing System’ (ByS) by Park et al. (2015). This approach exploits a simple design – a Ub thioester mimic in the form of UbMES (mercaptoethanesulfonate), permitting the direct transthiolation of the catalytic cysteine of the E3 ligase while eliminating the need for the E1 and E2 enzymes as well as ATP. Further development of this concept led to the generation of a fluorescent Ub thioester permitting the detection of both transthiolation and ligation activities of HECT E3 ligases (Krist et al., 2016). Given the facile detection method and the requirement for only the E3 enzyme and UbFluor, this mechanismbased reagent is well suited for high throughput screens (HTS) for Ub ligase inhibitors (Foote et al., 2017). What does the future hold? Unravelling the complexity of the highly sophisticated ubiquitination system is aided greatly by the development of numerous ABPs and reagents reporting on the dynamics and structural mechanisms of (de)ubiquitinating enzymes involved. Given the intrinsic role of Ub in the pathogenesis of a variety of diseases, most notably cancer and neurodegenerative diseases, enzymes involved in this system are emerging drug targets. The utility of these activity-based probes and reagents has been showcased by the discovery and validation of a USP7 inhibitor utilizing both Ub-AMC in the initial high-throughput screen and later Ub-VS in the validation studies (Reverdy et al., 2012; Lamberto et al., 2017). Without a doubt the next generation of Ub based tools will help increase our knowledge, ultimately leading to new diagnostic tools or therapeutics making it to the clinic. Although these recent advancements have helped gain insights into the functions of the engaged enzymes thereby facilitating more tailored solutions to interrogate their biology, it is becoming increasingly clear that these ABPs require innovation to address outstanding questions. The most pressing questions include dissecting DUB preference towards the Ub-linkage particularly

The Ubiquitin Toolbox |  27

of heterotypic and hybrid Ub chains; developing ABPs capable of capturing metalloprotease DUBs; advancing tools for specifically targeting distinct HECT and RBR-E3 ligases; and lastly, optimizing cell delivery methodologies for ABPs to enable incell enzymology. Customized tools – warranting study on a new complex layer of DUB recognition The advent of numerous ABPs and reagents for interrogating the different aspects of deubiquitinating enzymes, have enabled profound insights into the structural, biochemical and biological role of these ‘erasers’. More recently, the generation of tools specifically designed for dissecting the proteolytic processing of ubiquitin chains by DUBs have revealed profound differences among these proteases in their specificity. Adding to this complexity, the discovery of heterotypic and hybrid Ubiquitin chains warrants the development of customized tools in order to understand the regulatory roles of DUBs in this context. Given the recent insights that heterotypic Ubiquitin chains play a profound role in fine-tuning cellular responses (Xu et al., 2009), investigations into its biological and structural role need to be undertaken. To propel the study of their role, innovative ABPs recapitulating the structural and functional aspects of these mixed and branched Ubiquitin-chains need to be generated. Furthermore, the recent advances in synthetically obtaining Ubl proteins, permits the development of hybrid Ub/Ubl chains. Generation of such probes, especially for in-cell enzymology or proteomics context would be particularly conducive as the E3 ligases and DUBs regulating these heterotypic and hybrid Ub-chains are unknown (Xu et al., 2009). Furthermore, generating such complex linkages is a challenging feat as the E2/E3 enzymes generating these linkages in vitro are largely unknown and the known ones produce a mixture of linkage types that are difficult to separate by chromatography (Faggiano et al., 2016). Moreover, the modification of Ubiquitin or its Ubiquitin linkage by another PTM complicates the deciphering of the temporal order of events, which underlies the biological role of this modification. The urgent need for such ABPs and assay reagents is illustrated by the recently discovered MINDY DUBs, which preferentially cleave

K48 and K63 tetra-Ub linkages, raising the question whether they might display reactivity towards K48/K63 linkages (Xu et al., 2009; Ohtake and Tsuchiya, 2017). Since there are currently no ABPs recapitulating the mixed K48/K63 Ubiquitin linkage available, investigating this aspect is hampered. Currently, the metalloprotease DUBs have been neglected in the development of ABPs and reagents partly due to the difficulty of designing these tools. Unlike other deubiquitinating enzymes, metalloprotease DUBs do not have an active-site cysteine, but instead hydrolyse the isopeptide bonds of ubiquitinated substrates with a water-coordinated zinc ion. Designing chemical probes with potent and specific zinc-ion chelating reactive groups is prerequisite to generating an innovative toolkit for metalloprotease DUBs. Generally, metalloproteases are typically expressed as an inactive form (zymogen) inhibited by additional proteins and require proteolytic processing before rendering the active enzyme (Saghatelian et al., 2004). This additional layer of regulation, however, introduces another layer of complexity that must be taken into account when designing such reagents (Saghatelian et al., 2004). Introducing such innovative chemical probes would propel the study of these understudied deubiquitinating enzymes and enable the development of therapeutics. The quest for E3 ligase inhibitors – challenges and opportunities Given that E3 ligases are involved in the pathogenesis of a variety of diseases, most notably cancer, neurodegenerative diseases such as Parkinson’s, as well as numerous inflammatory diseases they are emerging drug targets (Goru et al., 2016; Uchida and Kitagawa, 2016). Although numerous assays, such as fluorogenic assays (Foote et al., 2017; Krist et al., 2017), FRET assays (Goldenberg et al., 2010), tandem ubiquitin-binding domains (Marblestone et al., 2012; Heap et al., 2017), bacterial or cellular two hybrid approaches (Levin-Kravets et al., 2016; Maculins et al., 2016), as well as biophysical methods (Regnström et al., 2013) have been reported, these approaches suffer from both low throughput, high number of false-positive or false-negative hits, and high costs. To overcome these shortcomings, a mass spectrometry-based assay using mono-ubiquitin to determine not only the E2/E3 enzyme activity facilitating highly sensitive and

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reproducible high-throughput inhibitor screening, was developed (De Cesare et al., 2018). Yet, one of the most challenging aspects to consider in such an undertaking is the lack of comprehensive prerequisite knowledge of the interacting E2–E3 enzyme pairs, which substantially modulate the biological outcome (De Cesare et al., 2018). Despite this progress, the current ABPs targeting the ubiquitin conjugating cascade utilize either a modular approach (e.g. E2–Ub probe conjugates) or are mechanistic-based relying on the active-site cysteine (Mulder et al., 2016) or the ATP-binding pocket (An and Statsyuk, 2013, 2016) thereby being limited to indiscriminately detecting HECT- and RBR- E3-ligases. This limitation could potentially be overcome by designing ABPs featuring increased selectivity for HECT/RBR-E3 ligases by utilizing specific Ub-variants generated by phage display (Zhang et al., 2016). Since some mechanistic aspects of E3 ligase-mediated catalysis is intrinsic to most E3 ligase probe designs, it excludes direct labelling of the scaffolding RING E3-ligases, which ironically comprise the vast majority of ligases that are pivotal in cancer development and progression (Wang et al., 2017). Yet, prerequisite for devising ABPs capable of selectively labelling RING E3 ligases is a priori knowledge of the specific interfaces between E2 and RING-E3 enzyme amenable to protein–protein interaction disruption. Probing ubiquitination in living cells Most ABP profiling experiments are performed using either recombinant enzymes or cell lysates, yet this does not recapitulate the activity of the enzymes in a cellular context. Since lysing cells results in disruption of the cellular compartmentalization as well as in dilution of the enzymes which might affect enzyme reactivity, delivery of DUB and ubiquitin ligase ABPs into intact cells is of critical importance. However, to achieve this, several methods including electroporation (Mulder et al., 2016) or the use of cell-penetrating peptides attached to the Ub-ABP (Gui et al., 2018; Hameed et al., 2018) have been reported. Additionally, the introduction of ABPs into living cells permit the visualization and in-cell enzymology of the ubiquitin cascade enzymes in a spatial and temporal context. The critical need for an intact cellular environment for proper enzymatic function of Ubiquitin enzymes arises from the interaction with protein complexes

as well as their substrates, but also the intrinsic regulation by cellular signalling events such as phosphorylation (Sowa et al., 2009; Heideker and Wertz, 2015). The significance of additional posttranslational modification, e.g. phosphorylation, of DUBs to enhance their proteolytic activity is highlighted by the necessity of serine phosphorylation of OTUD5/DUBA (Huang et al., 2012). Furthermore, cross-regulation of DUBs with E2 enzymes (Wiener et al., 2012) and E3-ligases (Heideker and Wertz, 2015) underscore the significance of studying the ubiquitin cascade in living cells. One notable example of aforementioned interactions is the well characterized deubiquitinating enzyme USP7, which binds to the E3 ligase MDM2 and its substrate tumour suppressor p53 through its TRAF-domain (Sheng et al., 2006). Considering the significance of an functional cellular environment for the enzymatic function of the ubiquitin enzymes, their biochemical study should be conducted in living cells thus meriting ABPs compatible with in-cell enzymology. Another facet necessitating in-cell enzymology using ABPs is the application in proteomics to access not only the functional consequence of these interactions, particularly in the context of pharmacological inhibition (De Cesare et al., 2018). Conclusion Since the first ABP targeting DUBs, the field has brought forth an assortment of tools for interrogating a wide scope of biochemical and structural questions. The ensuing course of development illustrates how the development of activity-based probes and assay reagents for DUBs led to the discovery of new DUBs subsequently spawning the innovation of specialized reagents. While a variety of tools are reported for DUBs, the complexity of sequentially targeting an enzymatic cascade hampered the development of analogous advancements for the ubiquitin activating, conjugating, and ligating enzymes. Although the first ABPs targeting the ubiquitin activating enzyme have been reported almost a decade ago, reagents for the downstream enzymes are now slowly starting to emerge. One ABP that stands out is UbDha, which has the unique capability of being sequentially transferred through the ubiquitin cascade in a manner reminiscent to native Ubiquitin. Conclusively, the current

The Ubiquitin Toolbox |  29

platform of reagents and ABPs have the potential to accelerate drug discovery efforts targeting all aspects of the ubiquitin cascade. Yet, the frontier of Ubiquitin activity-based probe and reagent development lies in the introduction of innovative technologies and unique concepts enabling the dissection of many enigmatic aspects of ubiquitination as well as accessing enzymes previously not targeted by conventional ABP designs. References

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Recent Highlights: Onco Viral Exploitation of the SUMO System Domenico Mattoscio1,2*, Alessandro Medda3 and Susanna Chiocca3*

3

1Department of Medical, Oral, and Biotechnology Science, University of Chieti-Pescara, Chieti,

Italy.

2Center on Aging Science and Translational Medicine (CeSI-MeT) ‘G. d’Annunzio’, University of

Chieti-Pescara, Chieti, Italy. Department of Experimental Oncology, European Institute of Oncology IRCCS, Milan, Italy.

3

*Correspondence: [email protected] and [email protected] https://doi.org/10.21775/9781912530120.03

Abstract Small ubiquitin-like modifier (SUMO)ylation is a crucial post-translational modification that controls functions of a wide collection of proteins and biological processes. Hence, given its pleiotropic role, viruses have developed many approaches to usurp SUMO conjugation to exploit the cellular host environment for their own benefit. Consistently, cancer cells also frequently impact on SUMO to force cellular transformation, underlining the importance of SUMO in health and diseases. Therefore, after a brief introduction to the multistep SUMOylation pathway, in this chapter we will focus our attention on several examples of strategies adopted by oncogenic viruses to hijack SUMOylation in order to promote infection, persistence and malignant transformation of host cells. Introduction The Small Ubiquitin-like Modifier (SUMO) proteins are involved in post translational modification (PTM) of target proteins (Matunis et al., 1996; Kamitani et al., 1997). The name SUMO comes from a structural similarity with ubiquitin and from the similar mechanism by which it is attached to target proteins (Mahajan et al., 1997). Indeed, both ubiquitination and SUMOylation

are reversible processes catalysed by a cascade of enzymes, namely E1, E2 and E3 proteins (Gong et al., 1997; Mahajan et al., 1997; Johnson and Gupta, 2001). SUMO proteins The expression of SUMO proteins is conserved among eukaryotes. Lower eukaryotes have only one SUMO, while higher eukaryotes express three or more SUMO paralogues. In particular, in humans five different isoforms of SUMO are present, differing for response to physiological or stress conditions, tissue-specificity, and the ability to form SUMO chains [recently reviewed in Yang et al., (2017)]. SUMO1 is 101 amino acids protein found almost always conjugated to targets, and therefore often associated to physiological processes (Shen et al., 1996; Yang et al., 2017). SUMO2 consists of 95 amino acids and shares 95% homology with SUMO3 (103 amino acids), differing for only three N-terminal residues, and showing the same molecular functions, therefore often referred as SUMO2/3. They show only 45% homology with SUMO1 but they present a very similar tridimensional structure. SUMO2/3 are conjugated mostly under stress conditions and they are able to form chains (Mannen et al., 1996; Lapenta et al., 1997). SUMO4 seems to be expressed only in lymph nodes, kidneys and

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spleen. SUMO4 has not been well characterized yet and its role still needs to be elucidated. It is probably non-conjugated under physiological conditions and it has been associated to diabetes (Wang et al., 2006). Finally, the expression of a fifth SUMO isoform has been recently reported (Liang et al., 2016). SUMO5 seems to be a conserved 84 amino acids protein whose mRNA is expressed at high levels in testes and peripheral blood lymphocytes, and at lower levels also in placenta, lungs and liver. Conjugation with this novel SUMO variant facilitates the formation of Promyelocytic Leukaemia Nuclear Bodies (PMLNB), structures rich in SUMOylated proteins that regulate a variety of cell functions. The SUMO machinery The SUMOylation process is carried out in different steps (Fig. 3.1). Initially, SUMO is processed by a protease (belonging to the SENP family, as described in more details below), that generates the mature form consisting of a C-terminal diglycine (Hickey et al., 2012). This motif is required for the following step, in which the SUMO E1 enzyme activates SUMO. There is one only SUMO

E1 enzyme expressed in mammalian cells, a heterodimer composed by SUMO-activating enzyme subunit 1 (SAE1) and ubiquitin-like activating enzyme subunit 2 (SAE2/UBA2) (Desterro et al., 1999). SUMO is adenylated by the E1 complex in an ATP·Mg2+-dependent reaction and transferred to the catalytic Cys of the UBA2 subunit by an E1~SUMO thioester bond. Then, the unique SUMO E2 conjugating enzyme, ubiquitin-like conjugating 9 (UBC9), receives SUMO on a conserved catalytic cysteine, forming an E2~SUMO thioester complex (Tong et al., 1997; Duan et al., 2009). The E2 enzyme can attach SUMO to substrates, with the formation of an isopeptide bond between the carboxy-terminal carboxyl group of SUMO and a ε-amino group of the substrate acceptor Lys residue. UBC9 can be itself modified by different PTMs which increase or decrease its activity and localization, and confer substrate specificity (Knipscheer et al., 2008; Su et al., 2012). UBC9 can interact directly with some SUMO substrates but more often it needs the help of SUMO E3 enzymes, ligases that are able to give specificity to the targets (Sachdev et al., 2001; Tatham et al., 2005). Opposite to the unique E2, there are different SUMO

Figure 3.1  The SUMO conjugation system. Ulp1/SENP proteases catalyse cleavage of the C-terminal domain of SUMO proteins, exposing a diglycine motif. Processed SUMO is transferred to a cysteine of the heterodimeric E1 enzyme Uba2/SAE1. SUMO is then conjugated to a cysteine of UBC9, the E2 enzyme and attached to a lysine residue of the consensus motif on target proteins. The conjugation is often facilitated by an E3-ligase, which enforces the interactions among the involved components. SUMOylation is a reversible pathway where the Ulp1/SENPs proteases dictate the de-SUMOylation process.

Oncoviruses and SUMOylation |  37

E3 ligases, selective for SUMO1 or SUMO2/3 [described in Mattoscio and Chiocca (2015)]. Some E3 ligases act by orientating the E2~SUMO thioester in an optimal conformation for catalysis without directly contacting the substrate, while others facilitate the release of SUMO from E2 [reviewed in Wilkinson and Henley (2010)]. As mentioned before, SUMOylation is a reversible process governed by the action of two families of proteases that deconjugate SUMO from substrates. They include Ubl-specific proteases and sentrinspecific proteases (Ulps and SENPs, respectively) (Li and Hochstrasser, 2000). SENPs The human SENP family is composed of seven members: SENP1, SENP2, SENP3, SENP5, SENP6, SENP7 and SENP8, even if SENP8 is not specific for SUMO but acts on Nedd8, another ubiquitin-like protein (Hickey et al., 2012). SENPs are cysteine proteases with a papain-like folded catalytic domain and specific N-terminal domains crucial for their own regulation and for substrate selection (Hay, 2007). SENP proteases regulate both the level of processed SUMO and the rate of substrate modification by counterbalancing SUMO conjugation [recently reviewed in Kunz et al. (2018)]. In the maturation process they hydrolyse a peptide bond close to the C-terminus of SUMO precursors, eliminating the very C-terminal amino acids from SUMO1, SUMO2 and SUMO3 and exposing two glycine residues. In SUMO1–3, the diGly motif is preceded by a glutamine (Q) and threonine (T), while SUMO4 exhibits a PTGG motif, in which the proline residue confers resistance to SENPmediated cleavage (Owerbach et al., 2005). In the deconjugation process, SENPs cleave an isopeptide bond that links SUMO moieties to the ε-amino group of lysine residues. The mechanism by which SENP1 and SENP2 exert their functions has been described by X-ray crystallography. In vitro protease assays with the isolated catalytic domains demonstrate the processing activity of SENP1 and SENP2 on all three SUMO precursors (Reverter and Lima, 2006; Shen et al., 2006a). However, they exert differential activities towards distinct precursors. In particular, SENP2 is most active on SUMO2, then SUMO3 and SUMO1, while SENP1 prefers SUMO1.

Probably these differences are due to the amino acid sequences of the C-terminal tail. SENP5 has been found to have a marked preference for SUMO2 cleavage, while SENP6 and SENP7 are not able to process SUMO for maturation. The SUMO system leaves a dilemma on how it is possible to achieve specificity on SUMOylation of a myriad of proteins with the small numbers of conjugating and deconjugating enzymes available. In some cases, specific biological processes are regulated by distinct deconjugation events. Though, in many cases, a single SENP may act on larger groups of SUMOylated proteins (Psakhye and Jentsch, 2012; Jentsch and Psakhye, 2013). Moreover, alternative splicing and PTMs of SENPs are important to determine their localization and their protease activity. To summarize, the special control of SENPs is a fundamental principle for deSUMOylation regulation (Kunz et al., 2018). The SUMO consensus motifs The main SUMO consensus motif existing in the primary structure of SUMOylated protein is ψKX(D/E), where ψ is a large hydrophobic residue, X is any amino acid and K is the acceptor lysine. These residues directly bind UBC9 and are crucial for a stable interaction between the E2 enzyme and the substrate (Rodriguez et al., 2001). In addition to the canonical four amino acid SUMO consensus motifs, longer sequences that include both SUMO consensus motifs and additional elements have been identified in some SUMO substrates (Gareau and Lima, 2010). Among these, phosphorylationdependent SUMO motifs (PDSMs) and negatively charged amino acid-dependent SUMO motifs (NDSMs) are present. PDSMs present a SUMO consensus motif located adjacent to a phosphorylation site, ψKX(D/E)XXSP. Phosphorylation increases SUMO conjugation levels because the phosphorylated Ser side chain interacts with a basic patch on the E2 surface, extending interactions with the E2 enzyme beyond recognition of the SUMO consensus motif. This mechanism is probably shared with proteins that contain NDSMs, which comprise negatively charged residues that are C-terminal to the SUMO consensus site in the place of the phosphorylation site of PDSMs, although NDSMs may interact with a different subset of Lys residues on the UBC9 surface (Yang et al., 2006; Mohideen et al., 2009). Recent studies

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revealed new motifs for SUMO conjugation, including inverted consensus motifs and motifs with an N-terminal hydrophobic cluster. These alternative motifs are probably important to give specificity to E2–substrate interactions through direct interaction with the E2 (Impens et al., 2014). SIM SUMO interacting motifs (SIMs) establish noncovalent hydrophobic interaction between SUMO and the target proteins. Canonical SIMs contain a core of hydrophobic residues that can be preceded or followed by negatively charged amino acids that take contact with hydrophobic groove on SUMO with following basic residues. This interaction is generally weak but can be increased by the binding of multiple SIMs to SUMO chains. Crucial hydrophobic and basic residues involved in SIM binding are conserved among the SUMO paralogues. However, the isoforms might differ in the placement of their hydrophobic groove suggesting that the arrangement of hydrophobic and acidic residues in SIMs might dictate their ability to bind specific SUMO isoforms (Hecker et al., 2006; Kerscher, 2007). STUbLs SUMO Targeted Ubiquitin Ligases (STUbLs) are Ubiquitin E3 enzymes able to recognize SUMOylated proteins and to interact with them through SIMs. They attach ubiquitin chains to SUMOylated proteins to target them for degradation by the proteasome. STUbLs constitute an important regulatory mechanism to control the levels of the SUMO conjugated form of a protein (Sriramachandran and Dohmen, 2014). The human RING Finger protein 4 (RNF4) is one of the best studied STUbL, containing at least three SIMs. These motifs mediate a similar non-covalent interaction with SUMO1 and SUMO2, with a preference for chains of a length of at least three SUMO moieties. RNF4 works as homodimer, in which the RING domains of both subunits take contact with a single ubiquitin-charged E2 (Sun et al., 2007). SUMO functions SUMOylation is involved in many different biological processes and can confer different properties to substrate proteins. A high number

of known SUMOylation targets are nuclear proteins, involved in DNA repair, regulation of transcription and chromatin structure. Many important nuclear targets of signalling pathways can be SUMOylated. SUMOylation is important for subcellular localization of proteins, competes with other PTMs, and also participates to protein– protein interaction. SUMOylation can also change the interaction between DNA and RNA, alter enzymatic activity and protein conformation, and modulate other modifications (recently reviewed in (Zhao, 2018). In the following paragraphs we will describe some paradigmatic example of how SUMOylation can impact on the activity of important selected targets. RanGAP RanGAP, the first protein shown to be SUMO modified, is important for nuclear import. Unmodified RanGAP is cytoplasmic, whereas SUMO-modified RanGAP is associated with the nuclear pore. SUMOylation of RanGAP increases its interaction with the SUMO E3 ligase Ran binding protein 2, a component of the nuclear pore complex. Localization of the RanBP2 SUMO E3 ligase at the nuclear pore could be important for a broad role for SUMO in regulation of nuclear trafficking (Matunis et al., 1996). Promyelocytic leukaemia protein Promyelocytic leukaemia protein (PML) is posttranslationally modified by SUMO and is localized in subnuclear structures named PML nuclear bodies, structures highly enriched in SUMOylated proteins. PML bodies host more than 150 proteins with a wide range of functions, such as DNA repair, stress response, senescence, anti-viral immunity, and tumour suppression. Notably, a variety of SUMO-modified proteins including transcription factors, chromatin modifiers, and proteins involved in genomic maintenance, are expressed in PML nuclear bodies together with SUMO E3 ligases and SUMO-specific proteases. SUMO-modified PML probably supports some protein–protein interactions important for assembly or stability of this subnuclear domain. Focused studies of cellular membrane-less structures suggest that proteins able to form inter-molecular multivalent interactions can constitute large oligomers and phase separate

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from the surrounding solution. These proteins can use their interaction domains or intrinsically disordered regions to recruit additional macromolecules even in high concentrations, maybe promoting certain biological processes (Shen et al., 2006b). SUMO modulation of chromosomes and chromatin SUMO deficiency drastically changes chromosome integrity and segregation. Indeed, SUMO enzymes are enriched at centrosomes, very important structures that support kinetochores for microtubules attachment during cell division (Lapenta et al., 1997). Topoisomerase II is recruited to centromeres on SUMOylation of its non-catalytic C-terminus, to uncoil intertwined DNA before anaphase and facilitate centromeric segregation. Centromeric histones and chromatin regulators are also regulated by SUMO conjugation. In particular, SUMOylated Orc2 recruits the histone demethylase KDM5A to demethylate H3K4me3 into H3K4me2, enhancing non-coding RNA synthesis from the locus and subsequent heterochromatin maintenance. Moreover, Aurora B kinase deSUMOylation facilitates its localization to the spindle mid-zone, essential step during mitosis [recently reviewed in Zhao (2018)]. The SUMO pathway does not regulate only centromeric regions. For example, the heterochromatin assembly factor HP1 is SUMOylated to promote its association with RNA transcripts located at these regions. In addition, SENP7 activity is important for HP1 regulation in order to retain it at heterochromatin, even if the molecular details are still unclear (Maison et al., 2012). SUMOylation also affects chromatin modifiers such as histone deacetylase 1 (HDAC1), an essential epigenetic regulator of a conserved family of deacetylases frequently involved in cancer progression (Ropero and Esteller, 2007). Indeed, in non-tumourigenic cells, SUMOylation of HDAC1 by SUMO1 promoted by the overexpressed PIASy triggers its ubiquitination and degradation in a proteasome-dependent manner, thus reducing HDAC1 expression. Conversely, in breast cancer cell lines, HDAC1 is preferentially conjugated by SUMO2 that protects HDAC1 from ubiquitin conjugation and degradation. Therefore, SUMOylation

significantly affects the expression and activity of an important chromatin modifier involved in breast cancer progression (Citro et al., 2013). In addition, SUMO plays a fundamental role in DNA double-strand breaks (DSB), where SUMOylation enables broken DNA ends to move outside to prevent illegitimate repair of repetitive sequences. Similarly, SUMO promotes movement of target eroded telomeres and DSB to nuclear periphery, in a STUbLs-mediated mechanism that interacts with SUMOylated DNA repair proteins leading to their proteasomal degradation [recently reviewed in Garvin and Morris (2017)]. SUMO in DNA damage SUMOylation is important in the DNA damage checkpoint pathway. In both yeast and human cells, SUMOylation of DNA damage proteins occurs in parallel with checkpoint mediated phosphorylation. Interestingly, changes in the checkpoint pathway can modify SUMOylation events: decreasing Ataxia telangiectasia and Rad3 related (ATR) checkpoint kinase increases protein SUMOylation. Moreover, Ataxia-telangiectasia mutated (ATM) checkpoint kinase is able to increase SENP2 transcription in particular contexts, but also ATM seems to promote SUMOylation in the absence of ATR. This phenomenon suggests a context-dependent crosstalk between these pathways (Munk et al., 2017). As described, SUMOylation affects almost all cellular activities, resulting as a key pathway regulating cells physiology. However, conversely, it is evident that alterations in normal SUMOylation could completely subvert cell functions [reviewed in Flotho and Melchior (2013)]. Therefore, SUMO pathway components are frequently altered in human diseases such as cancer (Mattoscio and Chiocca, 2015; Seeler and Dejean, 2017), and often exploited by viruses. Interestingly, oncogenic viral infections can also increase metabolic and proangiogenic markers through expression of a very specific domain that also controls SUMO enzymes expression (Pozzebon et al., 2013). Viral exploitation of SUMOylation has been recently detailed in elegant reviews (Mattoscio et al., 2013; Lowrey et al., 2017; Wilson, 2017), to which readers can refer. In the following sections we will provide some classic examples on how oncogenic viruses

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impact SUMOylation to increase their ability to infect, persist, and transform host cells. Oncoviruses exploitation of the SUMO pathway Infection with oncogenic viruses is also involved in cancer pathogenesis, accounting for about 15% of total malignancies in 2012 (Plummer et al., 2016). Seven viruses are associated with human cancers, including hepatitis B virus (HBV), hepatitis C virus (HCV), high-risk human papillomaviruses (HPV), Epstein–Barr virus (EBV), Kaposi’s sarcoma herpesvirus (KSHV, also known as human herpesvirus type 8 HHV-8), human T-cell leukaemia virus type 1 (HTLV-1), and the recently emerged Merkel cell polyomavirus (MCPyV) (Mesri et al., 2014; Spurgeon and Lambert, 2013). Hepatitis B virus HBV is a partially double-stranded circular DNA virus belonging to the Hepadnaviridae family. Persistent infection with HBV is associated with several liver diseases such as hepatocellular carcinoma (HCC), the most common cancer of the liver [reviewed in Di Bisceglie (2009)]. HCC pathogenesis is a combination of both indirect effects as a consequence of the chronic inflammatory condition due to the persistent HBV presence in liver cells, and directly through viral proteins expression. In particular, HBV X antigen (HBx), a viral product that acts as transcriptional cofactor during viral replication, is also able to promote cellular transformation altering crucial cellular pathways involved in cell growth, DNA repair, apoptosis, and cell cycle progression [recently reviewed in Xie (2017)]. Notably, several of these modifications are mediated by exploitation of SUMO pathway by HBx. Indeed, in HBV infected cells, HBx promotes deSUMOyation and relocalization of the host transcription factor Sp110, usually conjugated to SUMO1 and expressed inside PML-NBs. The detachment of SUMO1 moiety and the resulting Sp110 differential distribution in infected cells increases viral DNA load, decreases apoptosis and increases viability of hepatocytes, and markedly affects expression levels of genes involved in type I interferon pathway, a common response mechanism to viral infections. Mechanistically, HBx may

promote the formation of Sp110–SENP1–HBx complex able to catalyse SUMO1 removal from Sp110 and to translocate HBx to Sp110 gene promoters in order to reprogram host gene expression and to trigger viral proliferation (Sengupta et al., 2017). These findings highlight the importance of the SUMOylation and deSUMOylation switch in the infection lifecycle and tumorigenesis triggered by HBV. In addition, HBx expression in mice and human cell lines prompts cell growth altering the SUMOylation status of E-cadherin, a membrane protein crucially involved in epithelial-mesenchymal transition (EMT). Opposite to SUMOylation of Sp110, HBx expression promotes SUMO1 and 2/3-conjugation to E-cadherin, leading to E-cadherin degradation, EMT-transition, loss of cell-to-cell contact, and overgrowth of hepatocytes (Ha et al., 2016). Notably, SUMO1, SUMO2/3, SAE1/2, UBC9, and SENP2 are differentially expressed in HCC and play key roles in HCC pathogenesis [(Liu et al., 2015), as recently reviewed in Tomasi and Ramani (2018)], further underlining the importance of SUMOylation in liver cells transformation. However, if these alterations are directly mediated by HBV proteins or are a consequence of cancer growth is still an unresolved issue and will not be further described in this chapter. Hepatitis C virus Together with HBV, HCV is another important aetiological agent of HCC (Di Bisceglie, 1995). HCV is an enveloped, single-stranded RNA virus belonging to the Flaviviridae family. Similarly to HBV, HCV can promote HCC development as a consequence of the chronic inflammatory condition associated with its persistence in hepatocytes, or through direct effects mainly mediated by the viral core, non-structural proteins 3 (NS3), and NS5A, crucial players in viral replication and in alteration of the host gene expression landscape [reviewed in Irshad et al. (2017)]. In particular, NS5A affects cellular pathways involved in liver cell proliferation, apoptosis immune response, and DNA repair (Irshad et al., 2017), and requires SUMOylation to increase its stability in host cells and to promote HCV replication. Indeed, NS5A is SUMOylated in the context of HCV infection by both SUMO1 and SUMO2/3, perturbing ubiquitination occurring at the same target lysine, and suppressing NS5A

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proteasomal degradation. In addition to stability, SUMOylation also regulates the interaction of NS5A with NS5B occurring during replication complex formation (Lee et al., 2014), a key event for viral replication. Indeed, SUMO1 is overexpressed in infected Huh7.5 cells (Akil et al., 2016), and abrogation of SUMO conjugation by Ubc9 silencing markedly impairs HCV RNA replication (Lee et al., 2014; Akil et al., 2016), suggesting the importance of SUMOylation for HCV lifecycle in hepatocytes. However, on the contrary, a recent report showed that SUMO removal obtained by silencing of PIAS2 during HCV infection enhances stability and expression of NS3 and NS5A, and increases HCV replication, in a SUMO1-dependent manner (Guo et al., 2017). Reasons for these apparent discrepancies are currently unknown and future studies are therefore needed to better clarify whether HCV-mediated tumorigenesis benefits or is dampened by SUMO. Human papilloma virus High-risk HPV types (16, 18, 31, 33, 35, 39, 45, 51, 52, 56, 58, and 59) (Bouvard et al., 2009) are the aetiological agent of cervical cancer and are also associated with other anogenital malignancies, such as vulvar, vaginal, anal, and penile cancers, and with a significant proportion of oropharyngeal tumours [reviewed in (zur Hausen, 2009)]. HPVs are double-stranded DNA viruses that promote malignant transformation in chronically infected keratinocytes of epithelia mainly due to the viral oncoproteins E6 and E7 that, through degradation of tumour suppressors p53 and retinoblastoma (pRb), modify fundamental cellular pathways involved in cell cycle, apoptosis, DNA repair, and senescence [reviewed in Tommasino (2014)]. In addition to E6 and E7, HPV infection in keratinocytes entails the concerted and sequential action of other early non-structural proteins E1, E2, E4 and E5, and viral capsid protein L1 and L2 [reviewed in Woodman et al. (2007)]. Most of these viral proteins exploit SUMOylation to subvert cellular pathways and promote viral persistence in the host. E2 is a multifunctional regulatory protein that binds to viral DNA and interacts with cellular proteins to regulate viral gene expression, partitioning and replication, and to modify host transcriptome (reviewed in (McBride, 2013). HPV18 E2 is a substrate for mono SUMOylation in vitro, in an E. coli

expression system, and in HeLa cells after overexpression of HPV16 E2, Ubc9, and either SUMO1, 2, or 3, despite a preference for SUMO2/3. Notably, the defective E2 SUMO mutant shows defects in transcriptional ability, suggesting a crucial role for SUMOylation in mediating E2 activities during HPV-mediated transformation (Wu et al., 2008). In addition, SUMOylation at K292 increases E2 expression levels after exogenous overexpression of SUMO components and by endogenous elevation of SUMOylation obtained after heat shock, due to a SUMO-mediated inhibition of E2 ubiquitination and degradation (Wu et al., 2009). Combining these results with the observation that SUMO2/3 is progressively up-regulated during keratinocytes differentiation (Deyrieux et al., 2007), the emerging scenario depicts that the increased SUMO2/3 expression in suprabasal layer of epithelium stabilizes E2, increases E2 concentration and activity, and promotes viral production (Wu et al., 2007). The E6 and E7 viral oncoproteins drive malignant transformation mostly due to the degradation of p53 and pRb, respectively. However, in addition to these two well characterized pathways, a number of other cellular proteins are affected by E6 and E7 during viral infection and transformation [reviewed in Moody and Laimins (2010)]. Among these, clever strategies are adopted by HPV oncoproteins to hijack SUMO during infection and tumorigenesis. HPV16 E6 and E7 overexpression in the natural host of the virus, primary human keratinocytes, significantly increases the accumulation of UBC9 and SUMO1-conjugated species (Mattoscio et al., 2017). Notably, similar results were also found in human samples during the natural evolution of cervical and oropharyngeal cancer (Mattoscio et al., 2015, 2017), underlining the importance of SUMO alterations during HPV transformation. Mechanistically, HPV16 E6/E7 prevents the autophagy-dependent UBC9 degradation obstructing the final step of autophagic pathway in E6/p53-dependent manner (Mattoscio et al., 2017, 2018). The resulting increased UBC9 level confers apoptosis resistance to the infected keratinocyte (Mattoscio et al., 2017), thus extending HPV persistence in the host and triggering cellular transformation. However, this E6-mediated UBC9 accumulation seems to be a cell-specific mechanism dependent on the cellular background of analysed cells. Indeed, E6 overexpression in

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immortalized, p53 defective, cell lines drive UBC9 degradation through proteasomal-dependent pathway (Heaton et al., 2011) while in primary, p53 competent cells, E6 triggers UBC9 accumulation following autophagy defects. These and other results (Boggio et al., 2004, 2007) highlight a dual way to control UBC9 levels, further underlining the fundamental role of the SUMO E2 enzyme in cellular physiology. In addition to impact on UBC9 expression, E6 also co-opts SUMOylation to re-direct activities of cellular transcription factors. Indeed, the transcriptional co-activator hADA3 (human alteration/deficiency in activation3), is down-regulated by HPV16 E6 in cervical cancer cells. In contrast to HPV E2 that exploits SUMO-conjugation to protect its own ubiquitin-dependent proteasomal degradation, HPV16 E6 triggers SUMOylation to induce ubiquitin attachment and hADA3 degradation. SUMO-dependent hADA3 deprivation encourages cell proliferation, migration, and anchorage independent growth of cervical cancer SiHa cells, pointing to the important role of SUMOylation in malignant transformation (Chand et al., 2014). Similarly, in addition to contributing to UBC9 overexpression, E7 also usurps SUMOylation to regulate levels and activity of a transcription factor crucially involved in cell cycle progression, cell proliferation, and DNA damage response, Forkhead box M1b (FoxM1b). FoxM1 de-regulation occurs in a variety of malignancies [reviewed in Myatt and Lam (2007)], where its activity and expression are frequently modified by PTMs [reviewed in van der Horst and Burgering (2007)], including SUMOylation. Indeed, in vivo SUMOylation assays in HEK293T cells show that FoxM1 could be modified by all three SUMO paralogues after physical interaction with UBC9 and PIAS1. Similar results were obtained also in MCF-7 cells (Myatt et al., 2014). After SUMO conjugation, FoxM1 is rapidly degraded and re-localizes from nucleus to cytoplasm, suggestive of a negative regulatory loop mediated by SUMOylation to turn off its transcriptional activity (Myatt et al., 2014; Jaiswal et al., 2015). Notably, HPV16 E7 interferes with SUMO loading on FoxM1 by inhibiting its association with UBC9, in turn reducing FoxM1 SUMOylation and protecting it from re-localization and degradation ( Jaiswal et al., 2015). SUMOylated FoxM1

increases cell proliferation and delays mitotic progression (Myatt et al., 2014), indicating the importance of the E7-mediated SUMO manipulation in the context of HPV-mediated cellular transformation. Finally, SUMOylation of the late structural capsid protein L2 also plays crucial role in HPV infectivity and cellular transformation. Indeed, modification with SUMO2/3 increases L2 halflife and inhibits interaction with the other capsid protein L1, suggesting that capsid assembly could be modulated by SUMOylation during HPV infection (Marusic et al., 2010). Epstein–Barr virus EBV was the first virus clearly connected with human malignancies, since it was isolated in 1964 in cultured lymphoblasts from Burkitt’s lymphoma cells (Epstein et al., 1964). Since then, EBV infection was also consistently associated with a number of other malignancies such as nasopharyngeal cancer, Hodgkin’s and non-Hodgkin’s lymphomas, and a subset of gastric cancers [reviewed in Thompson and Kurzrock (2004)]. EBV is a double-stranded DNA Herpesvirus that could establish latent and lytic infection in lymphoblastoid cells, characterized by restricted viral gene expression and life-long persistence, and with virions production, respectively, in lymphocytes and epithelial cells. Several proteins are involved and expressed in lytic reactivation, to promote cell proliferation, virus production, and oncogenesis [reviewed in Tsurumi et al. (2005)]. Among them, the transcriptional activator Zta could be modified by both SUMO1 and SUMO2/3 (Adamson and Kenney, 2001; Hagemeier et al., 2010) at K12. SUMOylated Zta associates with and carries HDAC3 on its targeted promoters which, in this way, acetylates and exerts an inhibitory activity at Zta-responsive genes (Murata et al., 2010). Consistently with the role of SUMOylation in mediating repression of Zta, the SUMO defective mutant increases gene expression and re-activation of latent EBV. Notably, the SUMO-mediated repression of Zta in vivo could be reverted by both the viral encoded EBV kinase (EBV-PK) and RanBPM during infection, reducing Zta SUMO conjugation, promoting transcription of Zta genes and replication of the viral genome (Hagemeier et al., 2010; Yang, Y.C. et al., 2015). SUMOylation and activity of Zta are also

Oncoviruses and SUMOylation |  43

finely regulated by interaction of SUMOylated Zta with SIM motifs of the EBV protein kinase BGLF4. After SUMO-mediated Zta–BGLF4 interaction, the kinase activity of BGLF4 abolishes Zta SUMOylation, activating virus production (Li et al., 2012). Similar to Zta, the activity of the other EBV protein involved in lytic reactivation, Rta (Tsurumi et al., 2005), is crucially regulated by SUMO. Yeasttwo-hybrid screen identifies UBC9 and PIAS1 as binding partners of Rta, that is SUMOylated both in vitro and in vivo during the early stages after lytic induction of EBV infection. However, contrary to Zta, SUMO1 conjugation increases the transcriptional ability of Rta, suggesting a crucial role for SUMOylation for EBV lytic reactivation. Indeed, since Rta mediates Zta transcription (Adamson and Kenney, 2001), SUMO could both serve initially as activator of lytic phase by conjugating to Rta and promoting transcription of genes such as Zta, and then as modulator of EBV reactivation through SUMOylation-mediated Zta repression triggered by viral EBV-PK and cellular RanBPM. Latent membrane protein 1 (LMP1) is primarily involved in EBV oncogenesis due to its ability to mimic CD40 receptor and to constitutively transduce growth signals that trigger tumorigenesis in infected cells (Gires et al., 1997). LMP1 physically interacts with UBC9 to increase protein SUMOylation in latent infected cells (Bentz et al., 2011). Among SUMOylated proteins, LMP1–UBC9 interaction promotes SUMO conjugation of Interferon Regulatory Factor 7 (IRF7) to promote its nuclear localization and increases its stability in EBV infected cells. However, despite nuclear accumulation, SUMOylation inhibits IRF7 association with chromatin, thus reducing its transcriptional activity and the ability to induce innate immune response (Bentz et al., 2012). Moreover, LMP1 aids to preserve viral latency (Adler et al., 2002) and SUMOylation plays pivotal roles also in EBV lytic reactivation. Indeed, LMP1 triggers SUMOylation of the transcriptional repressor KRAB-associated protein-1 (KAP1). In EBV-transformed lymphoblastoid cell line, SUMOylated KAP1 associates with viral EBV lytic promoters OriLyt, ZTA and RTA, promoting the transcriptional repression that contributes to the maintenance of viral latency (Bentz et al., 2015).

In addition to LMP1, a recently reported genome-wide screening identifies other EBV proteins having global effects on host SUMOylation. In particular, overexpression of the transcriptional activator BRLF1 consistently decreased levels of both SUMO1 and SUMO2 conjugated proteins in transfected 293T and HeLa cells, while six EBV proteins up-regulated SUMOylation. Among them, expression of SM, an mRNA binding protein, increases levels of SUMO1 and to less extent SUMO2 conjugated proteins. This effect is due to the ability of SM to interact and bind UBC9 and SUMO, thus acting as an E3 ligase that promotes SUMO conjugation of cellular proteins such as p53. Consistently, SM depletion in AGS-EBV infected cells reduces global SUMOylation levels, suggesting the ability of SM to affect SUMOylation during viral lytic infection (De La Cruz-Herrera et al., 2018). In addition to proteins, EBV also encodes a variety of microRNAs (miRNAs) during viral infection and oncogenesis. Bioinformatic analysis based on miRNA target prediction identified 575 proteins of the SUMO interactome that could be potentially targeted and modulated by EBV miRNAs and a set of 14 predicted 3′ UTR were also experimentally validated in luciferase reporter assays. SUMO proteins targeted by EBV miRNAs are mainly involved in cancer-related functions such as proliferation, apoptosis, growth signalling, and intercellular communication, suggesting that miRNAs play fundamental roles during EBV carcinogenesis (Callegari et al., 2014). Accordingly, the EBV-encoded miR-BHRF-1 promotes accumulation of SUMO2/3 conjugated proteins during lytic infection due to down-regulation of RNF4 (Li et al., 2017). Kaposi’s sarcoma-associated herpesvirus KSHV is responsible for Kaposi’s sarcoma, a malignancy commonly occurring in AIDS patients. In addition to Kaposi’s sarcoma, KSHV has been detected in primary effusion lymphoma and in multicentric Castleman’s disease [reviewed in Ganem (2006)]. KSHV is a double-stranded DNA herpes virus that primarily infects endothelial and B cells that frequently exploits SUMOylation to promote its replication [recently reviewed in Chang and Kung (2014)]. Similar to EBV, KSHV infection

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cycle can be divided in lytic and latent phases. Viral reactivation can be triggered by a number of specific environmental stimuli and by the viral protein K-Rta. The lytic phase is characterized by a short period where viral genes are expressed, where during the latency there is the expression of a limited number of viral genes without production of viral particles (Aneja and Yuan, 2017). One of the most abundantly genes expressed during latent phase, the latency-associated nuclear antigen (LANA) protein, is a crucial regulator of dormant infections, viral reactivation and cellular transformation [reviewed in Uppal et al. (2014)]. To exert its functions, LANA extensively exploits SUMOylation machinery. Indeed, LANA contains a SIM to allow interaction with SUMO2 modified proteins such as KAP1 which is in turn recruited to specific chromatin sites to silence viral gene expression (Cai et al., 2013). Therefore, the LANA SIM motif plays a fundamental role in KSHV latency, even if data point to a direct binding of LANA with KAP1 independently of SIM (Sun et al., 2014). Furthermore, LANA is SUMOylated itself and its expression levels in KSHV infected SLK cells are regulated by a finely tuned deSUMOylation activity mediated by SENP6. Chromatin immunoprecipitation sequencing experiments identified that LANA binds SENP6 promoter, with subsequent repression of SENP6 expression. Given that SENP6 protease removes SUMO moieties from LANA to decrease its expression and to promote viral gene expression, these results suggest that LANA inhibits SENP6 to regulate its own SUMOylation and expression levels in infected cells, and to maintain KHSV latency (Lin et al., 2017). KHSV encodes two additional transcription factors, K-bZIP (KSHV basic leucine-zipper) and K-Rta that are crucially regulated by SUMO. K-bZIP is an early lytic gene rapidly expressed after acute infection or during reactivation from latency (Lin et al., 1999), that acts as transcriptional repressor through inhibition of the viral transactivator K-Rta (Izumiya et al., 2003). Similar to LANA, also K-bZIP needs SUMOylation to increase its activity, since expression of the SUMO specific protease SENP1 attenuates transcriptional repression of K-Rta. K-bZIP could be conjugated by both SUMO1 and SUMO2/3 and requires interaction with UBC9 at viral promoters to mediate its repression activity (Izumiya

et al., 2005). Consistently, ChIP-seq studies revealed deposition of SUMO2/3 throughout KSHV genome after viral reactivation, mirrored by decreased expression of KHSV genes (Yang, W.S. et al., 2015), suggesting that SUMOylation may be involved in chromatin remodelling during viral reactivation. Notably, K-bZIP also shows SUMO E3 ligase activity with specificity towards SUMO2/3 that catalyses SUMOylation of interacting partners such as p53 and pRb (Chang et al., 2010), and deposition of SUMO2/3 in chromatin locus enriched for SUMO2/3 (Yang, W.S. et al., 2015). Indeed, experimental KHSV reactivation in infected B lymphoma cell line is complemented by a specific increase of SUMO2/3 conjugation and inactivation of promoter regions of genes involved in immune response such as IRF-1, IRF-2, and IRF-7 (Chang et al., 2013). Collectively, these results suggest that SUMOylated K-bZIP interacts with UBC9, mediates SUMO2/3 modification of viral and cellular chromatin through its E3 SUMO ligase activity and shut-off of KHSV gene expression, and dampens the host immune activation, thus contributing to hide the virus from host responses during viral reactivation. Therefore, KHSV could regulate gene expression and viral replication manipulating SUMOylation. Indeed, modulation of global SUMO conjugation quickly occurs after induction of K-bZIP and K-Rta expression in chronically EBV-infected TRE × BCBL-1 K- Rta cell lines and is accompanied by modulation of viral gene expression (Wang et al., 2017). While K-bZIP promotes accumulation of SUMO-conjugated proteins, the viral activator K-Rta decreases global SUMOylation through its SUMO-targeting E3 ubiquitin ligase (STUbL)-like activity. STUbL proteins contain SIMs to interact with SUMO to ubiquitylate their targets (Perry et al., 2008). Indeed, K-Rta contains SIM domains able to bind SUMO moiety and to catalyse attachment of ubiquitin and proteasome-dependent degradation on targeted SUMOylated proteins. Among them, K-Rta promotes degradation of viral proteins like K-bZIP, and cellular proteins such as PML in order to create a conducive environment for viral replication (Izumiya et al., 2013). Therefore, KHSV expresses two different early genes acting as SUMO E3 ligases (K-bZIP) or STUbL (K-Rta) that differently affect SUMOylation status of infected cells in diverse phases of viral infection

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cycle. These examples are explanatory of how the dynamic and reversible alteration of SUMO conjugation represents a convenient strategy that oncogenic viruses exploit to alter and adapt host environment for viral purposes. Human T-cell leukaemia virus type 1 HTLV-1 is the aetiological agent of adult T-cell leukaemia (Poiesz et al., 1980). Transforming ability of HTLV-1 mainly relies on the oncoviral protein Tax, a transcriptional activator able to initiate T-cell proliferation ad differentiation ( Jeang et al., 2004). Activation of the NF-kB pathway, a crucial step towards transformation of a T-cell in a leukaemic cell, is finely regulated by concerted SUMO/ubiquitin conjugation steps that specifically shuttle Tax between cytoplasm and nuclear bodies. In particular, SUMOylated Tax is conjugated by ubiquitin through the STUbLs-mediated activity of RNF4 (Fryrear et al., 2012) and migrates to cytoplasm to allow interaction with the regulatory subunits of IkB kinase, NEMO, and the subsequent relocation of the Nf-KB subunit RelA to the nucleus. Then, deubiquitinated Tax translocates to nuclear bodies where it is SUMOylated on the same lysine residue by the resident UBC9 and SUMO, associates with RelA and NEMO, and starts the transcription of Tax-responsive genes mediated by NF-kB (Lamsoul et al., 2005; Nasr et al., 2006; Kfoury et al., 2011) Merkel cell polyomavirus MCPyV is the most recently emerged oncovirus, since it has been detected in about 80% of Merkel cell carcinoma, a neuroendocrine disorder of the skin frequently found in immune depressed patients [recently reviewed in (DeCaprio, 2017)]. MCPyV transforming ability mainly resides on the expression of Large T antigen proteins (Large- LT, and Short- ST) (Houben et al., 2010), even if molecular details driving MCPyV-mediated cell transformation are still not fully elucidated, although a role for ST is emerging (Shuda et al., 2011). Similarly, a role for viral exploitation of the SUMO pathway during Merkel cell carcinoma has not yet been investigated, even if a recent report shows that MCPyV replication depends on PML-NBs, suggesting a possible involvement of SUMOylation in regulating MCPyV transformation (Neumann et al., 2016).

Conclusions Post-translational modification by SUMO plays central roles during oncogenic viral infections. SUMOylation is a physiological pathway regulating proteins activity, altering localization, interaction with DNA and other proteins [reviewed in Wilkinson and Henley (2010)]. At cellular levels, SUMOylation regulates processes such as cell division, DNA replication and repair, cell signalling, chromatin remodelling, apoptosis and proliferation [reviewed in Wilson (2009)]. Since the wide impact on cell physiology, alteration of SUMOylation is a convenient way that oncoviruses frequently exploit to mediate persistence in the host. Indeed, with the exception of the recently discovered MCPyV, human oncovirus extensively manipulate SUMO to modify both viral and cellular proteins. Specifically, viral oncoproteins from HCV (NS5A), KHSV (LANA), and HTLV-1 (Tax), as well as viral structural and transcription factors such as HPV E2 and L2, EBV Zta and Rta, KHSV K-bZIP and K-Rta are all modified by one or more SUMO paralogues to alter their localization, transcriptional ability, protein-protein, protein–DNA interaction, and stability. Also, proteins from oncogenic viruses could act as specific SUMO E3 ligase to catalyse the addition of SUMO moiety to cellular target (K-bIZP) or as STUbL to clear SUMOylated proteins through the ubiquitin-mediated proteasomal degradation (K-Rta). Notably, these two apparently contrasting activities are mediated by the same oncovirus (KHSV) in different steps of viral infections, suggesting the importance of SUMOylation to quickly and completely revert cellular activities for virus purposes. Furthermore, several oncoviruses proteins, such as HBx, E6, E7, and LMP1, mediate SUMOylation of specific transcription factors to trigger or dampen expression of genes that finally promote virus infectivity and oncogenesis. Similarly, viral proteins and miRNAs could also globally affect SUMOylation in infected cells, both increasing (E6/E7, LMP1, SM, miRBHRF1–1) or decreasing (BRLF1) SUMOylation of specific SUMO paralogues. Strikingly, the ability of altering global SUMOylation could also be contingent on the manipulation of the sole SUMO E2-conjugating enzyme, UBC9, whose alteration could completely revert host functions to advantage virus endurance, as exemplified by HPV E6. Collectively, findings summarized here clearly

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suggest that SUMOylation is a pivotal pathway during infection and transformation triggered by oncoviruses. Therefore, strategies aimed at interfering with viral manipulation of SUMO components could be beneficial in the attempt to reduce cancer burden arising from viral infection. Acknowledgements Work in the S.C. laboratory related to the topics discussed in this review is supported by Associazione Italiana per la Ricerca sul Cancro (AIRC). D.M. is a recipient of the Fondazione Umberto Veronesi (FUV) fellowship. References

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Progress in the Discovery of Small Molecule Modulators of DeSUMOylation Shiyao Chen, Duoling Dong, Weixiang Xin and Huchen Zhou*

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School of Pharmacy, Shanghai Jiao Tong University, Shanghai, China. *Correspondence: [email protected] https://doi.org/10.21775/9781912530120.04

Abstract SUMOylation and DeSUMOylation are reversible protein post-translational modification (PTM) processes involving small ubiquitin-like modifier (SUMO) proteins. These processes have indispensable roles in various cellular processes, such as subcellular localization, gene transcription, and DNA replication and repair. Over the past decade, increasing attention has been given to SUMOrelated pathways as potential therapeutic targets. The Sentrin/SUMO-specific protease (SENP), which is responsible for deSUMOylation, has been proposed as a potential therapeutic target in the treatment of cancers and cardiac disorders. Unfortunately, no SENP inhibitor has yet reached clinical trials. In this review, we focus on advances in the development of SENP inhibitors in the past decade. Introduction Post-translational modification (PTM) of proteins is a crucial process for the regulation of biological growth and the stress response, and operates via extremely sophisticated mechanisms. There are at least 20 types of PTM in eukaryotes, such as ubiquitination, phosphorylation, methylation, glycosylation, and acetylation. Among them, a reversible modification process involving small ubiquitin-like modifier (SUMO) proteins, which is thus termed SUMOylation, has an indispensable role in various cellular processes, such as modulation of protein stability, subcellular localization,

protein–protein interactions, gene transcription, genome integrity, and DNA replication and repair (Wilkinson and Henley, 2010; Vierstra, 2012; Bailey et al., 2016). In 1995, Meluh and Koshland (1995) identified Smt3 in Saccharomyces cerevisiae, which is the earliest report within this filed. Two years later, based on the sequence similarity between ubiquitin and a new 11.5-kDa protein, ubiquitin/SMT3, the name SUMO was formally proposed for the first time (Mahajan et al., 1997). Although SUMO modification is closely related to the progression of various diseases, such as cancers and cardiac disorders, it has aroused increasing attention as a potential therapeutic target in recent years, especially concerning the Sentrin/ SUMO-specific protease (SENP), which is the key regulator of deSUMOylation. Unfortunately, no SENP inhibitor has yet reached clinical trials. In this review, we focus on advances in the development of SENP inhibitors within the past decade. The opportunities and challenges are discussed. SUMO modification and its cellular functions SUMO modification cycle SUMO is a class of small proteins that are highly conserved during evolution. Distinct from yeast and invertebrates, which have only a single SUMOencoding gene, there are at least four SUMO isoforms in vertebrate genomes, SUMO1–4. SUMO2 and SUMO3 are commonly referred to as

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SUMO2/3 because of their high sequence similarity (97%), whereas SUMO1 is less closely related to SUMO2/3 (almost 50%), requiring different activating and conjugating enzymes (Kerscher et al., 2006). Both SUMO1 and SUMO2/3 show certain preference for their substrates. For instance, ran GTPase activating protein 1 (RanGAP1) is a typical substrate of SUMO1, while Topoisomerase II is predominantly modified by SUMO2/3. Some other proteins, such as promyelocytic leukaemia (PML) seem to be insensitive to SUMO isoforms. Besides, their ability to form SUMO chains is also diverse. However, whether SUMO4 is capable of protein processing or conjugation remains unclear (Guo et al., 2004). In eukaryotic cells, all SUMOs are translated as immature precursors that must be transformed to the mature state before SUMOylation, which is initiated by SENPs through their SUMO peptidase activity (Park et al., 2011). Under the action of certain SENPs, about 10 amino acids at the C-terminus of SUMO precursors are removed and thus the crucial diglycine (GlyGly) binding site motif in SUMOs is exposed for the later conjugation ( Johnson, 2004) (Fig. 4.1). Similar to ubiquitination, SUMO modification of substrate proteins also requires a series of enzymatic reactions and results in the formation of an isopeptide bond between the SUMO C-terminal carboxyl group and the ε-amino group of a lysine

residue in the substrate. The first step of SUMO modification is catalysed by SUMO activating enzyme E1. In human cells, SUMO E1 is a heterodimer composed of two subunits, SUMO1 activating enzyme subunit 1 (SAE)1 and SAE2. The former can be further decomposed into SAE1a and SAE1b ( Johnson, 2004). During this step, an ATP molecule is hydrolysed for energy supplementation and an E1~SUMO high-energy thioester bond is formed between the glycine carboxyl group (SUMO C-terminal) and the sulfhydryl group (SAE2 cysteine) (Park et al., 2011). Subsequently, the activated SUMO is transferred to a cysteine residue in the SUMO conjugating enzyme E2 to form a new thioester bond E2~SUMO. It is worth noting that until now, only one SUMO E2, named ubiquitin conjugating enzyme E2 I (UBC9), has been identified, which is in sharp contrast with the tens of E2 enzymes that act in unique combinations. Although UBC9 can directly react with a certain portion of the substrates and transfer SUMO to the lysine residue in the target protein, thereby completing the SUMOylation process in absence of SUMO E3 ligase (Park et al., 2011), it has been proven that E3 ligase can enhance SUMO conjugation in two ways (Miura and Hasegawa, 2010). On the one hand, via interaction with the substrate, SUMO E3 ligase recruits the E2~SUMO thioester and the substrate into a complex, which narrows

Figure 4.1 The crystal structure of SUMOs and their evolutionary relationships. (A) SUMO1 (PDB: 1A5R). (B) SUMO2 (PDB: 2N1W) (C) SUMO3 (PDB: 1U4A). The flexible N-terminal extension is coloured grey. (D) Evolutionary relationship of SUMO family members.

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their distance and contributes to the specificity of transfer. On the other hand, the catalytic activity of E2 is up-regulated in the presence of E3 ligase, which contributes to the efficiency of SUMOylation. There are four E3 ligases in yeast, namely Siz1, Siz2, Mms21, and Zip3; whereas mammals possess ten, including three protein inhibitor of activated STAT (PIAS) family members, Methyl methanesulfonate sensitivity gene 21 (MMS21), RAN binding protein 2 (RanBP2), Pc2 (also known as Keratin 17) and OP1 binding arginine/serine rich protein (TOPORS). As mentioned above, the SUMO modification is a reversible process. To complete the SUMO modification cycle, deconjugation of SUMO from SUMOylated protein substrates is also indispensable and is catalysed by the SENP family. Apart from its function as a maturation enzyme, SENP is also capable of the cleaving the isopeptide bond formed between the C terminus of SUMO and the ε-amino group of the lysine residue in the target, thereby promoting the release of SUMO (Mukhopadhyay and Dasso, 2007). The members of SENPs were first discovered in Saccharomyces cerevisiae and named as Ulp1 and Ulp2, while the human SENP family was characterized later, including SENP1, SENP2, SENP3, SENP5, SENP6, and SENP7 (Li and Hochstrasser, 1999, 2000; Hickey et al., 2012). Interestingly, although the processing and deconjugation of SUMO is achieved by the same group of proteases, some SENPs act only as deSUMO enzymes and do not participate in

Figure 4.2  The SUMOylation and deSUMOylation cycle.

SUMO maturation (Saracco et al., 2007; Castro et al., 2016) (Fig. 4.2). SUMO interaction domains The recognition and conjugation of most SUMOylated substrates by SUMO E2 is achieved via a short consensus modification motif ψKx(D/E) characterized in multiple target proteins, in which ψ represents a large hydrophobic residue, whereas x can be any amino acid (Rodriguez et al., 2001). Acting directly on UBC9, these residues have a crucial impact on the stability of the interaction between SUMO E2 and the target. In the crystal structure of a SUMO consensus motifs-containing protein–UBC9 complex, the modification motifs adopt an extended conformation, which limits the acceptor lysine in the UBC9 hydrophobic groove. Electrostatic interactions as well as hydrogen bonding between UBC9 and the amino acids adjacent to the acceptor lysine also contribute to its recognition (Lin et al., 2002). In addition to the four-amino acidlength canonical consensus motifs, several variants have been identified, such as motifs with additional elements nearby. For instance, phosphorylation of certain sites, termed as phosphorylation-dependent SUMO motifs (PDSMs), promotes up-regulation of SUMOylation both in vitro and in vivo, and was first discovered in heat shock proteins. There are also negatively charged amino acid dependent SUMO motifs (NDSMs), which possess negatively charged residues. Reports of novel SUMO consensus motifs, such as inverted consensus motifs and

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motifs combined with an N-terminal hydrophobic cluster with distinct polarity, emphasizes the importance of detailed research into the diversity of SUMO conjugation sites (Hietakangas et al., 2003; Mohideen et al., 2009; Matic et al., 2010). Furthermore, such consensus motifs do not strictly follow the sequence order or geometrical requirements, for example, the lysine in the α-helix of ubiquitin conjugating enzyme E2 K (E2–25K) (Pichler et al., 2005; Knipscheer et al., 2008). Another SUMO-binding domain that has been studied in depth is the SUMO-interacting motif (SIM), which mediates non-covalent interactions between SUMO and proteins containing SIMs (Merrill et al., 2010). It is typically composed of a hydrophobic core with four consensus hydrophobic residues, V/I-x-V/I-V/I or V/I-V/I-x-V/I/L, where x can be any amino acid, flanked by acidic residues providing the necessary polarity (Merrill et al., 2010). When complexed with SUMO, SIMs adopt either a parallel or antiparallel β-conformation to the SUMO β-sheet, exposing the hydrophobic side chains to the hydrophobic pocket on the SUMO surface. Molecular dynamic simulations demonstrated higher stability of the complex with the antiparallel orientation together with better tolerance to sequence changes, possibly because of the establishment of more backbone-mediated interactions (Conti et al., 2014; Jardin et al., 2015). In addition, a subclass of SIMs possessing serine residues as phosphorylation sites, adjacent to the hydrophobic core, has been characterized in PML, exosome component 9 (EXO9), and PIAS proteins (Stehmeier and Muller, 2009). Phosphorylated by casein kinase 2 (CK2), these phospho-SIMs enhance their non-covalent binding to SUMO through electrostatic interactions between the negatively charged phosphorylated serine and positively charged lysine on the SUMO surface, which is also presumed to further affect the specificity of different SUMO isoforms. SIMs are found in many proteins, including SUMO substrates and binding partners, SUMOtargeted ubiquitin ligases (like Slx8-rING finger protein (rfp) in S. pombe and Slx5-Slx8 in S. cerevisiae), as well as all known SUMO E3s, serving as a crucial regulator in various cellular processes. Although found in ubiquitin-specific protease 25 (USP25), a substrate for SUMOylation, in which the SIMs were observed to contribute to

a modification preference for SUMO2/3, their structural determinants for such specificity remain to be verified and the measured affinity between SIMs and different SUMO isoforms did not appear to be significantly different (Meulmeester et al., 2008; Sekiyama et al., 2008; Kung et al., 2014). Human ZNF451, a SUMO E3 ligase, contains two SIMs separated by a Pro-Leu-Arg-Pro sequence in its catalytic region, which provides support for the effect of SIMs on E3 ligase activity (Cappadocia et al., 2015). The N-terminal SIM of ZNF451 maintains the donor SUMO in a closed conformation, whereas its C-terminal SIM combines with a second SUMO on the reverse side of UBC9, which ensures direct contact between certain residues and UBC9. ZNF451 itself is also a target of SUMO modification. SUMOylation occurs close to the catalytic module, which causes an increase in its activity as a SUMO E3 ligase (Hendriks and Vertegaal, 2016). Furthermore, it was proposed recently that the SUMO modification targets entire groups of interacting proteins rather than individual proteins, thus the presence of SIMs in the substrates has a key role in protein-group SUMOylation via multiple SUMO–SIM interactions ( Jentsch and Psakhye, 2013). Other types of SUMO interaction domains have also been reported, for instance the ZZ zinc finger domain that interacts with SUMO in a zinc-dependent manner and the zinc-independent MYM-type zinc finger domain (Danielsen et al., 2012; Guzzo et al., 2014). However, the latter seems to bind to the same site in SUMO as SIMs, such that the destruction of the SUMO–SIM interaction simultaneously affects the stability of the SUMO–MYM interaction (Guzzo et al., 2014). Cellular roles of SUMOylation To date, numerous studies on the essential role of SUMOylation in normal cell homeostasis have been carried out, the majority of which were based on the regulation of transcription. Early studies demonstrated that SUMOylation is closely related to transcriptional repression (Gill, 2005). One of the hypotheses attributes this to the recruitment of transcriptional corepressors, such as the histone deacetylase 1 and 2 (HDAC)1/2 complex (Yang and Sharrocks, 2004). SUMOylation might also be involved in the regulation of factors related to RNA polymerase Pol II, an emerging

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finding proposed more recently (Neyret-Kahn et al., 2013; Niskanen et al., 2015). As reported by Yu et al. (2018), SUMO and MYC have opposite effects on global transcription by controlling the level of SUMO modification of cyclin dependent kinase 9 (CDK9), which is the catalytic subunit of positive transcription elongation factor b (P-TEFb) complex (Yu et al., 2018). SUMOylation of CDK9 interrupts its binding to the regulatory subunit Cyclin T1, thereby causing a pause in the formation of the active P–TEFb complex, which ultimately blocks global gene expression. By contrast MYC antagonizes SUMOylation processing in combination with PIAS in a CDK9-competitive way, serving as a broad-spectrum promoter for cellular transcription. SUMOylation is also implicated in the modulation of cellular stress responses, like the endoplasmic reticulum (ER) stress response, viral infections, nutrient response, and especially, the DNA damage response (Enserink, 2015). Ubiquitin–SUMO crosstalk occurs extensively in signalling responses to double-strand breaks (DSBs). One example is ring finger protein 4 (RNF4), a SUMO targeting ubiquitin ligase (STUbLs) in mammalian cells. Containing tandem SIMs in its N-terminus, RNF4 is capable of recognizing poly-SUMOylated proteins and promoting K48-linked ubiquitination (Galanty et al., 2012). In addition, RNF4 also catalyses ubiquitin conjugating enzyme E2 N (UBC13)dependent K63-linked polyubiquitination (Yin et al., 2012). Phenotypically, cells lacking RNF4 show defective RAD51 recombinase (RAD51) loading, leading to blockade of chromosome homologous recombination, for which inefficient exchange of replication protein A (RPA) with RAD51 is caused by the decrease of SUMO-modified RPA turnover from single-stranded DNA (ssDNA) may be an acceptable explanation (Galanty et al., 2012; Yin et al., 2012). DeSUMOylation and SENPs As stated above, the members of the SENP family have a dual effect as maturation enzymes in precursor processing and SUMO deconjugases. In the body, SUMOylation and deSUMOylation are require for a dynamic equilibrium relationship, while the balance between SUMO and deSUMO modification of proteins in diverse cellular

compartments is mainly attributed to SENPs. Based on their sequence homology, substrate specificity, and subcellular localization, the six SENP isoforms in mammals, SENP1, SENP2, SENP3, SENP5, SENP6 and SENP7, can be classified into three sub-families: SENP1 and SENP2, SENP3 and SENP5, and SENP6 and SENP7 (Gong and Yeh, 2006). In terms of their evolutionary relationship, the first two families can also be classified as the Ulp1 branch, while SENP6 and SENP7 belong to the Ulp2 branch (Mukhopadhyay and Dasso, 2007). Structural characteristics Compared with the variable N-terminus, which contributes to the differences in spatial distribution and substrate specificity among distinct SENP isoforms, the C-terminal regions among all six SENPs seem to be more conserved, containing a cysteine protease catalytic domain that is approximately 250 amino acids in length (Hickey et al., 2012). Currently, only the crystal structure of the catalytic domain of SENP1 (2IYC, 2IY1, 2IYD, 2IY0) coupled with SENP2 (1TH0, 1TGZ, 2IO0, 2IO1, 2IO2, 2IO3), and SENP7 (3EAY) has been reported, either in apo form or in complex. Composed of the typical catalytic triad (Cys-HisAsp) (Cys603, His533 and Asp550 for SENP1; Cys-548, His-478, and Asp-495 for SENP2; His794, Cys-926 and Asp-873 for SENP7), which is analogous to other cysteine proteases, the crystal structures of these three SENPs are quite similar (Kumar and Zhang, 2015). The structures highlight that this catalytic domain is indispensable for the hydrolytic activity in precursor processing and deSUMOylation, such that the replacement of the active-site cysteine residue with serine in human SENP1 damaged to its function (Xu et al., 2006). Meanwhile, a narrow tunnel lined by Trp residues in SENP1 is critical for the positioning of the di-Gly motif, while the closing it contributes to the orientation of the sessile bond, thus forming an unstable kink in the linkage to the SUMO substrate proteins, which is believed to promote cleavage (Shen et al., 2006). Among the catalytic triads, cysteine, as the main catalytically active site, is the most commonly used target for the development of SENP inhibitors (Fig. 4.3). In particular, the structure of the catalytic domain of SENP7, consisting of amino acid

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Figure 4.3  Structures of SENPs. (A) Details of the SENP1 catalytic triad. (B) Details of the SENP1 catalytic triad and the Trp tunnel in the SENP1–SUMO1 complex. SENP1 is shown in blue and SUMO1 is shown in in purple (PDB: 2IY1).

residues 662–984, reveals its relationship between the SENP/Ulp protease family and other Cys-48 cysteine proteases. Consistent with its substrate specificity, SENP7 has a unique catalytic structure that is apparently different from SENP1 and SENP2, including the absence of the N-terminal α-helix, the insertion of four conserved loops, and the extension of several secondary structure elements (Lima and Reverter, 2008). Loop1 is highly conserved in SENP6 and SENP7, suggesting its potential contribution to catalytic activity, which was subsequently proved (Lima and Reverter, 2008). Compared with wild-type SENP7, mutants lacking loop1 show an apparent defect in enzyme activity of SENP7 for both precursor processing and SUMO-deconjugation. Besides, Alegre and Reverter (2011) identified that the position of Asp71, coupled with Asn68 of SUMO2, is close to loop1 of SENP7 in the crystal structure of their complex, indicating possible polar interactions. Further studies showed that these two key amino acids are directly responsible for the preference of SENP7 for SUMO2/3 through interaction with loop1. Substrate specificity SENP1 and SENP2 SENP1 and SENP2 show broad specificity for SUMO1/2/3. SENP2 has a similar activity to SENP1 when overexpressed; however, it prefers SUMO2 over SUMO1 for deconjugation and has a relatively poor effect on SUMO3. The mechanisms

of action of these two isoforms are distinct, for example, although both SENP1 and SENP2 can regulate c-Jun-dependent transcription, SENP1 works by deSUMOylation of p300 while SENP2 targets PML (Best et al., 2002; Cheng et al., 2005). Emerging research suggests that SENP1 regulates the stability of hypoxia inducing factor 1 (HIF1), while SENP2 does not, clearly indicating that both of them have their own specific substrates (Yeh, 2009). SENP3 and SENP5 SENP3 and SENP5 share high sequence identity and the same localization; it is reasonable to deduce that they may have similar substrate selectivity (Gong and Yeh, 2006). Compared with SUMO1, SENP3 and SENP5 show a prominent preference for SUMO2/3. The stability of SENP3 is regulated by CHIP, which is the carboxyl-terminus of heat shock protein family A (HSP70) member 8 (HSC70)-interacting protein through the heat shock protein 90 (HSP90)-independent ubiquitinproteasome pathway (Yan et al., 2010). However, in response to mild oxidative stress, SENP3 undergoes thiol modification, by which HSP90 is recruited, and subsequently its degradation is repressed. In liver cancer, SENP3 accumulates and accelerates disease progression by responding to the abnormal redox background (Yan et al., 2010). SENP5 is crucial in cell mitosis and/or cytokinesis, and the absence of SENP5 causes proliferation inhibition and abnormal nuclear morphology (Di Bacco et al., 2006).

Small Molecule Modulators of DeSUMOylation |  57

SENP6 and SENP7 Among the six human SENP isoforms, the catalytic domains of SENP6 and SENP7 are the most diverse, especially the insertion of loop1, which mediates the specific interaction with SUMO2/3conjugated substrates. As a consequence, SENP6 and SENP7 preferentially act on SUMO2/3, and moreover, are more effective in cleaving diSUMO2/3 or poly-SUMO2/3 chains attached to lysine residues (Alegre and Reverter, 2011; Lima and Reverter, 2008). SENPs fail to achieve the proteolytic processing of SUMO4 precursor molecule in vivo, which is indispensable for its maturation; therefore, the posttranslational modification of substrate proteins by SUMO4 has not yet been observed (Owerbach et al., 2005). In precursor SUMO4, Pro-90 replaces Gln in SUMO1–3, which causes a conformational restriction that might keep the peptide bond to be cleaved distal from the catalytic site of SENP and thus disrupt the maturation process (Békés et al., 2011). A P50Q single amino acid mutant of precursor SUMO4 made it amenable to SENP2 cleavage, as did another mutant, G63D (Liu et al., 2014). Besides, although all six SENP isoforms possess SUMO deconjugation/isopeptidase activity, only SENP1, SENP2, and SENP5 can carry out SUMO maturation. Cellular localization Different SUMO isopeptidases have characteristic subcellular distributions, which is closely related to the varied lengths and specificities of the N-terminal domains, which seems to contribute to substrate specificity. SENP1 consists of 644 amino acids, with a nuclear localization signal (NLS) and nuclear export-signal (NES) at its N-terminus and C-terminus, respectively (Bailey and O’Hare, 2004). The structure of SENP2 is quite similar. Interacting with components of the nuclear pore complex, SENP1 and SENP2, coupled with Ulp enzyme, which was identified in yeast, gather on the nuclear envelope and accumulate in distinct subnuclear structures (Goeres et al., 2011). Although they are excluded from the nucleolus, substantial amounts of SENP1 and SENP2 are observed in nuclear foci that partially overlap with PML bodies. During mitosis, SENP1 and SENP2 are redistributed from the nuclear envelope to the kinetochore (CubeñasPotts et al., 2013). Notably, measuring a series of its

mutants with interspecies heterokaryons indicated that SENP2 shuttles between the nucleus and the nucleoplasm, which can be inhibited by mutation of its NES or treatment with leptomycin B (LMB), in spite of its predominantly nuclear localization (Itahana et al., 2006). In addition, diverse splice variants of SENP2 show specific subcellular localizations (Hickey et al., 2012). Both SENP3 and SENP5 are compartmentalized in the nucleolus, the function of which is to act on proteins involved in the early stage of ribosome maturation (Haindl et al., 2008; Yun et al., 2008). SENP3, also known as SMT3IP1 or SMTB1 in mice, comprises 574 and 568 amino acids, respectively. SENP3 contains unique sequences in its N-terminus, including glutamate clusters (residues 74–86) and two regions rich in arginine (residues 119–122, 147 and 153), which may account for its nucleolar localization (Nishida et al., 2000). SENP5 comprises 755 amino acids with an extended sequence at its N-terminus, a truncated variant of which co-localizes with the PML (Di Bacco et al., 2006). In addition, subfractions of SENP3 and SENP5 are also found in the nucleoplasm and cytoplasm. In particular, SENP5 translocates to the surface of mitochondria before the rupture of the nuclear envelope during G2/M stage of the cell cycle (Zunino et al., 2009). By contrast, SENP6 and SENP7 are generally concentrated in the nucleoplasm (Table 4.1). Cellular pathways controlled by SENPs Cell cycle Considering the spatiality and temporality of SUMOylation in mitosis, it is easy to associate SENPs with cell cycle progression. In budding yeast, Ulp1, which acts on Smt3 and SUMO1-conjugated proteins, exhibits an essential role in the transition from G2 to M phase (Li and Hochstrasser, 1999). Knockdown of SENP1 causes the failure of sister chromatid separation and arrests progression at M phase; however, overexpression of SENP2 also decreases global SUMOylation, which leads to prometaphase arrest because of defects in targeting the microtubule motor protein centromere protein E (CENP-E) to kinetochores (Zhang et al., 2008; Cubeñas-Potts et al., 2013). Moreover, Mukhopadhyay and Dasso (2010) identified SENP6 as a key

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Table 4.1  Properties of SENP isoforms SENP isoform

Substrate preference

Cellular localization

Function

SENP1

SUMO1/2/3

Nuclear pore and nuclear foci

Deconjugation/isopeptidase; precursor processing

SENP2

SUMO2/3 > SUMO1

Nuclear pore and nuclear foci; cytoplasm

Deconjugation/isopeptidase; precursor processing

SENP3

SUMO2/3

Nucleolus

Deconjugation/isopeptidase

SENP5

SUMO2/3

Nucleolus

Deconjugation/isopeptidase; precursor processing

SENP6

SUMO2/3

Nucleoplasm

Deconjugation/isopeptidase; chain-editing

SENP7

SUMO2/3

Nucleoplasm

Deconjugation/isopeptidase; chain-editing

regulator of inner kinetochore assembly. Deletion of SENP6 directly led to the mis-localization of inner kinetochore proteins (IKPs) in Hela cells, which caused chromosome misalignment with missegregation, and subsequently delayed cell cycle progression. By antagonizing the STUbL pathway, SENP6 functions as a protector of IKP, keeping it away from S phase degradation (Mukhopadhyay and Dasso, 2010). The mitotic substrate specificity of SENPs remains to be determined. Transcription Among the known substrate proteins of SUMOylation, nuclear proteins occupy a considerable proportion, which participate in the transcriptional regulation of genes and chromatin dynamics. In most cases, the conjugation of core histones is associated with transcriptional silencing, and SUMOylated transcription factors or transcriptional co-regulators are thought to induce a decrease or even inhibition of gene activation (Wotton et al., 2017). Correspondingly, deSUMOylation, mediated by SENPs, facilitates transcription (Niskanen and Palvimo, 2017; Wotton et al., 2017). For instance, SENP3 affects the assembly of the MLL1/2 (also known as lysine methyltransferase 2A) histone methyltransferase complex on distinct homeobox (HOX) genes, including the osteogenic master regulator distalless homeobox 3 (DLX3) (Nayak et al., 2017). Via the deSUMOylation of RB Binding Protein 5 (RbBP5), SENP3 activates the recruitment of Ash2 (absent, small, or homeotic)-like (Ash2L) and menin subunits to DLX3 by complexing with MLL1/2, which is a prerequisite for promoting transcriptional activation of the HOX genes (Nayak et al., 2017).

However, the deSUMOylation of the complex of transducin β-like protein 1 (TBL1) and TBL1related 1 (TBLR1) by SENP1 decreases complex formation and subsequently inhibits β-cateninmediated transcription, serving as a suppressor of the Wnt signalling pathway (Choi et al., 2011). Macromolecular assembly The biogenesis of pre-ribosomal particles in eukaryotic cells is controlled by Ulp/SENPs, which was first found in S. cerevisiae via mutations in UBC9, Ulp1, and Smt3, where the export of the pre-60S ribosomal subunit was defective (Panse et al., 2006). Formed in the nucleolus, pre-60S and pre-40S ribosomes need to undergo a series of sophisticated modifications to transform them into a mature state. Subsequently, they are transported to the cytosol, in which SUMOylation plays a major role. The late steps of nucleolar maturation of pre-60S particles involves the formation of a complex comprising proline, glutamate and leucine rich protein 1 (PELP1), testis expressed 10 (TEX10), and WD Repeat Domain 18 (WDR18), the SUMOylation of which is carried out in a SENP3-dependent way (Finkbeiner et al., 2011). PELP1 shows a prominent sensitivity to SENP3, while its SUMO conjugation enhances the recruitments Midasin AAA ATPase 1 (MDN1) to pre-60S particles, functioning as a key step in pre-60S remodelling (Raman et al., 2016). Another instructive example is that of the PML protein, a scaffold protein of PML nuclear bodies (Uversky, 2017). Being the focal point of SUMO conjugation and deconjugation, the modulation of PML nuclear bodies involves multiple SUMO–SIM interactions (Raman et al., 2013). SUMOylated PML proteins can be self-assembled via their own SIMs or recruited with other SIM-containing

Small Molecule Modulators of DeSUMOylation |  59

proteins. Therefore, the degree of SUMOylation greatly affects the number and composition of nuclear bodies. The key role played by SUMO in promoting nuclear bodies assembly by providing multivalent interactions was highlighted recently, suggesting the possible effect on the dynamics of nuclear bodies of SENPs (Banani et al., 2016). The poly-SUMOylated PML protein itself is a substrate for SENP6; therefore, down-regulation of SENP6 expression directly induces the formation of the SUMO chain on PML, causing an increase in both the number and size of PML nuclear bodies (Hattersley et al., 2011). DNA repair The importance of SUMO-dependent recruitment to the sites of DNA damage sites in the doublestrand break (DSB) response is evident from the appearance of the 70-kDa subunit of the replication protein A complex (RPA70) being controlled by SENP6 (Dou et al., 2010). RPA plays a key role in DNA replication, as well in damage responses. Associated with RPA70 during replication, SENP6 limits the SUMOylation of RPA70 to a lower level in S phase. Double-stranded DNA damage induces the expression of the replication stressinduced factor Camptothecin (CPT), at which point SENP6 is dissociated from RPA70, thereby relieving the restriction of RPA70 SUMOylation, which involves SUMO2/3 (Dou et al., 2010). SUMOylated RPA70 recruits RAD51 to the DNA damage foci and subsequently initiates DNA repair through homologous recombination. In another example, SENP7 acts on the chromatin repressive KRAB-associated protein 1 (KAP1), facilitating the removal of its coupling to SUMO2/3 (Garvin et al., 2013). The deSUMOylation of KAP1 contributes to chromatin relaxation through interactions between chromatin remodeller CHD3 and chromatin, which establishes the permissive chromatin environment required for DNA repair. Mitochondrial dynamics Previously, many proteins involved in the control of mitochondrial dynamics in mammalian cells were identified, including dynamin-related protein 1 (DRP1), a mitochondrial fission GTPase that is a substrate of SUMO1. The overexpression of SUMO1 protects DRP1 from degradation and subsequently leads to increased mitochondrial

fragmentation (Harder et al., 2004). However, SENP5 can reverse this SUMO1-induced fragmentation, while silencing of its expression altered mitochondrial morphology and inhibited mitochondrial fusion (Zunino et al., 2007). Moreover, the translocation of SENP5 at G2/M also has a crucial role in the regulation of DRP1–dependent fusion during mitosis (Gong and Yeh, 2006). DeSUMOylation in diseases As one of the most dominant post-translational modification, the substrates of SUMOylation are involved in almost all pathological processes. Thus, abnormal SUMOylation, especially the alteration of SENPs expression under diseased states, may be closely related to the development of various diseases, such as cancers and cardiac disorders. For example, SENP2, which is one of the direct targets of the transcription factor NF-κB, accelerates the pathogenesis of tumours via inflammatory signalling (Huang et al., 2003; Lee et al., 2011). SENP3 coupled with SENP5 are notably overexpressed in oral squamous cell carcinomas, osteosarcoma, and hepatocellular carcinoma (Ding et al., 2008; Sun et al., 2013; Wang and Zhang, 2014; Jin et al., 2016). In addition, up-regulated SENP3/SMT3IP1 promotes epithelial ovarian cancer progression; thus, SENP3/SMT3IP1 up-regulation could be regarded as a novel biomarker for prognosis (Cheng et al., 2017). Moreover, the statistical relation between the expression level of SENP5 and prognosis in patients with breast cancer has been demonstrated (Cashman et al., 2014). In addition, recent studies have also associated SUMOylation with the development, metabolism, and pathology of the heart. Numerous key proteins in cardiac development have been shown to undergo SUMOylation, including myocardin, GATA-binding protein (GATA)-4, Nk2 homeobox 5 (Nkx2.5), myocyte enhancer factor-2 (MEF2), and T-box transcription factors-2 and -5 (TBX2/ TBX5) (Wang and Schwartz, 2010; Beketaev et al., 2014). Meanwhile, SUMO elements are indispensable to the entire cardiac physiology. For instance, the absence of SENP2 resulted in cardiac hypoplasia in mice, whereas its overexpression was associated with cardiac dysfunction, such as congenital heart defects, cardiomyopathy, and hypertrophy (Kang et al., 2010; Kim et al., 2012).

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Advances in the development of SENP inhibitors Small molecule inhibitors To date, relatively few inhibitors of SENPs have been reported, and they are mainly concentrated in inhibiting SENP1 and SENP2. Several methods have been used to identify SENP inhibitors. Scientists used the feature that the cysteine on SENP can react with electrophiles (Hemelaar et al., 2004), and developed covalent inhibitors for this series of protein. Some researchers simulated the structure of SUMO (Albrow et al., 2011; Ponder et al., 2011), while other simulated the combination between SUMO and SENP, and utilized computer-aided drug design to develop relevant inhibitors (Qiao et al., 2011). Moreover, with the advances in computer technology, more and more research groups began to use in silico techniques to find compounds with high activity and selectivity from large libraries of compounds (Shen et al., 2006; Chen et al., 2012; Madu et al., 2013; Kumar et al., 2014; Wen et al., 2014). Non-covalent inhibitors of SENP have also been found using this method (Chen et al., 2012). In 2011, Bogyo’s (Albrow et al., 2011; Ponder et al., 2011) and Zhou’s (Qiao et al., 2011) laboratories first identified small molecule inhibitors of SENPs. Bogyo’s group developed a series of compounds that simulated the structure of peptides, while Zhou’s group developed the first series of non-peptide inhibitors of SENPs. During functional studies of Plasmodium falciparum SENPs, Ponder et al. (2011) screened 508 irreversible cysteine protease inhibitors, and identified a PfSENP1 inhibitor 1. PfSENP1 inhibitor 1 displayed an IC50 of 17.9 μM for PfSENP1, but the values were only 9.0 μM and 4.7 μM for human SENP1 and SENP2, respectively. To improve the stability of the compound, as well as simplify its synthesis, the aspartic acid side chain of the original compound was removed to form compound 2. Compound 2 showed increased inhibitory efficiency. For PfSENP1, the IC50 value was 16.2 μM, while for human SENP1 and SENP2, the value values were 7.1 μM and 3.7 μM, respectively. AS part of Bogyo’s group, Albrow et al. (2011) designed a series compounds with acyloxymethyl ketone (AOMK), which were based on the structure of compound 2 and SUMO. Most of the compounds showed inhibitory activities on human

SENP1 and SENP2, among which, compound 3, with the QTGG specificity sequence, showed the best inhibition and IC50 values of 3.6 and 0.25 μM, for human SENP1 and SENP2, respectively. However, compound 4, which contains the sequence of ubiquitin, showed inhibitory activities on human SENP6 (IC50 = 4.2 μM) and SENP7 (IC50 = 4.3 μM) (Fig. 4.4). However, considering that the compounds with peptidyl moieties may perform poorly in pharmacokinetics, Qiao et al. (2011) developed a series of SENP1 inhibitors based on a benzodiazepine scaffold, which were the first designed and synthesized non-peptide inhibitors. According to the crystal structure of human SENP1 complexed with unprocessed SUMO1 (PDB: 2IY1) (Shen et al., 2006), they found that the core structure of benzodiazepine docked into the catalytic pocket and could simulated the natural combination, via its formyl group forming a covalent bond with Cys-603. The two most potent compounds 5 and 6 displayed IC50 values of 19.5 μM and 9.2 μM, respectively, for SENP1, and also showed inhibitory activity against prostate cancer cells in vitro, with IC50 of 13.0 μM and 35.7 μM, respectively. In a follow-up study, Zhao et al. (2016) found 11 series of SENP1 inhibitors with different scaffolds using virtual screening. By analysing the structures of these inhibitors and the patterns of their binding to SENP1, a series of compounds with new scaffolds was designed and synthesized from two representative compounds. Subsequently, their structure–activity relationships were identified. Among them, the most potent compound 7 displayed an IC50 of 3.5 μM for SENP1 (Fig. 4.5). Compound 8 can inhibit hypoxia inducible factor (HIF)-1α accumulation (Uno et al., 2009), as well as the growth of KEK293 cells (IC50 = 7.2 μM). However, its inhibitory mechanism has not been determined (Shimizu et al., 2010). Uno et al. (2012) used a biotin-tagged compound version of compound 8 to identify its target molecules using pull-down experiments. Fortunately, they observed an interaction between 8 and SENP1. Through structural optimization, compounds 9 and 10 were synthesized, which have more potent inhibitory activities against SENP1, with IC50 values of 39.5 μM and 29.6 μM, respectively (Fig. 4.6). In recent years, computational approaches have become important to identify small molecule

Small Molecule Modulators of DeSUMOylation |  61

11

2

33

44

Figure 4 Figure 4 Structures of compounds 1–4. Figure 4.4 

5

5

Figure 4.5  Structures of compounds 5–7.

inhibitors. Several approaches have been reported, such as virtual screening and docking, in attempts to find SENPs inhibitors. Based on the crystal structure of the SENP1– SUMO2-RanGAP1 complex reported by Hay’s group (PDB entry: 2IY0) (Shen et al., 2006), Chen et al. (2012) identified novel lead compound 11 as a SENP1 inhibitor by molecular docking of 180,000 compounds in the SPECS compound library using Glide version 4.5. According to the results of subsequent biological tests of the selected 38 compounds, compound 11 showed the best inhibitory activity, with an IC50 of 2.385 μM. A series of derivatives of 11 based on 2-(4-Chlorophenyl)-2-oxoethyl

6 6

7 7

4-benzamidobenzoate was then designed and synthesized, among which the IC50 of compound 12 reached 1.08 μM (Fig. 4.7). Madu et al. (2013) performed in silico screening of 250,000 compounds using the program GLIDE and obtained 40 candidates that exhibited inhibitory activities on SENP1, SENP2, and SENP7. According to the data from biological measurement and their structural features, a novel class of SENP inhibitors based on sulfonyl-benzene groups was proposed. The most potent compound 13 displayed IC50 values of 2.1 μM, 2.0 μM, and 2.7 μM, respectively, for SENP1, SENP2, and SENP7. Moreover, unlike the most common SENP inhibitors, which

62  | Chen et al.

8 (R1 = H, R2 = CH3); 9 (R1 = CH3, R2 = CH3); 10 (R1 = CH3, R2 = C2H5) Figure 6

8 (R1 = H, R2 = CH3); 9 (R1 = CH3, R2 = CH3); 10 (R1 = CH3, R2 = C2H5)

Figure Figure64.6  Structures of compounds 8–10.

11 Figure 7 Structures of compounds 11–12. Figure 4.7  11

12 12

Figure 7

covalently target the catalytic cysteine residue, this class of inhibitors are not covalently bound to SENPs. They have a non-competitive inhibitory mechanism and can combine with SENPs and the SENPs–substrates complex (Fig. 4.8). Kumar et al. (2014) screened out small molecules from a library of 400 million compounds using the ROCS and EON programs. The molecules were similar to the TGGK peptide at the SUMO1 C-terminus in terms of their structure and electrostatic characteristics. These compounds were docked to SENP2 catalytic pocket and then Figure 8 a quantitative biological test was performed on 13 the selected 49 compounds using oxadiazole. This Figure series 8of compounds showed inhibitory activities on both SENP1 and SENP2. Finally, the most potent compound 14 was found to show an IC50 of 3.7 μM on SENP2 and > 30 μM on SENP1, which indicated partial selectivity for SENP2 (Fig. 4.9). Wen et al. (2014) used two virtual screening programs, DOCK and AutoDock to docked ≈ 100,000 drug-like compounds, which were selected from a library comprising two million compounds. Finally, 117 compounds were selected to evaluate SENP1 FigureThe 9 most potent compound 15 displayed activity. 14 an IC50 of 1.29 μM for SENP1. It shows selectivity Figure 9

for SENP1, but weak inhibition of other cysteine proteases, like cathepsin B and D (Fig. 4.10). Biotinylated probes Before these small molecules were identified, scientists focused on covalent binding with the thiol group of cysteine in the catalytic site of SENPs. Hemelaar et al. (2004) first reported peptide SENPs inhibitors. Based on the mechanism of SUMOylation and the structural characteristics of SUMO, they used the synthesis strategy of intein 13 to link the vinylsulfone (VS) group at the end of SUMO to obtain the peptide SENP inhibitor 16. A Michael addition reaction occurred, and the inhibitor covalently bound to the catalytic cysteine residues of SENP2 and other related enzymes. To identify the key role of the cysteine residues in catalysis, pre-incubation of SENP2 with N-ethylmaleimide (NEM) was carried out, which disrupted the irreversible conjugation between VS and SENP2 (Fig. 4.11). Borodovsky et al. (2005) used a similar strategy and reported peptide compounds of the 14 ubiquitin-like proteins Nedd8, SUMO1, FAT10, Fau, and APG12 linked to a VS group. Among them, there are three C-terminal peptide chains

Small Molecule Modulators of DeSUMOylation |  63

Figure 8 4.8  Structure of compound 13. Figure Figure 8

Figure 9 Figure 9 4.9  Structure of compound 14. Figure

13 13

14 14

15 Figure 10 4.10  Structure of compound 15. Figure

of different lengths from SUMO1 (5 peptide, 9 peptide, 13 peptide) linked to the VS group that could bind to a series of proteins in the EL-4 cell lysate. Subsequent competition experiments showed that the peptide 17 was able to bind to at least one SUMO1 protease and was sufficient to establish selectivity. This study showed that only peptides of a few amino acids can specifically bind to SENPs (Fig. 4.12). 16 Figure 11

Dobrotă et al. (2012) designed and synthesized peptide 18. One terminal of the peptide contains a glycine-derived fluoromethylketone group that can covalently bind to the cysteine of SENPs. The results showed that peptide C binds to SENP1 and SENP2 and can compete for SUMO1 from the SENP1–SUMO1 complex, indicating that this compound binds to SENP1 more strongly than the SUMO1 molecule in nature (Fig. 4.13).

64  | Chen et al.

16 Figure 11

16 Figure 11 Structure of compound 16. Figure 4.11 

17 Figure 12Structure of compound 17. Figure 4.12  17 Figure 12

18 Figure 13Structure of compound 18. Figure 4.13 

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Yeh, E.T. (2009). SUMOylation and De-SUMOylation: wrestling with life’s processes. J. Biol. Chem. 284, 8223–8227. https://doi.org/10.1074/jbc.R800050200. Yin, Y., Seifert, A., Chua, J.S., Maure, J.F., Golebiowski, F., and Hay, R.T. (2012). SUMO-targeted ubiquitin E3 ligase RNF4 is required for the response of human cells to DNA damage. Genes Dev. 26, 1196–1208. https:// doi.org/10.1101/gad.189274.112. Yu, F., Shi, G., Cheng, S., Chen, J., Wu, S.Y., Wang, Z., Xia, N., Zhai, Y., Wang, Z., Peng, Y., et al. (2018). SUMO suppresses and MYC amplifies transcription globally by regulating CDK9 sumoylation. Cell Res. 28, 670–685. https://doi.org/10.1038/s41422-018-0023-9. Yun, C., Wang, Y., Mukhopadhyay, D., Backlund, P., Kolli, N., Yergey, A., Wilkinson, K.D., and Dasso, M. (2008). Nucleolar protein B23/nucleophosmin regulates the vertebrate SUMO pathway through SENP3 and SENP5 proteases. J. Cell Biol. 183, 589–595. https://doi. org/10.1083/jcb.200807185. Zhang, X.D., Goeres, J., Zhang, H., Yen, T.J., Porter, A.C., and Matunis, M.J. (2008). SUMO-2/3 modification and binding regulate the association of CENP-E with kinetochores and progression through mitosis. Mol. Cell 29, 729–741. https://doi.org/10.1016/j. molcel.2008.01.013. Zhao, Y., Wang, Z., Zhang, J., and Zhou, H. (2016). Identification of SENP1 inhibitors through in silico screening and rational drug design. Eur. J. Med. Chem. 122, 178–184. Zunino, R., Schauss, A., Rippstein, P., Andrade-Navarro, M., and McBride, H.M. (2007). The SUMO protease SENP5 is required to maintain mitochondrial morphology and function. J. Cell Sci. 120, 1178–1188. https://doi.org/10.1242/jcs.03418. Zunino, R., Braschi, E., Xu, L., and McBride, H.M. (2009). Translocation of SenP5 from the nucleoli to the mitochondria modulates DRP1-dependent fission during mitosis. J. Biol. Chem. 284, 17783–17795. https://doi.org/10.1074/jbc.M901902200.

Part II

Novel and Advancing Technologies

Identification of SUMOylated and Ubiquitinated Substrates by Mass Spectrometry

5

Francis P. McManus1 and Pierre Thibault1,2,3*

1Institute for Research in Immunology and Cancer, University of Montréal, Montréal, QC,

Canada.

2Department of Chemistry, University of Montréal, Montréal, QC, Canada.

3Department of Biochemistry, University of Montréal, Montréal, QC, Canada.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.05

Abstract Further understanding of the crosstalk taking place between protein SUMOylation and ubiquitination can provide valuable insights into the biological function and turnover of proteins. Recent advances in sample preparation and the development of sensitive mass spectrometers enabled a systemslevel view of this cross communication. Here, we highlight the evolution as well as the merits and limitations of the various workflows that have been created to monitor protein SUMOylation and ubiquitination. Furthermore, this chapter delves into mass spectrometry centred approaches to study the co-occurrence of SUMOylation and ubiquitination on proteins in a non-biased large-scale fashion using immunoaffinity enrichment that target either the co-modified proteins from cell extracts or the modified peptides of the corresponding proteins. Lastly, we provide a perspective on methods that will permit the global analysis of endogenous proteins modified by different ubiquitin-like proteins (UBLs). Introduction Proteins are subjected to a multitude of different post translation modifications (PTMs) that can alter their physiochemical properties, localization

and even function. Although the majority of these modifications are composed of relativity small chemical groups ( 60 000 ubiquitination sites in the human proteome alone. UbiSite strategy The UbiSite strategy (Fig. 5.2D) was developed to overcome some of the limitations of the ubiquitin remnant approach (Akimov et al., 2018). Indeed, the ubiquitin remnant approach cannot distinguish between true ubiquitination sites, ISG15ylation and NEDD8ylation since the same diglycine remnant is produced when all three of these UBLs are digested with trypsin. Furthermore, the monoclonal antibody used for the diglycine enrichment has been suggested to have a compositional bias for certain amino acid sequences around the modified lysine residue, limiting the identification of the whole ubiquitin proteome (Wagner et al., 2012). Lastly, the antibody is unable to identify N-terminal ubiquitination since the diglycine remnant is not located on the ε-amino group of a lysine residue, and hence cannot be recognized by the αK-GG antibody. This method relies on a similar peptide level immunopurification as with the ubiquitin remnant approach (Fig. 5.2C). The difference being that the proteins are digested with Lys-C, revealing a 13 amino acid remnant on the modified lysine residue. This large remnant is UNIQUE to ubiquitin. The ubiquitinated peptides are enriched from the proteome using the UbiSite antibody that recognizes this 13 amino acid remnant. This makes the purification specific for ubiquitin and also allows for the purification of N-terminal ubiquitinated proteins/peptides. Moreover, since

the epitope is larger the antibody relies solely on the remnant for recognition purposes, which is not necessarily the case with the αK-GG antibody. Workflow The pipeline for this method begins with the digestion of the protein extract with Lys-C. The peptides are then desalted by C18 chromatography. The peptide pellet is reconstituted in a native buffer that is supplemented with 0.1% Triton X-100. Although the use of detergents has been found to improve peptide immunopurifications in the past (Lamoliatte et al., 2017), the use of Triton X-100 in the buffer is still surprising considering that most peptide purifications are performed in the absence of detergent due to the lack of compatibility with the downstream LC–MS analysis. After extensive washing of the UbiSite beads with buffer lacking detergent, the peptides are eluted using an acidic buffer (0.1% TFA). The pH is subsequently neutralized with ammonium bicarbonate and the peptides further digested with trypsin to reveal the diglycine remnant that is MS friendly. The peptides can also be fractionated with a high pH STAGE tip to obtain a greater coverage of the ubiquitin proteome. Advantages and limitations There are several advantages to this method. This workflow uses endogenous ubiquitination and can be performed on tissue or patient samples. The main advantage of this method over the ubiquitin remnant approach (Fig. 5.2C) is it’s specificity for ubiquitinated peptides since the antibody used recognizes a 13 amino acid sequence that is not shared with other UBLs. This method also holds the advantage of identifying N-terminal ubiquitination, which has gathered some interest. A slight disadvantage is that the workflow is slightly longer to perform than the ubiquitin remnant approach due to the need for a second protein digestion step after the peptide level purification. Also, the 13 amino acid remnant on the modified lysine residue is too large and therefore difficult for the bioinformatics software to assign to a peptide sequence, thereby requiring that a second digestion be performed. For example, omitting the trypsin digestion at the end of the workflow lead to an 8-fold decrease in ubiquitin site identification.

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Anticipated results Over 41,000 ubiquitin sites were identified with this workflow (including 104 N-terminal ubiquitination sites) when starting from 50 mg of cellular extract and fractionating the trypsin digest mixture over 17 fractions with a basic reverse phase STAGE tip. Despite performing a digestion after the peptide level purification step the overall enrichment of ubiquitinated peptides in the final mixture is an astonishing 23%. While the perspectives of this method are promising, it is too early to tell if it will gain a wide acceptance as it appeared only in July of 2018 and has yet to be used by other groups. The method is promising, and only time will tell if other researchers will adopt this approach or will continue to use the well-established ubiquitin remnant strategy currently employed by most proteomic laboratories. Methods for large-scale SUMO site identification The identification of endogenous SUMOylation sites in the proteome comprises greater challenges than the identification of ubiquitination sites. There

are two major issues that make SUMO site identification so challenging. First, SUMO is less abundant than ubiquitin in the cell, this leads to lower levels of protein SUMOylation. As a result, enrichment procedures for SUMO site identification methods must be improved with respect to those developed for ubiquitin. Second, the native amino acid sequence of SUMO1, SUMO2 or SUMO3, are not amenable to the peptide level immunoprecipitation method that was highlighted and developed for ubiquitin. For example, digesting SUMO3 (Fig. 5.3A) with trypsin generates a 32 amino acid product. Moreover, the most C-terminus arginine residue itself is prone to missed cleavage and the most abundant tryptic product is actually a 34 amino acid peptide. This extremely large amino acid remnant that is left behind on the lysine residue of the target protein after digestion is extremely complicated to identify by MS/MS. This challenge arises due to the complexity of the MS/MS spectra since fragmentation can occur within the backbone or the side chain amino acids of the branched peptides. This mixed fragmentation makes the bioinformatic analysis extremely complicated since most tools have been developed for linear peptides. A few tools were

Figure 5.3  Endogenous SUMO3 amino acid sequence and the various constructs generated to identify SUMO3 sites by proteomics. Blue residues depict the remnant that remains on the SUMO modified lysine residue after the respective workflows. Amino acids that are modified in the constructs are shown in red. Affinity tags that are appended to the SUMO3 gene are shown in green. Filled arrows depict the most C-terminal cleavage site for selective proteases, a dashed arrow depicts the major most C-terminal cleavage site for proteases that are promiscuous.

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developed to aid with this challenge such as an automated recognition pattern tool (SUMmOn) or the creation of a ‘linearized branched peptides’ database (ChopNSpice) (Pedrioli et al., 2006; Hsiao et al., 2009). However, none of these tools have found routine use, suggesting that they are not the ideal solution to the problem. Since most of the large-scale proteomic studies have been conducted with SUMO3, this chapter will dive into methods that were developed for SUMO3 site identification, but the principals can be applied and modified as needed to fit the requirement imposed by the other SUMO isoforms. To overcome these issues various SUMO3 constructs have been developed, where arginine or lysine residues are placed at the C-terminus of the SUMO3 gene to create smaller remnants on trypsin digestion, these are highlighted in Fig. 5.3B–D. These ‘first generation’ methodologies required that the modified SUMO3 be ectopically expressed to include the variant of the protein. Although the CRISPR methodology is routinely in use in the present day to knock-out genes of interest, the knock-in method is still technically demanding due to its higher complexity and lower efficacy. Indeed, no one has yet to knock-in the SUMO3 variants into the genome of an organism. It is therefore of the utmost importance that the SUMO3 variant be expressed at near basal levels in the cell. Considering that SUMOylation has been shown to alter so many biological functions the unbalance created by high levels of SUMO3 in the cell has the potential to deregulate the biology of the system. Lastly, other groups have circumvented the poor MS/MS identification of long remnants by using other, less commonly employed, proteases that digest SUMO3 closer to its C-terminal, producing a smaller remnant that is compatible with regular LC–MS/MS (Fig. 5.3E and F). In contrast to the evolution of the ubiquitination field that started with protein level purification for site identification purposes, the SUMO site identification methods rely on the peptide level purification strategy, inspired by the diglycine remnant peptide level purification (Fig. 5.4). Although most SUMO peptide level purifications are antibody based strategies, the K0 method that will be described in detail later on in this chapter, relies on a second Ni-NTA purification. Albeit, many lessons were learned from the ubiquitin field and employed

for the SUMO site identification strategies that greatly aided in the method development process. While some groups have pioneered the field by transient over expression of SUMO proteins for the identification of SUMO sites in the proteome, this chapter will focus solely on methods that rely on near endogenous SUMO levels, so that biological functions be maintained (Impens et al., 2014). Also, this chapter aims at the unambiguous identification of SUMOylation sites. This requires that a remnant or diagnostic modification be present to truly classify the site as a bona fide target of SUMOylation. Therefore, methods such as the Protease-Reliant Identification of SUMO Modification (PRISM) that relies on identifying SUMO sites by monitoring non-acetylated lysine residues that are generated after chemical acetylation of the proteome followed by removal of SUMO with a SUMO protease, which has its own merits, will not be included in this chapter. SUMO K0 strategy As shown in Fig. 5.4A, for the K0 approach the SUMO3 construct has all of its lysine residues changed to arginine rendering the protein refractory to Lys-C digestion (Schimmel et al., 2008). Moreover, a poly-histidine tag is appended to the N-terminal of the protein for purification purposes. It should be noted that this K0 method is based on the same K0 approach that was previously created for ubiquitin proteomics, which found a great degree of success (Oshikawa et al., 2012). Workflow The plasmid that encodes the poly-histidine tagged K0 SUMO3 variant is inserted upstream of a GFP reporter protein that are expressed from the same RNA transcript as separate proteins by means of an inner ribosomal entry site located between the two genes (Hendriks et al., 2014). This construct permits for fluorescence-activated cell sorting (FACS) and selection based on cell GFP fluorescence. Moreover, since both GFP and the SUMO3 variant are expressed from the same RNA the level of ectopically expressed SUMO3 is directly correlated to cell GFP signal. Cell populations can therefore be selected with SUMO3 variant expression that are similar to endogenous SUMO3 levels. Cells that have been selected by FACS are expanded and lysed in a harsh denaturing buffer

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Figure 5.4  Methods developed to map SUMOylation sites by proteomics.

containing guanidine to eliminate all cellular function since SUMOylation is thought to be a highly dynamic process. The protein extract is loaded on Ni-NTA beads for a first round of SUMO purification by means of the poly-histidine tag that is appended on the SUMO3 construct. After extensive washing, the bead bound material is eluted with high concentrations of imidazole. The eluted sample is applied to a 100 kDa molecular

weight cut-off membrane to remove free SUMO from the sample. The retained material is subsequently digested with Lys-C. This breaks down all the proteins into shorter fragments, except for the K0 SUMO3 due to the absence of lysine residues. The digested material is then loaded once more on Ni-NTA beads, where a second purification step ensues. After extensive washing of the beads the material is eluted once again from the beads with

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imidazole. The eluate is then concentrated on a 10 kDa membrane and digested with trypsin prior to analysis by MS. This semi-peptide level purification is extremely beneficial since most peptides are not retained on the beads, leaving behind the target SUMOylated peptide attached to the solid support through the K0 SUMO3 protein. Advantages and limitations An advantage of this SUMO site identification methodology is that antibodies are not required. This makes this method less expensive since Ni-NTA beads are much more cost effective than antibody bound beads. This method is not without its drawbacks though. The method requires the samples to be passed through different molecular weight cut-off filters, which are renowned to lead to considerable sample loss. The major drawback of this method is the biological validity of the results. The method is not optimal for biological assays, but rather is reserved for the identification of SUMO sites on a large scale. Since there are no lysine residues on the SUMO3 construct, all other UBL conjugation (including SUMO itself) on K0 SUMO are abolished. This might be of importance since the crosstalk between different PTM on biological function have grown in scope, and it is clear that methods enabling the identification of UBL crosstalks will be of interest. For example, the poly-SUMOylation of PML is required for the subsequent ubiquitination of the protein by RNF4 and its eventual degradation (Weisshaar et al., 2008). Since PML nuclear bodies are a hub for protein SUMOylation, hindering its degradation will cause increased SUMOylation in the K0 SUMO expressing cells, which can alter the biology of the system. Anticipated results Nonetheless, this method paved the way for the identification of SUMO3 sites in human cells. This method garnered over 4300 SUMOylation sites (Hendriks et al., 2014). The results from this method were improved by deepening the depth of the analysis through basic reverse phase fractionation of the SUMOylated peptides and testing different cell lines under different stimuli to raise the number of SUMOylation sites to 40,765 (Hendriks et al., 2017). The final enrichment of identified SUMO peptides over the total number of peptides is around 50%, which is excellent.

SUMO αK-GG strategy This approach to SUMO site identification relies on introducing a specific protease cut site just before the pair of glycine residues that are located at the C-terminus of the protein (Fig. 5.4B). Since NEDD8, ubiquitin and ISG15 all terminate with an RGG motif, a typical tryptic digestion cannot be employed to unambiguously differential the different UBL modifications. To address this a T90K alteration is introduced, creating a KGG terminal (Tammsalu et al., 2014). The proteins are therefore digested with the Lys-C protease rather than the typical trypsin protease to selectively generate the diglycine remnant on the SUMO3 modified lysine residues, leaving the C-terminal of the other UBL that terminate with the RGG motif unaffected. The resulting diglycine modified peptides can then be immunopurified from other peptides with the commercially available antibody. Workflow Stable cell lines that express the poly-histidine tagged T90K variant of SUMO3 are expanded in regular DMEM and lysed in the same harsh denaturing buffer used in the K0 approach. The cell lysate is subsequently loaded on Ni-NTA and the beads washed extensively. The enriched proteins are then eluted from the beads with high imidazole concentrations. The eluted proteins are digested with Lys-C to reveal the diglycine motif on the SUMOylated lysine residues. Since Lys-C can only digest after lysine residues and not after arginine residues, some large peptides are generated that are not compatible with large-scale proteomic analysis on the high resolution Orbitrap instruments. To address this, the Lys-C digested peptides are desalted and applied to a 30 kDa molecular weight cut-off membrane. A diglycine immunoprecipitation can be performed on the small peptides that are not retained on the membrane. For the larger peptides that are retained on the molecular weight cut-off membrane, the sample is further digested with Glu-C. The doubly digested peptides are then passed through the 30 kDa molecular weight cut-off membrane and SUMOylated peptides immunoprecipitated from the remaining material with the αK-GG antibody. This means that two LC–MS injections are required per sample, as noted by the authors. The authors also indicated that the bioinformatics search is hindered when analysing the

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Lys-C only digested samples with both proteases in the search engine. Therefore, two separate bioinformatics searches are recommended. Importantly, during the optimization of this pipeline the authors uncovered an important MS parameter that can greatly aid in the identification process. Typically, the injection time for the MS/ MS scan is in the low ms range, usually 50–80 ms. This injection time is usually appropriate for extremely complex samples like whole proteomes or even phosphopeptide analysis where 20,000 or more peptides can be assigned within a 1 h LC– MS program. On the other hand, SUMO peptide samples are much less complex and typically only harbour several hundreds to a couple thousand peptides. More importantly, these SUMO peptides tend to be of low abundance and the automatic gain control (AGC) target is rarely reached within the injection time window. Therefore, increasing the MS/MS injection time improves the sensitivity of the analysis at the cost of speed. MS scan speed is not a major concern since the SUMO peptide chromatograms are of low complexity and longer duty cycles do not considerably impact the MS/ MS scan rate for newly sequenced peptides. When using a classic MS program with a 60 ms MS/MS injection time the authors identified 352 SUMO sites. On the other hand, when using a 1 second injection time the authors identified 596 SUMO sites. Moreover, the increased MS/MS injection time not only increased the number of SUMO sites but also augmented the reported enrichment level. This increase is due to the more complex nature of branched peptides and their lower abundance in the purified samples, requiring more time to collect enough ions for proper assignment. Advantages and limitations This method is practical since it uses already commercially available tools, such as the αK-GG antibody. Also, this variant of SUMO3 is biologically similar to the endogenous protein, making this tool applicable to study the biology of SUMOylation in different contexts. On the other hand, it is one of the longer methods since the samples must be applied to molecular weight cut-off membranes and digested twice. Also, two injection on the LC–MS are required per biological sample, in effect doubling the instrument usage time. One final drawback is that other UBL modifications cannot

be monitored since trypsin is not used in the workflow. The remnants produced on Lys-C or even Glu-C digestion for ubiquitin, ISG15 and NEDD8 are too large to be identified with conventional bioinformatics software. Anticipated results The hallmark of the first 1000 SUMO sites in a single study was attained using this method in 2014. A total of 1002 unique SUMO sites were identified from 286 mg of extract. Overall, this method provides valuable results with up to 45% of the identified peptides being assigned as SUMOylated peptides. This advanced enrichment is attributed predominantly to the αK-GG immunoprecipitation that was found to increase the overall enrichment by more than 600-fold. This method provided the first example of using an antibody for the peptide level enrichment for SUMOylation site studies, which is a common theme with the remainder of the methods discussed below. SUMO αK-NQTGG strategy Like the previous two methods, this SUMO site identification strategy requires the use of an exogenously expressed form of SUMO3, Fig. 5.4D (Lamoliatte et al., 2014, 2017; McManus et al., 2017). As with the previous methods, a polyhistidine tag (6 × HIS in this case) is appended to the N-terminus of the protein for the first enrichment at the protein level. At the C-terminal the final QQQTGG sequence is replaced with RNQTGG. The arginine that is introduced at the C-terminus produces a small epitope remnant of 5 amino acids (NQTGG) once the protein is digested with trypsin. This epitope is unique to the construct and is not endogenously found attached to lysine residues in the human proteome. The Q to N alteration that is found at position −5 with respect to the C-terminal was introduced purely to distinguish the SUMO2 and SUMO3 constructs produced by the same group. Indeed, three SUMO constructs, each for a different SUMO paralogue, is available for large-scale proteomics. All constructs harbour the N-terminal poly-histidine tag as well as the arginine residues at position −6 with respect to the C-terminus. In essence, the same method can be used with the various constructs to study the different SUMO paralogues. Lastly, the plasmids have a geneticin resistance gene allowing for the

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creation of stable cell lines. To improve the biological relevance of the results clones are isolated and tested for the level of expression of the SUMO construct to match that of the endogenous SUMO levels.

extensively washing the beads with PBS and eluting the SUMO3 modified peptides with an acidic buffer (0.1% TFA). The purified peptides can also be fractionated on an SCX STAGE tip to increase the coverage of the SUMO proteome.

Workflow The selected clone that expresses the SUMO3 construct at near endogenous levels is expanded in DMEM media supplemented with geneticin and the cells lysed in a denaturing buffer containing 6 M guanidine. The lysate is applied on a Ni-NTA resin for the protein level purification. The Ni-NTA beads are washed extensively with a chaotropic buffer containing 8 M urea. Rather than eluting the material from the beads with imidazole, like the two previous methods, the proteins are digested directly on the Ni-NTA beads prior to their desalting which improves peptide recovery. PolySUMOylation readily occurs on some proteins generating a branched network of proteins that can have several poly-histidine tags, hindering their elution from the beads. The on-bead digestion also saves time since the elution process with imidazole (up to 30 minutes) is bypassed. However, a drawback of this step is its lower selectivity. Trypsin digestion will cleave all proteins after lysine and arginine residues. On the other hand, eluting with imidazole is more specific since it competes directly with the poly-histidine tag for the Ni2+ ions on the solid support. Therefore, imidazolebased purifications do not elute proteins that have adsorbed or indirectly interact with the agarose or sepharose solid support. The lack of specificity from the trypsin digestion to effectively ‘elute’ the proteins from the beads is overcome by the second purification step that further decomplexifies the sample. The tryptic digestion of the SUMO3 variant reveals the NQTGG motif on the SUMO3 modified lysine residue. The peptide level enrichment in this method uses a new and highly specific antibody raised against the NQTGG remnant on the ε-amino group of lysine. As with the αK-GG antibody, the αK-NQTGG antibody is bound to protein A beads prior to their chemical crosslinking using dimethyl pimelimidate (DMP). The SUMOylated peptides are immunoprecipitated from the Ni-NTA enriched sample using the solid support bound αK-NQTGG by applying the peptide sample on the αK-NQTGG containing beads,

Advantages and limitations In contrast to the other SUMO site workflows only a single digestion with trypsin is required in the whole process. The other methods generally use other proteases that are less commonly used which can lead to larger peptides, rendering the peptide assignment by MS/MS more difficult. This method can be used for the other SUMO paralogues by simply transfecting the desired construct in the host cell. No alterations are required to the workflow to follow the other SUMOylation events that take place in the cell. As with the αK-GG approach, this method can be used to follow biological processes in the cell since the poly-SUMOylation of SUMO3 is not hindered. This method is also more efficient than the previous workflows, where the whole protocol can be performed in 2 days. The highest enrichment rates are obtained with this SUMO site identification methodology. These elevated levels of purity are attributed solely to the αK-NQTGG since this is the step that is unique to this method alone. Indeed, the αK-NQTGG antibody is extremely selective, in part due to the size of the epitope that spans 6 amino acids (when counting the lysine that is modified), which is closer to the 8–20 amino acid optimal antibody recognition length (Xu and Jaffrey, 2013). Most importantly, this method is capable of monitoring SUMOylation and ubiquitination events from the same sample to understand the crosstalk that prevails under different biological stimuli. The orthogonality that is provided by having different epitopes for SUMO (NQTGG) and ubiquitin (GG) allow the two PTM modified peptides to be isolated sequentially from the same sample. The sequential immunoprecipitation of SUMOylated and ubiquitinated peptides will be further discussed in greater detail later in this chapter, see Sequential Peptide Strategy (Lamoliatte et al., 2017; McManus et al., 2017). However, this method also has its own drawbacks. It relies on the purchase of the commercially available αK-NQTGG antibody and requires exogenous expression of a SUMO construct.

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Anticipated results A total of 1200 SUMO sites can be identified from a single LC–MS injection when starting from 4 mg of cell extract. The comprehensiveness of the SUMO proteome can also be expanded on by performing an offline strong cation exchange (SCX) fractionation where ≈ 10,000 SUMO sites can be identified from 16 mg of cell extract fractionated into 6 SCX fractions. This method also provides the highest purity rate of all methods, where enrichment levels above 70% are typically obtained. SUMO WaLP strategy This is the first method that was developed to identify endogenous SUMOylation sites and can be used in clinical samples or other tissue samples (Fig. 5.4D). Moreover, there is no need for cloning or cell transfection. This method relies on an atypical protease that is seldom used in proteomics. The wild-type Alpha-lytic protease (WaLP), which was first found in Lysobacter enzymogenes, cleaves preferentially after threonine, alanine, serine and valine residues (Meyer et al., 2014). The specificity of the enzyme is more relaxed than trypsin, which generates more peptides and increases the bioinformatic search space when trying to assign peptide sequences to the in silico library. This enzyme can be used for all SUMO paralogues since they terminate with a TGG motif. Therefore, digesting the proteome with this enzyme will expose the diglycine motif on the SUMOylated lysine residues. This technique therefore identifies all SUMO sites at once and not just one specific paralogue, which was not the case for the previous methods. Since the mature form of ubiquitin terminates with an RGG motif, the WaLP enzyme does not expose the diglycine remnant on ubiquitin, which is fundamental for the selectivity of this method. The selective generation of the diglycine remnant on SUMOylated peptides allows for the enrichment of these peptides with the commercially available αK-GG antibody. Workflow The workflow is simple and is reminiscent of the ubiquitin remnant method. Cells are grown in regular DMEM media or tissue samples can be used. Typically, 5–10 mg of cell extract/protein is needed per sample. The cells are lysed in a buffer containing 8 M urea. Or likewise, tissue samples

are homogenized in the same buffer. The cysteine residues are reduced with dithiothreitol (DDT) or Tris(2-carboxyethyl)phosphine hydrochloride (TCEP) followed by their alkylation with 2-iodoacetamide. DTT is added once again to quench the remaining 2-iodoacetamide. It is noteworthy that WaLP is a serine protease and therefore it is not essential to quench the 2-iodoacetamide. The buffer is diluted 4-fold with a Tris buffer at pH 8.0 to bring the urea concentration below 2 M. WaLP is added to the sample at a 1:100 ratio and allowed to react overnight at 37°C. The digestion is then terminated by acidifying the sample with TFA. The peptides are desalted on a reverse phase column and the diglycine modified peptides are enriched with the commercially available antibody from cell signalling technologies (αK-GG). The enriched peptides are ready for analysis by LC–MS. Advantages and limitations The most attractive advantage of this method is the ability to perform the workflow in native cells without the need for any molecular biology. This method enables SUMO site identification to be conducted with tissue samples or patient samples. Performing SUMO site identification and quantification in patient samples is of great importance since several novel biomarkers are actual PTM derived peptides (Andersen et al., 2010). The developed workflow is simple and efficient. It takes the same amount of time to perform this workflow as for the ubiquitin remnant approach, which can be done in a single day. Lastly, this method has the advantage or disadvantage of looking at all SUMO paralogues at the same time. The paralogue specificity is lost when using this workflow, but more biological information can be garnered since all paralogues are probed at once. This means that further experiments need to be conducted on selected targets to validate which SUMO paralogue was conjugated to the substrate. Unfortunately, the WaLP enzyme has ‘relaxed specificity’, as indicated by the authors. The enzyme is not selective for threonine but also cleaves after leucine and isoleucine to a certain degree. As a result, this protease can release a diglycine motif on lysine residues that are endogenously modified by FAT10 and FUBI. However, by using the SENP protease to remove the conjugated SUMO, the authors noted that 88% of the diglycine containing peptides reduced

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in abundance, suggesting that the vast majority of the identified SUMO sites are in fact correctly assigned. As a result of WALP’s lack of specificity, the bioinformatics search is not efficient and requires that no enzyme be selected and special software trained by machine learning for WaLP digested samples must be used. Moreover, electron transfer dissociation (ETD) fragmentation must be employed to improve MS/MS spectra assignment which limits its wider application in view of the widely used collisional activation regimes. Lastly, SUMOylation and ubiquitination cannot be studied sequentially from the same sample due to the need for different proteases for the different PTM. SUMO site identification relies solely on WaLP, while the ubiquitination studies require trypsin. Employing both proteases simultaneously is not feasible since the same diglycine remnant would be produced for both SUMOylation and ubiquitination sites. Anticipated results A modest number of SUMOylation sites were identified using the WaLP workflow. This could be due in part to the poor identification of non-tryptic peptides by the bioinformatics software as well as the relatively low abundance of native SUMO in the cell. A total of 1209 SUMO sites were obtained from 15 mg of cell extract when using a classic LC–MS setup. The depth of the SUMO proteome could have been improved had the authors used a 2D LC methodology, which has greatly expanded the SUMO repertoire in the past (Lamoliatte et al., 2017). SUMO Lys-C + Asp-N strategy This most recent method permits an in depth analysis of the endogenous SUMO2/3 proteome (Fig. 5.4E). Unlike the WaLP method discussed above, this workflow allows for the selective identification of endogenous SUMO2/3 sites (Hendriks et al., 2018). It is currently unable to identify SUMO1 nor SUMO4 sites, though changes to the workflow could be performed in the foreseeable future to allow for their analysis as well. This method makes use the 8A2 SUMO2/3 antibody that recognizes the C-terminus of SUMO2/3 (Zhang et al., 2008). The epitope of 8A2 (57-IRFRFDGQPI-66) is still present when SUMO2/3 is digested with Lys-C since

the last lysine residue on SUMO3 is at position 44. This digestion leaves a 48 amino acid remnant on the target lysine residues that were originally modified by SUMO2/3, which is not MS compatible for the same reasons that were eluded to earlier. As a result, after the peptide level immunoprecipitation with 8A2 antibody a second digestion is required to shorten the remnant to a size compatible with the commonly used search engines to facilitate the assignment of SUMOylation sites from the MS/ MS spectra. Therefore, the peptides are further digested with Asp-N to generate a nine amino acid remnant composed of DVFQQQTGG on the modified lysine residue. Surprisingly, the bioinformatics tool MaxQuant is capable of assigning these large branched peptides without the need for major alterations to the program. For proper assignment diagnostic ions are appended to the PTM that is added to the software’s search algorithm. It should be noted that the largest remnant that could be assigned by MaxQuant prior to this study were the 5 amino acid QQTGG and NQTGG sequences (Fig. 5.4A and C). Workflow As with all workflows, cultured cells are grown in regular DMEM media or tissue samples can be used. Typically, 120 mg of cell extract/protein is needed per sample. The cells are lysed and protein denatured in a buffer containing 6 M guanidine. After reduction and alkylation with TCEP and 2-chloroacetamide, respectively, the proteins are digested with Lys-C. The guanidine concentration is then reduced to 1.5 M, at which point a second Lys-C digestion is conducted. The peptide mixture is then applied to a C8 column. The smaller peptides are eluted from the column with low acetonitrile concentrations (25–35%). Since the SUMO2/3 remnant is 5.6 kDa, this large hydrophobic moiety is retained on the column. The SUMOylated peptides are then eluted with increasing acetonitrile concentrations (35–45%). After lyophilizing the C8 eluate, the SUMO modified peptides are enriched using protein G agarose beads that are functionalized with the 8A2 antibody. The immunoprecipitated material is subsequently digested with Asp-N to reduce the remnant on the modified lysine residues to a nine amino acid sequence of ≈ 1 kDa. The SUMO

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modified peptides are then fractionated by basic reverse phase using STAGE tips to enhance to coverage of the SUMO proteome. Advantages and limitations Although further improvements to the method are likely to appear in the future, this is currently the most efficient workflow for endogenous SUMO site identification. This method allows for the largest coverage of the SUMO proteome at the endogenous level. One drawback of the method is the need for a large search space during the bioinformatics analysis since the search is conducted with eight missed cleavages. This makes the bioinformatics search, extremely long and also affects the peptide score for the false discovery rate (FDR) cut-off, which may explain why the authors used a 2% FDR cut-off instead of the typical 1%. Unfortunately, this method is not compatible with ubiquitin site identification methodologies. Since the immunoprecipitation is performed after the Lys-C digestion the SUMO modified protein is already digested into its constituent peptides and ubiquitinated sites found on other locations of the protein are washed away during the 8A2 mediated purification. Alterations to the protocol could be employed to circumvent this issue, such as performing the 8A2-based purification at the protein level, prior to the Lys-C digestion. The purified material could then be digested with Lys-C and the 8A2 based purification repeated. The UbiSite purification (Fig. 5.2d) could be performed on the purified digest to identify ubiquitination sites and the flow through from the UbiSite purification digested with Asp-N for SUMO site identification. Anticipated results This workflow was repeated several times to garner ≈  14,000 endogenous SUMO2/3 sites in the human proteome. This method can yield ≈ 8500 SUMOylation sites when starting from ≈ 100 mg of cell extract. Although there is no mention of the enrichment ratio for this method, it is probably lower than those observed for the other methods. The reason being that several new peptides are generated when performing the Asp-N digestion on the immunoprecipitated material. Since there are no further purification steps after the digestion it is likely that the SUMO modified peptides constitute a small proportion of the final peptide pool.

Current methods to study ubiquitin and SUMO co-modified proteins The study of the crosstalk between the various UBL on target proteins or on a global biological level is in infancy. Currently, there are less than a handful of methods that allow for the global study of crosstalk events between UBLs. The methods that have been developed have focused on the interplay between ubiquitination and SUMOylation. However, new methods will likely be developed to improve our understanding of the crosstalk between other UBLs. Indeed, advancements in peptide level immunopurification, improvements in MS instrumentation along with novel proteases will clearly help in method development. For example, a new protease from the foot-and-mouth disease virus called Lbpro has been recently characterized and was shown to selectively cleave before the diglycine remnant of ISG15 (N-terminal to the diglycine), making it a prime candidate to study this modification (Swatek et al., 2018). Moreover, this selective cleavage can easily be used in tandem with other workflows to incorporate ISG15 crosstalk with other UBLs or PTM. Over the past decade, we witnessed important technological advances in the field of large-scale proteomic analysis of SUMOylation and ubiquitination that served as building blocks towards the creation of workflows to study proteins comodified by these PTMs. The methods that have been developed to study the crosstalk between ubiquitin and SUMO are mixtures of previously created pipelines. The challenge is therefore not the creation of new methodologies, but the process of combining and optimizing workflows together that can be used in a sequential fashion. The creation of these workflows relies on employing strategies that are compatible with each other biologically and technically. For instance, the K0 SUMO site approach (Fig. 5.4A) is not compatible to any ubiquitin approach in a biological context since the UBL polymerization on SUMO (Fig. 5.1B) is impeded. On the other hand, the αK-GG approach (Fig. 5.4B) is biologically compatible with the ubiquitin remnant approach (Fig. 5.2C) but is not technically suitable since both PTM will provide a diglycine motif on their respective lysine modified residues, rendering them indistinguishable by LC–MS/MS.

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Sequential protein strategy This method, which is highlighted in Fig. 5.5A, is the only method in this chapter that does not study SUMOylation of ubiquitination at the site level (Cuijpers et al., 2017). The method relies on the affinity tagged approach to study ubiquitination and SUMOylation (Fig. 5.2A). A flag tag is placed on the N-terminus of ubiquitin and a poly-histidine tag at the N-terminus of SUMO. The principle of the method is simple and is easy to perform. All reagents that are required for this approach are

easily accessible. The workflow relies on performing two enrichment steps at the protein level, each enrichment for the different UBLs. Although the workflow itself is simple, the data filtering and analysis is more complex. Several control experiments need to be performed to increase the validity of the results. For instance, the same workflow needs to be performed with untransfected cells to rule out possible false negative hits, since some proteins may not selectively bind to the solid supports used during the purification steps. Moreover, another set

Figure 5.5 Proteomic methods for global identification of proteins that are co-modified with SUMO and ubiquitin.

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of control experiments where the affinity tags are swapped (SUMO is Flag tagged and ubiquitin is 10 × His tagged) is highly suggested to reduce false positive identification. Workflow To begin, cells are transduced with the 10 × HisSUMO3-IRES-GFP plasmid and filtered by FACS to isolate cells that express GFP. These cells are transduced with Flag-Ubiquitin-Puromycin and cultured for 2 weeks under puromycin selection. This stable cell line expressing both constructs are then grown in standard DMEM growth media. Cells are lysed in a denaturing buffer containing 6 M guanidine. The cell lysate is then loaded on Ni-NTA beads for the first level of enrichment. The protein bound beads are then washed extensively under denaturing conditions. Proteins are eluted from the Ni-NTA resin with high imidazole concentrations. The harsh denaturing conditions are important to eliminate the co-purification of interacting proteins. This is extremely important for this workflow since SUMO and ubiquitin sites are not identified and the method relies solely on the identification of the purified proteins. The eluent is then concentrated on a 100 kDa molecular weight cut-off membrane. This concentration serves two purposes: (1) it allows for the eventual buffer exchange step that is needed for the Flag purification, and (2) the free 10 × His SUMO3 protein that was also retained on the Ni-NTA beads are washed through the membrane since it is only ≈ 13 kDa. After the sample has been concentrated, the proteins are diluted into a native buffer and filtered through a 0.45 μM membrane to eliminate precipitated proteins that did not renature properly. The filtrate is applied to α-Flag antibody functionalized beads to purify ubiquitin modified proteins. The solid support bound material is exhaustively washed with native buffer and the proteins digested on beads with trypsin. The resulting peptides are then analysed by LC–MS/MS. Advantages and limitations The main advantage of this method is that it can be used to look at proteins that are co-modified by SUMO and ubiquitin without the need for expensive antibodies. The α-Flag antibody that is already linked to agarose beads are readily available and are ≈ 5 times less expensive than specialty

antibodies that are typically used for peptide level purification. This makes the method inexpensive (with the omission of the MS usage time) with respect to most other workflows. However, there are several disadvantages with this method. It provides poor substrate overlap for unstimulated cells, probably due to the low stoichiometry of co-modified proteins. For this reason, cells need to be stimulated with a proteasome inhibitor to generate robust overlap between experiments. As eluded to earlier, this method requires several controls, greatly increasing the number of samples and manual labour. These extra controls also increase MS usage time, greatly increasing the cost of the overall experiment. The most important limitation of this method, which is also the cause of several of the aforementioned disadvantage, is the inability to identify the sites of SUMOylation and to some degree the ubiquitination sites. Half of the ubiquitination sites that were identified using this method were on SUMO1, SUMO2, SUMO3 or ubiquitin directly, highlighting that only the most abundant sites are identified using this methodology. Furthermore, it is documented that proteomics results are much more reliable and reproducible when looking at modification sites directly rather than at modified proteins as a whole (Hendriks and Vertegaal, 2016). Anticipated results This method allowed for the identification of 498 co-modified proteins when starting from ≈ 50 mg of cell extract per sample. Interestingly, an additional 1545 proteins were pulled down by the sequential protein purification workflow. Therefore, roughly a quarter of the proteins in the final samples were deemed as co-modified. This method could have benefited with a fractionation level prior to the LC–MS analysis, thus expanding the repertoire of ubiquitin and SUMO co-modified proteins. Sequential peptide strategy This approach is currently the only method that allows for the unambiguous identification of SUMO and ubiquitin co-modified proteins at the site level. The workflow presented in Fig. 5.5A relies on the αK-NQTGG procedure (Fig. 5.4C) as well as the endogenous identification of ubiquitination sites by the ubiquitin remnant approach (Fig. 5.2C) (Lamoliatte et al., 2017; McManus et al.,

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2017). These two workflows are biologically and technically compatible. Biologically, the SUMO3 variant that is employed (Fig. 5.3D) behaves just like the endogenous protein, where SUMO and ubiquitin polymers are formed readily. Technically, these two methods work well in concert. Trypsin is the sole protease that is required since it reveals the diglycine remnant for the ubiquitin peptide pull down as well as the NQTGG remnant for SUMO3 peptide purification. Moreover, the buffer that is used for the ubiquitin peptide immunoprecipitation can easily be supplemented with glycerol for the subsequent SUMO3 peptide enrichment. Hence, there is no need for a buffer exchange step in this workflow. Every step of the protocol was optimized to allow the whole workflow to be conducted in 2 days. This procedure unambiguously identifies the SUMOylation and ubiquitination sites on co-modified protein, alleviating the need for several controls as direct evidence of the modifications is observed. Workflow Cells that express the NQTGG SUMO3 variant (Fig. 5.3D) are expanded in regular DMEM media. After harvesting, the cells are lysed in a highly chaotropic buffer containing of 6 M guanidine. The poly-histidine tagged SUMO3 proteins are enriched from the total protein pool on Ni-NTA beads. After exhaustively washing the matrix with a denaturing buffer the proteins are directly digested with trypsin on the beads. The tryptic peptides are desalted on a C18 cartridge and lyophilized. The peptides are suspended in a phosphate buffered saline (PBS) solution and added to agarose beads that have been chemically crosslinked with the αK-GG antibody to immunoisolate the ubiquitinated peptides. After a 1 hour incubation the flow through is removed from the αK-GG beads. This flow through is supplemented with 90% glycerol in PBS to a final concentration of 50%. The glycerol containing sample is placed with αK-NQTGG antibody crosslinked to magnetic beads for a 1 hour incubation to enrich the SUMOylated peptides. The αK-GG beads are washed several times with PBS and a couple times with 0.1X PBS. The ubiquitinated peptides are eluted from the αK-GG beads with an acidic buffer and lyophilized for LC– MS/MS analysis. After the 1 hour incubation with the αK-NQTGG beads, the peptide containing

supernatant is removed and discarded or saved for further analysis. The beads are washed and eluted in the same way as the αK-GG beads. Lastly, the SUMOylated peptides are fractionated on an SCX STAGE tip by eluting peptides with plugs of increasing concentration of ammonium acetate. The fractions are then dried down in a speed vacuum prior to LC–MS/MS analysis. Advantages and limitations This is currently the only method that permits the study of the interplay between SUMOylation and ubiquitination at a site specific level. Site identification reinforces the results greatly and also provides important information for follow-up experiments. The workflow is simple, requires no specialty proteases but does require the αK-NQTGG antibody. The whole workflow requires 2 days to perform (excluding MS analysis), which is shorter than the other methods that only identify SUMOylation sites. In contrast to the Sequential Protein Strategy (Fig. 5.5A) there are no controls that are required as a result of the direct evidence of the modification sites on the proteins. The main disadvantage of this method however is the need for an exogenous SUMO3 variant that is compatible with the αK-NQTGG antibody. Anticipated results This method provides the greatest coverage for the ubiquitin and SUMO crosstalk. Starting from 16 mg of starting material more than 9000 SUMO sites and 4500 ubiquitin sites can be identified. Moreover, more than 1000 substrate proteins can be observed as directly co-modified, where ubiquitination and SUMOylation sites are observed to occur on the target proteins. On the other hand, several substrates were modified only by ubiquitin or SUMO, highlighting a site-specific recognition of UBL enzymes. These co-modified proteins stem from SUMO and ubiquitin polymerization that occur on a single lysine residue of the target protein (Fig. 5.1B, right panel). For example, ubiquitin is attached directly on the substrate protein and SUMOylation occurs on the ubiquitin moiety. As a result, the protein is pulled down in the Ni-NTA extract but the SUMOylation site that is identified for this specie is a SUMOylation event on a ubiquitin lysine residue.

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Concluding remarks and future perspectives The workflows presented in this chapter have evolved dramatically over the course of the past decade. The technology started with protein purification by appending tags on the UBL of choice followed by proteolytic digestion prior to LC–MS analysis. These methods, though ground breaking at the time, have become obsolete. Rather, peptide level purification has dominated the field of largescale PTM profiling. The peptide level purification is a vastly superior methodology that isolates only the modified peptide and not the whole protein. Moreover, the final purity of the sample is far superior since the isolation are performed on digested proteins, alleviating any sort of protein–protein interaction that can cause the co-isolation of unwanted species in the final sample, which is the case when performing purifications at the protein level. It is important to note that peptide level enrichments allow for the direct identification/ quantification of the actual site of SUMOylation or ubiquitination, which is beneficial for the robustness of the final data set and for future biological validation and functional follow-up experiments. There are some major drawbacks however to all the methods above. The inherent nature of bottomup proteomics makes it impossible to know the linkage of the UBL that are identified. Indeed, since the protein are digested only a small stub or remnant remains where the UBL was originally tethered. It is therefore impossible, with the workflows presented earlier, to determine if the protein substrates are mono- or poly-SUMOylated on a lysine residue or even if there are mixed chains at the specific site. This drawback is a current limitation that hampers our understanding of the full nature of the crosstalk between UBLs. Large-scale proteomic analysis is, however, the optimal tool to use in the discovery stage due to its unbiased, sensitive nature. To obtain a more in depth view of the crosstalk other methodologies can be employed. For instance, follow up experiments by western blot can be performed for specific target substrates that have been identified by proteomics to view the polymerization pattern, providing some complementary information on whether the protein is poly-SUMOylated (or polyubiquitinated). The proteomic workflows that are available to monitor the interplay between protein

SUMOylation and ubiquitination currently rely on the exogenous expression of either SUMO and/or ubiquitin with an N-terminal tag (Fig. 5.5). Considering the recent developments and tools available, we expect that a proteomic workflow capable of monitoring the crosstalk at the endogenous level will emerge soon. In light of the information contained in this chapter we propose the following workflow as a viable approach to a fully endogenous method that could reliably map the ubiquitination and SUMOylation crosstalk (Fig. 5.6). This workflow relies on a combination of the endogenous SUMOylation site identification method (Fig. 5.4E) and the UbiSite workflow (Fig. 5.2D) with an added SUMO protein immunopurification. These are the two newest methods that have recently emerged and appeared to be technically and biologically compatible for the isolation of endogenous proteins. To monitor the crosstalk either ubiquitin or SUMO proteins must be isolated from the whole cell extract prior to the isolation of the SUMOylated and ubiquitinated peptides. The 8A2 antibody would be a prime candidate to immunopurify SUMO proteins (Fig. 5.6 ‘SUMO IP1’) due to its selectivity and its use in the later stage of the purification, thereby reducing the cost of the work flow. Moreover, the 8A2 antibody has been exhaustively characterized and a selective peptide based elution is possible since the epitope has been mapped by the Melchior lab (Becker et al., 2013). A Lys-C digestion (Fig. 5.6 ‘Lys-C digestion’) can be performed subsequently on the SUMO enriched sample to release the UbiSite epitope on the ubiquitinated peptides and partially digesting the proteome to aid in the subsequent SUMO immunoisolation with the 8A2 antibody (Fig. 5.6 ‘SUMO IP2’). After the immunoisolation with the 8A2 antibody, the purified extract is further digested with Asp-N to unveil the DVFQQQTGG remnant on the SUMOylated peptides prior to LC–MS/MS analysis. The flow through from the second SUMO immunoisolation with the 8A2 antibody, which has not been digested with Asp-N, can be further processed for ubiquitin site identification/quantification. The UbiSite based purification can be performed on this flow through and the enriched peptides further digested with trypsin to release the diglycine motif on the ubiquitinated peptides, which is readily identifiable by LC–MS/MS.

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Figure 5.6  Proposed method for endogenous identification of proteins that are co-modified with SUMO3 and ubiquitin.

Although the study of protein SUMOylation and ubiquitination (and their crosstalk) have been the primary focus in the field of UBL proteomics, the study of other UBL are likely to follow suit. With the advent of the UbiSite methodology, similar methods could be used to monitor NEDD8ylation, ISG15ylation and FUBIylation with the production of appropriate antibodies (Fig. 5.7). The above mentioned UBLs all provide a short remnant on the modified lysine residue when digested with trypsin, thus facilitating their

subsequent identification and quantification by LC–MS/MS. As with the UbiSite method, the work flow starts with a digestion using Lys-C, creating large peptides (shown in red in Fig. 5.7) that can serve as epitopes for the peptide level immunoprecipitation once the proper antibody has been commercialized. The enriched peptides are then further digested with trypsin to expose the smaller remnant (shown in blue in Fig. 5.7) on the modified lysine residues (diglycine for NEDD8 and ISDG15; MLGG for FUBI).

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Figure 5.7  Endogenous amino acid sequence of selected UBLs. Red residues depict the remnant that remains on the UBL modified lysine on Lys-C digestion that can serve as an epitope for selective UBL modified peptide enrichment. Blue residues depict the remnant that is produced on the UBL modified lysine residue after trypsin digestion, allowing for easy analysis by LC–MS/MS. Filled arrows depict the most C-terminal cleavage site for Lys-C. Dashed arrows depict the most C-terminal cleavage site for trypsin.

Acknowledgements This work was carried out with financial support from the Natural Sciences and Engineering Research Council (NSERC 311598). IRIC proteomics facility is a Genomics Technology platform funded in part by the Canadian Government through Genome Canada, the Canadian Center of Excellence in Commercialization and Research, and the Canadian Foundation for Innovation. References

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Global Proteomic Profiling of SUMO and Ubiquitin Alla Ahmad, Ryan Lumpkin and Elizabeth A. Komives*

6

Department of Chemistry and Biochemistry, University of California, San Diego, CA, USA. *Correspondence: [email protected] https://doi.org/10.21775/9781912530120.06

Abstract In this chapter, we introduce the small ubiquitinlike modifier (SUMO) and describe how it is attached to target proteins. What is known about how SUMO attachment changes the function of a target protein is briefly reviewed. The main focus of the chapter is to introduce the various methods for identifying SUMO attachment sites on target proteins using proteomics and mass spectrometry. Finally, we present a future outlook for one newly discovered function of SUMO that will be quite interesting to study using the new proteomics approaches. The function of SUMO The Small Ubiquitin-like Modifier (SUMO) proteins are encoded in four genes and divided into three types: SUMO1, SUMO2/3, and SUMO4. Reversible attachment of the C-terminal carboxyl of SUMO proteins to a target protein’s free ε-amine of a lysine residue through a covalent isopeptidebond (Makhnevych et al., 2009) ‘tags’ a target protein with SUMO. This conjugation is enacted by a SUMOylation cascade that, similar to ubiquitin, involves an E1 activating enzyme, E2 conjugating enzyme, and E3 protein ligase (Capili and Lima, 2007). SUMO regulates target proteins by causing changes in their protein activity, intracellular localization, stability, and interaction partners (Fig. 6.1). Some of the processes that SUMO proteins regulate include the cell cycle (Azuma et al., 2001), heat

shock (Golebiowski et al., 2009), DNA damage (Morris, 2010), and phosphorylation (Uzoma et al., 2018). For example, in the case of base excision repair, SUMOylated TDG, Thymine DNA Glycosylase, loses its affinity for DNA and catalytic capabilities due to a conformational change that happens at the N-terminus on the conjugation of SUMO (Morris, 2010). Because SUMO is capable of functionally influencing a variety of downstream processes, viruses attempt to hijack this pathway to effectively control the cell (Wimmer et al., 2012). Adenovirus is said to use its E1B-55K and E4-ORF3 proteins to modulate SUMO activity in order to facilitate viral production (Higginbotham and O’Shea, 2015). In addition, the SUMOylation pathway itself is also regulated by various factors at the transcriptional, translational, and degradation levels of different components of the SUMO pathway (Gareau and Lima, 2010). The intricacies of the SUMOylation pathway makes it a very interesting cascade to study. Because SUMO contributes to and is influenced by a multitude of cellular functions, methods to determine SUMO targets have been an important focus in SUMO research. Identification of SUMOylated proteins The presence of SUMO conjugated to proteins is readily detected by immunochemistry taking advantage of commercially available antibodies specific to each SUMO form (Lim et al., 2014; Li et al., 2018). SUMOylated proteins are routinely

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Figure 6.1  Schematic showing the functional consequences and cellular localization of SUMOylated proteins.

imaged in situ and purified by anti-SUMO immunoprecipitation. The specificity of anti-SUMO antibodies makes this an attractive method for determining which SUMO isoform is attached to a particular target. SUMOylated proteins are difficult to identify from complex mixtures by western blotting due to the fact that the amount of the SUMO modification at steady state is low (Tammsalu et al., 2014). Western blotting does detect the SUMO modification from partially-purified samples (Hilgarth and Sarge, 2005). Until recently, identification of all proteins in a cell bearing the SUMO modification (the SUMOylation proteome) was perceived as a great challenge due to the low steady-state levels of SUMO modification, the presence of SUMO proteases in the cell, and the fact that tryptic digestion results in a large C-terminal remnant of SUMO being left at the site of modification creating a ‘cross-linked’ peptide that was difficult to identify by mass spectrometry (Eifler and Vertegaal, 2015). Despite these challenges, both exogenous and endogenous proteomic methods have been discovered to identify SUMO target proteins (Eifler and Vertegaal, 2015). An exogenous method involves ectopic expression of epitope-tagged SUMO allowing SUMO to be isolated and analysed by mass spectrometry (Vertegaal et al., 2004). To overcome the problem of the large

C-terminal remnant of SUMO being left at the site of modification, a mutated SUMO with an additional cleavage site near the C-terminus was introduced (Tammsalu et al., 2014). This method effectively circumvents the large C-terminus tryptic fragment issue. These exogenous approaches are capable of producing SUMO-interaction maps, but the introduction of exogenously modified SUMO may disrupt normal pathways of SUMO addition and removal and could lead to higher SUMOylation levels of target proteins as compared to endogenous methods. Furthermore, these exogenous methods are restricted to specific cell types and organisms. Endogenous methods typically use antibodies to that recognize SUMO2/3 or SUMO1 to purify SUMOylated proteins and to identify SUMO interacting proteins (Makhnevych et al., 2009; Becker et al., 2013). These approaches create interaction maps between SUMO and various proteins it is associated with, but do not allow for identification of the SUMO modification sites. The previously stated methods have been effective in identifying potential SUMO target proteins and SUMOylation sites, however a native method to identify large numbers of SUMOylation sites is preferred to extend experiments to a larger variety of cell types and conditions. Global SUMO profiling approaches Global ubiquitin-modification profiling was revolutionized by the development of a proteomic workflow that makes use of the -RGG sequence at ubiquitin’s C-terminus (Kim et al., 2011; Carrano and Bennett, 2013) (Fig. 6.2a). Ubiquitin, like SUMO, is attached via its C-terminal carboxyl group to the ε-amine of a lysine residue in the target protein. Trypsin will cleave after the R leaving a diglycyl-lysine remnant at the ubiquitination site, which can then be enriched using antibodies specific for the diglycyl-lysine, and identified by mass spectrometry (Kim et al., 2011; Carrano and Bennett, 2013) (Fig. 6.2b). As was mentioned before, SUMO does not have this convenient arginine residue close to the C-terminus, instead it has a threonine in place of the arginine (Fig. 6.2a). We discovered that wild type α-lytic protease, WaLP, has a preference for cleaving after threonine, leading to SUMO-specific cleavage to generate the same

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Figure 6.2  (a) Sequence alignment of ubiquitin and SUMO isoforms showing the C-terminal sequence that becomes attached to the target protein. (b) Schematic of the proteomic workflow for identification of ubiquitination sites in target proteins. (c) Schematic of the proteomic workflow for global identification of SUMOylation sites in target proteins that can be done in parallel with the workflow for identification of ubiquitination. (d) Schematic of the latest proteomic workflow for global identification of SUMOylation sites in target proteins.

diglycyl-lysine. WaLP generates peptides of the same average length as trypsin despite its relaxed substrate specificity (Meyer et al., 2014). Owing to the TGG C-terminal sequence of SUMO, WaLP was used to digest cell lysates to leave a SUMOremnant diglycyl-lysine (KGG) that was identical to that resulting from a trypsin digest of ubiquitinated proteins (Lumpkin et al., 2017) (Fig. 6.2c). That meant that the same workflow used to identify ubiquitin modification sites could be used to identify SUMO modification sites with just a change in whether the cell lysate was digested with trypsin or WaLP. A caveat of the method is that WaLP produces non-tryptic peptides requiring careful MSMS sequencing protocols such as MS-GF+ that do not bias against non-tryptic peptide fragment profiles (Kim and Pevzner, 2014). This method for identification is not only advantageous in that it can determine SUMO sites under completely native conditions, but it can also allow a sample to be split and subjected to parallel analysis of both Ub and SUMO modified proteomes. This method can be used to identify both Ub and SUMO sites in any sample and under any growth conditions. This method resulted in the discovery of 826 novel SUMO attachment sites. Of the 1209 unique

sites identified, only about 30% overlapped with the SUMOylation sites of Hendriks et al (Hendriks and Vertegaal, 2016). These sites were validated by the reduction of KGG-modified peptides when cells were treated with SENP1/2 which result in a reduction of SUMO1 and SUMO2/3 modified proteins. It was observed that 88% of sites identified were decreased on SENP1/2 treatment validating the method used. Conversely, the use of a deubiquitinating enzyme Usp2cc only resulted in a less than 2% decrease of KGG peptides produced. This shows that the sites identified are verified to be SUMOylation sites and not ubiquitination sites. A new approach to identify SUMOylation sites in endogenous conditions has been described by Hendriks et al. (2018) (Fig. 6.2d). In this paper, endogenous SUMOylation sites were identified by digestion of cell lysate with LysC. This was done to cleave proteins into peptides while retaining the fragment of SUMO2/3 from K45 to the C-terminus attached to the target protein peptide. By using the SUMO2/3 8A2 antibody which recognizes the SUMO epitope containing residues 57-IRFRFDGQPI-66 epitope, SUMOylated peptides were enriched. Subsequent digestion of the enriched peptides with Asp-N leaves the remnant

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85-DVFQQQ TGG-93 on a lysine of the target protein peptide. Using this method, 14,869 unique endogenous SUMO2/3 sites mapped onto 3870 SUMOylated proteins were identified. A positive aspect of this most recent method is the capability to distinguish the SUMO2/3 sites separately from the SUMO1 sites. However, their method does not propose a protocol to identify SUMO1 sites, which is essentially leaving out part of the SUMOylation story during proteomic analysis. In addition, it does not allow for a direct, side-by-side comparison between SUMOylation and ubiquitination, which in the former study illuminated the interplay between these two lysine modification events. It will be interesting to see whether creative combinations of the two methods could be devised that would provide even more complete proteomic information in the future. New discoveries of SUMO functions in biology Regardless of the identification method for SUMOylation sites, the new sites identified by current SUMOylation studies will help illuminate additional roles that SUMO could be playing in cells. One such role that SUMOylation may be playing is in the formation of membrane-less organelles in the cell. By fluorescently labelling a constitutive P-granule protein, Brangwynne and colleagues (2009) discovered that P-granules display liquid-like properties. The promyelocytic leukaemia tumour suppressor protein (PML) is highly SUMOylated and involved in the formation of nuclear bodies (Lallemand-Breitenbach and de Thé, 2010). Rosen’s group demonstrated the formation of membrane-less organelles in vitro by mixing a protein containing ten repeats of human SUMO3 (polySUMO) and a protein with ten repeats of the SUMO Interaction Motif (SIM) from PIASx (polySIM) (Banani et al., 2016). By adjusting the amount of each protein, the switch-like formation of the organelles could be demonstrated. These authors further demonstrated the requirement for PML SUMOylation for formation of membraneless organelles in cells. Another membraneless organelle in which SUMO seems to play a role is the stress granule. Stress granules are structures typically formed

during times of cellular stress, such as in response to arsenite or ionizing radiation, in order to sort mRNAs for storage, degradation, or translation so as to conserve energy during that stressor (Spriggs et al., 2010). Modification levels of eIF4A2 by SUMOylation was increased during times of arsenite and ionizing radiation stress and the SUMO modification was used to localize the eIF4A2 to the stress granules ( Jongjitwimol et al., 2016). Indeed, when a construct of eIF4A2 that does not contain the lysine for SUMOylation was expressed in cells, both the size and amount of stress granules was significantly decreased ( Jongjitwimol et al., 2016). This therefore shows that SUMOylation of proteins plays a role in causing the formation of stress granules. In addition, DDX6 is a human RNA helicase that has been linked to functioning with stress granules (Bish et al., 2015). It was determined that DDX6 and many of its binding partners, including TIF1β which is an E3 SUMO ligase, are either SUMOylated or have a higher than average amount of SUMOylation motifs than the general proteome (Bish et al., 2015). For this reason, it was hypothesized that SUMOylation may play a role in the prevention of aggregation for cytoplasmic granules because SUMO has been shown to decrease aggregation (Krumova et al., 2011) and the localization to P bodies and stress granules depend on aggregation-prone domains (Bish et al., 2015). Overall, it seems quite evident that SUMO plays a significant role in the formation of stress granules, whether by the localization of certain proteins to these structures or by stabilizing these structures. In the future, it will be interesting to utilize the proteomic approaches described here to obtain quantitative information about how P-bodies and stress granules form by measuring which sites on which proteins are SUMOylated over time during membrane-less organelle formation in cells. References

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Biotin-based Approaches for the Study of Ubiquitin and Ubiquitin-like Protein Modifications

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James D. Sutherland*, Orhi Barroso-Gomila and Rosa Barrio*

CIC bioGUNE, Bizkaia Technology Park, Bizkaia, Spain. *Correspondence: [email protected] and [email protected] https://doi.org/10.21775/9781912530120.07

Abstract Since its discovery, the high-affinity interaction between biotin and avidin has formed the basis of numerous tools and techniques in molecular biology and biochemistry. The post-translational modifications of cellular proteins via conjugation of ubiquitin (Ub) and ubiquitin-like (UbL) proteins contribute to crucial cellular processes, such as protein homeostasis and the DNA damage response, yet they are challenging to study due to their dynamic nature, scarcity, and sensitivity to removal by proteases. Understanding how the Ub/ UbL-modified proteome changes during development or in response to environmental insults or pathological conditions may yield new biomarkers or identify new drug targets. The use of biotinbased technologies for Ub/UbL studies is relatively new but has already contributed to the identification of new substrates and promises much more. In this review, we focus on two separate approaches: biotin-tagged Ub/UbLs to modify and capture target proteins in vivo, and BioID, a tool to facilitate the labelling and identification of proximal interactors of Ub/UbL enzymes. Coupled with ongoing advances in proteomics to increase sensitivity in peptide identification, and gene-editing techniques to avoid overexpression artefacts, biotin-based systems promise to reveal new information about the role of Ub/UbL modifications in development and disease.

Exploiting the biotin–streptavidin interaction for biotechnology Biotin is an essential vitamin and coenzyme that is required for all forms of life (Chapman-Smith and Cronan, 1999). From bacteria to mammals, biotin is added as a post-translational modification (PTM) to biotin-dependent carboxylases, where it facilitates the transfer of carboxyl groups between metabolites and mediates gluconeogenesis, catabolism of select amino acids, and energy transduction (Tong, 2013). The biotin molecule has an unusual dual-ring structure and requires both ATP and a biotin protein ligase (BPL) enzyme, such as the bifunctional ligase/repressor BirA, for carboxylase modification (Fig. 7.1A). Briefly, structural studies have revealed that biotin and ATP are likely recruited to a conserved pocket in the BPL, which undergoes conformational changes and traps an activated intermediate, biotinyl-5′-AMP (Wilson et al., 1992; Sternicki et al., 2017). The target carboxylases contain a conserved domain, called BCCD (biotin carboxyl carrier domain) or more generically BAP (biotin acceptor peptide). This can interact with the BPL holoenzyme, allowing it to be precisely positioned to allow biotin transfer to a specific lysine, with the release of AMP. Once modified, the carboxylases can execute a number of essential metabolic reactions. Like most PTMs, biotin can be removed. Biotinidases both liberate biotin from dietary sources for cellular needs and

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Figure 7.1  BirA-mediated biotinylation and the Ub modification cycle. (A) Biotin and ATP are recruited by the BirA biotin ligase into a conserved binding site, where biotin is converted to the reactive biotinoyl-5′-AMP. When a substrate is encountered and the BAP (Biotin Acceptor Peptide) is properly positioned, biotin is then transferred to a specific BAP lysine, with release of AMP. In the text, BAP is used interchangeably with the prefix ‘bio’, e.g. bioUB. Also mentioned is AviTag, which is an optimized BAP. (B) A schematic of the ubiquitination cycle. Similar cycles are followed for UbLs, with each type having their own unique E1/E2/E3 enzymes. Ub forms a thioester bond with an E1 activating enzyme in an ATP-dependent manner. This E1-Ub then passes Ub to an E2 conjugating enzyme, also forming a thioester bond. Among Ub and UbLs, there are many variations in the next steps, but a frequent outcome is depicted here: the Ub-E2 encounters an E3 ligase which can recognize both the Ub-E2 and the substrate, leading to substrate ubiquitination. Ub and the substrate can be recycled by the action of a DUB or other specific UbL peptidase. DUB, deubiquitinase; UB, ubiquitin.

recycle biotin from carboxylases (Hymes and Wolf, 1996). This exquisite specificity of a PTM for a specific peptide substrate has attracted the attention of cell and molecular biologists as a way to specifically label biomolecules with biotin, both in vitro and in vivo. Most studies have focused on the E. coli BPL BirA and its tightly controlled biotinylation of BCCP (biotin carboxyl carrier protein; 156 amino acids). BCCP is a subunit of acetyl-CoA-carboxylase (Knowles, 1989) and requires biotinylation for activity. BirA catalyses attachment of biotin to lysine within a conserved motif (AMKM) found in BCCP. A fragment of ≈ 75 residues encompassing the AMKM motif can be fused to a protein of interest to confer biotin labelling, in the presence of BirA, biotin, and ATP (Cronan, 1990). In an attempt to both understand the spatial and sequence requirements for BirA-mediated biotinylation and define a shorter BAP to use for fusion purposes, screenings led to identification and optimization of the AviTag, a 14 residue tag that can be used as

an N- or C-terminal fusion and can be biotinylated as efficiently as BCCP itself [GLNDIFEAQKIEW (Schatz, 1993; Beckett et al., 1999)]. This tag has been widely used with numerous proteins in biotechnological applications ranging from bacterial protein expression and purification to advanced applications in subcellular labelling and enrichment in mammalian cells and transgenic animals (Fairhead and Howarth, 2015). Why is a biotin label so interesting? Because avidin, a protein highly enriched in egg whites, and bacterially-derived streptavidin exhibit an extraordinary affinity for biotin (Kd 10–11 to 10–15, depending on the avidin variant), making this the strongest non-covalent interaction yet discovered (Green, 1975). The biotin–streptavidin interaction can withstand conditions that cause most other proteins to become denatured (8M urea, 6M guanidinium hydrochloride, 1% sodium dodecyl sulfate, heat), allowing tight and rapid binding of biotin-conjugates with streptavidin supports. This can be followed by stringent washing, which serves

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to remove proteins that interact non-specifically with the support matrix and proteins that may be indirect partners to biotin-labelled targets. On the other hand, this strong interaction has a drawback in that biotin conjugates bound to streptavidin matrices can be difficult to elute for downstream analysis. Harsh conditions are usually used [high heat, 2% SDS, reducing conditions (Rosli et al., 2008). Alternate protocols have been developed (Holmberg et al., 2005; Cheah and Yamada, 2017) but have not been extensively validated. Also, for purposes of peptide-based mass spectrometry (MS), direct digest using trypsin or other proteases of the biotin conjugates bound to the streptavidin support has been used to release peptides for analysis (Hesketh et al., 2017). Requiring optimization, this strategy also releases abundant streptavidin-derived peptides, and from bound proteins, primarily interstitial peptides that do not contain a biotin, as biotin-containing peptides can remain bound to any intact streptavidin. No comprehensive study has been published to compare and contrast these ‘release’ strategies and how they influence representation in final data sets. Work on streptavidin mutants and use of biotin-like molecules (e.g. desthiobiotin) may yield combinations that both bind tightly and elute more easily (Laitinen et al., 2006; Dundas et al., 2013; Lim et al., 2013; Lu et al., 2014). An emerging application for biotin– streptavidin pairing is imaging, especially super-resolution microscopy. Site-specific biotinylation of the AviTag-fusion by BirA minimizes perturbation to localization dynamics. The biotin label can be detected by fluorescent streptavidin. In cases where streptavidin size and its potential for aggregation (due to multiple biotin-binding sites in a tetrameric conformation) can affect results, the compact monomeric streptavidin (mSA; 12 kDa) can be used (Chamma et al., 2016). Recombinant mSA labelled with bright organic dyes can be used to detect biotin in fixed samples, and fusions of mSA to fluorescent proteins can be expressed in vivo to follow the dynamics of biotin-labelled proteins. A drawback to the use of biotin interactions in microscopy in low expression or low-light scenarios lies in the presence of endogenous biotinylated carboxylases, localized mostly at the mitochondria, that can give background labelling, so its use might be restricted to situations of medium-high

expression or analysis of other organelles or plasma membrane proteins. The success of biotin–streptavidin for biotechnological purposes can be attributed to many aspects, foremost being high affinity and programmability. Ease of use and wide availability of reagents has driven the development of novel applications for working with DNA, RNA, and proteins. Here, we will cover several ways in which the biotin-streptavidin and BirA-based systems have been applied to Ub/UbL studies. The complexity of the ubiquitin code Like biotinylation, the addition of Ub or UbL to a target protein can change its function or fate (Flotho and Melchior, 2013; Ciechanover, 2015). Inherent in its name, ubiquitination occurs in all eukaryotic cells, with the Ub molecule itself being highly conserved, showing 100% identity between yeast and human isoforms. Ub modification is most commonly associated with protein degradation but plays a role in practically all biological processes. Beyond Ub, there is an extended family of Ub-like proteins (UbLs) that share some sequence homology, and even higher structural homology with Ub (van der Veen and Ploegh, 2012). SUMO is perhaps the best-characterized UbL, and SUMOylation of targets often leads to changes in protein localization or creates a scaffold to recruit and assemble higherorder molecular complexes. NEDD8, UFM1, FAT10, ISG15 are other less-studied UbLs. Most cellular PTMs consist of adding a small molecular moiety, such as a phospho- or methylgroup, and are catalysed by single enzymes, such as a kinase or methylase. By contrast, ubiquitination and SUMOylation occur through a multi-enzyme cascade and consist of addition of a whole protein onto a target substrate. In general, Ub/UbLs share a similar biology in the way they are processed, added to substrates, and recycled (for schematic, see Fig. 7.1B). Ub and the distinct UbLs each tend to have a dedicated set of enzymes, with ample opportunity for regulation, feedback, and some crosstalk (Hershko and Ciechanover, 1998). They begin as precursor proteins, with short C-terminal extensions that must be cleaved by proteases. The abundant Ub is encoded by several genes in humans, either as fusions to ribosomal proteins (e.g.

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RPS27A, UBA52) or as polyubiquitin (e.g. UBB with three repeats, or UBC with nine repeats). Ub proteases, also known as deubiquitinases or DUBs, act on the polyubiquitin or the fusions to release Ub monomers. The UbL precursors are cleaved in a similar way, each with a distinct protease. The Ub/UbL is then passed to an ATP-dependent E1 activating enzyme, where it becomes coupled to the E1 by a thioester linkage between a C-terminal glycine residue in the Ub/UbL and a cysteine within the E1. Next, the Ub/UbL-charged E1 interacts with an E2 conjugating enzyme, and the Ub/UbL is passed from E1 to E2, forming a new thioester linkage. In some cases, the charged Ub/ UbL-E2 can interact with substrates and conjugate the Ub/UbL directly to them, but more commonly an additional step is required. E3 ligases can interact with both the charged Ub/UbL-E2 and specific substrates, to facilitate the transfer of the Ub/UbL to form an isopeptide bond with a substrate lysine. Addition of the Ub/UbL confers new properties to the target, including changes in conformation and activity, shifted subcellular localization, and creating or obscuring binding sites for interacting proteins. Eventually, the Ub/UbL can be removed and recycled by DUBs. Beyond the relative simplicity of the E1/E2/ E3 cycle lies a stunningly complex landscape populated by proteins modified by Ub/UbLs. To survey the numbers of implicated proteins found in human cells, there are relatively few E1 enzymes, up to 40–50 E2 enzymes, 100s of E3 ligases, and 1000s of substrates among the cellular proteins (Clague et al., 2015). Charged E2s can interface with a subset of E3 ligases, and E3s bring Ub/UbL modifications to a subset of targets. Specific recognition of substrates by E3s is likely accomplished through structural features, since conserved sequence features are often lacking. Aside from the addition of a single Ub/UbL to a target, multiple sites on the same target might be modified, called multi-monoubiquitination in the case of Ub. Ub/UbLs are added primarily to lysines, and since Ub/UbLs themselves have multiple conserved lysines that can be modified, formation of Ub/UbL chains greatly adds to the complexity (Komander and Rape, 2012). Best described for Ub, different types and lengths of Ub chains can dictate different fates for the substrate. Extension of Ub chains using K48 or K11 serves as

a signal for proteasome-mediated protein degradation, whereas K63 chains can trigger recruitment of substrates to sites of DNA damage. Head-to-tail concatemers known as linear Ub chains are crucial for immune responses (Rittinger and Ikeda, 2017). Chains with mixed linkage types or composed of mixtures of Ub and UbLs are infrequent but detectable by sensitive proteomic techniques. Ub/ UbLs can be phosphorylated or acetylated, leading to unique properties (Herhaus and Dikic, 2015; Gartner et al., 2018). Modifications by Ub/UbLs can serve to assemble larger complexes, with new topologies created by the PTM being recognized by short interaction motifs and domains contained within recruited proteins (Husnjak and Dikic, 2012). Just as E3 ligases have preferences for certain substrates or specialize in building Ub chains with specific linkages, the peptidases that remove and recycle the Ub/UbLs also exhibit specificities and preferences. Only a small fraction of the total pool of a particular substrate might be Ub/UbL-modified at a given time, but enough to lead to biological outputs. Due to the scarcity of these modifications and their dynamic nature, changing rapidly in response to nutrients, environmental insults, cell cycle or developmental stages, they can be challenging to study. Numerous methods have been designed with the Ub/UbL landscape in mind, several of which are based on the biotin-streptavidin system. Fishing for ubiquitomes using bioUb and BirA Small protein epitope tags can be fused to the N-terminus of Ub/UbLs and be used as molecular handles to follow expression and capture modified substrates. DNAs encoding these fusions can be introduced into cells or transgenic organisms. Tagged Ub/UbL conjugates can then be purified by affinity resins. Because a large number of proteins contain motifs and domains that interact non-covalently with Ub/UbL-modified proteins, stringent washing conditions ensure that these ‘passengers’ are released and that only true covalent conjugates are retained. Also, capture of some epitope tags requires mild lysis and binding conditions, which can allow DUBs and other peptidases to act on conjugates, so tags that can withstand stronger denaturing conditions are

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preferred. A popular tag is the poly-histidine tag (HIS), which is compact and binds well to metal affinity matrices under more stringent conditions. HIS-tagged versions of Ub/UbLs have been used to isolate conjugates from cells and from transgenic animals for peptide-based MS identification (Peng et al., 2003; Tirard et al., 2012; Akimov et al., 2014; Tammsalu et al., 2014). Some contamination can arise from proteins with internal poly-HIS stretches or non-covalent binders that survive the binding/ washing conditions, but this approach has generated a wealth of useful data. After the biotinylation pathway was characterized, and more specifically that bacterial BirA could catalyse the transfer of biotin to a specific peptide substrate, it was shown that the system could be used to label and purify recombinant proteins in bacteria (Cronan, 1990) and even transplanted to mammalian cells for in vivo site-specific protein labelling (de Boer et al., 2003). In this latter study, BirA alone or in combination with the GATA-1 transcription factor, N-terminally tagged with a 23-residue biotinylation tag, were stably expressed in mouse erythroleukemic cells. Using mild lysis and washing conditions, streptavidin pulldowns from nuclear extracts showed an efficient one-step purification for biotinylated GATA-1 and potential binding partners. Using BirA alone revealed background binding of the expected endogenously biotinylated carboxylases, and possible contaminants, such as abundant splicing factors and ribosomal proteins. Biotinylated GATA-1 was also used in a variant of the chromatin immunoprecipitation assay (ChIP), and even in transgenic mice. Characterization of the biotinylated GATA-1 pulldowns by MS revealed association of GATA-1 with both known and novel transcriptional regulatory complexes (Rodriguez et al., 2005). A similar strategy has been used for other transcription factors (Goardon et al., 2006; Meier et al., 2006; Rudra et al., 2012), secreted proteins (Predonzani et al., 2008), ribonucleoprotein complexes (Penalva and Keene, 2004), and for biotinylating virions for gene therapy use (Lesch et al., 2010). Stepwise improvements to in vivo biotinylation strategies have improved its efficiency and driven its wider adaptation. These include the aforementioned identification of the AviTag, the development of vectors for co-expressing biotinylation targets and BirA using internal ribosome entry sites [IRES

(Kulman et al., 2007)] or viral 2A ribosome skipping sequences (Pirone et al., 2017), and using a codon optimization strategy to generate an improved version of BirA for use in mammalian cells (Mechold et al., 2005). Another perhaps unexpected advance has been Addgene (Kamens, 2015), a non-profit plasmid repository that provides rapid access to biotinylation tools for the researchers, which in turn drives faster innovation. Recognizing that the biotin-streptavidin linkage could withstand very stringent binding and washing conditions, and that the necessary components could be genetically encoded, the system was adopted for studying ubiquitination. The fruit fly Drosophila melanogaster is a widely used, genetically tractable model organism for research into developmental processes, including neurogenesis and neurodegeneration. Mutations in certain Ub E3 ligases and deubiquitinases had been linked to neurogenerative phenotypes ( Jaiswal et al., 2012), but information regarding Ub substrates in the nervous system was lacking. At the time, most large-scale efforts to identify the whole ‘ubiquitome’ were carried out in human and mouse tumour cell lines, not primary cells or tissues, and especially not from flies. The biotinylated-Ub (or bioUb) approach was developed to allow tissue-specific expression of bioUb and BirA in the developing nervous tissue of fly embryos (Franco et al., 2011). In the bioUb flies, six repeats of N-terminally AviTag-Ub and E. coli BirA are expressed as a single polypeptide, which is subsequently cleaved by endogenous DUBs. This design mimics the processing of endogenous polyubiquitin. BirA catalyses the in vivo biotinylation of the bioUb, either before or after conjugation to substrates. Taking advantage of the flexible molecular genetic toolbox available for D. melanogaster, the bioUb/BirA protein was expressed in embryonic nervous tissues by virtue of the UAS-GAL4 system, in which a neuralexpressed GAL4 (elav-GAL4) drives expression of bioUb/BirA under the control of 5x upstream activation sites (UAS). The result was tissue-specific biotinylation of ubiquitinated proteins, that could later be captured on streptavidin beads, extensively washed with very stringent conditions (urea, guanidine, alcohols, SDS) to remove contaminants, and processed for MS. A generalized scheme is presented in Fig. 7.2, depicting the AviTag-Ub or bioUb as the more general BAP-UbL, since the

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Figure 7.2 The bioUbL system. Ub or UbLs are expressed in cells as fusions to a BAP, where they can enter the Ub/UbL cycle and become incorporated into substrates, most commonly via lysines. BirA mediates the biotinylation of the BAP-UbL, either on substrates (as depicted here) or before/during the Ub/UbL cycle. Cells are lysed and capture is performed in denaturing conditions for maximum solubility. Owing to the high affinity of the biotin– streptavidin interaction, stringent washes remove any non-biotinylated passenger proteins, and Ub/ UbL-modified proteins are identified by MS. BAP, biotin acceptor peptide; MS, mass spectrometry; UbL, ubiquitin like.

same scheme can be applied to Ub and other UbLs (discussed in sections below). The resulting peptide identifications revealed the efficiency of the bioUb method. As expected, Ub peptides were seen, as well as some peptides showing diglycine-containing branchpoints, consistent with different Ub-Ub linkages (K48, K63, K6, K11). Eleven distinct Ub carrier proteins (E1, multiple E2s, a HECT-type E3) were found. Peptides of SUMO were also found, perhaps present as a potential co-modifier of ubiquitinated proteins or a component of Ub-SUMO mixed chains. Of the additional 48 proteins identified, 18 were already known to have roles in synaptogenesis. A number of the substrates were validated by western blotting and existed as mono- or poly-ubiquitinated species in the streptavidin pulldowns. Importantly, many of these candidates were further validated in a followup study using an independent system to evaluate ubiquitination, specifically that candidates were coexpressed as GFP-fusions with Flag-Ub in a neural cell line from flies (Lee et al., 2014). Using the bioUb method, full-length Ubmodified proteins are captured and aliquots can be saved for western validation. Another popular proteome-scale method for identifying Ub-modified proteins relies on antibodies that recognize the branchpoint where the Ub/UbL diglycine (diGly) and substrate lysine are joined (Xu et al., 2010; Kim et al., 2011). This method cannot distinguish between the diGly remnants of Ub and the UbL proteins NEDD8 and ISG15. More recently, a novel antibody (called UbiSite) that specifically recognizes the C-terminus of Ub has been reported (Akimov et al., 2018). In both cases, after digest with trypsin or Lys-C proteases, peptides displaying these epitopes can be immunoprecipitated and identified by MS. While this is a great advantage for identifying the precise site of modification, with the diGly and UbiSite approach it is impossible to know whether the original substrate was monoubiquitinated or carried multiple Ubs (in chains or multi-monoubiquitinated). Western blotting after bioUb pulldowns can provide supporting evidence for this distinction. Identification of the ubiquitinated proteome in a given tissue at a given developmental timepoint reveals a snapshot, but the method gains power when used in comparative situations. By comparing bioUb datasets from embryonic versus adult

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neurons (Ramirez et al., 2015), a certain number of Ub conjugates, including some E1/E2/E3 carrier enzymes, were overlapping, suggesting that those modifications are found in most cells, or even most neural cells. However, a larger number of bioUb-modified proteins differed between the embryonic and adult neural tissues, perhaps linked to differentiation and specialization. In a separate study (Ramirez et al., 2018), the bioUb system was used to identify potential substrates for the E3 ligase UBE3A, the function of which is lacking in the neurodevelopmental disorder Angelman Syndrome. Positing that increased UBE3A levels/ activity would lead to increased ubiquitination of bona fide substrates, bioUb conjugates were analysed from flies overexpressing or not UBE3A. Almost 80 candidates were identified, with an enrichment in proteasomal subunits or regulators, such as the protein Rngo/DDI1. A focus on proteasomal dysregulation may lead to new ideas for understanding and treating Angelman Syndrome. A similar approach was used to identify candidate substrates for Parkin (Martinez et al., 2017), an E3 ligase which primarily acts at the mitochondria and is linked to familial Parkinson’s disease. Increased expression of Parkin in fly neurons led to increased ubiquitination of some mitochondrial targets (expected) and regulators of endosome trafficking (unexpected), suggesting that Parkin may have an expanded role in neural function. In all cases, the findings reveal Ub-modified candidates in a physiological setting and may provide new clues for understanding and exploiting the Ub system in development and disease. More than just flies: bioUb applications in mammalian systems After extensive validation in Drosophila, the bioUb system was adapted for use in mammalian contexts. In human cells, it has been applied to a key stage in the cell cycle, the mitotic exit (Min et al., 2014). Targeted proteolysis of mitotic cyclins via polyubiquitination and proteasomal degradation drives the transition from metaphase to anaphase. In this step, the majority of Ub conjugation is catalysed by the anaphase-promoting complex/cyclosome (APC/C), a large multi-subunit E3 ligase complex (Pines, 2011). Some substrates had been identified previously, but the full spectrum of Ub conjugates promised to reveal additional regulators. Using

synchronized U2OS cells, stably expressing an inducible form of bioUb and BirA, conjugates at metaphase and anaphase were isolated and compared. A large number of mitotic-exit-specific Ub conjugates were identified, many of them novel. Of note, during validation of selected candidates by western blotting, a dramatic switch from mono- to poly-ubiquitination was observed for the Aurora A kinase in the interval between 30 and 70 minutes after release from metaphase block. This switch would likely have been missed using the diGly antibody-based Ub remnant approach. While bioUb is able to conjugate efficiently to most substrates and can participate in Ub chain linkages through any of its lysines, there is one scenario where it is problematic: linear Ub chains. The covalent joining of one Ub through its C-terminal glycine to the N-terminal methionine of an existing Ub forms linear Ub chains. Since bioUb has an AviTag at the N-terminus, this blocks linear extension. If bioUb incorporates into an existing linear chain, it would likely serve as a terminator or capper. Linear Ub chains are particularly important for control of inflammation and the NFκB signalling pathway (Iwai et al., 2014). Finding more substrates for this chain type might reveal key inflammation regulators or give clues towards other pathways in which it is involved. This problem was addressed by placing a tag internally within the Ub sequence, to be located in a region of the protein that is accessible and rather inert, e.g. no lysine modification sites, and not recognized by Ub-binding proteins (Kliza et al., 2017). All seven internal lysines of Ub were also mutated, so that only the N-terminal methionine remains as a target for chain elongation. A short peptide called Strep-Tag-II was used instead of AviTag, which binds with high affinity to an engineered streptavidin called Strep-Tactin that does not require BirA. After extensive validation that the internally-tagged Ub was functional, it was used to enrich linear ubiquitinated substrates for MS identification. Among the candidates, the E3 ligase TRAF6 was further characterized and found to be essential for interleukin-induced NFκB signalling. Transgenic mice with ubiquitous expression of the original bioUb system (with N-terminal AviTag) have been described (Lectez et al., 2014), with validated isolation of biotinylated Ub conjugates from various organs (muscle, brain, heart, pancreas, and liver). An extensive analysis of the liver ubiquitome

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revealed peptides representing almost 400 proteins, which included many novel substrates. Ub carriers, coupled to Ub by a thioester bond (E1/E2/E3), were present. A protocol was described to allow elution of these carriers using the reducing agent DTT. Some unexpected proteins also eluted with DTT, namely a number of peroxisome-associated proteins (e.g. PEX5, ABCD3, PRDX1) that are likely ubiquitinated on cysteines rather than the more common lysine (Francisco et al., 2014). Whether a specialized machinery is needed for this variation, and moreover, how it affects function of these targets, is unknown. The bioUb mice provide a platform for analysing the ubiquitome of multiple organs in response to genetic perturbations, drug treatments, or tumorigenesis, and can serve as a source for primary cells of different types to facilitate Ub study in culture. Beyond ubiquitin: expanding the bioUbL toolbox Although ubiquitination is the most abundant and best-studied PTM of its kind, conjugation of other UbL proteins also contribute to the regulation of multiple cellular events, such as transcription, ribosome assembly, and responses to DNA damage and pathogens. SUMO in encoded by a single gene in Drosophila, and up to 4–5 genes in mammalian cells. Generally, the less abundant SUMO1 is the preferred isoform for SUMOylating RANGAP1, an important regulator of nucleocytoplasmic transport, whereas the more abundant SUMO2 and SUMO3 are rapidly conjugated to substrates under stress conditions, beyond their baseline levels. SUMO2 and SUMO3 are almost identical in sequence, and much like Ub, can form polymeric chains, primarily though K11 linkages. SUMO1 contributes less (if at all) to growing chains, likely acting as a chain terminator. Different strategies have been used to isolate and characterize SUMO conjugates, which remains challenging due to the scarcity and labile nature of this PTM. Overexpression of HIS- or HA-tagged versions of SUMO (Galisson et al., 2011; Tammsalu et al., 2014; Hendriks et al., 2015) or affinity purification of endogenous SUMOylated conjugates using anti-SUMO antibodies (Becker et al., 2013) are the most widely used approaches. Using these approaches, a recent meta-analysis of human SUMO proteomic studies catalogued more

than 3600 proteins as being SUMOylated (Hendriks and Vertegaal, 2016), revealing a substantial role for this modification in cellular homeostasis and stress responses. In Drosophila, a single SUMO isoform, Smt3, has many roles during development (Talamillo et al., 2008a; Cao and Courey, 2017). One example is its role in cholesterol uptake by the prothoracic gland, where it is converted to the steroid hormone ecdysone, which in turn mediates larval transitions and metamorphosis (Talamillo et al., 2008b). Although SUMOylation of the transcription factor FTZ-F1 contributes to cholesterol homeostasis through proper expression of membrane scavenger receptors (Talamillo et al., 2013), there are likely many other SUMO targets in the prothoracic gland that allow progression and proper completion of metamorphosis. A tissue-specific bioSUMO system, inspired by the bioUb system, would facilitate the characterization of a SUMO sub-proteome from this important, yet tiny organ. Because the system is bipartite, needing both AviTag-SUMO and BirA, a multicistronic approach was designed that depending on viral-derived 2A ribosomal-skipping peptides to separate the open reading frames (González et al., 2011; Luke and Ryan, 2018). This alternative design was successful and validated in both cultured Drosophila S2R+ cells and transgenic flies (Pirone et al., 2016), with the identification of > 1000 potential SUMO conjugates in cells, and ≈ 140 from heat-shocked transgenic larvae. The higher number in cells likely reflects that the SUMO E2 conjugase, Lesswright, was also expressed. Of note, the bioSUMO system fulfils physiological roles in the prothoracic gland. Specifically, RNA interference (RNAi) of SUMO in the prothoracic gland leads to a prolonged larval stage, since ecdysone levels do not reach the high threshold level needed for entering into metamorphosis. If an RNAi-resistant form of bioSUMO and BirA is simultaneously expressed with SUMO RNAi, a genetic rescue is observed and flies can enter and complete metamorphosis properly. Reducing endogenous SUMO also increases the chance that bioSUMO will be used for conjugates. This replacement strategy has also been used successfully for other tagged Ubs (Xu et al., 2009; Akimov et al., 2014). Besides Ub and SUMO, vectors that express AviTag versions of other Drosophila UbLs (NEDD8, UFM1, and URM1)

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have been tested successfully in the S2R+ cell line (Pirone et al., 2017). Many cell lines exist for the Drosophila model, but most published work is limited to a few easy-to-grow lines (Cherbas and Gong, 2014). By contrast, human and mouse cell lines show impressive diversity in tissue-of-origin, with many tumour-derived and transformed lines available and a growing number of immortalized primary cell types in use. To broaden the applicability of the bioUbL approach, expression vectors were developed for mammalian expression and the collection was expanded to encompass all the UbLs encoded by the human genome (Pirone et al., 2017). The human bioUbLs incorporate into substrates when tested in transfected cells, revealed by western blotting and immunofluorescence showing subcellular localization, an example being bioSUMO1 that targets PML nuclear bodies and inner nuclear lamina, both known to be sites of abundant SUMOylation. UFM1 (Ub-fold modifier 1) is a less-studied UbL that is implicated maintaining ER homeostasis and regulating haematopoiesis (Daniel and Liebau, 2014), although few substrates had been identified. Isolation and analysis of conjugates from cells expressing bioUFM1, BirA, and UFC1 (the UFM1 E2 conjugase) revealed ≈ 80 potential substrates. Several of the candidates were already known to be associated with each other in protein complexes, raising the possibility that UFMylation may act on multiple proteins within a given complex, which has been previously described for SUMOylation ( Jentsch and Psakhye, 2013). Similar to bioUb, the bioUbL system can be used in cell lines or in transgenic animals, to allow characterization of UbL subproteomes and, using quantitative proteomic methods, to see how conjugate representation and relative abundance changes in response to drug treatments, genetic manipulation, and developmental timepoints. Biotin-mediated proximity proteomics for exploring the Ub/ UbL network In the previous section, BirA was highlighting as an enzyme with exquisite specificity, only releasing its reactive biotinoyl-5′-AMP when it engages with a biotin acceptor peptide (BAP), found in endogenous carboxylases or appended to proteins of

interest for in vivo biotinylation purposes. In addition to the central catalytic domain that binds biotin and biotinoyl-5′-AMP, E. coli BirA has an N-terminal DNA-binding domain for the regulation of the operon encoding biotin synthetic enzymes and a C-terminal domain that likely binds to ATP and biotinylation targets (Sternicki et al., 2017). Based on the X-ray crystallographic structure of E. coli BirA (Wilson et al., 1992), previous genetic studies (Barker and Campbell, 1981; Buoncristiani et al., 1986), and sequence comparisons of diverse bacteria BirA orthologues, enzymatic studies showed that the conserved sequence 115GRGRXG120 is responsible for tight binding of biotin and intermediates (Kwon and Beckett, 2000). Specifically, the R118G mutation of BirA has about 100-fold reduced affinity, allowing reactive biotinoyl-5′-AMP to escape without the strict requirement for the BAP, where it can cause biotinylation of any primary amine (e.g. lysine side chain). When the BirA R118G mutant (hereafter called BirA*) is present in E. coli, promiscuous biotinylation is observed on proteins besides the BCCP carboxylase. When purified, BirA* can self-biotinylate and cause biotinylation of nearby interacting proteins in vitro (Choi-Rhee et al., 2004; Cronan, 2005). It is this permissive or promiscuous biotinylation property of BirA* that forms the basis of the BioID technique. When fused to a protein of interest and expressed in cells, BioID is capable of biotinylating itself, its fusion partner, and primary amines on any proteins that are in close proximity (Roux, 2013). It was first demonstrated using BioID with Lamin A, a quite insoluble protein that localizes to the inner nuclear lamina. Proximal biotinylated proteins were captured, analysed by mass spectrometry, and revealed a significant enrichment for nuclear lamina components, including some novel factors (Roux et al., 2012). Since this original report, BioID has been readily adopted and applied by cell and molecular biologists to examine interactions of proteins, complexes, and even organelles in a variety of organisms. Some examples include tight junctions (ZO-1; Van Itallie et al., 2013), claudin/ occludin (Fredriksson et al., 2015), E-cadherin (Guo et al., 2014; Van Itallie et al., 2014), the Hippo kinase growth control pathway (Couzens et al., 2013), pathogen and host interactions (Boucher et al., 2018; Coyaud et al., 2018; Khan et al., 2018), centrosome and cilia network (Firat-Karalar et al.,

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2014; Gupta et al., 2015), and mRNA-associated stress granules and P-bodies (Youn et al., 2018). Proximally biotinylated proteins may be direct interactors of the BioID fusion, but may also represent indirect associations, such as components of larger complexes in which the BioID participates, or highly expressed proteins that are in the vicinity. A list of proteins that have been identified in many BioID interactomes, i.e. potential false positives, has been published and should be consulted when prioritizing candidates for follow-up studies (Kim et al., 2014). In the same report, to estimate the radius of the promiscuous biotinylation effect, BioIDtagging was performed for various components of the nuclear pore, a megadalton protein assembly with known topology and relative distances. By analysing the spatial distributions of the interactors, an estimate of ≈ 10nm has been proposed. Because BioID relies on diffusion of the reactive biotin intermediate, many factors could potentially influence this value (e.g. biotin concentration, relative density of proteins in immediate environment, temperature, time of labelling period). It

may also change when different versions of the BioID are used (see Table 7.1 and text below). The method is capable of identifying weak and dynamic interactions that may be difficult to catch by other techniques, like co-immunoprecipitation. Also, many assays aim to detect interactions in protein lysates, but the interplay between protein solubility and buffer stringency can be an issue. BioID works in situ, biotinylating proximal proteins that can later be extracted with denaturing urea buffer, which solubilizes most proteins but is still compatible for binding to streptavidin supports. While BioID is very effective and has been used extensively since its recent introduction as a tool for exploring in vivo interactions, it has some shortcomings. The size of BioID (≈ 35 kDa) is slightly larger than GFP, another widely-used fusion partner, and therefore validation of localization and function are advisable to see whether the fusion is tolerated due to size or steric hindrance. Proximal biotinylation by BioID is dependent on adding exogenous biotin to cell culture, and biotinylated proteins accumulate over time, reaching

Table 7.1  Compendium of biotin ligase-dependent labelling methods for studying Ub/UbL pathways Tool

Enzyme

BioUb/ BioUBL1,2

Mutations

Action

Ub/UbL applications

WT Humanized E. coli biotin ligase (BirA)

BAP sequence biotinylation

Study of Ub/UbL conjugates3,4,5

BioID6

R118G Humanized E. coli biotin ligase (BirA)

Proximity biotinylation

Interactomes/substrates of Ub/UbL cycle enzymes7,8,9

BioID210

Humanized A. aeolicus biotin ligase

R40G

Proximity biotinylation

Interactomes/substrates of Ub/UbL cycle enzymes11

BASU12

Truncated B. subtilis biotin ligase

Δ1–65,R124G, E323S, G325R

Proximity biotinylation; Not yet reported increased activity; ID of RNA–binding complexes11

TurboID13

Q65P, I87V, R118S, E140K, Humanized E. coli biotin ligase Q141R, S150G, L151P, V160A, T192A, K194I, (BirA) M209V, M241T, S263P, I305V

Split-BioID

ReconstitutionR118G; two split versions Humanized E. coli biotin ligase reported: E256/G25714; E140/ dependent proximity biotinylation (BirA) Q14115

Sensitive and rapid proximity biotinylation

Not yet reported

Not yet reported

Details of each tool are listed, and they can be used to study Ub/UbL modifications themselves, the enzymatic cycles responsible for writing and erasing those modifications, and effectors that interpret them. 1Franco et al. (2011); 2Pirone et al. (2017); 3Ramirez et al. (2018); 4Lectez et al. (2014); 5Martinez et al. (2017); 6Roux et al. (2012); 7Coyaud et al. (2015); 8Yeh et al. (2015); 9Odeh et al. (2018); 10Kim et al. (2016); 11Hussain et al. (2018); 12Ramanathan et al. (2018); 13Branon et al. (2018); 14De Munter et al. (2017); 15Schopp et al. (2017).

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a saturation point at ≈ 24 hours. This time scale is relatively long, considering that the technique is usually carried out in growing cells which may complete a whole cell cycle in that time. Dependency on exogenous biotin may limit BioID use in whole animals due to tissue accessibility issues. In addition, BioID may also be less efficient at temperatures less than 37°C, perhaps limiting it use in some popular model organisms, such as yeast, nematodes, fruit flies, and zebrafish (Chen and Perrimon, 2017). Fortunately, these disadvantages have driven innovation and development of new improved versions of BioID (Table 7.1). Simply removing the N-terminal DNA-binding domain of BioID would be a way to reduce its size, but it also results in loss of biotinylation activity. However, some bacteria have BirA orthologues that are naturally lacking the N-terminal domain, but still act as biotin ligases. BioID2 (Kim et al., 2016) is one such example from Aquifex aeolicus. A single mutation (R40G), which lies in same position as R118 of E. coli BirA according to sequence and structure comparisons (PDB ID: 2EAY), also results in promiscuous biotinylation. When compared side-by-side with BioID, the smaller size of BioID2 may cause less steric hindrance in fusions. BioID2 also requires less exogenous biotin to achieve the same level of proximal biotinylation, so may be more efficient for in vivo applications. Use of BioID2 has identified novel inner nuclear envelope components, such as VRK2A (Birendra et al., 2017), and datasets exist for both BioID-LaminA and BioID2-LaminA to allow direct comparisons (Roux, 2013). The third generation of BioID has arrived with speed and efficiency. A new method to look for proximal interactors of RNA-binding proteins has been reported and relies on a novel enhanced promiscuous biotinylator called BASU (Ramanathan et al., 2018). It is derived from Bacillus subtilis BirA, which retains its biotinylation capacity when its DNA-binding N-terminus is removed. With additional mutations, it yielded an enzyme with smaller size and very fast kinetics, labelling in minutes to the same extent as the original BioID does in 18–24 hours. Yet another improved version was obtained by rational design and in vitro evolution, called TurboID (Branon et al., 2018). Using yeast surface display, FACS sorting, and libraries generated by error-prone PCR, screening consisted of multiple

rounds of mutation and selection and yielded a new extensively mutated version of BirA: TurboID. It is optimized for fast kinetics (detectable labelling in 10 minutes) and showed improved performance at lower temperatures, opening up in vivo applications in many model organisms. A second variant, miniTurbo, lacks the N-terminal DNA-binding domain but contains other changes that restores and extends the promiscuous biotinylation property. While smaller in size, miniTurbo has slightly reduced biotinylation efficiency when compared to TurboID. Benchmarking by TurboID creators to compare between BioID, BioID2, and BASU surprisingly showed that all three worked equivalently (Branon et al., 2018), contrary to claims of incremental improvements by BioID2 and BASU (Kim et al., 2016; Ramanathan et al., 2018). Moreover, all three were slower or less efficient than TurboID or miniTurbo. Independent comparisons, and perhaps with different fusion partners, would be very informative to decide which proximal biotinylation approach is best for a particular system or application. It is worth mentioning that there are other biotin-based proximity labelling methods described, also fast and efficient, that rely not on BirA, but on peroxidases. APEX (Rhee et al., 2013), and the second-generation APEX2 (Lam et al., 2015), are modified ascorbate peroxidases that can be used as fusions in a similar fashion to BioID for detecting proximal interactors. Using a modified biotin (biotin-phenol) and hydrogen peroxide, APEX catalyses the conversion of biotin-phenol into a short-lived reactive biotin-phenoxyl radical, which can diffuse and covalently modify nearby Tyr, Trp, His and Cys residues. The proximity biotinylation radius is smaller and labelling time is shorter, making it attractive for some applications. The biocompatibility of labelling reagents can restrict its utility in whole organisms, but it can be used in explanted tissues (Chen et al., 2015). Another emerging application is the BAR method (Biotinylation by Antibody Recognition; Bar et al., 2018), in which primary and horseradishperoxidase-coupled (HRP) secondary antibodies are used to label antigens of interest in fixed cells or tissue sections. Then, using biotin-phenol and hydrogen peroxide, as described for APEX, proteins proximal to the HRP are biotinylated, and can later be captured and identified by MS. As long as

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good primary antibodies are available, this method may prove useful for Ub/UbL pathway applications and beyond. Off to a good start: BioID meets ubiquitin BioUB and bioUbLs, as well as many other complementary methods, have facilitated the isolation and cataloguing of 1000s of potential substrates and even modification sites. But persistent questions remain: Which of the 100s of E3 ligases are responsible for adding Ub/UbLs to which substrates? Which are the proteins that interpret a given Ub/UbL modification, and how do they respond? Which of the many Ub/UbL peptidases (DUBs) are responsible for removing and recycling Ub/UbLs from modified substrates? Since the action of E3 ligases and DUBs is very dynamic and may not involve tight or stable interactions with the underlying substrates when adding or removing the Ub/UbL, identification of substrates can be particularly challenging. For E3 ligases, some methods have been reported that successfully identify interactors and substrates (Zhuang et al., 2013; O’Connor et al., 2015; Kumar et al., 2017). Application of BioID technology to the enzymes of the Ub/UbL pathways may reveal new regulators, cofactors, and especially substrates (Fig. 7.3). Some examples of BioID-E3s and BioID-DUBs have already been published, and with the development of faster, more efficient derivatives of BioID, this number is certain to grow. Certain ubiquitin E3 ligases are multi-subunit complexes that use an adaptor (Skp1-cullin) to bring together E2 conjugases and substrate-recognizing F-box proteins. Two such F-box proteins, β-TrCP1 and β-TrCP2, are regulators of developmental and disease signalling pathways (e.g. Wnt, NFκB, Hippo, Shh). Since only a few substrates were known, BioID was applied to β-TrCP1 and β-TrCP2 (Coyaud et al., 2015). When coupled with inhibition of the proteasome to enrich for potential ubiquitinated substrates, proximal interactors were obtained. Both known and novel substrates were identified (>50) and many were validated with other methods. In another study that utilized both the BioID and APEX approach, proximity biotinylation was used to study the regulation of ribosome quality control (RQC; Zuzow et al., 2018). Ltn1 is an E3 ligase that assists the core RQC to prevent accumulation of

errant translation products via rapid ubiquitination and proteasome-mediated degradation. BioIDtagging of Ltn1 revealed novel interactors that were further validated to be Ltn1 target substrates, as well as a non-degradative ubiquitination of ribosomal S6 kinase family members. Like Ltn1, RNF41 is a RING-finger-containing E3 ligase, but with multiple roles in intracellular trafficking. A multifaceted approach to identify RNF41 interactors included BioID-based proximity proteomics (Masschaele et al., 2018). A stabilized, ligase-defective mutant of RNF41 was used to increase expression levels and chances of identifying substrates. In addition to known RNF41 substrates (BIRC6 and USP8), novel protein interactors were identified. More detailed analysis of one candidate, AP2S1, showed that RNF41 was able to mediate a change in its localization and stability, but had no effect on AP2S1 ubiquitination. RNF41 might have an indirect role in stabilizing AP2S1, perhaps by acting on another unknown E3 ligase. This example serves to demonstrate the importance of independent validations in BioID experiments. Lastly, regarding regulators of E3 ligase activity, BioID was used to verify the in vivo interaction of the mitochondria-associated E3 ligase UBE3B with calmodulin, which is thought to mediate changes in its ubiquitination activity in response to calcium levels (Braganza et al., 2017). Several studies highlight the application of BioID to DUBs with the goal of identifying interactors and potential substrates for deubiquitination. There are several classes of DUBs, grouped by the structure and specificity of their catalytic domains. As mentioned, ubiquitin chains exist with different linkages (e.g. K48, K63, linear, etc.) and it follows that DUBs can have preferences for cleaving certain linkage types. DUBs also contain other domains and motifs that may recognize and confer specificity to certain types of substrates (Mevissen and Komander, 2017). Because they likely act rapidly on their targets, improvements in the speed of BioID action (i.e. using second and third generation versions) may also increase chances of identifying bona fide substrates. A DUB called USP37 has roles in chromosome segregation and mitotic progression, and BioID was utilized to identify potential interactors that mediate this role (Yeh et al., 2015). Known interactors of USP37 were identified including multiple proteins from SCF and APC complexes, but also novel

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Figure 7.3  The BioID system as applied to E3 ligases and DUBs. A promiscuously biotinylating mutant of BirA (termed BirA* or BioID) is fused to an E3 ligase or DUB, and then expressed in cells. While the E3 or DUB are executing their functions, biotin supplementation can drive biotinylation of proteins proximal to the E3/DUB, which may be substrates, E2s, adaptors, or other regulators. Cells are then lysed in denaturing conditions and biotinylated proteins are purified by streptavidin pull-down. The E3/DUB interactome is then identified by MS. For simplicity, the resulting pool of biotinylated proteins depicted here is common for E3-BirA* and DUB-BirA*, but the two pools would be unique, both in the proteins represented and conjugation status of the Ub and Ub chains. DUB, deubiquitinase; MS, mass spectrometry; Ub, ubiquitin.

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interactions were found with cohesin, a protein complex that holds together sister chromatids after DNA replication, as well as its regulator WAPL. Additional validation showed that USP37 likely deubiquitinates WAPL and stabilizes its presence on chromatin, which is necessary for proper cohesin function and chromosome segregation. Another USP-type DUB called USP12 was identified as one of several DUBs that are recruited to the cytoplasm of T lymphocytes during T-cell receptor (TCR) activation. To identify potential targets of USP12, BioID was employed and proximal interactors were identified ( Jahan et al., 2016). Among them were LAT1 and Trat1, two adaptor proteins that serve to stabilize TCR at the membrane. Further validation comparing WT and USP12 knockout cells showed that these proteins were mono-ubiquitinated and degraded through the lysosomal pathway. In another example, BioID2 was fused to the DUB UCHL1 for proximity proteomics (Hussain et al., 2018). The aim was to identify the mechanism by which this DUB can regulate the mTOR pathway, a key mediator of cell growth and proliferation. Among the potential interactors or substrates, they focused on the eIF4F translational initiation complex, which is a target of the mTOR substrate 4EBP1. Two of the three subunits of the eIF4F complex were biotinylated by BioID2-UCHL1, and the assembly of this complex was shown to be promoted by UCHL1 in a catalytic-dependent manner. Co-immunoprecipitation suggested a direct interaction with one subunit, but expression levels or stability of the subunits did not depend on presence of UCHL1, so it is unlikely to be a direct substrate for deubiquitination. Beside deubiquitinases, BioID has been used for the deSUMOylase SENP2 with the aim to identify proximal proteins as an indicator of subcellular localization (Odeh et al., 2018). An N-terminal domain of SENP2 was described that is responsible for targeting SENP2 to intracellular membranes, and the BioID-SENP2 interactome revealed novel associations with proteins related with the inner nuclear envelope, nuclear pore complex, as well as the ER and Golgi membranes. A list of 187 highconfidence interactions was reported, and although SUMOylation status was not the focus, our quick comparison to a comprehensive list of SUMO2/3modified human proteins [≈  3800 proteins (Hendriks et al., 2018)] shows that almost 40% of

the BioID-SENP2 candidates are potential SUMO targets. One should be mindful that the reactive biotinoyl-5′-AMP generated by bioID targets the primary amines of nearby lysines in proximal interacting proteins. As lysines are the primary targets for Ub/UbL modification, as well as other PTMs, bioID fusions and the resulting biotinylation could cause interference and have biological consequences for the cells in which they are expressed. Faster, more efficient derivatives of BioID and controlled expression of fusions should minimize this effect. In summary, the BioID method (as well as optimized derivatives) promises to be an effective tool to dissect the Ub/UbL pathway and reveal its breadth and specificity. Future perspectives Another use of biotin which has not been mentioned is its use as a molecular handle or traceable label for in vitro studies. The need for good biochemical tools in the Ub/UbL field will certainly grow since mechanistic studies are always needed for the new discoveries coming out of large-scale systems-level proteomics. Biotin can be used with many of the novel activity-based probes for Ub/ UbL pathway enzymes (e.g. de Jong et al., 2012; Mulder et al., 2016). Activated NHS-ester biotin enables the chemical biotinylation of purified proteins such as Ub or UbLs, which can be used for in vitro assembly or disassembly assays. Because this type of bulk chemical biotinylation is indiscriminate and modifies any primary amine, unwanted effects may arise due to blocked lysines. Using the BirA-AviTag system, biotinylation can be sitespecific and avoid these problems. This approach has been used for SUMO molecular traps (Da SilvaFerrada et al., 2013), by biotin-labelling the traps at the N-terminus to facilitate the capture of polySUMOylated proteins on streptavidin supports (Lang et al., 2016). Other biotinylated molecular traps that enrich for endogenous ubiquitinated proteins carrying chains of different linkage types (K48, K63, linear), are also commercially available, and are useful for applications with extracts from cells and primary tissues. The emerging development of new affinity reagents such as affimers or nanobodies for purifying Ub/UbLs may benefit from biotin labelling (Hughes et al., 2017; Michel et al., 2017). Also, it is possible to make recombinant

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proteins with either site-specific biotinylation or fusions to monomeric streptavidin (Lim et al., 2013), which may be combined to create new tools, more potent enzymes, and more innovative assays to apply towards Ub/UbL pathways. We have described the use of bioUbL/BirA systems to allow in vivo labelling of Ub/UbL conjugates for later characterization by MS. While powerful, these systems usually need to be introduced into cells or model organisms as transgenes, which raises concerns about expression levels and dominant effects. Additionally, the tagged Ub/ UbL needs to compete with endogenous versions, reducing the efficacy of recovery. Using RNA interference to silence the endogenous copies is one way to enhance the tagged versus endogenous ratio, but genome-editing approaches might be an alternative solution. The CRISPR-Cas9 system (Doudna and Charpentier, 2014) can be used to introduce tag-encoding sequences directly into Ub or UbL genes, aiming for 100% replacement. This will not only test the functionality of the tagged version, but also lead to maximal recovery of tagged Ub/UbL conjugates. In most in vivo biotinylation experiments, BirA is expressed constitutively in cells, where it shows a diffuse localization throughout the nucleus and cytoplasm. Experiments usually compare two cell lines, expressing bioUbL with BirA, or just BirA alone, to discern enriched versus background proteins. Switching to inducible forms of BirA or the bioUbL, either through transcriptional activation [e.g. Tet system (Das et al., 2016)] or by protein stability [e.g. via controllable degrons (Kanemaki, 2013)], might allow better control of BirA/bioUbL experiments since a single cell line, induced or noninduced, can be assayed. Also, it may be possible to express BirA in particular cell types within a tissue or target BirA to subcellular locations in order to have a more refined set of bioUbL conjugates. Similar approaches combining BirA and the AviTag have been used to isolate modified nucleosomes (Lau and Cheung, 2013), chromatin (Shoaib et al., 2013), and even nuclei (Deal and Henikoff, 2010; Amin et al., 2014). It is also recommendable to control expression levels in BioID-based approaches. Overexpression will likely lead to random biotinylation of abundant proteins, which will confound efforts to find real proximal interactors. In most BioID experiments

published to date, fusions are expressed as low levels, sometimes using inducible systems and low copy-insertion strategies, but BioID knock-in is possible using CRISPR-Cas9-mediated genome editing (Mulholland et al., 2015). Using this approach on E3 ligase or DUB genes may render more sensitive data. Another recent development that might find use for studying Ub/UbL modifications is Split-BioID (Table 7.1). If a protein is divided into two parts, neither of which retains the activity of the parental protein, then the two halves may refold and reconstitute the original activity if placed in proximity to each other. This concept, protein fragment complementation, has been used successfully for GFP (Cabantous et al., 2005), luciferase (Paulmurugan et al., 2002), and even Cas9 nuclease (Wright et al., 2015; Zetsche et al., 2015). Recent reports show that BioID can be split in two halves, such that each half can be fused to a protein of interest (De Munter et al., 2017; Schopp et al., 2017). The same concept has also been validated for APEX2 (Xue et al., 2017). When the two fusions are in close proximity, BioID or APEX2 is reconstituted and begins to biotinylate proximal interactors, some of which might be dependent on the formation of the complex between the two fused proteins. The Ub/UbL system is full of pair-wise and multi-component interactions that could benefit from such split-protein approaches, exemplified by a study examining E2 conjugase and E3 ligase interactions using splitGFP (Blaszczak et al., 2016). In summary, biotin is a versatile tool that has been used to label biomolecules for studies in biochemistry, cell and molecular biology. Here we have highlighted how biotin/streptavidin and BirA have been used to explore the Ub/UbL landscape, not only to survey what has been done, but also to inspire further development of biotin-based tools for this field and others. Acknowledgements RB and JDS are funded by grants BFU2011-25986 and BFU2014-52282-P (MINECO/FEDER, EU) and the Severo Ochoa Excellence Accreditation (SEV-2016-0644) and Consolider Programs (BFU2014-57703-REDC). Support was also provided from the Department of Industry, Tourism and Trade of the Government of the Autonomous Community of the Basque Country (Elkartek

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Research Programs) and from the Innovation Technology Department of the Bizkaia County. OBG is supported through the UbiCODE consortium (funded from the European Union’s Horizon 2020 research and innovation programme under the Marie Skłodowska-Curie grant agreement No 765445). References

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Screening Mammalian SUMOylated Proteins by Fluorescence Protein Reconstitution

8

Maki Komiya1,2, Mizuki Endo1 and Takeaki Ozawa1*

1Department of Chemistry, Graduate School of Science, The University of Tokyo, Hongo,

Bunkyo-ku, Tokyo, Japan.

2Present Address: Laboratory for Nanoelectronics and Spintronics, Research Institute of

Electrical Communication, Tohoku University, Sendai, Japan.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.08

Abstract SUMOylation is an essential post-translational protein modification in various cellular functions. To clarify the role of SUMOylation, numerous screening approaches have been reported for the discovery of novel SUMOylated proteins. However, the reversibility of SUMOylation and the highly varied SUMOylation levels among targets have made it difficult to detect infrequentlySUMOylated proteins, especially in mammalian cells. Here, we describe a newly developed screening system for mammalian SUMOylated proteins in living cells, which is based on split fluorescence protein reconstitution and fluorescence-based cell sorting technique. The experiments demonstrated that SUMOylation by SUMO2 was detectable as a fluorescence signal in living mammalian cells, which enabled exploration of SUMOylation candidates without cell destructive processes. The system successfully identified 2 reported SUMO2-substrates and 36 SUMO2-substrate candidates, of which Atac2 was shown as SUMOylated at a lysine 408. We summarized the applicability to other SUMO isoforms and various cell types, which will be able to contribute to broader exploration of the roles of SUMOylation in numerous biological phenomena.

Introduction A small ubiquitin-related modifier (SUMO) is a post-translational protein modifier, which is highly conserved protein in eukaryotes ( Johnson, 2004). In mammalian cells, at least three different SUMO isoforms, SUMO1, SUMO2, and SUMO3 are expressed, with different substrate selectivity. SUMOs are reversibly conjugated with the substrate proteins via covalent isopeptide bond between the C-terminal glycine residues of SUMOs and the lysine residues of substrate proteins. The reaction, called ‘SUMOylation’, is sequentially mediated by distinct enzymes, E1 (a SUMO-activating enzyme), E2 (a SUMO-conjugating enzyme), and E3 (a SUMO ligase) (Capili and Lima, 2007). The inverse reaction, called ‘deSUMOylation’, is induced by SUMO-specific peptidases (SENPs) (Hickey et al., 2012). The strict regulation of the balance between SUMOylation and deSUMOylation precisely maintains the SUMOylation level of each SUMO substrate, usually less than 1% ( Johnson, 2004; Geiss-friedlander and Melchior, 2007), which serves to their functional modulation. It has been reported that SUMOylation plays crucial roles in various biological processes, such as DNA repair, cell cycle, and signal transduction (Girdwood et al.,

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2003; Lee et al., 2008; Bergink and Jentsch, 2009; Ouyang and Gill, 2009), thereby highlighting the importance of novel SUMOylation substrate discovery. Various screening methods for novel SUMOylated proteins have been devised to clarify the roles of SUMOylation in diverse biological contexts. For instance, immunoprecipitation (IP)based proteomic methods have been mainly used for screening mammalian SUMOylated proteins (Zhao et al., 2004; Tirard et al., 2012; Filosa et al., 2013). In the IP-based methods, SUMOylated proteins were collected from cell lysates with antibodies against exogenously expressed SUMO, followed by proteomic analysis using mass spectrometry. However, due to the difficulty in complete inhibition of deSUMOylation after cell lysis, the IP-based proteomic approaches are susceptible to the bias caused by highly varied SUMOylation levels and tolerance to deSUMOylation, which possibly in turn makes it difficult to detect infrequently SUMOylated proteins with high deSUMOylation rate. To avoid cell destructive procedures, yeast twohybrid screening method was developed to identify SUMOylated proteins using living yeast cells (Hannich et al., 2005). Although the method enabled detection of SUMOylation in intact cells, it has several difficulties in the detection of mammalian SUMO-substrate proteins. First, the two-hybrid technique used in the system required nuclear translocation of candidate proteins to initiate gene reporter expression for the detection of SUMOylation. Therefore, it is difficult to examine proteins which were confined in different organelle compartments. Second, since yeast cells express only single SUMO isoform (Bylebyl et al., 2003; Takahashi et al., 2003), it cannot reflect complex SUMOylation properties in mammalian cells, which expresses at least three SUMO isoforms with different substrate specificity (Melchior, 2000; Saitoh and Hinchey, 2000). It was also reported that the substrate preference was affected by the expression pattern of E3 proteins (Tatham et al., 2005; Vertegaal et al., 2006). Third, mammalian SUMOylation patterns depend on cell types (Degerny et al., 2005; Ji et al., 2007), which is not explorable in yeast-cell-based approach. Taken altogether, a novel screening system was required to detect SUMO-substrate candidates in living mammalian cells.

Here, we introduce a novel system for screening mammalian SUMOylated proteins (Komiya et al., 2017). The system is based on fluorescence protein reconstitution for the detection of SUMOylation in living mammalian cells. Since the fluorescence protein reconstitution is an irreversible reaction (Shyu and Hu, 2008; Isogai et al., 2011), the designed system is useful for the detection of infrequent or transient SUMOylation in live-cell condition. Based on the fluorescence signal, the cells harbouring reconstituted fluorescence proteins were sorted by a fluorescence-activated cell sorter (FACS) in a high-throughput manner. The basic scheme is similar to the one demonstrated previously in the identification of mitochondrial proteins (Ozawa et al., 2003). The system screened SUMO2-substrates in living mouse cells, and successfully identified 2 reported SUMO2-substrates and 36 SUMO2substrate candidates. The biochemical analysis demonstrated that Atac2, one of the candidates, was SUMOylated by SUMO2 at a lysin 408. The whole procedure of the novel screening method based on fluorescence protein reconstitution in mammalian cells The whole procedure of our novel screening method using fluorescence protein reconstitution was summarized in Fig. 8.1. In the system, fluorescence signal generated by fluorescence protein reconstitution upon SUMO conjugation to the substrate was used for the detection of SUMOylation in living mammalian cells (Fig. 8.2). A fluorescence protein ‘Venus’, which emits bright yellow fluorescence (Nagai et al., 2002), was used for the reconstitution. The SUMO2 sequence was genetically fused to the sequence coding the N-terminal fragment (amino acids 1–158, named VN) of Venus with a GS linker (corresponding to Gly-GlyGly-Gly-Ser amino acids), named VN-SUMO2. Mouse cDNA libraries generated from mRNAs were enzymatically digested and genetically fused to the sequence encoding the C-terminal fragment (amino acids 159–240, named VC) of Venus, named VC-Library. The VC fragment sequence was fused with the library sequences via three linkers of different length, ggcggaggcgga, ggcggaggcggag, and

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Figure 8.1  Schematic of screening mammalian SUMOylated proteins based on the reconstitution of split Venus fragments. Library DNAs are inserted into retrovirus infection vectors with DNA of a C-terminal fragment of Venus (VC) and transfected into PlatE cells. The produced retroviruses harbouring VC-library DNAs are added to NIH3T3 cells stably expressing SUMO2 fused with a N-terminal fragment of Venus (VN). The fluorescent cells harbouring reconstituted Venus are sorted by FACS. The library DNA is extracted from each fluorescent cell. SUMOylated protein candidates are identified by an analysis of the extracted DNA sequences.

ggcggaggcggagg in consideration of the frame shift. If the VC-library proteins are SUMOylated with VN-SUMO2, the VN and VC fragments interact with each other, thereby generating fluorescence signal upon reconstitution. The plasmid DNA encoding VN-SUMO2 was transfected with a murine cell (NIH3T3 cell), and a VN-SUMO2 stable cell line was established by the selection marker, zeocin. The VC-library DNAs were converted into retrovirus libraries by transient transfection to PlatE cells (retrovirus packaging cells). The medium containing the produced retrovirus were added to the VN-SUMO2 stable cell line in the presence of polybrene. The infection efficiency was adjusted to achieve a condition that each single cell has at most one copy of VC-library DNA. The infected VN-SUMO2 stable cell line expressing VC-library proteins were trypsinized

and suspended in PBS. The cells with fluorescent signal generated by reconstitution upon SUMOylation of VC-library proteins were rapidly collected by a fluorescence-activated cell sorter (FACS) in a live cell condition, using the standard FACS procedure with an excitation wavelength of 488 nm and a measurement wavelength of 525 (± 15) nm. The sorting region was identified by a fluorescence intensity higher than the maximum intensity of autofluorescence. The collected fluorescent cells were spread on a culture dish with a cell density low enough to separate each cell. The single cell clones were isolated to different dishes and incubated. From each cell clone, cDNA was retrieved by PCR and the sequence encoding the VC-library was analysed. Finally, the SUMO2-substrate candidates were identified from the DNA analysis, with reference to the GenBank database.

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Figure 8.2  The probes and the principle for detecting SUMOylation in living cells. (A) Schematic DNA constructs of VN-SUMO2 and VC-library probes. VN: N-terminal fragment (amino acids 1 to 158) of Venus. VC: C-terminal fragment (amino acids 159 to 240) of Venus (VC). (B) Schematic of detecting SUMOylation under a live-cell condition using the probes.

Identification of SUMOylated proteins Evaluation of the probes using the reconstitution of split Venus The generation of Venus fluorescence upon SUMOylation in living mammalian cells was evaluated by using a famous SUMOylated protein called RanGAP1(Mahajan et al., 1997). The plasmid DNA encoding VC fragment fused with RanGAP1 (VC-RanGAP1) was introduced into murine VN-SUMO2 stable cell line. The confocal fluorescence microscopic analysis demonstrated that the Venus fluorescence signal was generated around the nucleus and partly in the cytosol (Fig. 8.3A). From the previous report that showed that modification by SUMO translocated RanGAP1 from cytosol to perinuclear region (Mahajan et al., 1997), the result suggested that the SUMOylation of RanGAP1 triggered Venus reconstitution

in living cells, without affecting protein-intrinsic localization profiles. Next, the fluorescence intensity was analysed for the SUMOylation-induced cells and the control cells. VN-SUMO2 stable cell line was infected with retrovirus harbouring VC-RanGAP1 DNA. As a negative control, DNA encoding VC fragment fused with a RanGAP1 deletion mutant (VC-Δ20aaRanGAP1), which lacked SUMOylation site K524 (Macauley et al., 2004) and the amino acid sequence required for SUMOylation (Matunis et al., 1998; Rodriguez et al., 2001), was also introduced to the cell lines by infection. The fluorescence intensities of the infected or non-infected cells were analysed by FACS (Fig. 8.3B). The fluorescence intensity histograms revealed that 43(±3)% of the cells expressing VC-RanGAP1 showed higher fluorescence intensities than the maximum fluorescence intensity of the control cells. In contrast, only 0.16 (± 0.03)% of the VC-Δ20aaRanGAP1

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Figure 8.3  Evaluation of the probes by using SUMOylated protein RanGAP1. (A) VN-SUMO2 stable cell lines expressing VC-RanGAP1 and H2B-EBFP imaged by confocal fluorescence microscopy. Scale bar: 10 μm. (B) Fluorescence intensity analysis of the VN-SUMO2 stable cell lines without infection (blue) and with infection of VC-RanGAP1 (left, red) or VC-D20aaRanGAP1 (right, green). Histograms were generated from five repeated measurements of 5000 cells. Dark coloured lines indicate the averaged intensities. Light coloured areas indicate the standard deviation. Dotted grey lines indicate the threshold where control cells no longer exist. Reprinted from Komiya, M. et al. (2017). Sci Rep. 7, 17443. Distributed under the terms of the Creative Commons CC BY 4.0. license.

expressing cells yielded higher intensities. These results suggested that the maximum fluorescence intensity of the non-infected stable cell line could be used as a threshold to discriminate cells harbouring SUMOylated proteins with reconstituted Venus. The above results indicated that the probes using the reconstitution of split Venus fragments fused with SUMO and its substrate enabled to detect the SUMOylation of target protein with fluorescence microscopy and FACS analysis in living mammalian cells. Isolation of the fluorescent cells that harboured reconstituted Venus with putative SUMOylated library proteins Based on the threshold criteria, the cells harbouring putative mammalian SUMOylated proteins with reconstituted Venus were sorted by FACS using various VC-library DNAs. The retrovirus solution harbouring the VC-library DNAs was added to VN-SUMO2 stable cell line at ≈ 30% infection efficiency. The efficiency was estimated

using GFP-infected cells, whose retrovirus was produced under the same condition. Fluorescence intensity analysis by FACS detected infected cells with fluorescence intensities higher than the autofluorescence (Fig. 8.4). The result indicated that the infected cells included cells with SUMO-substrate candidates. The cells with higher fluorescence intensities were sorted by FACS and incubated for a week. The cells were sorted again to reduce falsepositives. The sorting cycles were repeated for 3–4 times. The final cell populations showed clearly higher fluorescence than the non-infected cells. The result indicated that the cells with SUMO-substrate candidates were properly collected by FACS based on their reconstituted fluorescence intensities. Identification of the candidates of SUMOylated proteins by DNA analysis The DNA sequences encoding the library proteins were analysed and the SUMOylated protein candidates expressed in the FACS-sorted cells

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Figure 8.4  FACS isolation of the fluorescent cells. Fluorescence intensities of the VN-SUMO2 stable cell lines with or without infection with VC-library DNAs were analysed by FACS. (A) Intensity profiles before FACS sorting. In the region indicated with double-headed arrow, the infected cells have higher fluorescence intensities than control cells. (B) Intensity profiles after FACS sorting. The infected cells were repeatedly sorted by FACS for four times. The data show the fluorescence intensity of the finally sorted cells. Reprinted from Komiya, M. et al. (2017). Sci Rep. 7, 17443. Distributed under the terms of the Creative Commons CC BY 4.0. license.

were identified. The sorted cells were individually plated on a culture dishes to isolate single clones. Each DNA was extracted from the single clones, amplified by PCR, and subjected to agarose gel electrophoresis to remove contaminated materials. The DNAs detected in the gel was separated, purified, and subjected to direct DNA sequencing. With reference to the GenBank database, the SUMOylated protein candidates were identified. DNA sequence analysis identified 38 SUMOsubstrate candidates (Table 8.1). The previous reports showed that the identified candidates had a variety of subcellular localization patterns: Anxa5 was found in both cytoplasm and nucleus (Sun et al., 1992); Drosha localized in nucleus (Tang et al., 2010); Plscr3 in mitochondria (Liu et al., 2003); and Tuba1b in microtubules (Azakir et al., 2010). The screened proteins also showed diverse functions in living mammalian cells: Narf acted as an ubiquitin ligase (Yamada et al., 2006); Myof regulated membrane integrity in vascular endothelial growth factor signalling (Bernatchez et al., 2007); Arpc1b as a progression factor of cell cycle (Molli et al., 2010); Taz was involved in cell proliferation (Lei et al., 2008). These facts indicated that the developed screening method was able to detect proteins with different subcellular locations and various functions. From amino acid sequence analysis of the identified candidates, SUMO consensus recognition sites (Rodriguez et al., 2001; Johnson,

2004) and SUMO-interacting motifs (SIMs) (Hecker et al., 2006) were predicted. A SUMO consensus recognition site is a potential position for covalent SUMO conjugation, which is shown as Ψ-K-X-E/D (‘Ψ’: a hydrophobic amino acid, ‘K’: the SUMO-modified lysine residue, ‘X’: one of any amino acids, ‘E’: a glutamic acid, ‘D’: an aspartic acid). In contrast, SIM is a site for non-covalent SUMO interaction, one of which is shown as [V/I]-X-[V/I]-[V/I] (‘V’: valine, ‘I’: isoleucine). A computational prediction based on the SUMO consensus recognition sites (Xue et al., 2006; Zhao et al., 2014) showed that 17 proteins of the identified proteins harboured SUMO consensus recognition sites. As for SIMs, the algorithm (Zhao et al., 2014) predicted 24 proteins. These results implied that the present method could potentially detect both SUMOylation substrates and proteins with non-covalent SUMO interaction. As for SUMO2-specific SUMOylation, two of the identified 38 proteins, Lmna and Rpl37a, have been already reported as modified by SUMO2, which was shown by immunoblotting (Zhang and Sarge, 2008; Yun et al., 2008). The result supported that the present method detected SUMO2 modification in living mammalian cells. Among candidates, 14 proteins have not yet been identified as SUMOylated protein candidates, based on the previous report that compiled several MS-based screening results (Hendriks and Vertegaal, 2016). The result suggested the

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Table 8.1  The SUMOylated protein candidates identified by the screening method based on fluorescence protein reconstitution The detected protein species Rpl37a, Lmna, Rps9, Rpl32, Eif3e, Gsn, Stx12, Bgn, Drosha, Uqcrh, Plxnb2, Rpl18a, Atac2, Ermp1, Mrpl4, Tmsb4x, Rpsa, Lgals3, Pcolce, Tuba1b, Pbrm1, Myof, Dynlrb1, Fam63b, Taz, Rps3a, Myl9, Rpl6, Narf, Arpc1b, Psmb4, Polr1d, Rpl10, Fth1, Anxa5, Plscr3, Wisp2, Cops7a *The protein names in red indicate that the proteins have been previously reported as SUMOylated. Reprinted with modification from Komiya, M. et al. (2017). Sci Rep. 7, 17443. Distributed under the terms of the

Creative Commons CC BY 4.0. license.

scope of detectable protein species by the present reconstitution-based method was partially different from that of the previous MS-based screening methods. This fact indicated that the screening method has a potential to contribute to the discovery of novel SUMOylation candidates in a complementary manner with the MS-based approaches. Consequently, the developed method based on reconstitution of split Venus screened 2 reportedSUMOylated proteins and 36 SUMOylated protein candidates with diverse subcellular protein locations and functions, with distinct difference in the scope of detectable candidates from MS-based approaches. Validation of the candidate SUMOylation identified by the screening method based on fluorescence protein reconstitution The candidate proteins were subjected to the conventional biochemical analysis based on immunoprecipitation (IP) and Western blotting, and SUMOylation of the candidates was validated. Murine NIH3T3 cells were transiently transfected with the DNA encoding candidate protein labelled with a V5-epitope tag with or without DNA harbouring Myc-epitope tagged SUMO2 (MycSUMO2). Overexpression of both candidate and SUMO2 by transient transfection was supposed to contribute to the easier detection of SUMOylation by biochemical approach. On cell lysis, the V5-tagged molecules were immunoprecipitated with anti-V5 antibodies, followed by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) with protein denaturation. Since the covalent modification by SUMO2 was not destroyed during denaturation, the molecular weight of SUMOylated candidate was expected to become larger than that

of the unmodified form. The separated proteins were transferred on the nitrocellulose membrane and immunoblotted with either anti-V5 antibodies or anti-Myc antibodies. In the biochemical SUMOylation candidate analysis, Atac2 showed unique immunoblotting signal with anti-Myc antibody, multiple bands at 120 kDa and over 150 kDa (Fig. 8.5A) in a ladder manner. As for immunoblotting results with antiV5 antibody, the polypeptide bands were detected at 100 kDa and 120 kDa. The smallest band around 100 kDa was almost consistent with the molecular weight of Atac2, 92 kDa. From the detection of the band around 120 kDa in both immunoblots with Anti-V5 and anti-Myc antibodies, the band was assigned as Atac2 modified by Myc-SUMO2 molecule, whose size was around 12 kDa. Based on the SUMOylation site prediction algorithm (Xue et al., 2006; Zhao et al., 2014), several lysine residues as SUMOylation site candidates for Atac2, K305, K408 and K749, were predicted. Among them, point mutation at K408 completely abolished SUMOylation (Fig. 8.5B). These results demonstrated that Atac2 was a novel SUMO-substrate in mammalian living cells, SUMOylated at K408. Conclusion Our newly-developed screening method for mammalian SUMOylated proteins is based on fluorescence protein reconstitution and cell sorting by FACS. The method has several advantages in detecting SUMOylated proteins. Fluorescence protein reconstitution enables us to detect SUMOylation as fluorescence signal in living cells, which is not affected by their subcellular protein localizations or functions. Owing to the irreversibility of the reconstitution reaction, the fluorescence signal from the reconstituted fluorescence protein would

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Figure 8.5  Identification of novel SUMOylated protein Atac2 and its SUMOylation site. (A) Posttranslational modification of Atac2 by SUMO2. NIH3T3 cells transiently expressing indicated molecules were subjected to immunoprecipitation with anti-V5 antibodies. The immunoprecipitated samples were blotted with the indicated antibodies. IB, immunoblotting; IP, immunoprecipitation. The arrowheads show the predicted protein sizes. (B) Atac2 is modified by SUMO2 at K408. NIH3T3 cells transfected with indicated plasmids were subjected to immunoprecipitation followed by Western blotting. Reprinted from Komiya, M. et al. (2017). Sci Rep. 7, 17443. Distributed under the terms of the Creative Commons CC BY 4.0. license.

not decrease even after deSUMOylation, which would be suitable for the detection of transient or infrequent SUMOylation. Moreover, the signal from such proteins would not be overwhelmed by that from highly SUMOylated protein, since the protocol analysed candidate proteins individually expressed in different cells. The rapid cell collection by FACS and the candidate identification by genetic approach also simplifies the interpretation of screened results, which will enable us to explore broader protein candidates in a high-throughput manner. Conventionally, various attempts have been made to screen SUMOylated proteins, such as IP-based proteomic methods (Zhao et al., 2004; Tirard et al., 2012; Filosa et al., 2013) and yeast twohybrid methods (Hannich et al., 2005). IP-based proteomic methods have been widely used for the identification of novel SUMOylated proteins, SUMOylation site, and even for the simultaneous analysis with other modification processes, such as ubiquitination (Hendriks et al., 2014, 2017; Lamoliatte et al., 2014, 2017). The proteomic approach also enabled large-scale analysis of the peptides digested from total immunoprecipitated proteins. Therefore, the IP-based approach has a tremendous potential in the discovery of novel SUMOylated proteins. However, it still has some difficulties in SUMOylation detection. The IP-based proteomic

methods inherently required cell-destructive process to collect SUMOylated proteins, where complete inhibition of deSUMOylation is quite difficult. In addition, since the highly SUMOylated proteins would be preferentially collected by IP, the transiently or infrequently SUMOylated proteins might be missed during the IP process. In contrast, as discussed previously, our method using fluorescence protein reconstitution would effectively detect faint or rare SUMOylation in living cells without being overwhelmed by higher or more frequent SUMOylation in other proteins. The difference in the detected SUMOylation candidates might reflect the diversity in the detectable range of both approaches, which ensures complementary screening of unexplored SUMOylation candidates. Regarding the yeast two-hybrid methods, a cell-disruption process is not required to assess SUMOylation. However, the conventional twohybrid assay required translocation of target proteins into nucleus, which limited the scope of detectable candidates. Recently, multi-well plate assay method based on fluorescence protein reconstitution in yeast cells was reported (Sung et al., 2013). Since the method was free from the limitation in the protein subcellular localization, it greatly expanded the scope of analysable SUMOylated protein candidates in living cells. However, yeast cells only express single SUMO isoform. Considering

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the multiple SUMO isoforms expression in mammalian cells, the isoform-specific substrate preferences depending on the expression pattern of E3 proteins (Tatham et al., 2005; Vertegaal et al., 2006), and the cell type-dependent SUMOylation (Degerny et al., 2005; Ji et al., 2007), screening systems in yeast cells might be inappropriate for the discovery of mammalian SUMOylated proteins. In contrast, our method enabled the detection of SUMOylated proteins in mammalian cells. The detected SUMOylated candidates had a variety of subcellular protein localizations. Though not yet demonstrated, it is possible to apply our approach to explore modification by other SUMO isoforms or analysis of SUMOylation patterns in different mammalian cell-type with different stimuli. Of course, our method also has several disadvantages. The repeated procedure of the FACS sorting and cell incubation would increase the number of same cell clones, resulting in detection of same SUMOylated protein candidates. The linker length (Gly–Gly–Gly–Gly–Ser amino acids in this experiment) would be in some cases constraint for the VC and VN fragments reconstitution, depending on the fused proteins’ conformation, or location of a SUMOylation site. The preparation protocol for VC-Library DNAs generation using restriction enzymes would restrict the scope of insertable genes, since it is impossible to insert genes without the restriction enzyme sites. As further improvements, a more accurate cell sorter for the decrease in the number of the cell sorting and cell incubation cycles, or different linker lengths to tolerate the conformational constraint, and other gene transfer techniques would be beneficial to widen the scope of detectable protein candidates. In conclusion, our new screening method was devised for identifying mammalian SUMOylated proteins, using fluorescence protein reconstitution and a cell sorter. Our method enabled detection of SUMOylation in living mammalian cells without limitation in subcellular protein location. The fluorescent cells that harboured putative SUMOylated library proteins with reconstituted fluorescence proteins could be rapidly distinguished by the fluorescence intensities and automatically isolated by FACS. The SUMOylated protein candidates could be identified by genetic approach. In this method, murine cells (NIH3T3), mouse cDNA libraries, and SUMO2 were used as model for screening

mammalian SUMOylated proteins. Using different SUMO isoforms, mammalian cell types, and cDNA libraries, broader range of mammalian SUMOylated proteins will be explorable under various conditions. Also, by applying stresses such as UV, heat, and osmotic pressure before cell sorting, stress-induced SUMOylated proteins (Tempé et al., 2008) will be screened. By further alterations as described above, our method has a potential for the discovery of numerous SUMOylated proteins that have not been identified, contributing to a deeper insight into the roles of SUMOylation in diverse biological phenomena. Acknowledgements We are grateful for the support from the Japan Society for the Promotion of Science ( JSPS) and the Ministry of Education, Culture, Sports, Science, and Technology (MEXT) of Japan (Grants-in-Aid for Scientific Research S 26220805 to T.O.). We also appreciate the support from the international and interdisciplinary environments of the JSPS Core-to-Core Program ‘Asian Chemical Biology Initiative’. References

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and ubiquitylation crosstalk in human cells using sequential peptide immunopurification. Nat. Commun. 8, 14109. https://doi.org/10.1038/ncomms14109. Lee, J., Lee, Y., Lee, M.J., Park, E., Kang, S.H., Chung, C.H., Lee, K.H., and Kim, K. (2008). Dual modification of BMAL1 by SUMO2/3 and ubiquitin promotes circadian activation of the CLOCK/BMAL1 complex. Mol. Cell. Biol. 28, 6056–6065. https://doi.org/10.1128/ MCB.00583-08. Lei, Q.Y., Zhang, H., Zhao, B., Zha, Z.Y., Bai, F., Pei, X.H., Zhao, S., Xiong, Y., and Guan, K.L. (2008). TAZ promotes cell proliferation and epithelial-mesenchymal transition and is inhibited by the hippo pathway. Mol. Cell. Biol. 28, 2426–2436. https://doi.org/10.1128/ MCB.01874-07. Liu, J., Chen, J., Dai, Q., and Lee, R.M. (2003). Phospholipid scramblase 3 is the mitochondrial target of protein kinase C delta-induced apoptosis. Cancer Res. 63, 1153–1156. Macauley, M.S., Errington, W.J., Okon, M., Schärpf, M., Mackereth, C.D., Schulman, B.A., and McIntosh, L.P. (2004). Structural and dynamic independence of isopeptide-linked RanGAP1 and SUMO-1. J. Biol. Chem. 279, 49131–49137. https://doi.org/10.1074/ jbc.M408705200. Mahajan, R., Delphin, C., Guan, T., Gerace, L., and Melchior, F. (1997). A small ubiquitin-related polypeptide involved in targeting RanGAP1 to nuclear pore complex protein RanBP2. Cell 88, 97–107. Matunis, M.J., Wu, J., and Blobel, G. (1998). SUMO-1 modification and its role in targeting the Ran GTPaseactivating protein, RanGAP1, to the nuclear pore complex. J. Cell Biol. 140, 499–509. Melchior, F. (2000). SUMO – nonclassical ubiquitin. Annu. Rev. Cell Dev. Biol. 16, 591–626. https://doi. org/10.1146/annurev.cellbio.16.1.591. Molli, P.R., Li, D.Q., Bagheri-Yarmand, R., Pakala, S.B., Katayama, H., Sen, S., Iyer, J., Chernoff, J., Tsai, M.Y., Nair, S.S., et al. (2010). Arpc1b, a centrosomal protein, is both an activator and substrate of Aurora A. J. Cell Biol. 190, 101–114. https://doi.org/10.1083/ jcb.200908050. Nagai, T., Ibata, K., Park, E.S., Kubota, M., Mikoshiba, K., and Miyawaki, A. (2002). A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat. Biotechnol. 20, 87–90. https://doi.org/10.1038/nbt0102-87. Ouyang, J., and Gill, G. (2009). SUMO engages multiple corepressors to regulate chromatin structure and transcription. Epigenetics 4, 440–444. Ozawa, T., Sako, Y., Sato, M., Kitamura, T., and Umezawa, Y. (2003). A genetic approach to identifying mitochondrial proteins. Nat. Biotechnol. 21, 287–293. https://doi. org/10.1038/nbt791. Rodriguez, M.S., Dargemont, C., and Hay, R.T. (2001). SUMO-1 conjugation in vivo requires both a consensus modification motif and nuclear targeting. J. Biol. Chem. 276, 12654–12659. https://doi.org/10.1074/jbc. M009476200. Saitoh, H., and Hinchey, J. (2000). Functional heterogeneity of small ubiquitin-related protein modifiers SUMO-1 versus SUMO-2/3. J. Biol. Chem. 275, 6252–6258. Shyu, Y.J., and Hu, C.D. (2008). Fluorescence complementation: an emerging tool for biological

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Dissecting Complex SUMOylation Networks in Humans Ijeoma Uzoma1,2 and Heng Zhu1,2*

9

1Deparment of Pharmacology and Molecular Sciences, Johns Hopkins University School of

Medicine, Baltimore, MD, USA.

2The Center for High-Throughput Biology, Johns Hopkins University School of Medicine,

Baltimore, MD, USA.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.09

Abstract To continue improving our understanding of the physiological impact of SUMO modification in humans at a global level, dissecting enzyme/ SUMO/substrate relationships within the complex SUMOylation network is essential. This effort requires multifaceted proteomic approaches that can capture comprehensive data on SUMO substrates, E3 ligase–substrate specificity, SUMO paralogue specificity, and SUMO-binding proteins. The HuProt™ protein microarray contains > 20,000 purified human proteins, providing an ideal platform for identification of substrates that can be covalently modified by SUMO-1 or -2, as well as proteins that recognize SUMO substrates through non-covalent interactions. Protein microarray studies successfully identified > 2500 covalent SUMO substrates, linked several E3 ligases to hundreds of their protein substrates, established SUMO paralogue preference for substrates and E3 ligases, and identified hundreds of SUMO-binding proteins. These large-scale protein microarray datasets were integrated to construct a multidimensional SUMO network that can be used to connect substrates to upstream E3 ligases and to predict SUMO-dependent protein–protein interactions. By enhancing our knowledge of the architecture and regulation of the enzyme–substrate network, we can strengthen our understanding of the functional outcomes of SUMO modification in human cells.

Complexity of SUMOylation cellular networks In the field of proteomics, systematically charting the biochemical properties and functional interactions of all expressed proteins has been a major undertaking. In large part, generating protein interactomes has focused on developing networks based on protein–protein interactions with technologies, such as the yeast two-hybrid and mass spectrometry (Vertegaal et al., 2006; Golebiowski et al., 2009; Makhnevych et al., 2009; Tammsalu et al., 2014). As the field of proteomics has evolved, we no longer consider the proteome to be comprised of a finite set of genetically encoded proteins. With the discovery of numerous protein posttranslational modification (PTM) systems, which can regulate protein activity, stability, subcellular localization, etc., it has become apparent that the complexity within protein networks is much greater than originally appreciated (Matunis et al., 1998; Ban et al., 2011; Yang et al., 2012). Therefore, new proteomics technologies are required to capture the biochemical information needed to expand our understanding of functional networks. In the field of protein SUMOylation, little is known about the architecture and dynamics of the SUMOylation network. Compared to other enzyme substrate networks, SUMOylation exhibits additional layers of complexity and uncertainty

136  | Uzoma and Zhu

( Johnson, 2004). The E1/E2/E3 enzymatic cascade, poorly defined role of E3s, multiple SUMO paralogues, and low stoichiometry highlight the challenges that must be tackled to construct multidimensional SUMOylation networks. The area of SUMO proteomics has focused on developing high-throughput mass-spectrometrybased approaches that mitigate the inherent and experimental challenges associated with SUMO and mass spec, to enable large-scale studies, which have identified hundreds and thousands of substrates and sites of modification (Matic et al., 2008; Golebiowski et al., 2009; Hendriks et al., 2014; Tammsalu et al., 2014; Hendriks et al., 2017), respectively. While substrate identification and modification site mapping are critical, large-scale mass spectrometry studies alone cannot integrate the many layers of complexity required to illustrate a comprehensive SUMOylation network. If we consider regulation of SUMOylation in the context of single generic target, we know that the assembly of the SUMO E1/E2/E3 enzymatic cascade precedes covalent modification of the target by SUMO1, SUMO2, or SUMO3. In cell-based studies, E3 ligases are considered to be critical for mediating substrate specificity, however, there is lack of consensus on the number of E3 ligases and limited information on the number of substrates linked to each purported E3 ligase. Ultimately, a target may be modified by multiple E3 ligases or by multiple SUMO paralogues, which may result in different downstream functional outcomes. Constructing a SUMOylation network including specific E3 ligase-SUMO-substrate connections would be critical for functional analysis of E3 ligase–substrate relationships and associated cellular functions. Further, consequences of SUMOylation are mediated, in part, by non-covalent SUMO interaction. To enhance the value of the E3 ligase–substrate relationships, comprehensively identifying SUMO binding proteins that can interact with SUMO modified substrates in a SUMO-dependent manner, can also help elucidate the downstream effect of SUMO modification. Based on the complexity describe above, it is clear that within the SUMOylation system there are multiple layers of data which need to be integrated to construct an interaction network that captures

the features that contribute to SUMO specificity and function. Protein microarray technology High throughput technologies strive to provide an unbiased platform for charting enzyme–substrate relationships at the proteome scale. Mass spectrometry and protein microarray technologies are two major platforms that are well suited for proteomic analysis of various PTMs. Mass spectrometry is ideal for high throughput identification of substrates and PTM site mapping of highly expressed proteins from cell extracts, however, determining E3–substrate relationships with mass spec is not a practical approach. Conversely, protein microarrays are an ideal tool that provides a versatile platform for characterizing enzyme–substrate relationships in a parallel, high-throughput manner. Functional protein microarrays Functional protein microarrays are constructed by immobilizing thousands of individually purified proteins in discrete spatial locations on a glass slide (Smith et al., 2005; Tao et al., 2007). Once fabricated, functional protein arrays are ideal for comprehensively interrogating biochemical properties and activities of the collection immobilized proteins. Functional protein microarrays have been successfully employed to chart numerous binary protein interactions including protein–protein, protein–lipid, protein–antibody, protein–small molecule, protein–DNA, protein–RNA, lectin– glycan, and lectin–cell interactions. As the name implies, functional protein microarrays are also a useful tool for identifying substrates or enzymes involved in phosphorylation, ubiquitination, acetylation, and nitrosylation, as well as to profile the immune response (Oh et al., 2007; Foster et al., 2009; Lin et al., 2009; Merbl and Kirschner, 2009; Jeong et al., 2011; Newman et al., 2013). Previous studies have utilized protein microarrays for elucidating post translational modification networks and enzyme–substrate specificity including ubiquitination (HECT ligases), phosphorylation (human kinome), and lysine acetylation (histone

Dissecting Complex SUMOylation Networks in Humans |  137

acetyltransferases) (Lu et al., 2008; Lin et al., 2009; Newman et al., 2013). A major advance in the protein microarray field was the construction of the first human proteome microarray (i.e. HuProt™) containing over 17,000 full-length proteins ( Jeong et al., 2012), which is the world largest available functional protein array (Fig. 9.1). The latest VerIII HuProt™ includes 20,240 human proteins and represents great discovery potential due to the large number of individually purified proteins, covering 80% of the human proteome, miniaturized onto a single glass. With respect to SUMOylation, the protein microarray platform can accommodate the complexity of the features of such as E3 ligases, multiple SUMO moieties, and SUMO binding proteins.

Comprehensive SUMOylation network construction using HuProt array technology As indicated above, construction of a multidimensional SUMOylation network requires multiple layers of high-throughput data: 1

2 3 4

Identification of a comprehensive set of SUMO substrates (mass spec. datasets, studies of individual substrates, protein microarray data) E3 ligase-substrate specificity mapping SUMO paralogue (e.g. SUMO1, SUMO2, SUMO3) specificity SUMO binding (‘non-covalent’ SUMO interaction)

HUPROT VERSION III: 20,240 HUMAN PROTEINS

Figure 9.1  The HuProt Array Ver III. A total of 20,240 full length GST-fusion proteins purified from yeast were printed on glass slides in duplicate. The protein microarray was incubated with anti-GST antibody, and printed spots were identified by probing with an anti-rabbit Alexa Fluor 555 conjugate. The endogenous functional distribution of target proteins is shown.

138  | Uzoma and Zhu

Global analysis of SUMO E3 ligase specificity using a protein microarray platform: A recent study, Uzoma et al. (2018) employed the HuProt™ array and set out to identify a comprehensive set of the SUMO substrates to establish connections between E3 ligases and their targets, and to link the E3 ligases and substrates to the preferred SUMO paralogue (Uzoma et al., 2018). By connecting these features of SUMOylation, the intricate SUMO network will begin to emerge. Assay development The authors developed a novel method to identify E3 ligase specific SUMO substrates, using the protein microarray platform, by carrying out biochemical reactions on the surface of the protein microarray with an optimized mixture of recombinant purified E1, E2, E3 enzymes, and Alexa Fluor-labelled SUMO. To truly query SUMO E3 ligase specificity in a protein microarray substrate discovery assay, the proportions of E1:E2:E3 enzymes must be titrated such that the true activity of the E3 can be assessed, as originally described

by Rogers et al (2003). The in vitro SUMOylation reaction was adapted and optimized for the protein microarray platform using a mixture of E1, E2, fluorescent-tagged SUMO-1 or-2, and ATP (see reaction scheme in Fig. 9.2.) To establish reaction conditions suitable for the E3 ligase activity assay, protein microarray reactions were first conducted on a pilot array using E1 and E2 at increasing concentrations to identify a minimal concentration of enzymes such that only control proteins such as RanGAP1 will be modified (See titration scheme in Fig. 9.3A). By introducing an E3 ligase into a reaction mixture containing minimal concentrations of E1 and E2, the true activity of the E3 ligase can be assessed (Fig. 9.3B). The authors aimed to purify all of the proteins reported to have SUMO E3 ligase activity in the literature (13 were reported at the start of the study). Those that were successfully purified were then tested for activity against their reported substrates. RanBP2, TOPORS, and PIAS1–4 were the E3 ligases that demonstrated the ability to enhance SUMOylation of their substrates when E1 and E2 concentrations were limiting and could not carry-out modification

Figure 9.2 Protein microarray SUMOylation reactions. Protein microarrays were blocked to prevent nonspecific interactions. Chips were incubated with the SUMOylation enzyme reaction mixture containing the E1 and E2 enzymes, fluorescently - tagged SUMO-1 or -2, and ATP. The reaction was carried out at 37°C for 30–60 minutes. To quench the reaction and to remove non-covalent protein interactions, the protein microarrays were washed with denaturing wash buffer containing TBST and 1% SDS. The protein microarrays were scanned with a fluorescent chip-reader and analysed.

Dissecting Complex SUMOylation Networks in Humans |  139

(A)

(B)

Figure 9.3  (A) E1 and E2 enzyme titration on pilot array. (B) E3 ligase SUMOylation activity and specificity on pilot array. A pilot chip was fabricated, composed of 82 SUMO substrates and conjugation enzymes printed in duplicate. Fluorescent controls (e.g. RanBP2) were also printed on the array. Titration reactions were carried out on the surface of the pilot array with increasing concentrations of E1 and E2 enzymes, while the SUMO concentration was constant, to determine the relationship between the enzyme concentrations and the targets that were modified. Substrates labelled on the pilot array are underlined in the main text.

of the substrates, in the absence of the E3. The activity of PIAS4 and RanBP2, on the pilot array, is pictured in Fig. 9.3B. The pilot chip was incubated with increasing concentrations of E1 and E2 enzymes and used to determine the appropriate concentration for high and low concentration controls for the human proteome global screening assays (Fig. 9.3A). At 1X only Uba2 and RanGAP1 are SUMOylated, with the exception of RanBP2, which has autoSUMOylation activity, and fluorescent controls. At 5X RUVBL2, SUMO4, H2B, H3, H4 became modified. At 10X many additional targets are modified including SAE1, NAP1L1, GTF2IRD2, p53, PCNA, and ANXA11. Substrate proteins were spotted on the array in quadruplet. In parallel assays, SUMOylation of these targets was assayed by incubation with high concentrations of E1 and E2 and low (limiting) concentrations of E1 and E2. E3 ligases were added to the low concentration E1 and E2 mixture to determine ligase activity and specificity. PIAS4 and RanBP2 activity is pictured. E3 ligases were able to rescue SUMOylation of targets modified under high E1 and E2 conditions as well as modified E3-dependent targets that were not modified by high concentrations of E1 and E2 alone (Fig. 9.3B).

Global profiling of E3 ligase mediated SUMOylation By employing optimal concentrations of E1, E2, and E3 enzymes, a systematic global E3 ligase-substrate screen was performed using the Human Proteome chip (Fig. 9.4). Protein microarrays incubated with high concentrations of E1 and E2, and SUMO1 or SUMO2, were used as a positive control for SUMOylation. To compare the activities of the six active E3 ligases, low concentrations of E1 and E2, and SUMO1 or SUMO2, were spiked with each E3 ligase. In a parallel reaction, E3s were omitted as a control for ligase activity. Chips incubated with reaction buffer and SUMO1 or SUMO2 served as negative controls. Including all experimental permutations, a total of 18 reactions conditions were carried out. All reactions were performed in triplicate to ensure a high quality, reproducible dataset. For more details on methods see the following papers (Cox et al., 2015; Neiswinger et al., 2016). Global SUMOylation results A total 3640 SUMO1 and SUMO2 substrates were identified (“Layer 1” see Table 9.1 for summary of data). By removing the hits in the high concentration E1 and E2 experiment from the E3 ligase assays,

140  | Uzoma and Zhu

Figure 9.4  Global study design schematic. HuProt™ microarrays were incubated with low concentrations of E1 and E2, plus PIAS1–4, RanBP2, TOPORS, high concentrations of E1 and E2, or no enzyme controls. The concentration of ATP and Alexa-555 SUMO-1 and -2 was constant in all 18 experimental conditions.

Table 9.1  Summary of SUMO E3 ligase substrate data Reaction

Total

Unique

ΨKXD/E motifs

SIMs

Phospho-protein

Kinases

ZINC fingers

High E1/E2 S1

2346

---

194

240

1286

81

80

High E1/E2 S2

1933

---

162

205

1073

72

62

PIAS1 SUMO1

767

184

62

105

445

32

39

PIAS1 SUMO2

853

366

74

99

478

34

43

PIAS2 SUMO1

329

48

37

44

224

12

15

PIAS3 SUMO1

70

0

2

9

47

4

4

PIAS3 SUMO2

1092

470

96

136

601

32

55

PIAS4 SUMO1

3

0

0

1

3

0

0

PIAS4 SUMO2

249

26

38

37

181

10

11

RANBP2 SUMO1

181

33

8

17

116

7

2

RANBP2 SUMO2

99

7

4

7

51

4

1

TOPORS SUMO1

197

46

13

15

111

6

10

TOPORS SUMO2

73

2

5

8

50

7

3

the E3 ligase-specific or -dependent targets were revealed. Following removal of high concentration substrates, 2150 were substrates of E3 ligases, which represents the largest dataset and only the systematic study of E3 ligase substrates (“Layer 3”). Following detailed bioinformatic analysis, the data revealed that in hundreds of cases where proteins were unable to be modified by high concentrations of E1/E2, they were readily modified when the correct E3 ligases was introduced to the low E1/E2 reaction. Substrates that demonstrated E3 ligase-dependent modification were considered ‘unique’ substrates. The number of unique targets for each E3 ligase/SUMO pairing ranged from only 3 to 1092 for PIAS4 SUMO1 and PIAS3 SUMO2,

respectively. Some substrates were modified by multiple E3 ligases, however, for over 1000 substrates identified, modification only occurred under 1 specific condition of the 18 that were carried out. Strong preference for SUMO paralogues was exhibited by PIAS3 (94% SUMO2), PIAS 4 (99% SUMO2), RanBP2 (62.8% SUMO1), TOPORS (73% SUMO1), whereas PIAS1 did not show paralogue preference (“Layer 3”). The specificity and connectivity of SUMO E3 ligase–substrate relationships are illustrated in Fig. 9.5. Dogma in the literature indicates that the presence of the dominant SUMO consensus motif (ΨKXD/E) is a major factor in SUMO substrate lysine modification, however, the motif only was

Dissecting Complex SUMOylation Networks in Humans |  141 FAM177A1

PPM1B

GDI2

PRKCSH

GABPA

HSP90B1

PAGE2

GRB2

DIXDC1

GYG2

CALR

SGCG

CLIC4

FMO5

SH3BGR

GYG1

ASMTL

FGF7

SART3

MTUS2

NASP

GPR119

ATG4C

UACA

C1orf21

UBXN6 TFE3

PIAS4/SUMO2

PIAS4/SUMO1

CRIP1

LOC284297

DBNDD1

PYCR2

DDC

QDPR DNAJB2 FAH

LAMTOR3

ACOX1

NT5C3

FABP1

IL37

ALG13

PGD

FMR1

HSD17B7

ALKBH7

PFKFB3

GEMIN6

BCL2L2

CALCOCO1

GSTA3

GABARAPL2

ATXN7L1

SCP2

MT1X

TPRKB

NCALD C12orf60

NDRG1

TRIM27

BID

URM1

SSBP2

RanBP2/SUMO1

TRIM28 PGRMC2

C9orf9RBM3 SPRR3S100B

ATP6V1E1

C16orf3

CRYZL1

CYGB

GNPDA2 HSD3B2

DCTD

RIPPLY1

RFESD

AMACR

ALDH1L2

ADH6

C6orf195

CACNB3

C9orf95

CD48

CD14

CCBL2

CBLN4

CERS4

CEP250

CENPN

CDV3

ARNTL

BAT1

BTRC

BCL2L11

BHMT2

BCL6

C15orf26

DKK3

CTSL1

DAZAP1

GLTP

FTH1

LOC286016

MAPK1IP1L

PFDN4

PYDC1

PHOSPHO2

STK16

SLC7A1

SIX5

ZNF281

CROCCP2

TPMT

GSTA1

COPZ2RAB4A

S100A8

ANKS1A APBB1 C4orf22 CBS

POP7WDR12

GTF2IMASP1PDCL2 NFKBIA MTL5TBCELSTMN4STAM 4-Sep SDCCAG3 ZNF397 XAGE2 USP5

EEF1A1

EIF6

GRK6

LOC113386

SVOP

CALM2

ASL

LGALSL

TMED6

RIPK2

PRSS1

CAPRIN2

KIAA1598

PSPC1

RPP25

MAP1LC3B

C3orf37

CCT5

CCDC97

DFFA

DCTN2

EEF2K

EEF1B2

CHCHD4

CLUAP1

GMEB1

FEZ1

GLRX3

LRRC23

LOC554202

IFIT3

PBK

PDCD2

P4HB

C6orf115

PHYHIPL

FRMD8 CTBP1

FAM92B

NUDCD2

NPM1

RUVBL1

S100A14

PTK2B

MYL6

MTUS1

PIGP

RGS19

RMI1

RPSA

PIM2

PKNOX1

TERF2IP

SPA17

SMYD3

SEC13

UBA5

TXNDC3

PSCA LOC51136

ABCF3

SLC9A3R1

IL12A

TCP11

ZNF274

SYAP1

SUGT1

SNX16

SCG3

SAE1

ZWINT

WDYHV1

VPS37A

UBE2Q2

TSR2

TOM1

TMOD4

ZNF257

AKR1C4 CSRP3

BDH2

NFATC3

GDPD5

COMMD9

CRTAC1

FGFR4

FAM134B

LRRC42

MGC23270 SND1

ABHD16A

ZNF557

FEZF2

PGM3

LRRC25

PRMT8

ALS2DLEU1 CYTH2CYTH1

CAPZA2 C17orf75

ABHD10

SMOC1

RXFP3 PPP1R7

IGFBP2

IFT81

S100A11

TNS1

PVR SNN

IDS

ACD

FECH

LOC51233

HS3ST1

YWHAZ

PSAT1 S100P

FGR

LOC112703

IGHG4

HPCA

HPD

UCHL3

RWDD3

PPP4R1L

ICAM1

GSN

GTF2H3

GTF2B

RanBP2/SUMO2

CCDC102B APOBEC2

BEGAIN

ERMN

GAS2

NFU1

ZNF333

AGFG1

SLC7A8

Klkbl4 HSPE1

HAND2

GRWD1

C10orf118

ERP27 ZSWIM1

FAM83F LASP1

HABP2

GRIK2

HNRNPD

AMPH

ATP1A3

MCFD2

FCER1G

HSPA2

HAVCR2

GPATCH3

EDG2

ANKHD1 nd

FAM13A

AES

MAP3K7

HSPA13

HAPLN3

H1F0

GOLM1 HCLS1

HNRNPUL1

HOGA1

MT1H

NR2C2AP

MT1F

GALE

ADAM22

SLC30A5

LACTB

HINFP

HEY2

HELQ

EDN3

EBAG9

POLK

CTNNBIP1

EXOSC5

ACTR10

SLC27A2

RPL37A

EFEMP1

EBNA1BP2

POLR2D

YKT6

ATXN3

LYPLA2 PTGES3

HSFX1

PCMTD1 MRPL12

GTF2H5

NAT1 TAGLN

CAMK1D

PSMA5

ROD1

PSMC1

LNX1 LDHAL6A

EIF1AY PLXNA2

PLEKHG2

PLGLB1

SRP9

ZCCHC8

TPM3

THG1L

NIT2

LMO1

HHLA3

HGSNAT

MTERFD3

MSH2

EVL CKS2

ITGB8

PAAF1

ZNF85

FAM189A2

ACSM5 NIPSNAP1

OR2T35

MRPS10

ILKAP

SSBP1

THAP4 FAM175B

ACTB

MSRB3 OGN

PCMT1

MRPS5

PSMA3

TSSC1

SYTL2

SLC25A45

PCOLCE

OR3A3

MTHFD1L

STAT4

SCHIP1

SLC25A24

KLK1

PMP2

OLFML3

PARS2

PCDHGB6

FKBP8

MAGEA2B

PATE2

MKNK1

PRAME

TRIML1

C14orf129

RPA1

QKI KCTD18

KCNV1 KIAA1456 KIAA1143

NLRP2

BCKDK

GRM3

FGF1

FLJ25328

ILK

KCTD17

TCAP

SERPINB8

TSGA10IP

MAPK11 RAB27A

CD83

AXL

GALNS

IGL@

LAS1L

PROS1

SEZ6L2

UBOX5

SKP1

RRAS

NF2

PLAC9

PIN4

PIP5K1B

CDK17

BET1

GSG1L

HAUS6

FSIP1

RABL3

MOCS3

TCEB1

SHMT2

TYW1

ANKRD39 AKR1C1

SLC16A4

NECAB3

PLA2G12A

NMT2

CDK20

BAK1

HEMGN

GBA2

LAMP3

MMP23B

POLDIP2

SLC2A6

SCIN

NDRG2

PNKP

CDK5R1

CD8B

CRX

HIPK4

MAP2K5

KYNU

RAB3IP

POLH

SOX8

TUBA4A

TGFBR2

PTS

NOTUM

NKD2 OLFM1

CDK5RAP3

TTBK2

QARS

MREG

RAB23

PMS2

WDR77

PLSCR4

GNA12

AMD1

NEIL2

PLA2G16 PIR PITHD1

PARVA

BATF3

NECAP1

NRN1L NSMCE4A

NSUN3

NUMBL

UTP18

NBPF3

NCK2

RGS5

NXF2

PARD6A

USF1

GATA5

MAGEC2

KTI12

NADSYN1 MPHOSPH8

SPOCK1

TMEM128

PIN1

SPRY2

GLB1L3

ALS2CL

RNASEK

SCAF4

ARSA

TTLL9

HOMER2

MGC16075

KREMEN2

NCLN

RUNDC3A

ZNF215

TMEM174

SLAIN1

SCGB2A2

CCDC155

CRELD1

HOOK3

FAM102B

MECR

KPNA2

NSUN7

RMND5B

ZNF205

TNFAIP1

ARF4

SLAMF6

ROBO2

RRP9

ARL8B

CCDC130

UBXN2A

TMEFF1

GIMAP6

ALKBH1

SIRT6 RNF41

RNF25

RERGL

PARP11

UHRF1BP1L

RNF113A

PTGS2

VPS11

TLE6 GAPDHS

SCML4

ARL5B

BCL2

WFS1

MTMR2 TLX3

GALNT6

ALG2 SIRT7

SCO2

UBL7

UBE2T

OPTN

SCG2

STARD4

GCFC1

SHARPIN

RNF208

VAC14

HENMT1

PAIP1

GATS

ASGR1

CCM2

CD2BP2

C17orf57

MFAP3L

FUZ

ALB

SF3B3

SCRN2

CD40

CAMK1

AZIN1 TAX1BP3

GAGE1

AKD1

ALKBH8

ATG5

UBE2S

COL6A2

FAM113A

MEMO1

GSTZ1GSTA4

ACBD6 TCEAL1 GABBR1

TLE3

CABP4

HSPA8

PDHX

TCEAL3

C4BPB

ARHGDIG

SESN2

SEPP1 SDR16C5 SEC24C

CARD14

HP

FAM83A

LRRC57

ISG15 TCEAL5

C3orf52

APH1A

APOA1BP

APOA5 ANTXR1 AP3M1

ANKRD5

UCP3

CLEC18C

GNAI2

FAM63A

SUSD4 STK36

C2orf43

C3orf21

C9orf25 C9orf47 C8orf82 C9 C6orf72

C8orf16

CAPN3

CAPN1

UBTD1

CIB1

GPR85

FBRS

MAGEA6 TADA3

TAF15

TACO1 SPATA7 SPINT1 SPON2

SPRR1A

SPTLC1

SRF

SRPRB

SSRP1

ZNF37A

PREPL ANKMY2

CEP290

OR8D4

PCYT1B

C11orf49

DYNLT3

MRFAP1L1

GPRASP2

HLA-DOA

POGZ

TTC16

HK1

MDM2

TUBA3E

CHD1L

NAP1L2

PDXDC1

OR5T1

MLX

NAAA

2-Mar

TMEM237

EIF3E

CMIP

TRIP6

RASSF9

ERC1

C1QTNF9 C20orf4

C18orf1

WBSCR28

WBP5

GTF2IRD2

SUMO2

ZSCAN20

COPS7A

CROT

DOCK7

DMD

GSTM5

C7orf30

FETUB

SPRYD4

RAB17 SSBP4 C11orf9

SLC20A1 EAF1

TOPORS/SUMO1 C1orf212

OTUB1

NAP1L3

NAPB

SMPX

KRT15

SLA

UCK1 DDX19B

HBZ

SENP1

LOC285141

MCM7

GSK3B

IDO1

RAB39B

GALK2

ASAP3

UBE2O

DNAJA2

SGK1

TGM4

PACSIN2 BNIP2

TIMM17A

VPS25

LETM1

ABTB1

STRADB

MAPKAPK3

2-Sep

HSF1

UFM1

EFHC1

XRCC4

PIAS3/SUMO2 WDR25

SPR

MED1 DPP3

YEATS4

PMPCA

BBOX1

TST

LOC113179

ATG4B

RPS6KL1

WDR5B

TRMT6 FAM131A PYCR1

CARD9

WNT5B

WSB1 WWOX

TMEM163

CDC42SE1

PSMB1 LYPLAL1

GSTM4

PSMB6

XRCC6BP1

TMEM126A

LTF

ARL1

THOC1

DNAJB12

TMEM59

RFXANK

DUSP23 TDP1

ASNS

FAM107A CPSF3

TMEM54

RDH11

MAGEA12

SAE2

WWC2

FABP5 CPA3

TMEM40

CELA2A

SNPH

NSUN5P1

PSMA2

ARFRP1

PCBP2

ESYT2 EXTL2

TMEM39B

CLDN11

SLC44A5

NQO1

IGHV4-31

PFKP

NME5

ALDH3B1

ZC3HAV1L

VIT

CORO1C

AKR1C2

NME2 NIP7

ACOT13

HSPB8

ATF6

ETV7

CSAG1 C15orf57

COLEC11 TRIP10

CLIC1

SF3B4

MYO1B

HJURP NUP54 LPAR4

IGKC

RAB24

THUMPD1

ZNF193

CSAG2

COMMD8 TRPM8

CNOT6 CSTF2

ADH5

CAPN6NSDHL

GUK1

CIAPIN1

PPID

APTX

ZNF174 EPHA10

CSNK2B

RBM17

CYB5R2

STAC

LOC374395

GCLM ZFPL1

EIF4H CTNNA3

RBM33 NXF3

TES

MAPK3 EIF4EBP3

CTHRC1

RABGEF1

RCHY1 RBPMS NUP62CL

SERPINB2

PGPEP1

GCK TAF9

PFKM

ZFP28

CTSB RAB7A

S100A6

NUDT3 C1QTNF6

AGPAT2 DNAJA4

ATCAY

COL4A3BP

RABEPK RTP4

SART1 METTL1

ZFAND1

FLII CXorf27

RTCD1

MAT2B

MEPCE

PIEZO1

BTN2A2 CNBP

TNNT2

APOPT1

RAD51

MCCC1 NAA40

MEOX2

PHB

CLTA

TPTE

TRIM69

HNMT

TRMT1

DBN1

MIPOL1 NAA50

MED22

PGM2

C10orf47

AMDHD1

INPP5A ZMYM3 FLJ14107

RAGE RTN2

PEX19

BRK1 CLASRP TRAP1

MPST HRAS

DENND1B

MORN1

CNOT7

VCY

ZMPSTE24

FOXRED1

RAP2B S100A10

METTL21B

OXCT2

OVOL2

CAPS

HAAO RNPEP

METTL7A

IQCK

P2RX7 PPFIBP2

BMPER CLDN6

TRAPPC3

HLA-DMB

PPM1A

FOXRED2 ACSL6

OXNAD1

IRGC DOK3

PNLIPRP2 NARF

MDH1

RARRES1

RXRA

ISCA2 DPPA2

NAPSA

PDIA6 BMP7

EIF4A2 FNDC3B

MGMT PAH

KCNJ13

GPSM1

GPRIN2

DSTN PNLIP

PDE12 C13orf44

UBD CIB3

SCCPDH

ZFYVE9

GPSM3

DPH5

MPI CHURC1

GDI1

FCHSD2

ACP5

PRAP1 KCNMB2

PNMA1

TSPYL6

SOD2

MICU1

HSD17B8

HLA-E

PPBP

C12orf4

S100A13

IDH3G

MUM1 MYL2

DYSFIP1 MRE11A

PDCD6

CHST9-AS1

TRIM44

ZBED1 NAGK

PRKCZ

ANKRD20A5P

CENPM

DEM1

MGC16291

METTL21A

KIAA1147

DNAJB8

RAB37

SDS

WDR69

VSTM2L

ADH1B

PSEN1

WNK1

TRAPPC1

EGFLAM

NDP G6PD

TRMT12

UBE3A

HS1BP3 FAM9C

ANXA11

KCNAB2

AK3

RPLP1TWF2

GOT1

EZH1

TET3 MAGEA4

KIAA1257 WDR31 SUMO1 TDP2

C16orf42

EPC1

CXorf48

H1FOO

RAB5B

SNX8

N.D.

UFSP2

C14orf93

PCK2 ADAL

ERCC3

CRK

SYT17

AKR7A3 WDTC1

PEF1

ARL2BP

CPNE2 HSFY2

DOK1

TUBA1B BRPF1

SPARC SAAL1

FNTB

C9orf103

C19orf10

EIF2S2

CLP1

COPS2

GPAA1

FAM108A1

GFOD2

METTL16

MEF2C

IFI44

NRBP2

ETS1

PDCL

EIF2S1

RAB32

MYL1

CKMT1B

RAB5A

PLEK2

ADSL

COL9A1

NME1

PLK1 PSMA8

SF3B5

ACOT9

DDA1

ITM2C

PDPK1

APEX1

AFAP1L2

ACTR1A

ADD1

ACSL4

ARL4A

ARHGAP15

CAPN2

CAB39

CA12

C9orf150

CCDC51

DOK4

PIPOX

TFF2

SIRPG

ZNF69

WDR1

TRIM16

PRMT2

CARS

CTR9 CAST

BCL2L13

CMBL

DBNL

HSD17B14

LOC440295

ANKRD40

KCTD5

NBPF22P

RABL5

CORO6

COX19

COX7A1

CYP4F11

PLLP

TBC1D21

STUB1

SPAG16

FARSB

FCHSD1

FAM154A

FAM120B

SHD

SERBP1

SDF4

AHNAK2

CDS2

CDCA3

CDC16

DGCR8

DDX41

DDX20

DIRAS2

DNMT3A

C9orf86 MAPK8IP2

DYRK2

KLHL10 PLEKHJ1

ARIH2

LRRC48

SUMO3

ZRANB2

YES1

TRUB1

TK1

POLR3DTFPT DDX4

EEF1D

EP400

EPB42

ELOVL2

EPM2A

EZR

GPI

HOMER3

GFRA1

GGH

ETHE1

CES3

HN1

HMG20A

LGTN

LGALS3

CLCF1

FGF12

LEMD1

CNIH4

CNOT2

FIP1L1

FGFBP1

LECT1

LOC51035

ZMYND19 UGGT2

DTNBP1

CRBN

BOD1P

MDH2

MAGEB4

MCC

MAPK7

JAM2

KBTBD6

JKAMP

KCNIP3

NR4A2

NRGN

OR5AN1

OSCP1

OMG

NHEJ1

MYO3A

MYF6

RGS14

RNASE6

PROSC

PRPF40A

PSTPIP2

PPIH

PON2

PPYR1

IL8

IRF3

IP6K1

HSH2D

KCNK10

KCNS3

P2RX4

PAPSS2

PGRMC1

PGK1

OLIG2

OR2A12

OR10H5

OPRL1

NMRAL1

RPP40

RPL11

RPL27

RAD51AP1

RBFA

TCF4

PHF17

HUS1

IFIT2

PLCD4

TCTN2

TBCD

SUMO4

STXBP6

STAC3

SSX3

SMU1

SMAD3

SKA3

SETDB1

SCD

SCFD1

ZNF280D

ZNF554

ZNF625

ZNF658

ZC3H14

WDR55

VSTM2A

WDFY3

TWSG1

TTC4

TNFRSF14

TNFSF4

THYN1

TMEM8A

GOPC

FBXL5

MED7

IDH3A

PIP

POLA2

SIPA1L2

UBR7

ATP5J

BBS4

FAM131C

IGHG1

RGP1

SYK

CDC123

TOPORS/SUMO2

MAPK10

PIAS2

PAGE5

hCG_1778643 TUBD1

WDR4 TDRD3

MTCP1 PICK1

GSTM3

ZBTB46

MLIP

ARAF

CDKN2D

ASB17

AKAP1

IGHM

KCNAB1

PIAS3

TDG EIF2B3

EYA2

IST1

PLRG1

TARS2

C10orf81

PIAS3/SUMO1

CCIN

CRYM

NR0B2 FAM131B

GCAT

LSP1

SH2B1

MIS18BP1

ATP1B1

GSDMB WDR45

MED8

IFI35

NUB1

NIPA1

EIF3G

GLOD4

L2HGDH

NFKB1

PPP2R1B

SLC35F5

ZNHIT6

ZCCHC7

ZKSCAN3

VIPAR

TPRA1

ZNF655

RUVBL2

RAD54B

THUMPD3

SIRT1

PRSS12

PPP3R2

PTGER3

PPP2R3C

QPCTL

EYA4

P4HTM

SEMA3D PCCA

SGK2

TTLL7 MGEA5

PUF60

SFMBT1

LUM

MAP3K11

LOC196463

MAD2L1 NOL3

EID1

RWDD2B

RAB28

POLR3E

SHMT1

ZCCHC4

SRRM2 RAP1GDS1

SPINT2

SLC22A15

CPA1 CREB3L1

SPIN1

CPB1

RAD51D

PPM1K

OSBPL9

NUF2 CUL4A

KLK7 LPL

FKBP6 ALDH1A1

LAD1 EGLN2

MBIP

LINC00471

CYP2W1

SPATA2L

ALKBH4 SZT2

EPHB3

ACTR6

CLEC7A AP3B1

COL2A1

CETN1

MLF2

SLC3A1 STIP1

KCTD4

ADRA1A

ANKMY1

AK8

ALDH1A3

AGA

AARSD1

CCDC11

CBWD1

CASQ1

ARPC3

ARRDC4

C11orf74

DSN1

DNAJC10

ELL3

CPA4

CREM

CSRP2BP

CSNK2A1

HNRNPAB

GDF3

LOC595101

LZTFL1

LEPREL1

LOC197350

RFC2 COX6B1

C1orf114

DMWD ANKRD50

CHD2

BCS1L

MMP19

SLC25A13

PLCD1

PITPNB

ATF6B

H2AFZ

NUDT9P1

ZXDC

VPS26A

FBP2

FBXW11

FAM40B

MAPK1

MAGEB1

MAP1LC3C

MRM1

GATA1

IDH1

OTUD7B

IGFBP5

KPNA1

KPNA4

MZF1

NHP2L1

MITF

RNF5

QTRT1

RAB3IL1

TEAD4

COG2

IVD

NT5E

ZUFSP

KHK TCN2 ACTRT2

C3orf20

ATP2B3 FAM124A

ZNF446

AMPD2

MMP13

RASSF4

RBBP5

SPHK2

SOX5

SNX9

SMEK1

SLC22A2

ZNF323

ZNF213

WT1

XRCC3

UNC45A

TUBB2B

TTC27

TSTD2

TOR1A

TREX1

MRPL24

RBMS1

CDKN1B

TPT1

ICAM3

PSMB8

TESC

BAG5

PMS2CL

POLR3C

SNX10

ZBTB33

WIPI2

USP53

MAL2 HCFC2 RFX3

MAGOHB RFX5 ERLIN2

HES1

THAP1

DEPDC1B

DUOXA1

HNRNPC

ESCO1

FAM57A OR4C15

DOK6 WFDC6

MFSD2A ADCK4

C19orf43 C12orf5

HORMAD2

DIMT1

ATP6V1B1

GMPS

TRIP13

OR56A4

CDK15

CIDEC

WDR91

GABBR2

FAM194A

VDAC2

MGC16025

RFX6

C17orf49

B9D2

FBXO38 CHEK2

PIAS2/SUMO1

MAX HADHB

RIOK3

ABHD16B

ZNF280A

ACYP1

FBXO25

UQCRFS1

SLC25A40

C14orf169

C12orf41

TOMM40

MFN1

DNAJB4 ADAMTS12

OTUB2

DNAJB14

ATP5D

WDR85

GOSR2

GNAO1

GALNTL2 GABRG1 FAM161B

UBAC1

C12orf44

XPR1

OSR2 C8orf76

C1QB

ABLIM3

C1QTNF1

C18orf25

C11orf54

DDX59

VTA1

C1orf27

BMF

C1orf96

ALCAM

CA10

TNS4

FNDC8

BRCC3

FOXP1

FSD1L

TNIK

FXN

ANKRD13A

C4orf37

SFXN2

SERPIND1

MESDC2

CALCOCO2

NTAN1

P2RY8

RAB33A

COL6A5

COX6A1

CCT8L2

DHX16

CDKN1A

APPL1

EAF2

EPM2AIP1

DCAF5

GPCPD1

ASXL2

EPHA7

CTSC

HBG1

FGL2

GAL3ST1

FPGT

PCYT2

OSBPL11

OPHN1

NACA2

PPIL6

PRUNE2

PTPN2

PLA2G6

ZNF496

ZBTB25

WIPI1

XPNPEP3

TP53

TP53I3

TRAK2

PPM1D

SKIL

SETD4

VGLL2

WDFY2

WBP2NL

UBE2I

GNB3

LRFN5

NOSIP

PDZD11 GSDMC PGBD3

CALHM1

PANX1

KIAA0226L

SKAP2

TLE4 LOC441046

PEX14

LXN

GSDMD

PEPD SDCCAG8

HPX

KCNJ3

RAB14

NIT1

VGLL3 LOC138046

GRAP2

PDIK1L

CCDC106 TIMM10

SDR39U1

EXOC5

HSD3B1

ICAM4

STAT5B

GPN1

GPR182

SEC31A ITPA

OR7E91P

NUDT4

CSTF1

EPHA4

ERCC8

ENG TMED8

SERPINB9

LCAT PIP5K1P1

F2

GLYATL2

METAP2 CXXC5

CASC4

ACCN1

ACRC

ACP1

TPM2

LAPTM5 PRPF4

ACCN4

GPN2

CAPNS2

C3orf32

ACOX3 TMEM133

TP63

OR8D1

APCDD1

CAMK2G

C3orf45 FAM122A

TMEM14C

SHKBP1

ANUBL1

ANO6

FAHD2A

TMIGD2

TSPAN4

C6orf165 C5AR1

FAM19A5 TTC39A

SH3BP5

LAMTOR2

METTL19

DCAF6 CASQ2

GBP2 FAM21A

SH2D3C KRTAP19-7

PRSS35

GPN3

OSBPL10

C20orf26

FLOT2

TNIP2

TSGA13

SIRPB1

PPARA

PRDM1

DCST1 CELA2B

VSIG4 GC

FAM81A

TXNDC8 SHPK

KRT4

C12orf57 ACTN2

ATP6V0A2 ARMCX5

TONSL SLC16A10

KLRG1

PP2D1

PSMD2

ALDH5A1

METTL6 SLC16A1 KLK6

PPARD

KEAP1

NUDT2

PSMD3

ALPL

C16orf59

CDK5

BTG3

MSTO1

C1QTNF7

PDGFD

PDE1A

PCDHGC3

THOC3

EMILIN1

DUSP26 ZNF280B

BEND7

C11orf1

SOCS3

RAB3B

SEH1L

PIAS1/SUMO1

LMOD3

RGS1

TCEAL8

EIF2S3

ERLIN1

ZNF385A

C11orf70 C11orf55

ZMYM5

CDCA4

KLHDC3 SLC26A1

MCM8 RNF145

TDGF1

FADS3

ABI1

ZNF587

ZCCHC12

ZFYVE19

FEN1

MRPS30

KLHL36

MAPRE2 LMO2 RMI2

TECR

GNL3

EXOSC3

A4GALT

DUSP6 ZNF585A YPEL5

COG3 ARHGEF16

ATG16L1

TRIM34

MTFMT

STMN1

LINC00319

RPL36AL

CDKN2B

CLDN7

CNOT8 ARHGAP29 CSRP1

CSNK1D

FBXL8

ZNF771 SLC25A19

TAF7L

NPR2 RQCD1

TDRD1

GFPT2

CDK7

CMTM1

CSRP2

SAMSN1

SAP30BP

S100A3

SLC25A18 KLHDC2

PDIA5

TRIM24

RAB2B

GOLGA5

CNST ARMC8 ARRB2 CLSTN3

SPINLW1

SPDEF

MYEF2

UBTD2

UBE2H

KBTBD12

GPR63

NDUFV1 LDHAL6B RPS21

GET4

COMMD7

CRISPLD1

CROCCP3

APP SLX4

SLC4A1

SLC25A20 ZNF680

PPPDE2

COPB2

COX6B2

FLJ45256

SAT1

IGFBP4

IFT52

MPDU1 KLF4

LOC144097

OGFR

PGBD1

TMEM151A SNX27

GNB1L HAPLN1

PDK4

TOLLIP

TLE1

TMEM74

ARRB1

TBC1D23

GIMAP7

FLJ42258 SRP19

SPEM1

SOD1

MOBKL2A

MORN3

TSSK3 PSME2 MAPK12 MYL3 MYL4

MIA

GHDC

GATAD1

STRADA PHYHD1

SNX3

IFIT1 UBE2G1

MAPK14

HDGF2

CALB1

RAB34

TBC1D7

GK

IFT43

MND1

PTPN1 ISY1

GLTPD1

ARMC1

TBCCD1

STK32B RECK

MPPED2 NPM3

KIAA1967

GLRX5

CINP

AKT1S1

COPE

SUSD2

TAF6L STK40

REC8

ECE2

FHL3 CFHR3

ADI1

SCML1

ATP5H

MRPS23

RAB3C

IKZF1

RUNX1T1

RDX

MRPS18A POLR1D

EIF3I

RCAN1

ISM2 CCDC28A

CHGB

CYTH3

SLC6A18

SF3A3

HDAC8

PELI3

RAB18

SLC9A6

SMYD2

TIMP1 TIMM44

ADHFE1

CXorf21

FAM129A

MPP1

TNFSF18

TMEM106A VNN1 WDFY1

AK2

FAM186B

GMDS CDC20B

CSF3

HARS

SACM1L TBC1D16

ZBTB5

C16orf73 AAMP FBXO44

HDAC6 ACSBG1

DPCD

NDOR1 IKZF2

TBRG4

YWHAE CENPP CDC25B C16orf58

DGKE

HIC2

GSK3A

NARFL

IL10

SAR1B

CDK6

FBXL4

DDOST

FBXO39

MTMR4

NAE1

IL32

RAET1G

TCEB2

GNAI3

BRE C14orf132

KNG1

GPS1

C1GALT1C1 MRTO4

PLK1S1

10-Sep

NPC2

IRF4

RASD2

RBBP7

SLFNL1

CFB

C21orf63

FAM109A MMP1

MUC7

PSMC2

TRPM3

NOC4L

IRF9

INPP5E ITGB1 ITGB3BP

PGM5

RPRD2 DLGAP1

OAS2 MUM1L1

NEK11

MYOT

PPP6C

SH3YL1

ACTA1

NLK

NLGN3 NIM1 NFE2 NEUROD2

MRO

SLC41A3

CDCP1

ATPAF2

GNPTG

IL17RA

KIAA1958 IQUB

NR3C1

ZNF238

TRIM43 PPIL3

STAT5A

RNASEH2A ADAM32 DHDH

RSPH9

CUEDC2

PHEX

ERF

KIAA1539

C1orf64 HNRNPK

SNX1

SNCB

PILRA

ERBB3

DPT ATRX

BRD9

HNRNPR

C21orf91

RFX2

SULT2B1 CPNE6

CLEC3A

DCAF8

LPIN1

C4orf19

SLC35E1

RGNEF

AQP7

ECHS1

IKBKB

CCDC120 DNMT3L RNF40 FAM118A

PSMA1

ODF3L2

USP48

NKIRAS1

MRPL1

IL12RB1

PSMB7 CCT3

MTMR12

SH2D2A RASAL1

NR1D1

NPFF NEK10 LOC554235

RTN4

OTOGL

LOC387793

LST1

LOC91461

PLG CEP85

LUZP1

PAF1 GTPBP3

PLSCR1

LZTS2 KCNMB4 POLR1C

EDNRA KCNJ8

LOC554207

EIF2A KCNRG

POLR3F

EDIL3

EIF2B5

LOXHD1

PIAS1/SUMO2 POC1B

LOH12CR1

PNMA6A

GTF2H2

LRRC14

Figure 9.5  SUMO E3 ligase substrate specificity network from HuProt™ array study. A network showing the connections between each E3 ligase/SUMO paralogue pairing and the modified substrates was generated using Cytoscape. The coloured edges depict the connection to upstream E3 ligase. Many substrates are connected to more than more one E3 ligase/SUMO pairing, thus revealing the overlap and redundancy between E3 ligases.

present in only 20.2–56.62% of the E3 ligases substrates, depending on the reaction pairings. It is critical to note that protein microarray studies do not provide any information about the sites of SUMO attachment therefore, only correlative connections to SUMO consensus motif conjugation can be made. To gain deeper insight into the biological functions SUMOylation may regulate, the authors conducted gene ontology analysis on the total collection of SUMO substrates, as well as the individual substrate sets for each E3 ligase– SUMO pairing. Many of the enriched biological categories were consistent with roles of SUMOylation reported in the literature such as response to stress, DNA damage, DNA recombination, protein localization, and protein transport. Of note, many of the significant molecular function terms were related to enzymatic function (e.g. kinase activity, MAP kinase activity, GTP binding, transferase activity and hydrolase activity). There is emerging evidence for roles for SUMOylation regulating

kinase activity and GTP binding. Phosphorylation was also an enriched biological process within SUMOylation targets. There is an increasing body of literature pointing to potential interplay between the two PTMs on a systems level, therefore the authors investigated the possibility for crosstalk. Crosstalk between protein phosphorylation and SUMOylation A comparative analysis with an outside kinome substrate dataset was conducted (Newman et al., 2013) to ask whether SUMOylation and phosphorylation share a significant number of common targets. Indeed, the analysis revealed that over 1200 proteins were mutual substrates of the two PTMs. Deeper analysis of enrichment of SUMOylation across different kinase families indicated that although kinases, in all groups, were targets of SUMOylation in the screen, the CMGC group showed the strongest enrichment with 19 of 49

142  | Uzoma and Zhu

One major impact of SUMOylation is thought to arise from its ability to temporally regulate the protein–protein interactions of a SUMO target. Isolated examples that demonstrate the phenomenon of SUMOylation mediating the interaction between a substrate and binding partner have been characterized. For example, the interaction between promyelocytic leukaemia (PML) protein and thymine DNA glycosylase (TDG) demonstrates how SUMOylation can promote protein–protein interactions (Takahashi et al., 2005). In this context, both proteins are SUMO modified and have SUMO binding activity. In cell-based studies, PML bound to SUMO1-modified TDG 2.5-fold stronger than to the unmodified form, and TDG associated more strongly with the wildtype form of PML compared to the SUMOylation-deficient mutant. Ultimately, the function of SUMOylation is coordinated by the upstream enzymes regulating covalent SUMO modification and the proteins that specifically interact with SUMO-modified

members modified by SUMO (p-value 4.62E-08) (Fig. 9.6). The mitogen-activated protein kinase (MAPK) family is included within the CMGC family providing an additional layer of evidence of biological relevance of SUMOylation in the MAPK family. Successful cell-based validation of MAPK SUMOylation by E3 ligases identified in the screen demonstrated the validity of the E3 ligase substrate specificity from the human proteome array. Regulation by non-covalent interactions with SUMO It has been proposed that SUMO is a molecular linker that mediates non-covalent interactions between SUMOylated substrates and SUMObinding proteins (Takahashi et al., 2005). At the biophysical level, SUMOylation may alter accessible protein binding surfaces, thereby creating the potential for new physical interactions with other proteins (Geiss-Friedlander and Melchior, 2007).

PRKX PR KY

DDR1

SGK49

TRIB 3

SP TTN EG

NE N K9 NE EK8 NEK6K7 NEK3

MYLK

LCK L FG YN R FY N

TK

C SR S1 YE

EPHA 2 NTR EPHA10 K3 NTRK2 DDR2

RE T PT PT K2B K2

BLK

X BMTK B

K FR

ITK K CS K L MAT AX O3 TYR MET

IGF1R

EP FG HB FR EP 3 F GF 2 HB R1 EP EPH 1 HA A 7 4 EPHA 3

STK17A STK17B NEK11 NEK4 NEK2

AC TR 2

MA PK 1

GC

3 0 PK K3 K1 MA CD 5 AP CDK K7 M P A MA 4 PK8 AD A K M R B CD 9 MAPK11 ST AD 6 K PK R CDK NL MA MAPK14 ST 1 MAPK12 SR OX M APK K 15 K 13 14 MAPK ICMA K 3 8 MAPK6A1 AP 3K M MAP 3K3 K2 1 P L3 CSN K2A2 CLK CDKL5 MA P3K5 N S K4 C CL CDK MA CLK23 GSK3A CLK GSK3B PK1 SR SRPK2 SRPK3 DYRK2 DYRK1B DYRK3 HIPKDYRK4 STK3PRPF4B HIPK 4 6 U 1 ULK2 LK4 KA AUR AURKB ULK3 AURKC PLK1 PLK2 PLK4 PLK3 BRSK2 PRKAA1 SN PRKAA2 RK MARK2 NIM PI TS MARK3 1 NNUAK1 M UA 2 SK TS SIK K2 3 T SK SIK 1 SS 1B 2 K2 A 1 K2 D M M K2 PI 3 M K2 A M C L PI CAK2B G K1 DC MK K2 M CA KK2 CA CAM APK2 CAM K MAP 1 K3 K1 2 MAPKAP ST K1 KG PH E MAPKAPK5 CH L MKNK1 1 IK MKNK2 PD 33 CHEK2 DAPK2 K 2 ST PRKD

AA K1

ABL2 FES

4 HB EP

PTK6

4 FR

STK40

BM PR 2

K TT

K P2 BM GAK 16 STK orf96 C9 PIK3R4

UH M IR GSG TE PINK K1 2 IR AK 1 S BUB1 IRA AK34 LIMK K2 K 2 TP53RK TNN NPR1 I3K SGK19 LRRK1 6 2 MAP3K ILK MAP3K17 MLKL 3 DSTYK MAP3K11 KSR2 RIPK3 ARAF RIPK2 BMPR1B 1 BMPR1A L1 STYK VR R1 AC ACV R1B ACV VR1C TYK2 AC 2 2 K R JA FB ZAP70 TG 2B VR SYK TNK2 AC K TE 3 BB FR ER EG T RB KI LT1 GF F PD FRA G PD

FG

17 CDK K18 CD

NE K1 0

K3 VR

TBCK

CK

1

L2 SCY 3 YL 1 SC YL SC

TKL

1 PK PD

G AD R R AD BK K5 RB 2 K1 N NR RBP BP 2 1 W NK V 1 VR RK TTK1 2 BK 2

CS N CS K1 CS NK G3 CS NK 1G2 1 N E K CS NK 1D 1A CS NK 1L 1A1

CM

PRKG1 DMPK K6 GR

W EE EIF PB 1 EIF 2AK K 2A 2 K1

STE

LATS 1 4 M STK M AST S STK3 R 3 AS 1 T IKB PXKPS6K STK 2A K3 8 T2 L1 32 8L TBK1 KB B MAP2K1 MAP2K5MAP2K2 MAP2K3 MAP 2K7 MAP2K MAP2K6 4 MAP4K2 TA OK MAP4K5 3 STK MST4 1 PA0 STK24 MY PAK1 S STK O3 S TK3 25 PAK2 T T A PA NIK K4 K K6 4 C D K1 CDK10 9 CD CD CD K9 K7 K20 CDK 14 CD K15 C D K1 6

N3 PK

SG K PKN SG 2 1 SG K3 AK K1 T AK 3 T1

STK31

C AG

PRKACG PRKACB ACA RPS6KB2 PRK 1 RPS6KB KA5 RPS6

6KA6 RPS 2 3 6KA RPS S6KAA1 RP K S6 CZ RP PRK RKCI P Q KC PR D KC PR CH K PR

PR KC G PRK CB PRKCA

D C C LK A C 1 MK AM CAMK1 V K4 PN CK CAM K1 CAM K1D G

CAMK

CMGC 4.62E-08 CAMK 9.01E-03 STE 1.12E-01 TK 4.79E-03 TKL 2.47E-02 CK1 2.24E-01 Other 7.51E-03 AGC 8.57E-02

Figure 9.6  Phylogenetic kinase tree overlaid with SUMOylation enrichment. The amino acid sequences of the kinase domains of all human kinase proteins have been annotated by Manning et al. (2002) with the Hidden Markov Model (Tamura et al., 2011). Sequences were collected from kinase.com and built the phylogenetic tree by Mega 5 (Tamura et al., 2011). Kinase families were outlined by distinct colours and the SUMOylated kinases were annotated with a red circle. The enrichment analysis indicated that SUMOylation kinases are significantly enriched in CMGC family (p-value 4.64E-8).

Dissecting Complex SUMOylation Networks in Humans |  143

targets. Deciphering the myriad of functions of SUMOylation relies on systematically linking SUMO-modified targets to downstream SUMOinteracting proteins. Similar to the efforts in the identification of covalent SUMOylation targets, there are challenges to identifying SUMO binding proteins in cell-based systems due to low levels of covalent SUMO modification of targets and high activity of SUMO deconjugating enzymes. Therefore, limited numbers of bona fide SUMO binding proteins have been identified using techniques such as yeast 2 hybrid and MS-pulldowns. Moreover, little is understood about non-covalent recognition of SUMO by cellular proteins or how proteins preferentially recognize SUMO-1 or SUMO-2. To address these global questions, Cox et al. (2017), developed methods using protein microarray technology to systematically identify novel SUMOinteracting proteins (Layer 4). SUMO binding study using HuProt™ array Purified SUMO1 and SUMO2 monomers, as well as SUMO1 trimer and SUMO2 trimer proteins, were used to probe the HuProt microarray. A total of 457 proteins bound to the four SUMO binding probes: SUMO1 monomer – 183 total hits, SUMO1 trimer–197 total hits, SUMO2 monomer – 306 total hits, and SUMO2 timer – 139 total hits. Pairwise comparisons of the hit lists for each SUMO probe revealed that, while there was a large degree of variation in the specificity profile of each SUMO moiety tested, there were subsets of proteins that bound to both SUMO1 and SUMO2, and that 39 proteins bound to all four probes. These results suggest that the protein microarray platform allows for detection of novel SUMO1- or SUMO2-specific proteins but it is also unlikely that the targets are random due to the degree of overlap between the binding targets of each probe. Integration of covalent and noncovalent SUMO networks To construct the multidimensional SUMO network, the authors integrated the systematic, large-scale protein microarray SUMO binding data, SUMO E3 ligase-dependent substrate data, and protein–protein interaction data from publicly

available databases. A total of 2910 SUMOylation substrates, 489 SUMO binding proteins, and 6121 reported protein interactions for proteins that are either SUMOylation substrates or SUMO binding proteins, were input into the network. The integrated dataset was searched for network motifs that were statistically overrepresented which illustrate interaction scaffolds that are relevant to SUMOylation. Specifically, each network is comprised of three proteins, including at least one node that represents SUMO and at least one node represents a SUMO binding protein or a SUMOylated protein. The proteins are connected by edges that represent non-covalent SUMO-binding, covalent SUMOylation, or protein–protein interactions identified from published databases. Analysis of the dataset resulted in over 800 predicted networks which included a SUMOylated protein and a SUMO-binding protein which were previously shown to interact, however, prior to this study, the common association with SUMO was unknown (Fig. 9.7). This network motif suggests that the previously identified protein–protein interactions may indeed be mediated through SUMO, where SUMOylation of the substrate enhances the non-covalent interaction with the SUMO binding protein. The authors investigated a predicted network that consisted of the INO80 chromatin remodelling complex subunits INO80E and TFPT, and SUMO2. According to the protein microarray dataset, INO80E bound to the SUMO2 monomer and the SUMO2 trimer, and TFPT was modified by SUMO2 in the presence the SUMO E3 ligases PIAS1 and PIAS3. Subsequent in vitro and cell-based experiments confirmed that INO80E preferentially bound to SUMO2 versus SUMO1. Generating a SUMO- binding deficient mutant was critical for interrogating the importance of SUMO for the interaction between INO80E and TFPT. Therefore, INO80E constructs with point mutations introduced to the SIM domain, and a SIM domain truncation mutant, were generated to determine whether SUMO2 binding was mediated through the SIM domain. Binding studies with both INO80E SIM domain mutants showed dramatic reduction in the interaction with SUMO2 relative to the wildtype version. Cell-based SUMOylation studies with TFPT were able to recapitulate that TFPT is SUMOylated

144  | Uzoma and Zhu

Figure 9.7 SUMOylation and SUMO binding mediated protein–protein interaction motif. Schematic of an enriched network motif that was identified 828 times in the integrated dataset analysis.

by SUMO2 and that E3 ligases, PIAS1 and PIAS3, can enhance SUMOylation of TFPT. A SUMOylation null mutant was generated by mutating lysine 216 to arginine (K216R), which is necessary for characterizing the importance of SUMOylation for the non-covalent TFPT–INO80E interaction. Now that both the SUMO binding and SUMOylation activity of INO80E and TFPT, respectively, were confirmed in cell based validation studies, it was reasonable to hypothesize that the INO80E SIM may mediate the interaction between INO80E and SUMO2-modified TFPT. To test the validity of the predicted SUMO mediated interaction, a series of in vitro, cell based SUMOylation and binding studies were conducted. First, SUMOylation reactions were carried out to generated mixture of unmodified and SUMOmodified TFPT. The mixed TFPT population was then subjected to binding studies with INO80E in a pulldown study. INO80E was recovered from the mixture and it was determined that INO80E preferentially bound to SUMO2-modified TFPT over the unmodified species. In a similar study, using the INO80E SIM deletion mutant, there was a reduction in the recovery of SUMO2-modified TFPT, however, binding to SUMO1-modfied TFPT was not significantly reduced. The INO80E SIM deletion mutant did not show any difference in the binding to unmodified TFPT compared to wildtype INO80E. Collectively, these data suggest the C-terminal SIM of INO80E is necessary for binding to the SUMO2-modified form of TFPT. Conclusions Protein microarrays enable discovery of SUMO E3 ligase substrates due to the versatility of enzymatic

reactions that can be carried out in parallel to query thousands of proteins per assay. With this method, a novel SUMOylation network was constructed which links E3 ligases and SUMO to specific cellular substrates. In additional to identifying over 2000 E3 ligase substrates, valuable information on SUMO paralogue selectivity and specificity was collected. Bioinformatic analysis of this dataset shed light on the global dynamics of the SUMOylation system related to the number of substrates that contain the SUMO consensus motif, subsets of proteins that are modified by single or multiple E3 ligases, as well as enriched biological function categories that SUMO may regulate. To deepen our understanding of the functional outcomes of covalent SUMO modification, it is critical to identify the proteins that specifically recognize SUMOylated proteins to propagate cellular functions. By using HuProt™ microarrays to identify non-covalent SUMO binding proteins and integrating these data with SUMO E3 ligase-dependent SUMOylation substrates, a multidimensional SUMO data network was developed. When integrated with additional publicly available protein-protein databases, the SUMO network was successfully used to predict hundreds of SUMOdependent protein–protein interactions. Understanding how SUMOylation modulates protein–protein interactions is paramount deciphering the functional role of the modification globally. Here, we have described how protein microarray technology has greatly contributed to the construction of an informative, multidimensional SUMOylation network. By taking a multilayer, multidisciplinary, network construction approach, the fine-tuned functions of SUMOylation will begin to emerge.

Dissecting Complex SUMOylation Networks in Humans |  145

References

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Makhnevych, T., Sydorskyy, Y., Xin, X., Srikumar, T., Vizeacoumar, F.J., Jeram, S.M., Li, Z., Bahr, S., Andrews, B.J., Boone, C., et al. (2009). Global map of SUMO function revealed by protein-protein interaction and genetic networks. Mol. Cell 33, 124–135. https://doi. org/10.1016/j.molcel.2008.12.025. Manning, G., Plowman, G.D., Hunter, T., and Sudarsanam, S. (2002). Evolution of protein kinase signaling from yeast to man. Trends Biochem. Sci. 27, 514–520. Matic, I., van Hagen, M., Schimmel, J., Macek, B., Ogg, S.C., Tatham, M.H., Hay, R.T., Lamond, A.I., Mann, M., and Vertegaal, A.C. (2008). In vivo indentification of human small ubiquitin-like modifier polymerization sites by high accuracy mass spectrometry and an in vitro to in vivo strategy. Mol Cell Proteomics 7.1, 132–144. Matunis, M.J., Wu, J., and Blobel, G. (1998). SUMO-1 modification and its role in targeting the Ran GTPaseactivating protein, RanGAP1, to the nuclear pore complex. J. Cell Biol. 140, 499–509. Merbl, Y., and Kirschner, M.W. (2009). Large-scale detection of ubiquitination substrates using cell extracts and protein microarrays. Proc. Natl. Acad. Sci. U.S.A. 106, 2543–2548. https://doi.org/10.1073/ pnas.0812892106. Neiswinger, J., Uzoma, I., Cox, E., Rho, H., Jeong, J.S., and Zhu, H. (2016). Posttranslational modification assays on functional protein microarrays. Cold Spring Harb. Protoc. 2016,. https://doi.org/10.1101/pdb. prot087999. Newman, R.H., Hu, J., Rho, H.S., Xie, Z., Woodard, C., Neiswinger, J., Cooper, C., Shirley, M., Clark, H.M., Hu, S., et al. (2013). Construction of human activity-based phosphorylation networks. Mol. Syst. Biol. 9, 655. https://doi.org/10.1038/msb.2013.12. Oh, Y.H., Hong, M.Y., Jin, Z., Lee, T., Han, M.K., Park, S., and Kim, H.S. (2007). Chip-based analysis of SUMO (small ubiquitin-like modifier) conjugation to a target protein. Biosens. Bioelectron. 22, 1260–1267. Rogers, R.S., Horvath, C.M., and Matunis, M.J. (2003). SUMO modification of STAT1 and its role in PIASmediated inhibition of gene activation. J. Biol. Chem. 278, 30091–30097. https://doi.org/10.1074/jbc. M301344200. Takahashi, H., Hatakeyama, S., Saitoh, H., and Nakayama, K.I. (2005). Noncovalent SUMO-1 binding activity of thymine DNA glycosylase (TDG) is required for its SUMO-1 modification and colocalization with the promyelocytic leukemia protein. J. Biol. Chem. 280, 5611–5621. Tammsalu, T., Matic, I., Jaffray, E.G., Ibrahim, A.F.M., Tatham, M.H., and Hay, R.T. (2014). Proteome-wide identification of SUMO2 modification sites. Sci. Signal. 7, rs2. https://doi.org/10.1126/scisignal.2005146. Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., and Kumar, S. (2011). MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol. 28, 2731–2739. https://doi. org/10.1093/molbev/msr121. Uzoma, I., Hu, J., Cox, E., Xia, S., Zhou, J., Rho, H.S., Guzzo, C., Paul, C., Ajala, O., Goodwin, C.R., et al. (2018). Global Identification of Small Ubiquitinrelated Modifier (SUMO) Substrates Reveals Crosstalk

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between SUMOylation and Phosphorylation Promotes Cell Migration. Mol. Cell Proteomics 17, 871–888. https://doi.org/10.1074/mcp.RA117.000014. Vertegaal, A.C., Andersen, J.S., Ogg, S.C., Hay, R.T., Mann, M., and Lamond, A.I. (2006). Distinct and overlapping sets of SUMO-1 and SUMO-2 target proteins revealed by quantitative proteomics. Mol. Cell Proteomics 5, 2298–2310.

Yang, F., Yao, Y., Jiang, Y., Lu, L., Ma, Y., and Dai, W. (2012). Sumoylation is important for stability, subcellular localization, and transcriptional activity of SALL4, an essential stem cell transcription factor. J. Biol. Chem. 287, 38600–38608. https://doi.org/10.1074/jbc. M112.391441.

TULIP: Targets of Ubiquitin Ligases Identified by Proteomics Román González-Prieto* and Alfred C.O. Vertegaal*

10

Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, the Netherlands. *Correspondence: [email protected] and [email protected] https://doi.org/10.21775/9781912530120.10

Abstract Ubiquitin is a small protein that can be attached to thousands of substrates via an enzymatic cascade involving an activating enzyme (E1), a conjugating enzyme (E2) and a ligase (E3). Over 600 different human E3 ligases have been found. The identification of specific targets for ubiquitin E3 ligases is challenging. So far, most of the approaches aimed to identify E3-specific substrates rely on indirect evidence. Here, we would like to introduce TULIP (Targets of Ubiquitin Ligases Identified by Proteomics) methodology, which enables the identification of ubiquitin E3-specific targets in a direct manner. The rationale behind this strategy is that upon construction of a linear fusion between an E3 of interest and ubiquitin, this ubiquitin moiety will be employed by the linked E3 to modify its substrates. Subsequently, trapped substrates are purified in denaturing buffers to remove non-covalent interactors. Starting from the cDNA sequence of an E3 ligase of interest and finishing with the identification of the ubiquitination targets by mass spectrometry-based proteomics, the whole process takes between 4 to 6 weeks. The description of the methodology includes a discussion of potential pitfalls and specific recommendations. Introduction The ubiquitination cascade is performed by the so-called E1, E2, and E3 ubiquitination enzymes. While there are only two E1 enzymes, and up

to several dozens of E2 enzymes, E3 enzymes, which are the ones that provide target selectivity, are counted by hundreds (Skaar et al., 2014). The number of ubiquitination targets reaches several thousands. The development of mass spectrometry-based proteomics has allowed to identify a huge amount of both ubiquitination targets and acceptor lysines within the target. So far, using state-of-the-art mass spectrometry equipment, more than 40.000 ubiquitination sites have been identified at endogenous levels (Akimov et al., 2018). However, identifying substrates for specific E3 ligases is still challenging. Mainly because most of the strategies for determining E3-specific targets rely on indirect evidence. One indirect approach is the purification of the total ubiquitin proteome after knockdown of a specific E3 ligase and subsequent identification by mass spectrometrybased proteomics. Searching for differences in the ubiquitination status of different proteins after knockdown of a specific E3-ligase tends to fail due to the complexity of the ubiquitin proteome and the fact that a specific target can be modified by multiple E3-ligases in a redundant manner. Moreover, E3 ligase cascades exist, therefore, indirect approaches are unable to link E3-ligases to substrates directly. The TULIP methodology provides a workflow for the determination of E3-specific ubiquitination targets in a direct manner. It is based on a previously described technique, termed UBait (O’Connor

148  | González-Prieto and Vertegaal

et al., 2015). The rationale behind this technique is that if a linear fusion between a specific E3 and ubiquitin is made, the E3 will be prone to use this ubiquitin to conjugate it to its ubiquitination target, leaving the E3 covalently bound to its target, allowing the subsequent purification of the E3, together with its ubiquitination target, which can be identified using mass spectrometry-based proteomics. The main pitfall of the UBait approach was that the purification of the conjugates was based on epitope–antibody interaction, which excluded the possibility of using harsh denaturing buffers. Therefore, it was unable to distinguish between ubiquitination targets and other potential strong interactors of the E3. TULIP methodology employs 10xHIS Nickel-based purification, which allows the use of very harsh denaturing buffers, solving this drawback. Moreover, TULIP uses lentiviral-based inducible constructs, which allows the generation of stable cell lines and the modulation of the expression levels, avoiding overexpression artefacts. Note that a stretch of histidines can mimic a nuclear localization signal, therefore, it is important to verify the subcellular localization of your tagged E3. The UBait approach was initially designed to identify targets of HECT-type E3 ligases. Employing the TULIP methodology, we showed that this strategy was also useful for the identification of targets for RING-type E3 ligases (Kumar et al., 2017). In this chapter, we explain the TULIP methodology step-by-step to enable the reader to determine the specific targets of an E3-ligase of interest in a direct manner. TULIP methodology The TULIP methodology comprises the whole process from inserting a given ubiquitin E3-ligase of interest (E3OI) in the TULIP–expression construct until obtaining a list of putative ubiquitination targets for such an E3OI. The whole process can be divided in five steps: (1) generation of the TULIP cell lines, (2) cell culturing and lysis, (3) TULIP conjugates purification, (4) concentration and trypsin digestion, and, finally, (5) the identification of TULIP conjugates by mass spectrometry-based proteomics (Fig. 10.1). The whole process can take between 4 to 6 weeks, depending mainly on the

time required to grow the cells in sufficient amount once the cell lines are generated. Generation of the TULIP cell lines The TULIP methodology relies on Gateway® cloning. This makes the process very fast and straightforward. Generation of a Gateway™ donor construct To perform Gateway® cloning, you need to have your E3OI in a Gateway® entry clone. In case you already have it available, you can go directly to the section ‘Generation of the TULIP constructs’. However, in case you do not have your E3OI of interest available in an appropriate entry clone, you will have to obtain it. There are several repositories where different ORFs can be purchased in different pDONR plasmids, such as DNASU (Seiler et al., 2014) or the CCSB Human ORFeome project (Lamesch et al., 2007) which in version 7.1 covers 18,414 ORFs. You may also want to construct a Gateway® donor plasmid yourself using your cDNA of interest. In that case you will have to amplify your cDNA by PCR using primers with the specific Gateway sequences to perform a BP recombination reaction. For this purpose, you can use the Gateway™ BP Clonase™ II Enzyme mix according to vendor instructions. Note that in every case your donor plasmid should contain the ORF from your E3OI without stop codon in order to allow the linear fusion with ubiquitin. Moreover, in principle, the ubiquitin attached to your E3OI can also be used by another ubiquitin E3-ligase to modify its targets, therefore, additionally, you may also want to include a catalytically dead mutant, or another kind of mutant, to differentiate your E3OI-specific targets from unspecific TULIP conjugates due to the use of the attached ubiquitin moiety by another endogenous E3-ligase. Generation of the TULIP construct Once you have your E3OI into a Gateway® donor plasmid. You simply have to perform an LR reaction using the Gateway™ LR Clonase™ Enzyme mix between you E3OI entry clone and the TULIP plasmids according to vendor instructions. The

Targets of Ubiquitin Ligases |  149

A

B

C

LTR’

ro Pu

R LT ’

tet:

D

E

F

Puromycin selection

Culture cells

24h

Induce with doxycycline

G

H

I

Ni-NTA purification

Lyse

J

K

i

Im

L

e

ol

z da

Ni-NTA

Ni-NTA Trypsin

Elution

LC-MS/MS

Figure 10.1 Workflow of the TULIP methodology. Once you have obtained your TULIP plasmids (A), use them to produce lentivirus (B) and subsequently infect cells (C). Use puromycin to select the TULIP construct containing cells (D–E). After puromycin selection, grow your cells in sufficient amounts to secure sufficient material for mass spectrometry analysis (F). Induce the expression of the TULIP construct with doxycycline (G), and lyse the cells (H). TULIP conjugates are then purified with Ni-NTA beads (I) and eluted with imidazole (J). After trypsin digestion (K), peptides are identified by LC–MS/MS.

150  | González-Prieto and Vertegaal

TULIP plasmids (Kumar et al., 2017), pTULIP and pTULIP-ΔGG can be freely obtained from our research group on request. TULIP plasmids contain the Gateway® cloning cassette under the control of the TRE promoter, followed by a linker containing 10xHIS and activated ubiquitin. Additionally, they contain the rtTA-VP16-2A-puro under control of the PGK1 promoter (Fig. 10.2). Therefore, they can be used as an all-in-one doxycycline-ON system and enable selection of infected cells by puromycin. Relevant sequences are flanked by LTR repeats, which allows the packaging into lentiviral particles. TULIP plasmids are large and quite unstable due to their lentiviral nature. The presence of viral LTR repeats makes them prone to recombination and lose the E3OI sequence. Therefore, we recommend the utilization of STBL2™ competent cells (ThermoFisher Scientific) for the generation, expansion and maintenance of the constructs. All the required material to generate TULIP constructs are listed in Table 10.1. Generation of the stable-inducible TULIP cell lines The TULIP plasmids are third generation lentiviral, so in principle, any strategy used to generate third generation lentiviral particles should work in order to generate TULIP-expressing cell lines. Nevertheless, in this book chapter, we will describe the

HIV-1 Psi

method we employ to produce TULIP constructcarrying lentiviral particles. The whole process takes 5 days: The DNA used to transfect cell must be of high quality and purity. To secure that these requirements are met we use the MAXI Prep Kit from Qiagen. Kits from other vendors should also be fine if similar quality standards are met. Polyethylenimine (PEI) should be diluted in water at 1ml/ml final concentration and pH adjusted to 7.4 with HCl. To reach maximum transfection efficiency, PEI solution should be frozen/ thawed at least 10 times before use. Day 1: • Seed HEK293T cells in a T175 flask at 30% confluency. As culture medium use 16  ml DMEM + 10% Fetal Bovine Serum (FBS) total culture volume. Day 2: • Prepare sterile filtered 150 mM NaCl • Prepare transfection mixture in 2ml sterile filtered 150 mM NaCl: –– pMD2.G (VSV-G envelope) 7.5 µg –– pMDLg-RRE (gag/pol) 11.4 µg –– pRSV-REV 5.4 µg –– TULIP plasmid 13.7 µg • Vortex the mixture for 10 seconds • Add 144 µl of PEI solution • Vortex the mixture for 10 seconds • Let the mixture stand for 15 minutes at room temperature. • Add the mixture to the cells. Day 3: • Replace the culture medium for fresh DMEM + 10% Fetal Bovine Serum + penicillin/ streptomycin.

TULIP plasmid 9630 bp

Figure 10.2  Map of the TULIP plasmid.

Day 4: • Seed your cell line to transduce at 10% confluency in a 15 cm dish (2 million cells). Day 5: • Collect the cell culture medium in a 50 ml tube. • Centrifuge 5 minutes at 500 g. This will pellet any debris of floating cells. • Pass the medium through a 0.45 µm syringe filter. The 0.45 µm filter will let the viral particles

Targets of Ubiquitin Ligases |  151

Table 10.1  Reagents required for the construction of a TULIP cell line for a given E3OI Material

Vendor

Catalogue number

ORF in pDONR

DNASU; CCSB Human ORFeome Collection

Depending on ORF

Gateway™ BP Clonase™ II enzyme mix

Thermo Fisher Scientific

11789020

Gateway™ LR Clonase™ enzyme mix

Thermo Fisher Scientific

11791019

TULIP plasmids

Freely available from Vertegaal lab on request

MAX Efficiency™ Stbl2™ competent cells

Thermo Fisher Scientific

10268019

DNA Maxi Kit

Qiagen

12163

Lentiviral packaging plasmids

Addgene

#12259 #12251 #12253

PEI (Polyethylenimine), linear, MW ≈ 25,000

Polysciences

23966

DMEM, high glucose

Gibco

11965–092

Fetal bovine serum

Gibco

10500064

Penicillin-streptomycin

Gibco

15140122

Acrodisc syringe filter 0.45 µm

Pall Corporation

PN4184

Polybrene, hexadimethrine bromide

Sigma-Aldrich

H9268

Puromycin dihydrochloride

CalBioChem

540411

HIV Type 1 p24 antigen ELISA

ZeptoMetrix Corporation

0801200

pass through, while any remaining debris from cells will stay in the filter. Take a small aliquot (≈ 20 µl) to determine viral content. Add Polybrene to your viral suspension at 8 µg/ ml. Remove the culture medium from the cells to transduce and replace it for the 16ml of viral suspension with Polybrene. NOTE: You may want to titrate your viral concentration yourself by using a p24 ELISA test (ZeptoMetrix Corporation) and adjust your amounts accordingly. In our hands, we usually obtain a viral protein p24 concentration on average of 200 ng per ml). This corresponds to a Multiplicity of Infection (MOI) of 2 for 4 million HeLa cells.

Day 8: • Replace the medium for fresh DMEM + 10% FBS + Pen/Strep + puromycin 3µg/ml. In order to remove dead cell debris, you can keep culturing and sub-culturing your newly generated TULIP cell lines.

Day 6: • Remove the viral suspension medium and replace it with fresh DMEM + 10% FBS + pen/ strep.

Cell culture and cell lysis Once you have obtained the TULIP-expressing cells for your E3OI, you need to grow your cells in sufficient amounts to enable purification of sufficient TULIP conjugates for identification by mass spectrometry analysis. In our previous project to identify targets of the SUMO Targeted Ubiquitin-Ligase RNF4 (Kumar et al., 2017), we lysed five subconfluent 15 cm dishes of U2OS

• • • •

Day 7: • Add puromycin at 3 µg/ml to the transduced cells. Non-transduced cells will die while transduced cells will keep growing.

TULIP plasmids encode puromycin resistance as selection marker. Multiplicity Of Infection (MOI) and puromycin concentration required to obtain TULIP-expressing cells might differ from one cell line to another. In our hands, a MOI of 2 and puromycin concentration of 3µg/ml was sufficient to secure TULIP construct-expressing U2OS cells. We recommend selecting in parallel a nontransduced negative control to check for efficient puromycin selection.

152  | González-Prieto and Vertegaal + Doxyclycine

2 days

subculture

Confluent TULIP cells 15 cm ∅ dish

5 x 15 cm ∅ dishes (15 % confluency)

24 h

5 x 15 cm ∅ dishes (40-60 % confluency)

Scrape cells

Figure 10.3  An example for a scheme of growing cells for TULIP. Starting from a confluent 15 cm diameter dish of a U2OS-TULIP cell line, subculture it at 15% confluency in five 15 cm diameter dishes. After 2 days of culturing, plates will be around 50% confluency. Add doxycycline for 24 hours to induce the expression of the TULIP construct and then lyse the cells.

cells (circa 100 × 106 cells), which were induced with 1 µg/ml doxycycline 24 hours prior to lysis (Fig. 10.3).

111 -

71 -

IP UL

-T OI

E3OI-TULIP + Conjugates

210 -

E3

kDa

E3

OI

-T

UL

IP

-∆ GG

Validation of the TULIP cell line Prior to performing large-scale cell culturing to prepare samples for mass spectrometry-based proteomics, you may want to check the expression and functionality of your TULIP construct, which can be visualized by immunoblotting. A functional TULIP construct will present a smear up from the

E3OI-TULIP E3OI (endogenous)

Figure 10.4  Visualization of functional TULIP and TULIP–∆GG constructs by immunoblotting. Functional TULIP construct will produce a smear up from the TULIP construct band corresponding to high molecular weight TULIP conjugates.

TULIP construct band which corresponds to the TULIP construct covalently attached to its target proteins (Fig. 10.4). We recommend titrating the doxycycline concentration for the induction of expression in a way that your TULIP construct is expressed at near to endogenous levels of your E3OI. This way you may avoid overexpressioninduced artefacts, although overexpression might also lead to a higher recovery of TULIP conjugates on purification. Consider that the presence of a ubiquitin moiety at the C-terminal part of the TULIP construct tends the TULIP construct to be more susceptible to poly-ubiquitination and subsequent degradation by the 26S proteasome. Therefore, in some cases, proteasome inhibition will be required in order to detect the TULIP constructs by immunoblotting or mass spectrometry-based proteomics. An efficient proteasome inhibitor is MG132 (Sigma-Aldrich, Cat. No. C2211). We use MG1232 at 10 µM concentration. The length in time of the treatment may vary from one E3OI– TULIP construct to another. Culturing and inducing your TULIP cell line At this point you just need to grow your cell line in large scale to perform your purification. five confluent or near to confluent 15cm dishes should be a sufficient amount. Scaling up even more will increase the amount of TULIP conjugates purified and subsequently increase the identification rate of peptides by mass spectrometry.

Targets of Ubiquitin Ligases |  153

Add doxycycline at the concentration determined in previous step 24h before harvesting the cells. In case you need to inhibit the proteasome, do it after 24h of doxycycline induction and lyse the cells after the treatment. Additionally, while the TULIP cells are growing, you can prepare stock solutions (Table 10.2)

that you will need to prepare the lysis buffer and the buffers needed for the HIS- purification (Table 10.3). It is highly important that all your solutions are detergent-free. Therefore, use clean glassware or new and clean 50ml polypropylene tubes. In many institutions, the glassware is cleaned in central facilities, and, although apparently clean, it may contain

Table 10.2  Suggested stock solutions for HIS- purification Stock solution

Notes

6 M Guanidine-HCl pH 8

The pH of the solution should be very carefully adjusted, a pH slightly over 8 (8.1 and above) will result in unspecific binding of proteins that will compete with the HIS-tagged proteins. pH below 8 (7.5–7.7) will produce a cleaner pulldown but may also result in a reduced yield.

1 M Tris-HCl pH 8

To be used in wash buffer 2.

1 M Tris-HCl pH 7

To be used in elution and urea buffers.

1 M Tris-HCl pH 6.3

To be used in wash buffers 3 and 4.

1 M NaH2PO4 1 M Na2HPO4

Solution precipitates at room temperature, keep at 37°C.

9 M urea

Prepare fresh on the same day as performing the HIS-pulldown.

5 M Imidazole pH 8

To be used in Guanidine lysis buffer and wash buffers 1 and 2. Set the imidazole solution to pH 8 using concentrated HCl.

5 M Imidazole pH 7

To be used in wash buffer 3 and elution buffer. To reach pH 7, the imidazole must be dissolved entirely in 6M HCl. The process is very exothermic, so it should be performed in clean glassware, very slowly, while mixing and on preferentially on ice.

10% SDS

To be used in the SNTBS lysis buffer.

10% NP-40

To be used in the SNTBS lysis buffer.

All stock solutions should be kept in clean glassware free of any soap contamination.

Table 10.3  Buffers required for the HIS- purification Buffer

Recipe

PBS

150 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4

SNTBS lysis buffer

2% SDS, 1% NP-40, 50 mM TRIS pH 7.5, 150 mM NaCl

Guanidine lysis buffer

6 M Guanidine-HCl, 93.2 mM Na2HPO4, 6.8 mM NaH2PO4, 10 mM Tris, pH 8.0. sterile filtered

Wash buffer 1

6 M Guanidine-HCl, 93.2 mM Na2HPO4, 6.8 mM NaH2PO4, 10 mM Tris, 10 mM imidazole, 5 mM 2-mercaptoethanol, pH 8.0

Wash buffer 2

8 M urea, 93.2 mM Na2HPO4, 6.8 mM NaH2PO4, 10 mM Tris, 10 mM imidazole, 5 mM 2-mercaptoethanol, pH 8.0

Wash buffer 3

8 M urea, 21.6 mM Na2HPO4, 78.4 mM NaH2PO4, 10 mM Tris, 10 mM imidazole, 5 mM 2-mercaptoethanol, pH 6.3

Wash buffer 4

8 M urea, 21.6 mM Na2HPO4, 78.4 mM NaH2PO4, 10 mM Tris, 5 mM 2-mercaptoethanol, pH 6.3

Elution buffer Urea buffer

7 M urea, 58 mM Na2HPO4, 42 mM NaH2PO4, 10 mM Tris, 500 mM imidazole, pH 7.0 8 M urea, 93.2 mM Na2HPO4, 6.8 mM NaH2PO4, 10 mM Tris, pH 8.0

PBS and lysis buffers can be prepared and stored indefinitely. Wash, elution and urea buffers should be prepared fresh on the same day of the HIS- pulldown.

154  | González-Prieto and Vertegaal

traces of soap or other polyethylene glycol (PEGs) polymers. We recommend flushing the glassware twice with deionized water, twice with methanol and then twice with deionized water again, in order to remove any PEG traces prior to use. The presence of PEG will reduce or disable the identification of TULIP conjugates by mass spectrometry. Lyse the cells • • • • • • • • • •

• • •

•

Place your cell culture dishes on ice. Remove culture medium. Wash two times with 10 ml of ice-cold PBS. Add 2 ml of ice-cold PBS to each plate. Scrape the cells. Transfer cell suspension to a 50 ml centrifuge tube. Centrifuge 5 minutes at 300 g Remove supernatant. Resuspend the cells in 5 ml ice-cold PBS. At this point you may transfer your cells to a 15 ml tube to facilitate handling. Take a 50 µl aliquot to a 1.5 ml microcentrifuge tube. Centrifuge 2 minutes at 1000 g, remove supernatant and add 100 µl SNTBS lysis buffer. This will serve as input sample. Centrifuge 5 minutes at 300 g Remove supernatant. Carefully add guanidine lysis buffer (10–15 pellet volumes), while vortexing. As a guideline, 2 ml of guanidine lysis buffer per full 15 cm plate should be sufficient. After that, close the tube and agitate violently to make sure all the cells are lysed. Snap freeze in liquid nitrogen and store indefinitely at −80°C. Storing at −20°C will

cause the samples to thaw and proteins will crash out of solution. Once lysed, samples can be stored at −80°C while performing biological repeats. Later on, you will need at least three replicates for proper statistical analysis of the data. Doing four or five replicates is recommended in order to strengthen the statistical power of the analysis. The different biological repeats should be processed together in the next step in order to diminish experimental variability in the handling of the different samples. HIS-pulldown to purify TULIP conjugates For the purification of the TULIP conjugates we follow our previously described protocol to purify SUMO conjugated proteins (Hendriks and Vertegaal, 2016). Materials required for the subsequent steps are listed in Table 10.4. Equalize samples • Take out all the lysates from the −80°C freezer and let them thaw at room temperature. Rotating on a roller or a rotating wheel is recommended. • Sonicate the lysates in order to homogenize them using a microtip sonicator. –– 2 to 4 pulses of 10 seconds at approximately 30 W should be sufficient to shear the DNA and reduce the viscosity of the lysate until it becomes pipettable with a 20 µl tip. Leave the samples at room temperature between rounds of sonication to avoid overheating.

Table 10.4  Other material needed Material

Vendor

Catalogue number

Pierce BCA protein assay Kit

Thermo-Fisher Scientific

23227

Microtip Sonicator

Not relevant

NiNTA beads

Qiagen

30210

LoBind tubes

Eppendorf

Z666505-100EA

UltraFree 0.45 µm centrifugal filter

Millipore

UFC30HV00

Vivacon 500, 100,000 MWCO spin column

SartoriusStedim

VN01H41

Sequencing grade modified trypsin

Promega

V5111

Empore SPE C18 disks, 47 mm

Sigma-Aldrich

66883-U

Targets of Ubiquitin Ligases |  155

Clean the sonicator microtip with 70% ethanol between samples. • Measure protein content of the samples using a BCA kit following vendor instructions. As a guideline, for the Pierce BCA protein assay kit, putting 3 µl of lysate in 50 µl of the BCA mix should provide a value in the linear range of measurement. • Equalize samples to the protein content of the lowest concentrated sample. –– For this, remove lysate from the tube and add back Guanidine lysis buffer. It may be the case that the protein content of one of the samples is very different (in terms of low amount) from the rest of the samples. In that case, first, repeat the BCA measurement to confirm this result. In case the difference stays true, discard this replicate, snap freeze all the samples again and obtain an extra biological replicate. Then repeat the process again. HIS-pulldown • Add 2-Mercaptoethanol to a final concentration of 5  mM and Imidazole pH 8 to a final concentration 50 mM. • Prepare 20 µl of NiNTA beads (40 µl of 50% slurry) per ml of lysate. Equilibrate the beads by washing them four times with at least five volumes of Guanidine lysis buffer supplemented with 5 mM 2-mercaptoenthanol and 50 mM imidazole pH 8. • Add the equilibrated NiNTA beads to the lysates (20 µl beads/ml of lysate). • Incubate overnight at 4°C in a rotator mixer. • Centrifuge for 5 minutes at 500 g • Remove supernatant and pipet the beads to an Eppendorf 1.5 ml LoBind tube with 5 to 10 bead volumes of wash buffer 1. • If your bead volume is higher than 200 µl, you may need LoBind tubes of a bigger volume. • Centrifuge 2 minutes at 500 g • Remove supernatant • Wash with 5–10 volumes of wash buffer 2 • Centrifuge 2 minutes at 500 g • Remove supernatant • Wash with 5–10 volumes of wash buffer 3 for 10 minutes while rotating. • Centrifuge 2 minutes at 500 g • Remove supernatant

• Wash with 5–10 volumes of wash buffer 4 for 10 minutes while rotating. • Centrifuge 2 minutes at 500 g • Remove supernatant • Wash with 5–10 volumes of wash buffer 4 for 10 minutes while rotating. • Centrifuge 2 minutes at 500 g • Remove supernatant • Add 1 bead volume of elution buffer, incubate for 30 minutes mixing by rotation. • In the meantime, wash a 0.45 µm UltraFree centrifugal filter unit per sample by passing through 200 µl of elution buffer. Centrifuge 1 minute at 10,000 g • Centrifuge the samples 2 minutes at 500 g. Place the UltraFree centrifugal unit in a LoBind tube, and pass the elution supernatant through the filter. Centrifuge for 1 minute at 10,000 g • Note that the LoBind tubes lid will not close after placing in the centrifugal unit. This might cause the lid to break apart during centrifugation. Therefore, make sure you label the tubes on the side. • Transfer the filtered elution to a new intact LoBind tube. • Repeat elution twice. In the third elution transfer the whole bead-elution buffer mix into the centrifugal filter unit to make sure all the elution volume is recovered. • At this step, the HIS-pulldown is finished. You may want to take a small aliquot to check the pulldown efficiency by immunoblotting together with the input samples. You may also now proceed to the next step, trypsin digestion, or snap freeze the samples in liquid nitrogen to proceed later. Concentration and trypsin digestion Concentration Before digesting the samples with trypsin, we will concentrate the samples and perform a reductionalkylation treatment. • Per sample, add 200 µl of urea buffer to a Vivacon 500 100,000 MWCO filter column. • Centrifuge at 8000  g for 10 minutes in a temperature-controlled centrifuge at 25°C. –– Washing the filters will remove surfactants

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•

• •

• • • •

that are used to keep the filter matrix stable, not washing the filter will likely produce contamination of the samples with detergents that will interfere with the mass spectrometry analysis. Additionally, long centrifugation times at high speed will heat up the samples, which is highly undesirable in urea buffer, producing lysine carbamylation of the samples. Remove the flow through and add the sample to the filter (400 µl max). –– Although your E3OI may be smaller than 100  kDa, TULIP conjugates under denaturing conditions will result in long branched peptides that will not be able to pass through the filter [also see Hendriks et al., 2014, 2015)]. In any case, you may want to keep an aliquot of the flow through to check that this holds true for your E3OI. Centrifuge at 8000  g for 10 minutes in a temperature-controlled centrifuge at 25°C. Remove the flow through and add the remaining amount of sample to the filter (if any, 400 µl max). Reconcentrate by centrifugation at 8000 × g for 10 minutes in a temperaturecontrolled centrifuge at 25°C. Remove the flow through and add 200 µl of urea buffer to wash the sample. Reconcentrate by centrifugation at 8000 g for 10 minutes in a temperature-controlled centrifuge at 25°C. Repeat the washing step once. After the concentration steps, your sample volume should be around 5–10 µl. Recover the sample by placing the filter upside down in a LoBind tube in a bench top open centrifuge. Spin down for 10 seconds at 1000 g. Rotate the filter 180° and spin down again. –– Be aware that the filter unit will not fit perfectly in the LoBind tube, if not placed carefully, the filter unit may dislodge during centrifugation resulting in loss of the sample. This is highly undesirable apart from being also potentially dangerous. Therefore, wear protective glasses.

• Add 2.5 µl of a freshly prepared 1M Ammonium Bicarbonate (ABC) solution, resulting in a final concentration of 50 mM. Vortex to mix. • Add 0.5 µl of a freshly prepared 100 mM DTT (Dithiothreitol) solution, resulting in a final concentration of 1 mM. Vortex to mix. • Incubate 30 minutes at room temperature. • Add 0.5 µl of a freshly prepared 500 mM CAA (Chloroacetamide) solution, resulting in a final concentration of 5 mM. Vortex to mix. • Incubate 30 minutes at room temperature. • Add 2.5 µl of a freshly prepared 100 mM DTT (Dithiothreitol) solution, resulting in a final concentration of 6mM. Vortex to mix. • Incubate 30 minutes at room temperature. • Add 200 µl of 50 mM ABC. • Add 500  ng of sequencing grade modified trypsin dissolved in 50 mM ABC. Vortex to mix. • Incubate overnight, still and in the dark at room temperature. • Inactivate trypsin by adding trifluoroacetic acid (TFA) to a final concentration of 2%. –– TFA is highly corrosive and toxic. Therefore, employ TFA in the fume hood wearing protective glasses, lab coat and gloves. Desalting of the peptides (stage tipping) Buffers required for StageTipping are listed in Table 10.5. • Prepare StageTips according to previously published instructions (Rappsilber et al., 2007). Stack 3 Empore SPE C18 disks to maximize peptide recovery. • Activate the StageTips by passing through 100 µl of methanol by centrifuging at 1000 g. • Do not leave the C18 matrix dry, instead, leave 1–2 mm fluid over the C18 material. • Pass through 100 µl of buffer B to condition the StageTips. Do not leave the C18 matrix dry. Table 10.5  StageTip buffers Buffer

Recipe

Buffer A

0.1% formic acid

Reduction alkylation

Buffer B

80% acetonitrile, 0.1% formic acid

Buffer C

60% acetonitrile, 0.1% formic acid

• Increase the sample volume up to 50 µl with urea buffer.

Prepare buffers fresh on the day of the experiment. Use clean 15 ml tubes. Use HPLC grade reagents.

Targets of Ubiquitin Ligases |  157

• Equilibrate the StageTip by passing through 100 µl of buffer A (Table 10.5). Do not leave the C18 matrix dry. • Load your sample in 100 µl cycles until the whole sample has been loaded. Do not leave the C18 matrix dry. • Wash the StageTip with 100 µl of buffer A twice. Centrifuge until the matrix is completely dry. • Make a whole in the lid of a LoBind tube by puncturing. Small size scissors are ideal for this purpose. Make the hole big enough so the StageTip can be placed in reaching down to the 500 µl mark in the tube. • Add 25 µl of buffer C (Table 10.5) to elute the sample. Centrifuge for 3 minutes at 1000 g. • Repeat the elution once. • Transfer the whole elution to a new, clean, nonpunctured LoBind tube. • Vacuum dry the peptides the peptides until complete dryness in a SpeedVac. • Samples can now be stored indefinitely at −20°C. • Resuspend the peptides in 10 µl buffer A by sonication for 2 minutes in a water bath. Briefly spin down the peptide solution and store them at −20°C or −80°C until they are ready to be analysed by LC–MS/MS. Analysis and identification of the TULIP conjugates by LC–MS/MS In this section, we will make some general recommendations on how to analyse your TULIP conjugates by mass spectrometry giving some examples based on the conditions used in our previous project on RNF4 targets (Kumar et al., 2017). However, we highly recommend consulting with mass spectrometry experts at your institution or facility, as conditions and settings may vary depending on the equipment you have available and the characteristics of your E3OI. In our case, we use an Easy- nLC1000 HPLC system coupled to a Q-Exactive mass spectrometer (Thermo Fisher Scientific). We make use of the freely available software packages MaxQuant and Perseus for the identification and statistical analysis of the TULIP targets, respectively. Standard workflows have been previously published (Tyanova et al., 2016a,b). Regardless of the characteristics of your E3OI and your LC–MS/MS equipment, in order to get

an accurate label-free quantification and matching between runs, we highly recommend running the samples on the same machine, same column, with the same gradient, and in a consecutive manner. Nevertheless, especially with large sample sets, this is not always possible (i.e. chromatography column block). The MaxQuant match between runs feature aligns identified peptides by retention time and m/z. Similar samples measured in different columns with the same chromatography gradients should align properly. Therefore, although suboptimal, samples can also be run in different sessions. Settings • First, perform a diagnostic run of the sample to determine peptide amount and complexity. This gradient can be short (30 minutes) including a small amount of the sample (1% approx.). An E3OI with a lot of different targets will produce more complex chromatography profiles than other more specific ones. • Based on your diagnostic run you (or your mass spectrometry expert) will have to decide the amount of sample to run, and the length of the chromatography gradient. In our research group, the chromatography gradients we perform for TULIP conjugates vary between 1 and 2 hour gradients from 0% to 30% Acetonitrile in 0.1% formic acid, reaching up to 95% Acetonitrile and 0.1% formic acid at the end of the gradient. In Fig. 10.5, an example of a diagnostic and a definitive run of the same sample is shown. • If possible, in case you have enough material to do multiple runs, it will be advantageous to perform technical replicates. Analysing RAW data by MaxQuant How to use MaxQuant has already been explained in detail (Tyanova et al., 2016a), and the default settings are sufficient to secure the identification and quantification of the TULIP conjugates. The software can be freely obtained after registration from the URL: ‘http://www.coxdocs.org/doku. php?id=maxquant:start’. The explanations written here correspond to version 1.5.3.30. At the moment you want to perform your analysis, the MaxQuant version available in the website will be a more recent one. This might affect the position

158  | González-Prieto and Vertegaal

A

Relative intensity

100% 80% 60% 40% 20% 0%

0

10

20

30

Time (min)

B Relative intensity

100% 80% 60% 40% 20% 0%

0

30

60

90

120

Time (min)

Figure 10.5  Example of both (A) a diagnostic and (B) a definitive run of an RNF4 TULIP-conjugates sample, corresponding to 1% and 20% respectively of the total TULIP conjugates obtained from five subconfluent 15 cm dishes of U2OS cells corresponding to approximately 100 million cells.

of the different tabs and labels in the Graphic User Interface of the software. Since, at the current moment, only the most recent version is available to download from the website, you will have to search through internet forums to obtain older versions from MaxQuant. Otherwise, we recommend exploring the software and search for the equivalent option to the one hereunder described in your MaxQuant version. In any case, we will list some configuration settings which we introduce in our searches that differ from the default settings: • Load your RAW files into MaxQuant. • Provide a different ‘Experiment name’ to each of your biological replicates, technical replicates should have the same ‘Experiment name’. • Set the number of threads to the maximum your computer is able to handle. • In ‘Group-specific’ parameters tab: –– Digestion → set maximum missed cleavages to 4. –– Modifications →  Variable modifications → Add GlyGly (K). This will potentially allow you to identify the ubiquitination sites in

your E3OI targets, although, in practice, a very small number of sites is identified. –– Label-free quantification →  Select LFQ → untick Fast LFQ. • In ‘Global parameters’ tab: –– Add a FASTA file corresponding to the full proteome of your model organism. Full proteomes can be downloaded from Uniprot. Make sure your FASTA file is included in the databases file in the Andromeda Engine configuration. –– Adv. Identification → tick match between runs. –– Protein quantification → untick ‘Use only unmodified peptides and…’ • Press Start. The calculations of MaxQuant will take several hours depending on your computer hardware and the size and number of your RAW data files. When the calculations are done, MaxQuant will generate a series of tables in: Your directory/combined/txt. Among them, you will find the proteingroups. txt file, which you will need for further analysis in Perseus software. It contains all the relevant

Targets of Ubiquitin Ligases |  159

information about every protein identified. Additionally, the summary.txt file, will give you information about the number of peptides identified per raw file and the percentage of MS/ MS spectra that could be identified as a peptide. We aim for identification rates between 20–30%. In case your identification rate is lower than 10%, we recommend optimising the chromatography and/ or mass spectrometry methods. Analyse MaxQuant output in Perseus The Perseus workflow has also been explained previously (Tyanova et al., 2016b). Perseus software can be freely downloaded from the URL: http:// www.coxdocs.org/doku.php?id=perseus:start after registration. Here we will describe a brief workflow for the determination of the TULIP conjugates. As previously explained for MaxQuant, these instructions correspond to Perseus version 1.5.5.3., and the positions of the different options within the software might differ from the ones in the latest version at the time you perform your analysis. • • • •

• • • • •

Load you proteingroups.txt file into Perseus Move FASTA headers to Text Columns. Set LFQ intensities as Main Columns. Filter Rows based on categorical columns leaving out’ –– Contaminants –– Only identified by site –– Reverse In categorical annotation rows → Include every different biological repeat of a condition in the same group. In Basic →  Transform →  Do a log2(x) transformation Filter Rows → based on valid values → set the minimum number of values to the number of biological repeats in at least one group. Imputation → Replace missing values by normal distribution →  Default values →  Total Matrix mode. You can start performing statistical tests between groups of samples. –– Permutation based FDR is a very stringent way to determine differences between sample groups. For an E3OI with a very large subset of ubiquitination targets, it is a very efficient way of filtering out real targets.

However, in most of the cases, the number of targets compared to the background binding proteins is very low and the FDR correction might be too stringent. You will also need extremely low technical variability. –– To compensate for this, we recommend using just the p-value for truncation in the t-tests and look for average differences higher than 1 (log2). Passing statistical tests for different conditions will provide extra statistical significance as a potential target of ubiquitination (i.e. it is statically enriched comparing wild type TULIP both with the TULIP-ΔGG and a catalytically-dead TULIP construct, while there is no statistical difference between the TULIP-ΔGG and a catalytically-dead TULIP construct). • After completing the analysis, export the tables. –– The Perseus Output tables are not very comprehensive and intuitive, as a personal subjective opinion, and does not make it straightforward to browse through the data for specific proteins of interest. Exporting the tables to Microsoft Excel or other similar spreadsheet software may facilitate further formatting and evaluation of the data. Also, this enables producing reader-friendly tables. Further confirmation of the E3OI targets After performing the TULIP methodology to identify your E3OI substrates you may want to confirm a subset of your E3-specific ubiquitination targets with a different approach. These can be confirmed directly using your TULIP samples for immunoblotting, but you should consider that the added molecular weight corresponding to the TULIP construct and, probably ubiquitin chains, might make it difficult to detect conjugates by immunoblotting as your target might reach molecular weights more difficult to transfer from gel to blotting membranes. Additionally, the knockdown of your E3OI might also produce a change in the ubiquitination status of its targets. Purifying the ubiquitin proteome and immunoblotting against your target protein might also provide information about the validity of your target protein of interest.

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References

Akimov, V., Barrio-Hernandez, I., Hansen, S.V.F., Hallenborg, P., Pedersen, A.K., Bekker-Jensen, D.B., Puglia, M., Christensen, S.D.K., Vanselow, J.T., Nielsen, M.M., et al. (2018). UbiSite approach for comprehensive mapping of lysine and N-terminal ubiquitination sites. Nat. Struct. Mol. Biol. 25, 631–640. https://doi. org/10.1038/s41594-018-0084-y. Hendriks, I.A., D’Souza, R.C., Chang, J.G., Mann, M., and Vertegaal, A.C. (2015). System-wide identification of wild-type SUMO-2 conjugation sites. Nat. Commun. 6, 7289. https://doi.org/10.1038/ncomms8289. Hendriks, I.A., D’Souza, R.C., Yang, B., Verlaan-de Vries, M., Mann, M., and Vertegaal, A.C. (2014). Uncovering global SUMOylation signaling networks in a sitespecific manner. Nat. Struct. Mol. Biol. 21, 927–936. https://doi.org/10.1038/nsmb.2890. Hendriks, I.A., and Vertegaal, A.C. (2016). A high-yield double-purification proteomics strategy for the identification of SUMO sites. Nat. Protoc. 11, 1630– 1649. https://doi.org/10.1038/nprot.2016.082. Kumar, R., González-Prieto, R., Xiao, Z., Verlaan-de Vries, M., and Vertegaal, A.C.O. (2017). The STUbL RNF4 regulates protein group SUMOylation by targeting the SUMO conjugation machinery. Nat. Commun. 8, 1809. https://doi.org/10.1038/s41467-017-01900-x. Lamesch, P., Li, N., Milstein, S., Fan, C., Hao, T., Szabo, G., Hu, Z., Venkatesan, K., Bethel, G., Martin, P., et al. (2007). hORFeome v3.1: a resource of human open reading frames representing over 10,000 human genes.

Genomics 89, 307–315. https://doi.org/10.1016/j. ygeno.2006.11.012. O’Connor, H.F., Lyon, N., Leung, J.W., Agarwal, P., Swaim, C.D., Miller, K.M., and Huibregtse, J.M. (2015). Ubiquitin-Activated Interaction Traps (UBAITs) identify E3 ligase binding partners. EMBO Rep. 16, 1699–1712. https://doi.org/10.15252/ embr.201540620. Rappsilber, J., Mann, M., and Ishihama, Y. (2007). Protocol for micro-purification, enrichment, pre-fractionation and storage of peptides for proteomics using StageTips. Nat. Protoc. 2, 1896–1906. https://doi.org/10.1038/ nprot.2007.261. Seiler, C.Y., Park, J.G., Sharma, A., Hunter, P., Surapaneni, P., Sedillo, C., Field, J., Algar, R., Price, A., Steel, J., et al. (2014). DNASU plasmid and PSI:Biology-Materials repositories: resources to accelerate biological research. Nucleic Acids Res. 42, D1253-1260. https://doi. org/10.1093/nar/gkt1060. Skaar, J.R., Pagan, J.K., and Pagano, M. (2014). SCF ubiquitin ligase-targeted therapies. Nat. Rev. Drug Discov. 13, 889–903. https://doi.org/10.1038/nrd4432. Tyanova, S., Temu, T., and Cox, J. (2016a). The MaxQuant computational platform for mass spectrometry-based shotgun proteomics. Nat. Protoc. 11, 2301–2319. https://doi.org/10.1038/nprot.2016.136. Tyanova, S., Temu, T., Sinitcyn, P., Carlson, A., Hein, M.Y., Geiger, T., Mann, M., and Cox, J. (2016b). The Perseus computational platform for comprehensive analysis of (prote)omics data. Nat. Methods 13, 731–740. https:// doi.org/10.1038/nmeth.3901.

Part III

Cellular Processes

Regulation of p53 Family Members by the Ubiquitin and SUMO Modification Systems

11

Viola Calabrò and Maria Vivo*

Department of Biology, University of Naples Federico II, Naples, Italy. *Correspondence: [email protected] https://doi.org/10.21775/9781912530120.11

Abstract The p53 family includes, in addition to the wellknown tumour suppressor p53, two additional proteins, p63 and p73. These proteins are encoded by two different genes, each of them subjected to different activation modes. All family members have an essential role in either tumorigenesis or morphogenesis. The high degree of identity among the three protein sequences is mirrored by the existence of a common modular protein structure. All of them present a transactivation (TA), a DNA-binding (DBD) and an oligomerization (OD) domain, with a high level of sequence identity. Each gene gives rise to multiple isoforms due to differential promoter selection and alternative splicing at both 5′ and 3′ ends of the mRNA. Despite the homology, p53, p63 or p73 gene inactivation in mice gave rise to different phenotypes indicating that the proteins encoded by these genes play different roles. While p53 has the central function of tumour suppressor, both p63 and p73 are actively involved in development and differentiation. A complex set of post-translational modifications such as phosphorylation, acetylation, ribosylation, glycosylation, ubiquitination and SUMOylation, with often-intertwined modes of action regulates p53 family members functions. Ubiquitination and SUMOylation appear to affect transactivation ability, localization and stabilization of these transcriptional factors and to confer their timely regulated role during differentiation and

development. In this chapter, the role of p53 family members will be described as well as the impact of ubiquitination and SUMOylation on their functions. Moreover, other ubiquitin-like proteins have also been shown to regulate p53 family members activity. The interaction of the Ub/SUMO system with the complex regulation pathways of both tumour suppression and development guaranteed by the p53 family members constitutes an example of critical control machinery regulating cellular fate. Introduction: the p53 family The p53 gene encodes the well-known oncosuppressor p53, with a central role in the cellular response to oncogenic stimuli and cytotoxic stress. Its activation in response to these events determines the transcriptional induction of several target genes involved in apoptosis, cell cycle arrest cellular senescence and DNA repair mechanisms (Lane, 1992; Vogelstein et al., 2000; Riley et al., 2008) In 1997 two new members of the p53 family were identified: p73 and p63 (Kaghad et al., 1997; Yang et al., 1998). These two new transcription factors show a high percentage of identity with the p53 gene sequence. Alignment of the three gene sequences revealed the existence of an ancestral proto-gene similar to TP63 and TP73, from which the gene coding for the transcription factor p53 would subsequently evolve in the higher organisms. All family members have a substantial impact

164  | Calabrò and Vivo

in tumorigenesis and morphogenesis. In addition to a high degree of identity, the three proteins display a typical modular protein structure (Yang et al., 2002a; Yang and McKeon, 2000). In p53 this modularity includes a transactivation domain (TA), located in the N-terminal portion, a DNAbinding domain (DBD), in the central part of the protein, and an oligomerization domain (OD), in the C-terminal portion, responsible for the tetrameric structure of the proteins (Fig. 11.1). Due to their partial homology in the oligomerization domain, the p53 family members can form hetero-tetramers, with important implications in the regulation of their functionality. The most conserved region is the DBD, which shows 63% identity between p73 and p53 and 60% identity between p63 and p53 (De Laurenzi and Melino, 2000). Although sharing high levels of amino acid identity with p53, certain isoforms of both p63 and p73 differs considerably from p53, with additional domains not present in p53 (Figs. 11.2 and 11.3). By binding to known p53-responsive elements they can transactivate p53 target genes and induce cell cycle arrest and apoptosis ( Jost et al., 1997; Marin et al., 1998; Osada et al., 2005). Nevertheless, these

proteins transcriptionally activate specific target genes (Levrero et al., 2000) such as PERP for p63 (Ihrie et al., 2005), Aquaporin 3 for p73 (Zheng and Chen, 2001), and JAG1/2 for p63/p73 (Sasaki et al., 2002). Structure and functions of the p53 family members The TP63 gene, located on chromosome 3q27–29, includes 14 exons and different promoters (Fig. 11.2). The gene bears two alternative transcription start sites (Yang et al., 1998). Transcripts originating from the first site give rise to the so called ‘TA isoforms’, containing a transactivation domain (TA) similar to that of p53, while transcripts originating within exon 3 give rise to the so called ‘ΔN isoforms’ that are devoid of TA region. Furthermore, due to alternative splicing events at the 3′ end of the gene, 3 different isoforms are generated named α, β and γ. Combining the carboxy- and amino-terminus variability, six different isoforms are thus produced from the p63 gene (Fig. 11.2). Unlike β and γ isoforms, α isoforms have a much more extensive C-terminal region including two additional domains, called SAM (Sterile Alpha

Figure 11.1 Structure of the TP53 locus and modularity of p53 isoforms. (A) Boxes indicates exons. UTR regions are indicated in black, while region encoding the different protein domains within exons are indicated with pink (transactivation domain, TAD), blue (DNA binding, DBD) and light blue (oligomerization domain, OD). Arrows indicate alternative transcription start sites. Dot lines indicate splicing options. Same colours code in (B) where the combinations deriving from both 5′ and 3′ end different usage are indicated by the scheme. The three N-terminal options can be combined through a unique DNA binding domain to three different 3′ ends. The alpha isoform is the known tumour suppressor p53 whose regulation through SUMO or Ub conjugation is described in the text. Adapted from: Watson and Irwin, 2006.

p53 Regulation by Ubiquitin and SUMO |  165

Figure 11.2 Structure of TP63 locus and modularity of p63 isoforms. (A) Exons are differently coloured as reported in Fig. 11.1. In p63 the second transactivation domain in the C-terminus is indicated in green (TA), the sterile alpha motif (SAM) in purple and Transactivation inhibitory domain (TID) in yellow. In grey are indicated regions that do not belong to the mentioned domains or that are not trascribed due to stop codons arising on alternative splicing. Arrows indicate alternative transcription start sites, while dotted lines above and below the exons indicate splicing options. (B) Combination deriving from both 5′ and 3′ end different usage is indicated by the scheme, showing the potential isoforms that can be produced by the locus. Two different 5’ends give rise to TA or DN isoforms that can be combined through the unique DNA binding domain to three different 3′ carboxy-terminus domains of the alpha, beta or gamma options thus producing six different isoforms. Adapted from: Watson and Irwin, 2006.

Figure 11.3  Structure of TP73 locus and modularity of p73 isoforms. (A) Exons are indicated as in the previous figure. Exon 12 can be translated differently depending on the messenger in which it is retained. Arrows indicate alternative transcription start sites, while dotted lines above and below the exons indicate splicing options. (B) Combination deriving from both 5′ and 3′ end different usage is indicated by the scheme, showing the potential isoforms that can be produced by the locus. Five different N-terminal domains can be combined through the unique DNA binding domain to six different 3′ carboxy-terminus domains. Adapted from: Watson and Irwin, 2006.

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Motif) and TID (Trans Inhibitor Domain) that are absent in p53. The SAM domain is required for interactions with proteins that modulate p63 activity. The TID domain, on the other hand, appears to be involved in the intra-molecular interaction with the trans-activation domain (TA) at the N-terminus determining the decrease in transcriptional activity of the TAα isoforms on the p53 activated promoters (Ghioni et al., 2002; Serber et al., 2002). TA isoforms exert the function of transcriptional activators of p53 target genes such as p21/waf and Mdm2, inducing cell cycle arrest and apoptosis. ΔNp63 isoforms are able to antagonize full-length isoforms of p63 and also other p53 family members and act like dominant negative transcription inhibitors. This happens because they lack the N-terminal transactivation domain. Nevertheless, they still maintain the ability to activate the transcription of their target genes thanks to the presence of another small transactivation domain located between amino-acids 410 and 512 (Ghioni et al., 2002). The structure of the p73 locus is more complex than that of p63 (Fig. 11.3). Seven C-terminal isoforms generated either by alternative splicing (α, β, γ, δ, ε, and ζ) or by alternative termination of translation can be produced. In addition, the p73 gene encodes four distinct N-terminal isoforms termed ΔTAp73 or ΔNp73 generated as a result of several mechanisms: transcription from an alternative promoter within intron 3 (ΔNp73), translation from an alternative initiation site (ΔNVp73), and alternative N-terminal splicing (DEx2p73 and DEx2/3p73). The locus would thus express more than 30 mRNA variants encoding multiple proteins; however, only 14 have been described (Ishimoto et al., 2002; Watson and Irwin, 2006). The two major forms are the α and β isoforms containing 636 and 499 amino acids, respectively (Kaelin, 1999). ΔNp73 is the predominant isoform in the murine foetal nervous system and its loss leads to enhanced apoptosis in cortical and sympathetic ganglia neurons resulting in either the absence or the loss of specific populations of neurons. This effect seems to be the result of p73-mediated inactivation, at least in part, of full-length pro-apoptotic p53 family proteins (p53, TAp63, and TAp73). Originally, p63 and p73 were assumed to function similarly to p53. Now, both p63 and p73 have also been shown to play developmental roles. In fact, despite the homology, p53, p63 or p73

gene inactivation in mice gave rise to different phenotypes indicating that the proteins encoded by these genes play different roles. P53–/– mice show a normal development but a strong increase in susceptibility to spontaneous tumorigenesis. Otherwise, mice deprived of p73 exhibit multiple defects in the development of the nervous system while p63–/– mice show abnormalities in the epithelial, craniofacial and limb development but no susceptibility to the tumour (Mills et al., 1999; Yang et al., 1999, 2000). After the discovery of the p63 and p73 locus organization, it became clear that p53 presented a gene structure similar to its cognate genes (Fig. 11.1). The p53 locus encodes different p53 mRNA variants through both the use of alternative splicing and the existence of an internal promoter in intron 4 (Bourdon et al., 2005). These p53 isoforms are expressed in a wide range of normal tissues but in a tissue-dependent manner, thus witnessing different levels of regulation taking place at both transcriptional and mRNA level. It is unclear what sub-fractions of total p53 are active, and above all how they are regulated by post-translational modifications, given that p53 abundance is not necessarily associated with p53 transcriptional activity. In any case, the different p53 protein isoforms are less abundant than full-length p53 protein. These isoforms are differentially expressed in several human cancer types and were shown to exhibit several biological functions, modulating p53 transcriptional activity and tumour-suppressor functions. The complex structure of the p53 gene, more similar to the p63 and p73 genes than previously thought, thus reveals an unforeseen even complex regulation. Moreover, the observation that the alternative promoter is conserved through evolution suggest that this peculiar gene structure plays an essential role in the multiple activities of the p53 family members, in which the interplay between p53 isoforms and p53 on specific targets may play a major role in controlling the activity of p53-related proteins. The tumour suppressor p53 p53 is not essential for completion of the cell division cycle, but disruption of its functions is central to the life history of most, if not all, cancer cells. P53 is a transcriptional factor that negatively regulates cell cycle progression following cellular stresses,

p53 Regulation by Ubiquitin and SUMO |  167

such as DNA damage, telomeres erosion, dNTPs depletion, hypoxia, and oncogene activation(Lane, 1992; Bates and Vousden, 1999; Woods and Vousden, 2001). The functions of this tumour suppressor include developmental processes, differentiation, DNA repair, senescence, ageing, and angiogenesis and are accomplished by both transcription-dependent and transcription independent mechanisms (Rufini et al., 2013; Comel et al., 2014). Under normal conditions the p53 protein is kept at low levels by its rapid turn-over. Signal transduction pathways activated in stressed cells lead to its stabilization and to the initiation of a p53 dependent transcription program that arrests cell proliferation or, more dramatically, induces cell suicide. For instance, activation of the p53 target genes CDKN1A (p21) and GADD45 plays a key role in p53-induced cell-cycle arrest, whereas the BH3-only encoding target genes BBC3 (PUMA) and PMAIP1 (NOXA) are critical players in p53-mediated apoptotic cell death. A transcriptional target of p53 is the MDM2 (Mouse Double Minute 2) gene, which plays a central role in regulating p53 functions (Haupt et al., 1997; Honda et al., 1997; Kubbutat et al., 1997). The two proteins are part of a negative feedback loop that keeps p53 levels low during normal growth and development. Activation of p53 in response to cellular stresses is mediated at least in part by inhibition of MDM2 functions. Inhibition of the MDM2 gene expression or post-translational modifications of both proteins that tend to weaken or inhibit the binding between p53 and MDM2 (Ashcroft et al., 1999; Ashcroft and Vousden, 1999). Role of ubiquitination in p53 functions A complex set of post-translational modifications such as phosphorylation, acetylation, ribosylation, o glycosylation ubiquitination and sumoylation, with often-intertwined modes of action regulates p53 functions providing an explanation for the versatile role of this protein. Depending on the cellular signal, several mechanisms of p53 stabilization (and destabilization) have been described. Among these, regulation of p53 ubiquitination has been studied for long time given its role in mediating p53 degradation through the proteasomal pathway. p53 was first shown to be regulated by ubiquitination mediated pathway based on the discovery that

the human papilloma virus type 16 (HPV-16) E6 protein induces degradation of p53 (Everett et al., 1997). Ubiquitination is the first identified posttranslational modification consisting in a protein that can modify another protein. The role of ubiquitination on the fate of a protein targets depends on the length of the Ub chains, with polyubiquitination having a role in protein degradation in the majority of the cases. Ub chains can be further modified by ubiquitination on several lysine residues within ubiquitin amino acid sequence giving rise to different chain linkages, that can have different functions. These different chains differ by the position of the modified lysine residue. In addition to proteasomal degredation, modifications with ubiquitin and ubiquitinlike proteins serve various functions in the cell. Indeed, while K48-linked chains have been related to proteasomal degradation, K63 chains, as well as mono-ubiquitination, regulates protein involvement in DNA repair, kinase activation, trafficking, localization and transcriptional regulation (Pickart and Fushman, 2004). Most of these functions are common to the tumour suppressor p53 and are involved in the control of p53 tumour suppressor stability. Role of MDM2 in p53 ubiquitination The importance of the regulation of p53 stability in controlling its activity has led to the discovery of several E3 ubiquitin ligases and other associated factors that directly affect p53 levels, sub-cellular localization, and activity. Many E3 ubiquitin ligases have been found to target p53 for degradation. The oncoprotein MDM2 actually plays a major role in regulating p53 stability. Interestingly, MDM2 is a transcriptional target of p53 and the two proteins are involved in a negative feedback loop altered in several human cancers. Consistently, the human homologue of MDM2, HDM2, is up-regulated in 7% of human cancers in which it causes a p53 deficiency. The E3 ubiquitin ligase MDM2 interacts with and ubiquitinates p53 through its RING (Really Interesting New Gene) domain, shared by many E3 ubiquitin ligases (Boyd et al., 2000). It has been extensively shown that Mdm2-mediated ubiquitination of p53 induces its nuclear export and degradation (Haupt et al., 1997; Honda et al., 1997; Kubbutat et al., 1997; Freedman and Levine,

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1998; Roth et al., 1998; Stommel et al., 1999; Geyer et al., 2000). Several lysine residues in the protein sequence of p53 are target of ubiquitination, both in the DNA binding domain and in the oligomerization domain (Fig. 11.4). In particular, it has been shown that MDM2 preferentially ubiquitinates 6 key lysine residues located in the p53 C-terminus domain (Rodriguez et al., 2000; Lohrum et al., 2001). Experiments with p53 mutants in which these lysines were replaced by arginines (K to R mutation) that cannot be modified, showed that they are not sufficient to induce p53 degradation. This observation led to the discovery of another group of lysines, in the DNA binding domain, which are required to induce p53 degradation by the proteasome (Chan et al., 2006; Chao, 2015). Mutation of these additional sites decrease both stability and overall p53 ubiquitination. However, as their mutation affects the stability of MDM/p53 interaction, the specific role of these sites in p53 ubiquitination is challenging to assess. Although nuclear export was first considered to be compulsory for p53 degradation, later

studies demonstrated that it can also occur in the nucleus (Geyer et al., 2000; Lohrum et al., 2001; Xirodimas et al., 2001b; Shirangi et al., 2002; Stommel and Wahl, 2004). Interestingly, the extent of p53 ubiquitination is suggested to dictate p53 fate. It appears that Mdm2 can differentially catalyse mono and poly-ubiquitination of p53, in a dosage-dependent manner. In particular, monoubiquitination would induce nuclear export while poly-ubiquitination nuclear degradation (Li et al., 2003). In the cytoplasm, p53 exerts transcriptional independent activities, such as apoptosis induction or autophagy by interacting with proteins of the Bcl family including Bcl-Xl and Bcl2 (Fontana and Vivo, 2018; Mrakovcic and Fröhlich, 2018). This means that p53 ubiquitination can be seen as a way to shut down its nuclear activity by inducing cytoplasmic relocalization and/or degradation. At the same time, increasing the concentration of p53 in the cytoplasm can be seen as a tool to trigger transcriptional independent functions of p53 thereby explaining how this protein exerts multiple functions. This aspect implies that the role of

Figure 11.4  TA Alpha isoforms of p53, p63 and p73 are depicted together with the relative positions of the lysine residues subjected to ubiquitin (red squares) or SUMO conjugation (green circles). K386 in p53 is also subjected to acetylated in cells treated with trichostatin A/nicotinamide as assessed by mass spectrometry (Tang et al., 2008) Asterisks indicate cryptic sumoylation sites. Lysines at position 193 and 194 of DNp63a were previously shown to be necessary for Itch ubiquitin ligase-mediated degradation of p63 (Rossi et al., 2006) and their substitutions partially protect DNp63a from MDM2 or FBW7-mediated degradation (Galli et al., 2010). Adapted from: Watson and Irwin, 2006.

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ubiquitination in regulation of p53 functions is not as simple as was thought before. The critical role of Mdm2 in p53 degradation is best illustrated by studies carried out in mice, where inactivation of p53 was shown to completely rescue the embryonic lethality caused by loss of Mdm2 function ( Jones et al., 1995; Lozano and Montes de Oca Luna, 1998; de Rozieres et al., 2000). The bulk of evidence about the role of MDM2 in determining p53 functions led to the known scenario in which under physiological conditions p53 levels are kept low by MDM2 mono-ubiquitination that mediates translocation of p53 to the cytoplasm. Upon stress signals p53 levels rise in order to allow DNA repair and cell response. As a result, also the expression level of Mdm2 in stressed cells is increased by p53 itself, providing a mechanism for poly-ubiquitination and degradation of p53 after repair has occurred. Interestingly, also MDM2 can undergo ubiquitination and sumoylation that results in the attenuation of its negative effect on p53. Regulation of p53 functions must be finely regulated to allow a response. At the same time, this control must be timely regulated to allow cellular growth in normal condition. This fine regulation mechanism is orchestrated by the Mdm2/MdmX complex. MdmX is a protein homologue to Mdm2 that negatively regulates p53 transcriptional functions independently from degradation. MdmX can stabilize p53 when overexpressed, as demonstrated by the accumulation of its poly-ubiquitinated forms into the nucleus ( Jackson and Berberich, 2000; Stad et al., 2001). However, it does not possess an in vivo ability to ubiquitinate and degrade p53. It has been shown that Mdm2 and Mdmx form heterodimers, through the interaction between the RING domain of one partner and the C-terminus of the other. This interaction constitutes the active and principal ubiquitin ligase that induces p53 poly-ubiquitination and proteasomal degradation. The outcome of the interplay between the MDM2/MDMX complex and p53 does not always results in p53 degradation but also to p53 relocalization to the cytoplasm, depending on Ub chain length and linkage. It has been proposed that, MDM2/MDMX complex could recruit only selected E2 enzymes that dictate the ubiquitin chain length and linkage, and thus the fate of the target protein (Christensen et al., 2007; Mace et al., 2008; Ye and Rape, 2009; Wade et al., 2010). Upon DNA damage, both MDM2 and MDMX

are phosphorylated. MDM2 loses the ability to bind p53 but is still able to bind MDMX. Growing evidence shows that upon DNA damage or cellular stress the ubiquitin ligase activity of the protein complex is redirected to MdmX. In this case, the increased p53 stability is not due to inhibition of MDM2-dependent ubiquitination, but rather to a switch of target, from p53 to MDMX. This results de facto in a block of Mdm2 E3 ligase activity towards p53 and its stabilization (Hock and Vousden, 2014; Chao, 2015). p53 is target of ubiquitin ligases with diverse modes of action Degradation of p53 in the absence of Mdm2 suggested that other mechanisms exist to mediate p53 degradation. Besides MDM2 a number of E3 ligases targeting p53 have been described, including RING domain E3 ligases (e.g. Pirh2, Cul4a/DDB1/Roc, synoviolin, COP1), HECT domain E3 ligases (e.g. ARF-BP1, Msl2/WWP1), U box domain E3 ligases (e.g. CHIP, UBE4B), and other undefined domain ligases usually referred as E4 ligases (e.g. p300/ CBP, E4F1, Ubc13) (Brooks and Gu, 2006; Lee and Gu, 2010) (Table 11.1). Some of these ubiquitin ligases ubiquitinate p53 without targeting it for degradation. These include MSL2 and WWP1, which drive cytoplasmic localization of p53 (Laine and Ronai, 2007; Kruse and Gu, 2009) and E4F1, which ubiquitinates lysine residues (K319–320–321) distinct from those targeted by MDM2 and promotes the ability of p53 to drive cell cycle arrest (Le Cam et al., 2006). ARF-BP1/Mule/HectH9 (ARF binding protein 1) is a direct molecular partner of p53 and induces its ubiquitination independently from MDM2 (Chen et al., 2006; Qi et al., 2012). ARF (known as p14ARF in humans and p19ARF in mouse) was originally identified as an alternative transcript of the INK4a/ARF tumour suppressor locus that suppresses aberrant cell growth in response to oncogene activation, at least in part, by inducing the p53 pathway (Kamijo et al., 1997; Quelle et al., 1997; Sherr, 2001; Sharpless and DePinho, 2004). p53 induction by ARF is mediated through inhibiting the activities of Mdm2 (Chin et al., 1998; Kamijo et al., 1998; Stott et al., 1998; Zhang et al., 1998; Zhang and Xiong, 2001) and, similarly, the functions of ARF-BP1(Chen et al., 2005). Growing evidence showed that down-regulation of ARF-BP1

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Table 11.1  p53 ubiquitin ligases and de-ubiquitinating enzymes Enzymes

Class

Domain

Effect

MDM2

E3

RING

Block transcriptional activity, nuclear export, degradation

TRIM

E3

RING

Degradation

MDMX

E3

RING

Block transcriptional activity, MDMX/MDM2 complex induce degradation

CULLIN

E3

RING

Degradation (Cul 1–4a–5 Block transcriptional activity (Cul 7) Cytoplasmic retention (Cul 9)

Hades

E3

RING

Degradation

Pirh2

E3

RING

Degradation

Topors

E3

RING

Degradation

Synoviolin

E3

RING

Nuclear export, degradation

COP1

E3

RING

Degradation

CHIP

E3

U-box

Degradation

UBE4B

E3

U-box

Degradation

MSL2/WWP1

E3

HETC

Nuclear export

ARF-BP1

E3

HETC

Degradation

P300/CBP

E4

Cytoplasmic degradation

E4F1

E4

Degradation

Yin Yang 1

E4

Degradation

Gankyrin

E4

Degradation

Hausp

DUB

Stabilization of p53, stabilization of MDM2 and MDMX

USP10 USP29 USP42

DUB

Stabilization through recycling cytoplasmic ubiquitinated p53

expression extended the half-life of p53, leading to the transcriptional activation of the p53 targets such as p21Waf1 and BAX, and to the p53-dependent apoptotic response. Thus, ARF activation on oncogenic signals induces p53 stabilization by interfering with p53 ubiquitination through either MDM2 or ARF-BP1 inhibition. Among ubiquitin ligases causing p53 polyubiquitination of degradation, there are the chaperone-associated ubiquitin-ligase as the protein CHIP (Carboxy-Terminus of Hsc70-Interacting Protein). The peculiarity of these proteins is that they need additional molecular players in order to mediate ubiquitination of their targets. Interestingly, it has been shown that the ubiquitin ligase activity is switched on CHIP binding to the chaperon protein (Narayan et al., 2015). In normal conditions, HSP90 chaperones regulate the protein levels of proteins they interact by directly recruiting ubiquitin ligases and presenting them for proteasome-mediated degradation. The chaperonedependent E3 ligase CHIP binds to Hsp70 and is a

resident part of the HSP90 complex. Both MDM2 and CHIP primarily function as the E3 ligases for mutant p53, although CHIP seems to be the more active one. In cancer cells, mutant p53 is trapped in stable interactions with up-regulated and activated HSP90. MDM2 and CHIP activity that also trapped in this complex in an inactive state, thus leading to the aberrant stabilization of mutant p53 molecules. Treatment of cells with pharmacological inhibitors of HSP 90 restore CHIP and MDM2 ability to induce p53 ubiquitination and subsequent degradation (Li et al., 2011; Narayan et al., 2015). A novel group of protein with ubiquitin ligase properties are E4-ubiquitin chain-assembly factors (Benirschke et al., 2010). These proteins reinforce E3 ligase functions, by increasing ubiquitination. The E4 UBE4B can physically interact with both p53 and MDM2 (Wu and Leng, 2011; Wu et al., 2011a). It is able to block p53 mediated transcription and apoptosis by inducing p53 degradation by promoting its mono-ubiquitination. When expressed in association with MDM2 it can induce

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poly-ubiquitination of p53, but only through MDM2 binding. UBE4B belongs to the group of U-box proteins, characterized by the presence of RING like domain indispensable for its function an E4 ligase towards p53. It has been shown that this protein is able to extend the poly-ubiquitin chains already assembled on p53. In addition to these proteins, also CBP and its paralogue p300 have been shown to mediate ubiquitination of p53 and to be part of the E4 ligase (Shi et al., 2009). This function appears to be exerted by cytoplasmic localized CREB and p300, while the nuclear counterparts induce acetylation of p53. It should be underlined that both acetylation and ubiquitination compete for the same C-terminal lysine residues on p53 (Ito et al., 2002; Li et al., 2002b). In normal condition, ubiquitination of these residues could serve to block p53 transcriptional activation. This evidence suggests that these activities take place thanks to the interaction with other molecular partners, probably in larger protein complexes or specialized cytoplasmic domains. Interestingly, p300 can induce poly-ubiquitination only of p53 species previously monoubiquitinated by Mdm2. Another family of E3 ligase is the Cullin-RING ubiquitin ligase (Dove and Klevit, 2017). These

enzymes are composed of several subunits that comprise an E3 ligase, a protein of the Cullin family members, a substrate specific receptor and often an adaptor protein. By functioning as scaffold these ligases recognize numerous substrates and participate in a variety of cellular processes thus constituting an ‘ubiquitination factories’ (Fig. 11.5). In particular, Cullin 7 is predominantly localized in the cytoplasm and binds directly to p53 without causing its re-localization. Cullin 7 induces monoor di-ubiquitination of p53 and thus antagonize its function promoting cell cycle progression (Andrews et al., 2006). Cullin-associated E3 ligases are involved in numerous human diseases, and multiple cancer types (Li and Xiong, 2017). p53 functions are regulated through the modulation of the activity of deubiquitinating enzymes An interesting level of p53 regulation is played by a group of proteins called deubiquitinases (DUBs). Ubiquitination of many proteins, as well as p53, can be reversed by these enzymes. The discovery of this additional layer of regulation of ubiquitination confirmed that deubiquitination of p53 results in p53 stabilization (Fu et al., 2017; Kwon et al., 2017).

Figure 11.5 Cullin-RING ubiquitin ligase complex constitutes a factory for protein ubiquitination. They are composed by a RING E3 ligase family member that binds to a protein of the Cullin family. The presence of an adaptor proteins such as Skp1, Elongin protein, DDB1 or sometimes unknown protein mediates the binding to a substrate recognition protein that tethers the target protein to the complex. Among the substrate recognition protein identified so far, F-bos, VHL box, BTB proteins DCAF, Fbw8 and SOCS-box. Figure adapted from Lijun Jia and Yi Sun, Cell Division 2009 4:16, https://doi.org/10.1186/1747-1028-4-16.

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HAUSP (Herpesvirus-Associated Ubiquitin-Specific Protease or called USP7) is a de-ubiquitinase specifically targeting p53 (Li et al., 2002a) and by its interaction stabilizes p53 protein levels. In contrast with this it was reported that complete knockdown or genetic knockout of HAUSP stabilized p53 protein levels (Cummins et al., 2004). Interestingly, HAUSP also interacts with and regulates Mdm2 as well as Mdmx ubiquitination (Li et al., 2004) thus inducing its stabilization in a p53 independent manner. In HAUSP-ablated cells, selfubiquitinated-Mdm2 becomes extremely unstable, leading to indirect p53 activation (Meulmeester et al., 2005). Therefore, ablation of HAUSP destabilizes Mdm2 leading to a subsequent decrease in ubiquitinated-p53 in favour of the unmodified and stable p53 species (Tavana and Gu, 2017). As p53 competes with MDM2 for HAUSP binding (Hu M, (2006)), the expression levels of the two players interferes with p53/Mdm2 binding. Upon stress signals, post-translational modification of HAUSP by ATM-dependent phosphorylation relieves Mdm2 inhibition, thus causing p53 stabilization (Meulmeester et al., 2005; Brooks et al., 2007). Being prevalently nuclear, HAUSP participates in this feedback loop in this cellular

compartment. The USP10 de-ubiquitinase instead, recycles cytoplasmic ubiquitinated p53, thus reversing Mdm2-mediated p53 nuclear export (Lee and Gu, 2010; Yuan et al., 2010). Interestingly, this protein translocates in the nucleus upon DNA damage, thus suggesting that it can contribute to p53 activation also in the nucleus (Yuan et al., 2010). The role of ubiquitination on p53 functions is summarized in Fig. 11.6. Role of sumoylation in p53 functions Sumoylation proceeds via an enzymatic pathway that is mechanistically analogous to ubiquitination but requires different enzymes that catalyse the covalent bond between a glycine residue of the SUMO moiety end a lysine residue of a protein substrate. One of the first SUMO targets to be identified has been p53 (Gostissa et al., 1999; Rodriguez et al., 1999). Unlike ubiquitination, that can take place on every lysine residue of the protein, sumoylation requires a consensus sequence the well-known ‘ΨKXE’ motif surrounding the modified lysine residue of most SUMO substrates. It has been extensively demonstrated that sumoylation can dramatically alter protein functions, by modifying protein-protein or protein–DNA interaction,

Figure 11.6  Role of p53 ubiquitination on p53. Chain length and chain linkage can have different effects on p53 functions and localization within the cell.

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subcellular localization, nucleo-cytoplasmic shuttling, enzymatic properties and protein stability. The majority of proteins that regulate the cell cycle and differentiation are sumoylated and in fact cancer, infections and neurodegenerative disorders are often associated with an alteration of sumoylation and cells over expressing SUMO 2 and 3 show signs of premature senescence (Gill, 2004). Role of Sumoylation on p53 mediated transcription Sumoylation affects p53 transcriptional activity, stability or subcellular trafficking (Stehmeier and Muller, 2009). A single lysine residue, K386 falling within a sumo consensus, has been found in p53 sequence. In vitro and in vivo assays further confirmed that this is the only sumoylation site present in the protein. The SUMO-1 conjugating enzyme Ubc9, that represents the first enzyme of the sumo reaction, plays an important role in substrate recognition as well as in substrate modification. Both Ubc9 and the two members of the PIAS family, PIAS 1 and PIAS 1β bind the C-terminal domain of p53 (Stehmeier and Muller, 2009). These interactions, that in part involve the p53 oligomerization domain, are required for p53 sumoylation. Given the low amount of sumoylated p53 within the cell, it has been challenging to understand the role of such modification on p53 functions. Indeed, only less than 5% of cellular proteins are sumoylated. This peculiarity of the sumoylation process is ascribed to SUMO specific isopeptidases called SENP that, as the DUBs for the ubiquitin, actively remove the SUMO molecules from the modified proteins. This made difficult the analysis of the biological role of such modification in vivo, especially in the case of p53 whose role in transcriptional activation is very complex. The role of sumoylation appears to be highly cell context dependent. While in some studies sumoylation increases the level of p53-mediated transcription (Gostissa et al., 1999; Rodriguez et al., 1999) in others it seems to have no effect (Kwek et al., 2001). The sumoylation-deficient K386R p53 mutant, when expressed in p53-null cells, exhibits higher transcription activity and binds better than the wild-type protein to the endogenous p21 gene promoter. Moreover, there are evidences in literature indicating that ectopic expression of PIAS appears to inhibit rather than induce p53-mediated

transcription (Schmidt and Müller, 2002). One explanation for these controversial data are that the majority of assays on p53 sumoylation have been performed under ectopic expression of SUMO (or other members of the sumoylation cascade) that can modify many proteins within the cell. Remarkably, several p53 molecular partners are themselves SUMO targets, such as MDM2 (Momand et al., 2000; Buschmann et al., 2000), or the kinase HIPK2, that causes p53 activation by phosphorylation on Ser 46 (Kim et al., 1999; Hofmann et al., 2002). The possibility to purify high amount of sumoylated p53 gave the possibility to analyse in detail the role of p53 sumoylation in p53 binding to chromatin and its effect on transcription through in vitro assays (Wu and Chiang, 2009). The tetramer is the predominant form of nuclear p53, and also the main substrate for post-translational modifications. In vitro studies, however, show that p53 sumoylation occurs preferentially on only 2 subunits of the tetramer. Sumoylated p53 cannot bind chromatin as well as induce p300-mediated acetylation of chromatin thus interfering at epigenetic levels with transcription. Although p300 binding to p53 is not affected by SUMOylation, the sumo moiety inhibits the accessibility to lysine residues adjacent to the sumoylation site, thus inhibiting p53 acetylation. Interestingly, while p53 sumoylation at K386 prevents its subsequent acetylation by p300, acetylated p53 remains permissive for sumoylation at K386 thus suggesting that K386 is not a major acetylation sites for p53 in vivo. It appears that sumoylation fails to disengage prebound p53 from DNA when acetylation already took place. These data also show how p53 sumoylation influences allosteric changes in p53 DNA binding domain with no effect on tetramerization of free p53 molecules. As sumoylated p53 binds DNA but it is transcriptionally impaired, sumoylation might have a role in the recruitment of transcriptional co-repressors, such as mSin3A whose DNA binding is increased when p53 is sumoylated. Components of either the nucleasome remodelling or the deacetylate and demethylase complex are also similarly recruited to the DNA with higher efficiency (Stielow et al., 2008; Ouyang et al., 2009). This specific mechanism could explain the negative role of p53 on target genes that have a role in pluripotency, such as AFP and Nanog (Lin et al., 2005).

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Role of sumoylation of p53 localization In addition to having a role in transcription, sumoylation appears to play a role in p53 subcellular localization. A recent study confirmed the negative role of sumoylation in p53 transcriptional activity and described how it affects p53 re-localization to the cytoplasm. The p53 protein shuttles between the cytoplasm and the nucleus. In particular, sumoylation appears to have a role in promoting the dynamic interaction of p53 with the exportin protein CRM1 (Santiago et al., 2013). CRM1 (Chromosomal region maintenance 1) is nuclear export receptor belonging to the karyopherin-β family of transporter proteins. It allows diverse cargoes, including proteins, small nuclear RNAs, and ribosomal subunits, to be delivered alternatively to the cytoplasm or to the nucleus (Cook and Conti, 2010; Güttler and Görlich, 2011). A genetic construct in which the sumo polypeptide was fused in frame with the C-terminal domain of p53 showed that SUMOylation was strictly required to dictate p53 localization to the cytoplasm. Two putative nuclear export sequences (NES) are present within the p53 protein sequence, one at the N-terminal and the other within the oligomerization domains (Stommel et al., 1999; Zhang and Xiong, 2001). Both of them are masked in the p53 tetrameric status. Interestingly, sumoylation does not affect p53 ability to form tetramers that are the only p53 species that can effectively bind the CRM1. By using several p53 and CRM1 point mutants, authors depicted a model in which p53 bind CRM1 during its travel through the pore complex. Sumoylation of p53 facilitates the disassembly of the transporting complex and the release of the cargo, in this case sumoylated p53, to the cytoplasm. While the described mechanism appears to be ubiquitin-independent, it has been shown that p53 shuttling can be synergically regulated by ubiquitination and sumoylation. Mono-ubiquitination seems to enable nuclear export of p53 as ubiquitin unmasks the NES and increases levels of monomeric p53 (Li et al., 2003). It has been shown that by increasing NES exposure, mono-ubiquitination promotes PIAS4-mediated p53 sumoylation (Carter et al., 2007; Carter and Vousden, 2008). In line with this, previous work has shown that a p53 mutant deficient for MDM2 binding is poorly sumoylated (Chen and Chen, 2003). The suggested molecular model describes that Mdm2-mediated

ubiquitination can unmask a nuclear export signal and facilitate the recruitment of PIAS4. Subsequent sumoylation of p53 also appears to be required for MDM2 release after mono-ubiquitination, to allow nuclear export (Stehmeier and Muller, 2009; Hock and Vousden, 2010). This model reveals an interesting novel aspect in SUMO-dependent regulation of p53 and in particular strengthens the concept of interconnections between sumoylation and ubiquitination and the intricacy of p53 regulation by post translational modifications. Interestingly, although sumoylation of K386 blocks ubiquitination of this site, it does not seem to interfere with the ubiquitination of other lysine residues that are freely accessible. The picture of p53 post translational modification by sumo is further complicated by the observation that poly-sumoylated chains can be assembled on this protein. It has been shown that p53 interacts with the protein TRIM2 that targets p53 for SUMO-2 modification. The higher eukaryotes have three different SUMOs that are encoded by three distinct genes and which are named, respectively, SUMO1, SUMO2 and SUMO3. SUMO 2 and 3 show an identity equal to 96% because they differ only for three amino acids at the N-terminal end; instead, they have an identity equal to 46% with SUMO (Watson and Irwin, 2006). SUMO 1 is prevalently present as a conjugated form to other proteins while, SUMO2 and 3 are mostly in free form. In particular, SUMO2 and SUMO3, in analogy with ubiquitin, have internal lysines that allow the formation of poly-sumoylation chains; these chains have a role in proteasome degradation of the target proteins as evidenced by treatments with the proteasome inhibitor MG132 (Vertegaal, 2010). Interestingly, external stimuli such as thermal or oxidative stress can increase the levels of SUMO 2 and 3. The study shows that high levels of TRIML2 corresponds to higher levels of expression of those p53 target genes induced upon prolonged DNA damage and apoptosis (Kung et al., 2015). In presence of TRIML2, high molecular weight p53 immuno-reactive band corresponding to poly-sumoylated p53 species become apparent. Interestingly, in human immortalized keratinocytes also p63 appears to undergo to sumoylation. Moreover, SUMO-2 conjugation takes place with higher efficiency respect to SUMO 1 (Pollice et al., 2008; Vivo et al., 2009). SUMO 2 and 3 increase

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during epithelial differentiation thus suggesting that in physiological conditions, during cellular differentiation sumoylation can be a mean to downregulate p63 expression. Besides nucleo-cytoplasmic shuttling, a subfraction of nuclear p53 molecules move from the nucleoplasm to the so called ‘nuclear bodies’, proteinaceous aggregates in which the PML (Pro-Myelocytic Leukaemia) protein is enriched (Bernardi and Pandolfi, 2007). These structures have a role in ensuing post translational modifications of several proteins but they also have a storage functions. In particular, PML bodies nucleation is primed by PML sumoylation that consequently triggers and PML multimerization through Sumo Interacting motif (SIM) present both in the protein itself and SUMO (Shen et al., 2006). During senescence or stress, it has been shown that p53 is recruited to nuclear bodies, where it possibly can be subjected to additional post translational modifications (Stehmeier and Muller, 2009; Marcos-Villar et al., 2013). Studies in yeast and Drosophila propose a model in which p53 and PML co-localization in nuclear bodies depends at least in part on sumoylation of lysine 386 (Di Ventura et al., 2008; Mauri et al., 2008). However, conflicting experimental evidence did not clarify the exact role of sumoylation in p53 recruitment to NBs as its localization does not appear to be regulated by K386 sumoylation. The ARF/MDM2/p53 triangle Of particular interest has been the discovery that hyper-proliferative stimuli can induce stabilization of p53 by enlisting the activity of ARF. ARF role in p53 stabilization has been studied in detail, given the essential role of their functional interaction in cellular tumour suppression mechanisms. ARF is expressed at very low levels in normal cells and acts as a sensor of hyper-proliferative signals emanating from oncoproteins such as E1A, Ras and inducers of S-phase entry like Myc. When proliferative signals that are normally required for cell proliferation exceed a critical threshold, a p53 dependent oncogene checkpoint gated by ARF is activated, ARF triggers growth arrest and, in the presence of appropriate collateral signals, sensitizes cells to apoptosis. Interestingly, ARF suppresses aberrant cell growth in response to oncogene activation, at least in part, by inducing the p53 pathway through MDM2 binding and inhibition of MDM2-mediated p53

degradation allowing p53 to accumulate in the nucleus (de Stanchina et al., 1998; Kamijo et al., 1998; Palmero et al., 1998; Sherr, 1998; Stott et al., 1998; Zhang et al., 1998; Vivo et al., 2015). However, ARF also displays pro-proliferative functions in different cell contexts (Fontana et al., 2018). ARF can also promote the conjugation of the small ubiquitin-like protein SUMO-1 to its binding partners and among these, p53 and MDM2 (Xirodimas et al., 2001a; Xirodimas et al., 2002; Chen and Chen, 2003; Vivo et al., 2017). In particular, the interaction between ARF and HDM2 is essential to determine HDM2 sumoylation and ARF binding to MDM2 is required in order to achieve p53 sumoylation. In line with the hypothesis that both proteins cooperate in the process, simultaneous MDM2 and ARF expression increases protein levels of sumoylated p53. Accordingly, sumoylation experiments performed in vitro show no increase in p53 sumoylation when only ARF is expressed. While the MDM2 RING domain is not involved in p53 sumoylation, ARF domain located in the region of the protein encoded by exon 2 appears to be required for this function. This region of ARF is very conserved between human and mouse and has been correlated to ARF ability to relocate p53 in the nucleolus. In particular, the performed experiments led to the conclusion that, while the exon1-encoded domain is required to inhibit MDM2 ubiquitin ligase activity towards p53, the regions located within the exon 2 of ARF are instead required for sumoylation of both p53 and MDM2. Sumoylation of p53 can be seen as a way to inhibit the ubiquitination and thus degradation of p53. In line with this SUMO-1, E1, and E2 mainly localize within the nucleus and in vivo sumoylation requires nuclear localization of the substrate (Rodríguez, 2014). It would be interesting to understand how nuclear compartments and nuclear structure dynamics can interfere with these mechanisms. These evidence supports the idea that p53 localization might have a role in determining its own PTM, by increasing the sumoylation rate or through protection from ubiquitin mediated proteolytic degradation. In line with this, ARF ability to induce sumoylation has been related to its ability to antagonize ubiquitination, causing protein stabilization (Wang et al., 2015). Sumoylation experiments in which both ARF and MDM2 were expressed ectopically in human cells confirmed that, although the major

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sumoylation site in p53 is K386, weak signal of sumoylation was still detectable on the p53K386 mutant after co-expression of both ARF and MDM2 (Xirodimas et al., 2002). This suggests the existence of alternative or cryptic sumoylation sites within the protein sequence. This has been proved to be the case of MDM2, in which two putative lysine residues have been mapped, neither in a consensus sequence for sumoylation. MDM2 mutants in which these sites were mutated demonstrates that alternative lysine can be modified once the major is mutated or modified by ubiquitination or acetylation. This behaviour suggests and confirms how the functionality of a protein can be modified by a complex set of PTM not simple to decode. It is still unclear whether ARF and MDM2 act by stimulating the rate of p53 sumoylation, by inhibiting desumoylation, by stabilization of sumoylated p53, or a sum of these events. It should be emphasized that ARF role in sumoylation has been related to its ability to block the SUMO-2/3 deconjugating protease Senp3. In particular, the p19Arf protein triggers the sequential phosphorylation, poly-ubiquitination and rapid proteasomal degradation of Senp3 (Kuo et al., 2008) The transcription factor p63 in development and tumorigenesis The p63 protein, unlike p53, plays a fundamental role in the developmental processes of the limbs, the skull, the epithelium and its derivatives (glands, hairs and teeth). Knockout of the TP63 gene in mice is lethal in the post-natal phase and determines the lack of stratification of the epidermis. This causes dehydration and death a few hours after birth. The p63–/– mice show severe epithelial defects such as absence of hair, skin, salivary and mammary glands and absence of the prostate. Moreover, the lack of epidermal stratification generates defects in limb development. Limb originates from the apical ectodermal ridge in the limb sketch. It is composed of a pseudo-stratified epithelium absent in these knockout mice. In KO mice, the hind legs are completely absent, while the forelimbs are underdeveloped and lack some parts. The balance of the different p63 isoforms of appears to have a relevant role to guarantee the correct process of terminal differentiation, both in embryogenesis, during the development of

the epidermis and its derivatives, and in the adult organism. Primary mouse keratinocytes differently express the various p63 isoforms (ΔN and TA) during differentiation. In the adult, ΔNp63α is abundant in the cells of the basal layer and is almost completely absent in the upper layers of the epidermis (Truong et al., 2006). It has been shown that this isoform is required for the maintenance of the proliferative potential of basal keratinocytes that continuously replenish the skin (Yang et al., 1998). By blocking calcium-induced differentiation ΔNp63α keeps cells in active proliferation (Truong et al., 2006). In particular, in the basal layer, ΔNp63α represses the transcription of genes required for keratinocyte terminal differentiation such as p21Waf and 14-3-3 σ, by promoter interaction (Westfall et al., 2003). In the upper layers of the epidermis, the expression of ΔNp63α decreases in favour of ΔNp63γ and TAp63 isoforms, thus allowing the expression of genes required for terminal differentiation (Carroll et al., 2006). In fact, in the mature epidermis the expression of TA isoforms seems to be linked to skin protection mechanisms triggered by cellular stresses such as exposure to UV rays, that by increasing the intracellular levels of TA isoforms triggers apoptosis (Yang and McKeon, 2000). Thus, alternation of various p63 isoforms, characterized by different trans-activating capacities on distinct promoters, is crucial for driving correct stratification and differentiation of embryonic and adult skin. As TAp63 and ΔNp63 isoforms play contrasting roles in the process of tumorigenesis, defining the role of these isoforms has been a difficult task. TP63 is rarely mutated in human cancer, but p63 activity is often increased. ΔNp63 is supposed to behave as oncoprotein and is up-regulated in squamous cell carcinomas of the cervix, ovaries and lung (Yang et al., 1998) and triple negative basal-like breast tumours (Troiano et al., 2015; Holcakova et al., 2017). It also plays roles in a variety of pathways that are implicated in CSC properties, reviewed in (Nekulova et al., 2011). ΔNp63 increases the expression of Wnt receptor Frizzled 7 thereby enhancing Wnt signalling which promotes stem cell activity of normal mammary and tumour initiating activity in the basal-like subtype of breast cancer (Chakrabarti et al., 2014). On the other hand, TAp63 shares the abilities of the ‘guardian of the genome’ p53 to induce cell cycle arrest and

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apoptosis and thus may act as tumour suppressor. In contrast, TAp63 is the predominant isoform expressed in haematological malignancies (Alexandrova and Moll, 2012), and its overexpression leads to increased tumour progression of head and neck squamous cell carcinoma. It is also expressed in colon carcinoma (Nylander et al., 2002). Interestingly, Su et al. (2017) have recently shown that TAp63 is crucial for the transition of mammary cancer cells to tumour initiating cells. The ability of ΔNp63 to function as oncogene is probably due to its ability to antagonize p53 and TA isoforms not only through DNA binding competition but also through the formation of transcriptionally inactive hetero-oligomers (Yang et al., 2002b). Therefore, it is reasonable to postulate that in some tumours an imbalance between the ‘oncosuppressor’ TA and ‘oncogenic’ ΔN isoforms can cause the de-regulation or more in general an alteration of p63 activity. Role of ubiquitination in p63 functions Both TA and ΔNp63 are tightly regulated at protein level. ΔNp63 isoforms are very stable compared to TAp63, which are expressed at low levels, have a relatively short half-life and display pro-apoptotic activity. Ubiquitination is a common pathway for p63 regulation, usually via negative regulation of p63 isoforms through the ubiquitin-proteasome system (Armstrong et al., 2016). Several E3 ubiquitin ligases regulate the p63 protein. A combination of biochemical and embryological approaches in HEK293 cells and zebrafish embryos demonstrated that ΔNp63α protein is destabilized by ubiquitination, partly mediated by the HECT-type E3 ubiquitin ligase Nedd4. Using a yeast-two-hybrid screening system, Nedd4 and the SUMO-conjugating enzyme Ubc9 were found to bind to distinct sites in the C-terminal region of ΔNp63α. The WW domains of Nedd4 interact with a proline rich domain (PPPY) of ΔNp63α upstream of the SAM domain. Thus, a single point mutation in this proline-rich domain (Y449F) of ΔNp63α abolishes the interaction with Nedd4 thus resulting in ΔNp63α stabilization. The physical interaction with Ubc9 and Nedd4 lead to ΔNp63α ubiquitination and Sumoylation, resulting in vulnerability of ΔNp63α to proteasomal degradation (Bakkers et al., 2005). In zebrafish, high expression of ΔNp63α is restricted to the dorsal region of the

embryo where Ubc9 and Nedd4 are expressed at low levels. Mutant versions of ΔNp63α unable to bind Nedd4 (Y449F) or Ubc9 (Q634X) exhibit a more uniform and ubiquitous expression when expressed upon mRNA microinjection (Bakkers et al., 2005). Mdm2, in analogy with p53, also mono-ubiquitinates p63 but it is unable to cause its degradation (Kadakia et al., 2001). Mdm2 interaction with p63 is capable of interfering with its transactivation function, likely by exporting p63 protein from the nucleus into the cytoplasm. The effect of Mdm2 on p63 functions remains controversial likely because of difference in the cell contexts where experiments have been performed. One study found Mdm2 unable to inhibit p63 function (Little and Jochemsen, 2001), another found that Mdm2 actually stabilized p63, increasing both its expression and its function (Calabrò et al., 2002), while yet another found no interaction between Mdm2 and p63 (Wang et al., 2001). Although Mdm2 is unable of targeting p63 for degradation, it can cooperate with the F-box ligase Fbw7 to poly-ubiquitinate ΔNp63α and target it for proteasome degradation (Galli et al., 2010). MdmX is an E3 ligase related to Mdm2, but it does not have the ability to target p63 for degradation or interfere with its functions (Kadakia et al., 2001). P63 was also found to be targeted for degradation by a HECT E3 ligase known as Itch/ AIP4 (atrophin-1 interacting protein 4) (Rossi et al., 2006). Sumoylation dependent regulation of p63 Among the targets of sumoylation there is the ΔNp63α isoform. Inspection of the p63 sequence revealed the presence of the tetrapeptide IKEE, centred on lysine 637 within the p63 transcriptional inhibitory domain TID of p63 (Ghioni et al., 2005) (Fig. 11.4). Ghioni and collaborators have shown that SUMO is able to regulate the protein levels of the ΔNp63α isoform inducing its degradation via proteasome and that the sumoylation site of p63 is precisely the lysine 637 in the post-SAM domain of the α isoforms. Mutant p63 in which lysine 637 has been replaced by an arginine (K637R), is not sumoylated nor degraded by the proteasome; this indicates that lysine 637 is necessary for the conjugation of SUMO and consequent de-stabilization of the ΔNαp63 protein.

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ΔNp63α, but not β and γ, is sumoylated in vitro and in vivo at the K637 residue, in the post-SAM domain of the α isoform. Indeed, neither p63β nor p63γ lacking the C-terminal TID domain are sumoylated, in vitro and in vivo. SUMO conjugation appears to exert a negative regulatory function on p63. Remarkably, natural p63 mutants showing alterations in their sumoylation capacity showed an overall clear increased transcriptional activation ability compared to the wild type isoform: on the other hand, whether p63 is sumoylatable or not sumoylatable resulted in very minor differences in transcriptional repression (Ghioni et al., 2005). Deletion of the eight C-terminal residues of p63α that contains the SUMO attachment site enhances its activity in reporter gene assays (Serber et al., 2002). This appears to be mainly due to the loss of sumoylation, since the non-sumoylatable natural mutants TAp63 αK637R and ΔNp63αK637R associated to the human Split Hand and Foot Malformation (SHFM IV) exhibit a much stronger activation potential than their wild type counterpart on a reporter system with the luciferase gene under the control of an artificial p53-responsive promoter (Ghioni et al., 2002). These data support the general idea that sumoylation of p63α limits its transcriptional activity and leads to functional inactivation. The inhibitory effects of SUMO-1 on p63 transcriptional activity does not seem to be directly involved in suppressing the intrinsic transcriptional activity of p63 rather it seems to act indirectly by controlling p63 intracellular level. Regarding the control of p63 protein level, the SUMO system is tightly interconnected to the ubiquitin system. In the simplest scenario, attachment of SUMO to a distinct lysine residue directly opposes ubiquitination by shielding the residue from ubiquitination (Gill, 2004). Several groups, instead, have delineated a conserved pathway, in which sumoylation and ubiquitination cooperate in protein degradation (Schimmel et al., 2008). The core of this pathway is a family of SUMO-targeted RING-type ubiquitin ligases (STUbL), which contains SUMO interaction motifs (SIM) and are therefore preferentially recruited to a SUMO modified substrate. The prototypic member of this family in mammalian cells is RNF4 (Kumar et al., 2017) that contains 4 SIMs in the N-terminal region and a RING domain in the C-terminal. Proteins of this family are recruited to the previously sumoylated

proteins that are substrate for the attachment of ubiquitin chains. The C-terminal region of p63α also includes the binding consensus site for Ubc9, a SUMO-1 conjugating enzyme that has been previously shown to bind a conserved site in the C-terminus of p73α. SUMO-1 induces p63 protein instability and is counteracted by the proteasome inhibitor MG132 (Ghioni et al., 2005). Moreover, expression of SUMO1 destabilizes wild type p63, but not the sumoylation-deficient variant p63K637R in a proteasome-dependent way. Accordingly, ΔNp63K637R is less efficiently ubiquitinated than wild type ΔNp63α. Association between p63 and SUMO-1 is completely abolished by a K637E mutation and the same mutation leads to a dramatic increase in TAp63α transcriptional activity (Straub et al., 2010). The ΔNp63αE639X mutant, also isolated from a patient affected by SHFMIV, is neither sumoylated nor ubiquitinated. The E639X mutation, indeed, disrupts the recognition sequence required for proper SUMO-conjugation of ΔNp63α, although the SUMO acceptor site at lysine 637 remains intact. Further insights on the crosstalk between sumoylation and ubiquitination in the control of ΔNp63α was provided by the generation of an artificial ΔNp63α mutant protein carrying K193 and K194 substitutions into glutamic acid (K193E/ K194E). Lysine residues at position 193 and 194 of ΔNp63α were previously shown to be necessary for the Itch ubiquitin ligase-mediated degradation of p63 (Rossi et al., 2006) and are located in the DNA binding domain (Fig. 11.4). Moreover, their substitution partially protect ΔNp63α from MDM2 or FBW7-mediated degradation (Galli et al., 2010). The artificial ΔNp63α mutant protein carrying K193 and K194 substitutions into glutamic acid (K193E/K194E) was not sumoylated thus clearly indicating that reduction of p63 ubiquitination has a negative impact on sumoylation. In conclusion, inefficient p63 ubiquitination impairs p63 sumoylation and degradation thus providing evidence that SUMO and Ub modifications are not redundant and both are required to guarantee efficient ΔNp63α degradation (Ranieri et al., 2018). TAp63α protein are regulated to a lower extent compared to ΔNp63α by exogenous SUMO-1, possibly because of the intramolecular masking that is known to occur in this isoform between the

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transcriptional activation domain TA and the TID domain that could prevent sumoylation (Ghioni et al., 2005). On the other side, the level of β and γ isoforms, both TA and ΔN, lacking the consensus for SUMO-1 site, were unaffected by SUMO-1 co-transfection. The evidence that several p63 mutants associated to human hereditary ectodermal dysplasia syndromes lost SUMO-dependent regulation indicates that regulation of p63 protein by the SUMO machinery is an essential step during development. The natural AEC Q540L mutation is associated with the Hay Wells syndrome, a rare hereditary ectodermal dysplasia. The Q540L mutation is a missense mutation that prevents the intramolecular folding that normally masks the SUMO-1 site. As consequence, the TAp63Q540L protein is destabilized by SUMO-1 overexpression while the wild type protein is SUMO-resistant. The ΔNp63Q540L is instead SUMO-1-sensitive as the wild type protein, suggesting that this mutation targets mainly the TAp63α isoform (McGrath et al., 2001). Finally, data from our group have revealed that p14ARF oncosuppressor gene targets ΔNp63α for proteasomal degradation in keratinocytes by preferential SUMO-2 conjugation (Vivo et al., 2009). ARF is able to bind p63 (both TA and ΔN) and repress the transcriptional activity of some isoforms (Calabrò et al., 2004). During calciuminduced keratinocytes differentiation, the ARF protein levels increase. Moreover, in vitro assays have shown that ARF induces p63 sumoylation. ARF and SUMO co-expression increases the efficiency of ΔNαp63 sumoylation and its subsequent degradation by the proteasome complex. A possible explanation of this phenomenon is that ARF acts as a molecular adapter between Ubc9 and p63. ARF and SUMO-mediated p63 degradation occurs only if the consensus sequence and the sumoylation site are intact; in fact, neither ARF nor SUMO are able to induce the sumoylation and degradation of the ΔNαp63E639X and ΔNαp63K637R mutants (Vivo et al., 2009). Sumoylation and ubiquitination interplay in human hereditary syndromes It has been observed that inhibition of sumoylation in cells of the epidermal basal layer does not cause changes in keratinocytes growth and morphology.

In contrast, when differentiation is induced and sumoylation is inhibited, cells undergo morphological alterations. During keratinocyte differentiation components of sumoylation machinery such as SAE1, SAE2, SUMO 2 and 3 are induced with the consequent sumoylation of their target proteins (Deyrieux et al., 2007). These results suggest that sumoylation has, therefore, an important role in epithelial differentiation. In human, hereditary syndromes involving limb development and/or ectodermal dysplasia have been identified due to missense, non-sense or frameshift mutations of the TP63 gene. These human syndromes are the EEC, (ectrodactylydysplasia ectodermica-labiopalatoschisi), the AEC/RHS (ankyloblepharon-ectodermal dysplasia-clefting/Rapp Hodgkin syndrome), ADULT (acro-dermato-ungual-lacrimal-tooth), LMS, (Limb Mammary Syndrome), and the SHFM (split-hand/foot malformation). All of these disorders are the consequence of mutations at a single p63 allele and, if one can extrapolate from the mouse knockout model, are not the results of haploinsufficiency. The pattern of mutations linked to p63 reveals a remarkable specificity of the molecular defects in this gene and the clinical consequences. The analysis of rare patients affected by these syndromes has brought to light a significant genotype-phenotype correlation; in fact, the localization of these mutations determines what are the functional changes of p63 and therefore the clinical consequences. However, the fact that this protein exists in different isoforms with distinct and sometimes contradictory biological activities makes it difficult to establish the role of p63 in the pathogenesis of ectodermal dysplasia. The SHFM is a limb malformation involving the central rays of the autopod (hand/foot). SHFM may present with syndactyly, median clefts of the hands and feet, and aplasia and/or hypoplasia of the phalanges. In severe cases, the hands and feet have a lobster claw-like appearance. SHFM is genetically heterogeneous with several identified locus: the autosomal dominant form (locus SHFM1), the X-linked (locus SHFM2) and the recessive form (locus SHFM3). Many families with SHFM, however, do not map in any of these known chromosomal regions. These mutations map to a previously un-mapped region in 3q27-28

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(locus SHFM4) and p63. P63 mutations in SHFM included two missense mutations in lysine residues that are target of ubiquitination such as K193E and K194E. Moreover, the other two non-sense mutations, one affecting a glutamine at position 634 (Q634X) and the other a glutamic acid at position 639 (E639X), are both located in the C-terminal domain of p63α isotypes. Moreover, although leaving intact the SUMO acceptor site at K637, the E639 mutation falls in the SUMO-1 φKXD/E motif thus disrupting the recognition sequence required for proper SUMO-conjugation (Ghioni et al., 2005). Natural p63 frameshift mutants showing alteration in their sumoylation capacity are also associated to LMS (Duijf et al., 2002). The artificial K193E/K194E double mutant and the sumoylation-defective ΔNp63αΕ639X mutant were shown to be both ubiquitination defective thus implying that SUMO-conjugation is required for efficient ΔNp63α ubiquitination (Ranieri et al., 2018). Moreover, compared to wild type ΔNp63α, both ΔNp63αE639X and ΔNp63αK637R mutants were less ubiquitinated under enforced expression of ubiquitin thus confirming that ubiquitin conjugation cannot properly occur if p63 sumoylation is impaired (Ranieri et al., 2018). Remarkably, the ubiquitination defective K193E/K194E p63 mutant also displays impaired ability to undergo sumoylation and is resistant to both SUMO and ARF-mediated degradation thus suggesting that inefficient p63 ubiquitination impairs not only p63 degradation but also its ability to undergo proper sumoylation. Taken together these findings indicate a tight intertwining between ubiquitination and sumoylation in the control of p63 protein stability (Fig. 11.7). Role of p73 in neurological development and cancer The TP73 gene maps to a chromosomal locus (1p36.3) often deleted in neuroectodermal human cancers such as neuroblastomas (Kaghad et al., 1997). Mice in which the locus has been artificially deleted present significant neurological abnormalities (De Laurenzi et al., 2000; Yang et al., 2000; Alexandrova et al., 2013). ΔNp73 is the predominant isoform in the murine foetal nervous system with pro survival properties (Ishimoto et

Figure 11.7  SUMO and Ubiquitin conjugation mediated degredation of p63. In absence of sumoylation p63 is inefficiently ubiquitinated. Moreover, ubiquitination is not sufficient to trigger p63 proteasome mediated degradation. It is not clear if p63 sumoylation can take place only on p63 sumoylation sites or on polyubiquitin chains or if both mechanisms occur.

al., 2002; Dulloo et al., 2010). Its loss likely leads to enhanced apoptosis in cortical and sympathetic ganglia neurons resulting in either the absence or the loss of specific populations of neurons. This effect seems to be the result, at least in part, of uncontrolled apoptosis mediated by the full-length proapoptotic p53 family proteins, p53, TAp63, and TAp73 of which ΔNp73 is an inhibitor (Vossio et al., 2002). In fact, the ΔNp73 protein can function as a dominant negative towards p53 and TAp73, similarly to ΔNp63 (Lee et al., 2004). p73 mutations, as well as p63’s, are rarely observed in human cancer, but several studies have shown that ΔNp73 can enhance transformation by oncogenes such as Ras. TAp73 is induced by a wide variety of chemotherapeutic agents (Agami et al., 1999; Gong et al., 1999). Blocking TAp73 function promotes survival and leads to enhanced chemoresistance (Bergamaschi et al., 2003; Irwin et al., 2003; Rocco et al., 2006). Although the loss of p63 and p73 does not make mice tumour prone, accumulating evidence suggests that the

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relative balance between the TA and ΔN isoforms may contribute to tumorigenesis. The full-length TA isoforms of p63 and p73 have pro-apoptotic tumour suppressor-like functions, while the ΔN isoforms behave as oncogenes. Specifically, ΔNp73 expression increased in breast, ovarian, hepatocellular, prostate, colon cancer, and neuroblastoma and has been associated with poor prognosis in patients. Given its dominant effect towards p53 and TAp73, ΔNp73 overexpression inhibits apoptosis and enhances chemoresistance (Concin et al., 2005; Domínguez et al., 2006). Interestingly, in HNSCC cells a complex pattern of regulation of the different isoforms takes place. ΔNp63α overexpression enhances cancer cell survival by inhibiting TAp73-dependent apoptosis by at least two different mechanisms, by physical interaction with TAp73 and competition for the binding of canonical p73 target promoters (Rocco et al., 2006). Data from mouse models and human tumours suggest that the balance between the expression of p53, p63, and p73 and their distinct TA and ΔN isoforms likely affects the outcome of signalling pathways leading to apoptosis or survival. Therefore, understanding the regulatory mechanisms, such as posttranslational modifications, that differentially modulate TA and ΔN isoform activity and stability are of particular interest because therapeutic modulation of the proapoptotic and antiapoptotic isoforms of the p53 family has potential therapeutic benefits in treating human cancers. Regulation of p73 functions by ubiquitination Preliminary assays with proteasome inhibitors suggested that the ubiquitin–proteasomal pathway regulates p73 stability (Bálint et al., 1999). Moreover, ubiquitinated p73 proteins strongly accumulate in cells expressing exogenous ubiquitin after treatment with proteasome inhibitors (Bernassola et al., 2004). Upon DNA damage and chemotherapeutic treatment ΔNp73 isoform is rapidly ubiquitinated and degraded by the proteasome, while TAp73 and p53 are not affected (Maisse et al., 2004). The selective down-regulation of ΔN isoforms can thus lead to increased apoptosis induced by TA or p53. Ubiquitin-mediated regulation of the stability and activity of the various ‘tumour suppressor-like’ TA and ‘oncogenic’ ΔN isoforms of p63 and p73 may

play a role in cancer development and response to chemotherapy. The first identified E3 ligase promoting p73 ubiquitination is NEDL2, a NEDD4-related HECT domain protein (Miyazaki et al., 2003). This protein directly interacts with p73. Interestingly, this region is also a binding site for another NEDD4-related E3 ubiquitin ligase, Itch, that also bind p63 (Rossi et al., 2005, 2006). Both Itch and NELD2 contain WW motifs that mediate protein–protein interactions. Although promoting p73 ubiquitination, NEDL2 interaction results in stabilization and increased TAp73 transcriptional activity. In contrast, Itch promotes ubiquitination and subsequent degradation of both ΔN and TAα isoforms of p73. The observation that, upon DNA damage also Itch is down-regulated may explain how TAp73 is stabilized following treatment with chemotherapeutic agents but leave the open question on how ubiquitination and degradation of ΔNp73 isoforms occur. The authors suggest that Itch could plays a role in maintaining both TA and ΔN isoforms at low levels under normal unstressed conditions, while other specific mechanisms take place on stress. In addition to Itch, also the F-box protein FBXO45 controls p73 ubiquitination and the proteasome-dependent degradation of p73 in normal conditions in order to maintain p73 expression at low levels (Rossi et al., 2005; Peschiaroli et al., 2009). Following DNA damage, both ITCH and FBXO45 are down-regulated leading to p73 accumulation and to p53-independent cell death (Wu and Leng, 2015). The imbalance between TAp73 and ΔNp73 protein levels appears to be of great importance in both tumorigenesis and resistance to chemotherapy. TAp73 rapidly accumulates in response to genotoxic stress. This is due to two different mechanisms, reduced degradation along with increased stabilization by acetylation, tyrosine phosphorylation, and PML interaction (Bergamaschi et al., 2003). When activated in response to DNA damage, TAp73 binds to p53-responsive elements located in target genes that induce cell cycle arrest, senescence, or apoptosis. In contrast, ΔNp73 isoforms are instead preferentially degraded. Upon microarray analysis to identify the transcriptional targets of p73, a ubiquitin ligase named PIR2 was identified that specifically targets ΔNp73 inducing its ubiquitindependent degradation. The expression of Pir2 is

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thus able to increase the TA/ΔNp73 protein ratio. Thanks to this differential function, co-expression of PIR2 along with ΔNp73 reliefs the inhibitory effect of ΔNp73 on TAp73 mediated apoptosis (Sayan et al., 2010). Finally, several other molecules such and c-Jun and Yes-Associated Protein (YAP) appear to critically regulate the balance between TA and ΔN p73 isoforms through mechanisms that are ubiquitin and proteasome independent to induce TAp73-mediated apoptosis (Toh et al., 2004; Danovi et al., 2008; Dulloo et al., 2010). MDM2, although a binding partner of both p63 and p73, does not play a role in their degradation. Overexpression of MDM2 appears to stabilize p63 and p73 (Ongkeko et al., 1999; Zeng et al., 1999). MDM2 is transcriptionally activated by p73 and represses the functions of p73, including p73-dependent transactivation and growth suppression. In particular, the ability to inhibit p73-dependent apoptosis and cell cycle arrest is strictly dependent on the E3 ubiquitin ligase properties of Mdm2. It has been recently shown indeed that Mdm2 mainly utilizes K11, K29 and K63linked chains to mediate p73 ubiquitination in vivo and in vitro (Wu and Leng, 2015). Interestingly, MDM2 mediated ubiquitination is not sufficient to induce p73 degradation, but the subsequent binding of ubiquitinated p73 with Itch is the signal that triggers proteasomal degradation. Analysis of the role of MDM2 in p53 degradation by mutational and comparative studies demonstrated that following MDM2 binding to p53, a 20 amino-acids stretch within the p53 sequence (aa 92–112) is required for p53 destabilization, thus functioning as degradation signal. This sequence is missing in p73 protein, thus explaining why MDM2 cannot induce p73 degradation. Moreover, this observation confirms that ubiquitination is not sufficient to induce protein degradation, as has also been shown for p63 (Ranieri et al., 2018). Pirh2, a RING finger E3 ubiquitin ligase, physically associates with TAp73 and promotes TAp73 poly-ubiquitination and proteasomal degradation. Pirh2, is transcriptionally regulated by p53 and in turn targets p53 for degradation thus participating in a negative autoregulatory feedback loop analogous to MDM2 (Leng et al., 2003). Differently from MDM2 that down-regulate p53 in normal conditions, Pirh2 inhibits p53 activation following stress, while Ser15 phosphorylation of p53 alleviates

MDM2 inhibition. Several pieces of evidence suggested that Pirh2 may promote tumorigenesis in both p53-dependent and independent manner. It functions as E3 ubiquitin ligase for TAp73, and its depletion restore TAp73-mediated growth inhibition in p53-deficient cancers ( Jung et al., 2011b). Finally, ectopic expression of Pirh2 repressed p73-dependent transcriptional activity. In contrast with the previous study, the author did not observe destabilization of p73 in their experimental conditions thus suggesting that ubiquitination per se does not imply degradation (Wu et al., 2011b). Interestingly, it was shown that while in vitro Pirh2 can induce mainly K63 ubiquitination of p73, in vivo a complex pattern of ubiquitination takes place with other lysine residues of ubiquitin involved. Moreover, a different ubiquitination pattern also takes place on different p73 isoforms. Differences in the pattern of ubiquitination between p73α and p73β have been reported; while Pirh2 primarily used Lys-63 to promote p73α ubiquitination, it uses multiple lysine residue to mediate the ubiquitination of p73β. The majority of the studies on p73 mediated apoptosis focused on transcriptional dependent apoptotic mechanisms. The observation that p73 can be recruited to the mitochondrion seems to suggest that it exerts additional nuclear-independent functions to induce cell death, in a way similar to p53. Several evidences suggested that p73 interacts with the E3 ubiquitin ligase Hades. This protein belongs to the RING domain family of ubiquitin ligase and localizes specifically to the outer membrane of mitochondria, facing the cytosol (Min et al., 2015). It has been identified as a molecular partner of p53 and to induce its poly-ubiquitination by targeting the N-ter lysine 24 in the TA domain of the protein. Its interaction with p53 results in the inhibition of cell death pathway triggered by p53 in the cytoplasm ( Jung et al., 2011a). Recently, it has been shown that Hades also interacts with p73, and upon apoptotic stress, the two proteins co-localize to mitochondria. Here, by increasing p73 polyubiquitination, Hades mediates p73 degradation through the proteasome thus blocking apoptosis. Regulation of p73 functions by sumoylation The p73 protein also appears to be regulated by sumoylation (Minty et al., 2000). SUMO-1

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covalently modifies both TA and ΔN isoforms of p73α at the conserved K627 thus excluding from the modification the shorter C-terminal isoforms that lack this lysine residue. The authors reported that sumoylation of p73α does not affect its transcriptional activity, but instead alters its subcellular localization to the nuclear matrix. In analogy to p53, also p73α binds Ubc9 and PIAS1. Differently from p53 in which the binding domains are partially overlapping, in p73 they are located in the C-terminus and in the oligomerization domain, respectively. Therefore, while Ubc9 can bind only to p73α isoforms, PIAS is able to interact with all p73 isoforms in the nucleus (Munarriz et al., 2004). As for p63, p73 sumoylation promotes but is not required for degradation. Indeed, it appears that sumo modification potentiates, but does not trigger p73 degradation (Minty et al., 2000). SUMO-1 modification may thus induce conformational changes potentiating ubiquitination or may influence protein degradation via modulation of others E3 ubiquitin ligases. A similar behaviour has been shown for p63 (Ranieri et al., 2018). In particular by blocking p63 sumoylation by Gam-1 (a viral protein that inhibits Ubc9 functions), p63 ubiquitination is similarly reduced. PIAS1 was also shown to stabilize p73α, but this stabilization was, in fact, independent of its sumoylation function. PIAS1 also inhibits TAp73α transcriptional activity, and this effect is dependent on the sumoylation function of PIAS1. In particular, PIASy overexpression inhibits p73α-mediated transcription of p21waf, causing a reduction of cells in G1 and cell cycle reentry (Zhang et al., 2010). The TAp73β isoform conversely, cannot be sumoylated and this could explain its higher basal transcriptional activity. It has been also proposed that, as it happens for p53, the covalent binding of a sumo moiety at the C-ter domain of p73 could interfere with p73 binding to transcriptional activators, such as the c-Abl tyrosine kinase or to chromatin remodelling complex, such as the histone deacetylation complex (White and Prives, 1999; Yuan et al., 1999). Conclusions Ubiquitination and sumoylation both play important roles in regulating the p53 family, and perturbations in these pathways have implications in both tumorigenesis and development.

The analysis of the molecular pathways inducing sumo and ubiquitin modification of the p53 family raises many questions about the role of these modifications. It appears that both in cancer progression and during development, the balance among the multiple isoforms of each locus is extremely important and thus finely regulated. The peculiar gene structure, although playing an essential role in the functions of the p53 family members raises the question of the regulation of the different isoforms. This regulation occurs at the transcriptional level through different promoter usage but also cotranscriptionally through regulation of splice sites selection or by alternative initiation of translation and is accompanied by an orchestra of post translational modifications such as phosphorylation, acetylation, ubiquitination, neddylation, sumoylation, and methylation that finely modulates the activity of p53 family members during the life of the cell. In the absence of stress signals, p53 protein is present at low levels, due to a dynamic and finely tuned balance between transcription and degradation. This dynamic equilibrium of p53 levels allows cells to maintain genetic stability by regulating different processes, such as cell-cycle arrest, DNA synthesis and repair, programmed cell death, and energy metabolism. Post translational modifications not only control the activation of p53 protein, but more importantly, its subcellular localization, degradation and protein partners. Stabilization of p53 is a critical step to guarantee stress response and thus limit cancer development. The ubiquitination process is a critical regulatory system in the p53 pathway. Although frequently inactivated in cancer, wild-type p53 is found in half of all human tumours. Indeed, many p53-specific E3 ligases are amplified in human cancers, thus allowing cells to escape p53 response. Cancer cells thus continue to proliferate and survive despite their exposure to various forms of oncogenic stress, including oncogene activation, hypoxia and DNA damage. Increasing p53 levels can lead to tumour regression (Ventura et al., 2007). The stabilization of p53 in cancers that retain wild type p53 is therefore an attractive strategy for therapy. This observation led to an in-depth study on the entangled network of E3 ligases mediating not only p53 ubiquitination and sumoylation but also other modifications. MDM2 is a relevant target for cancer therapy as described (Vassilev, 2007). Similarly, other molecular players involved in the p53

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stability regulation pathway might be employed for treatment. Clinical strategies aiming to overcome chemoresistance by preventing p53 degradation are being explored, together with the developing or isolation of chemical or natural compounds that inhibits p53 proteasomal degradation by targeting the ubiquitination pathway. Also, the observation that p53 ubiquitination can be reversed by deubiquitinating enzymes offers new insights in this field and support the strategy of cancer treatment by pharmacological reactivation of p53. As described, in contrast with the TA isoforms, the ΔN of both p63 and p73 are anti-apoptotic and pro-proliferative. The balanced expression of the different p63 and p73 isoforms can thus be a barrier to tumorigenesis. Similarly to p53, many chemotherapeutic compounds induce both p63 and p73 ubiquitination and subsequent degradation. This led to a great interest in elucidating the molecular pathways that differentially regulate the activity and stability of TA versusΔN isoforms. As for p53, regulation of either TA or ΔN isoforms stability through specific E3 ligases may have important therapeutic implications. An intriguing question is how the E3 and E4 enzymes are coordinated in response to both extracellular and intracellular signals to accomplish the task of p53 homeostasis. This is explicated by the existence of the Cullin–RING complex. A similar consideration can be done for the sub-nuclear structures such as the PML bodies in which enzymes of the sumo cascades seem to be enriched, also in response to stress signals. Even more compelling and intricate appears the regulation of the p53 family members during embryogenesis, and how do they respond to developmental cues. Mutations in p63 has been isolated and characterized in a number of human syndromes characterized by developmental abnormalities. Some of these mutations appears to affect amino acid composition of SUMO consensus sequences, thus confirming that p63 ubiquitination or sumoylation are critical tools that regulate level of p63 during development and in adulthood. Interestingly, similarly to p53, both the spatial and temporal location of ubiquitination and sumoylation can have a profound impact on p53 family regulation. It is to underline that p63 itself has been found in PML nuclear-bodies in vivo where it

interacts with PML. It appears that this interaction activates p63 transcriptional ability to transactivate the p53-responsive elements of the GADD45, p21 and Bax promoters (Bernassola et al., 2005). While the role of ubiquitination and sumoylation serve different purposes in p53 regulation, for p63 these modifications play an apparent redundant role as both of them induce p63 proteasome dependent degradation. Interestingly, analysis of the stability and of the sumoylation/ ubiquitination potential of several natural mutants of DNp63, showed that each modification per se is not sufficient to induce protein degradation. In line with this, p73 protein is not degraded following ubiquitination. This suggests that other molecular players are required to dictate the protein fate. While for p53 sumoylation has a role in dictating the transactivation ability of the protein, as regard p73 sumoylation appears to determine its relocalization in detergent-insoluble nuclear fraction, namely in the nuclear matrix. Sumoylation contributes to protein homeostasis through its ability to cooperate with, complement, and balance the ubiquitin system. The phenomenon of competition of post-translational modifications for the same lysine residues, in which sumoylation can compete with ubiquitination or acetylation, is extensively reported. Mass spectrometry analysis showed that almost a quarter of SUMO-acceptor lysines are also used for ubiquitin conjugation (Hendriks et al., 2014; Tammsalu et al., 2014). The observation that many ubiquitin ligase or de-ubiquitinase are subjected to sumoylation further underlines the functional interaction between the two modification systems. Similarly, also ubiquitin ligase can contact their target once these have been sumoylated, thanks to the action of STUbL proteins. More that the single type of modification, the study of p53 and p53 family members activity and functions demonstrates that a code of post-translational modifications dictates the fate of a protein, the choice and the effect of any PTM depending on both the subcellular localization and the molecular environment of the target protein. A detailed understanding of SUMO and ubiquitin dependent mechanism of p53 family members could lead to novel treatment options for both cancer and human syndromes in which these proteins are involved.

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Interplay between the Ubiquitin Proteasome System and Mitochondria for Protein Homeostasis

12

Mafalda Escobar-Henriques1*, Selver Altin1 and Fabian den Brave2

1Institute for Genetics, Cologne Excellence Cluster on Cellular Stress Responses in Aging-

Associated Diseases (CECAD), Center for Molecular Medicine Cologne (CMMC), University of Cologne, Cologne, Germany. 2Department of Molecular Cell Biology, Max Planck Institute of Biochemistry, Martinsried, Germany. *Correspondence: [email protected] https://doi.org/10.21775/9781912530120.12

Abstract Eukaryotic cells are subdivided into membranebound compartments specialized in different cellular functions and requiring dedicated sets of proteins. Although cells developed compartmentspecific mechanisms for protein quality control, chaperones and ubiquitin are generally required for maintaining cellular proteostasis. Proteotoxic stress is signalled from one compartment into another to adjust the cellular stress response. Moreover, transport of misfolded proteins between different compartments can buffer local defects in protein quality control. Mitochondria are special organelles in that they possess an own expression, folding and proteolytic machinery, of bacterial origin, which do not have ubiquitin. Nevertheless, the importance of extensive crosstalk between mitochondria and other subcellular compartments is increasingly clear. Here, we will present local quality control mechanisms and discuss how cellular proteostasis is affected by the interplay between mitochondria and the ubiquitin proteasome system.

Introduction In order to fulfil their biological function, proteins must fold into their native three-dimensional structures and organelles need to function properly. The factors controlling protein homeostasis processes are collectively termed the proteostasis network (Klaips et al., 2018). In sum, cells must ensure either proper protein folding or -if this failsundertake efficient elimination of malfunctioning proteins or damaged organelles. A prominent role in proteostasis is ensured by ATP-dependent cellular machineries dedicated to proper protein folding, called chaperones (Hartl et al., 2011). In turn, the central proteolytic components of this network are the ubiquitin proteasome system (UPS), a soluble machinery, and the lysosomes, organelles that enclose peptidases with a powerful and non-specific lytic capacity (Bard et al., 2018). Consequently, the UPS is the main proteolytic pathway of the cell for cytosolic substrates, being the lysosomes generally responsible for the clearance of membrane proteins, entire organelles and large protein aggregates (Kerscher et al., 2006; Amm et al., 2014). Both processes

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rely on the ear-marking of the desired protein with ubiquitin. In addition, autophagy requires the ubiquitin-like proteins or Atg8(yeast)/LC3(mammals). Organelles, despite being functional units with dedicated roles, must respond to their cellular environment. Mitochondria – the cellular energy powerhouse – are special semi-autonomous organelles, which evolved through a symbiotic event of alpha-proteobacterium at the origin of a eukaryotic cell (Zimorski et al., 2014). Mitochondria possess an own DNA and the machineries allowing its replication, transcription and translation, originating from their bacterial ancestors, thus resembling the ones from free living prokaryotes (Falkenberg et al., 2007; D’Souza and Minczuk, 2018). During evolution of this endosymbiotic process, most of the mitochondrial genetic information was transferred to the nucleus. This means that most proteins located at mitochondria need to be imported from the cytoplasm, rendering mitochondrial quality control processes fundamentally important for the biogenesis of this essential organelle (Pfanner et al., 2019). Therefore, the presence at mitochondria of specific chaperones and proteases is not surprising. As mentioned, these are closely homologous to their bacterial relatives and make up for local protein quality control (Voos et al., 2016). Perhaps for this reason, it was long assumed that mitochondrial proteostasis was ubiquitin-independent. Instead, it is now clear that mitochondrial stress is also engaging cytosolic and lysosomal proteostasis networks, including ubiquitin and the UPS but also Atg8/LC3 and the lysosomes (Germain, 2008; Escobar-Henriques and Langer, 2014; Topf et al., 2016; Braun and Westermann, 2017; D’Amico et al., 2017). The most prominent example is certainly the mitophagy process, where damaged mitochondria are selectively degraded (McWilliams and Muqit, 2017). Reciprocally, mitochondria also sense and regulate external stress, clearly impacting on cellular homeostasis and longevity and certainly relevant for neurodegeneration (Chung et al., 2018; Guaragnella et al., 2018; Ruan et al., 2018). In this chapter, we describe novel insights on how mitochondria crosstalk and bi‑directionally cooperate with their cellular environment to deal with proteotoxic stress (see Fig. 12.1). After a general overview on quality control – cytosolic and mitochondrial – we then describe emerging

Figure 12.1  Crosstalk of cytosolic and mitochondrial proteostasis. Protein aggregates or malfunctional proteins activate different proteolytic pathways for their clearance. Mitochondria as well as the cytosol harbour their own proteolytic machineries providing clearance of damaged proteins. Nevertheless, an extensive interplay between mitochondria and their environment is key to maintain cellular homeostasis.

cross-functional concepts. First, we focus on the role of mitochondria in coping with excessive cytosolic proteostasis. These findings illustrate the proteolytic power of mitochondria, which does not depend on ubiquitin. Second, we describe several cytosolic and ubiquitin-dependent pathways engaging on mitochondria. We present integrated cellular responses, requiring ubiquitin and the proteasome or the autophagic marker LC3 and lysosomes, which contribute to alleviate mitochondrial stress. Finally, we present the dual role of the peptidyl-tRNA hydrolase Vms1 (yeast)/ANKZF1 (mammals) in ribosomal quality control and mitochondrial proteostasis, being both processes regulated by ubiquitin. Principles of protein quality control – cytosolic and mitochondrial Aberrant folding or unfolding does not only compromise the affected protein but is also accompanied with a great risk of disrupting the functionality of other proteins, by undergoing nonspecific protein–protein interactions. Especially metastable proteins with disordered regions (up to 30% of the mammalian proteome) are prone to undergo unwanted interactions and form toxic protein aggregates, which are associated with

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neurodegenerative diseases (Dunker et al., 2008). This underlines the broad importance of proteome surveillance. In addition, the vast majority of proteins are synthesized by cytosolic ribosomes, followed by post- or co-translational transport of proteins to their final destination (Dudek et al., 2013). Thus, the cytosolic quality control machineries are essential for the integrity of the entire cellular proteome. Quality control components in the cytoplasm Protein folding Chaperones are central players in protein quality control, which support other proteins in acquiring their functional conformation, without usually being present in the final structure (Hartl, 1996). Unfolded proteins expose hydrophobic residues, normally buried inside their three-dimensional structure, being such non-native regions recognized by chaperones. Chaperones promote folding by ATP dependent cycles of binding and release of their substrate proteins, till they reach their native state. Thus, by assisting in protein folding, chaperones prevent unspecific interactions and protein aggregation and refold stress-denatured proteins. However, if encountering terminally misfolded proteins, chaperones also cooperate with proteolytic machineries in their degradation (Tyedmers et al., 2010; Balchin et al., 2016). Hsp70 chaperones Hsp70 chaperones and their co-factors constitute major components for protein quality control (Kityk et al., 2015). Hsp70 binds to substrates in an open, ATP bound conformation. The substratebinding pocket is closed on ATP hydrolysis and release of ADP results in substrate release. In addition to refolding soluble proteins, Hsp70 supports protein import into cellular compartments such as the endoplasmic reticulum and mitochondria, where the proteins have to pass membranes in an unfolded state, through an import channel (Craig, 2018). Moreover, Hsp70 support protein degradation machineries requiring soluble and at least partially unfolded proteins, like the 26S proteasome (Fernández-Fernández et al., 2017). The binding of Hsp70 shields hydrophobic regions in non-native

proteins, thereby preventing non-specific interactions, until proteins reach their native state and/or final destination. The intrinsic activity of Hsp70 alone is low and therefore folding requires the help of additional factors. On the one hand, efficient Hsp70 function requires one of several structurally unrelated nucleotide exchange factors (NEFs), which promote the exchange between ADP and ATP. On the other hand, ATPase activity is stimulated by Hsp40 chaperones, also called J-proteins, which bind the substrates and deliver them to Hsp70, thus avoiding their aggregation. Therefore, substrate specificity of Hsp70 is mainly determined by Hsp40 chaperones (Kampinga and Craig, 2010). These often contain substrate-binding domains themselves and mediate the transfer of substrates to Hsp70, depending on their J-domain. In addition, several specialized Hsp40s lack a substrate-binding domain but localize Hsp70 within the cell to the vicinity of certain substrates. For instance, the Hsp40 Zuo1 targets cytosolic Hsp70 to the ribosomal exit tunnel to aid in folding of nascent proteins (Yan et al., 1998; Gautschi et al., 2001). In sum, Hsp40 chaperones, in conjunction with NEFs, are responsible for the versatile functions exerted by the Hsp70 system (Fig. 12.2). Protein ubiquitination and turnover by the UPS When proteins cannot reach their native conformation, due to mutations or exogenous stresses, they might interfere with the function or folding of other proteins, and thus have to be separated from the rest of the proteome. This can be achieved by sequestration into inclusions or through proteolytic breakdown (Fig. 12.2). The main machinery degrading soluble proteins, in the cytosol and in the nucleus, is the ubiquitin proteasome system (UPS) (Kerscher et al., 2006; Amm et al., 2014). Turnover of proteins generally requires them being tagged with ubiquitin, a highly conserved small protein of 76 aa. It occurs by the covalent attachment of ubiquitin to lysine residues in target proteins (termed ubiquitination). Substrate ubiquitination is mediated by an enzymatic cascade, involving an ubiquitin-activating enzyme (E1), ubiquitin-conjugating enzymes (E2), and ubiquitin protein ligases (E3). Specificity towards individual

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Figure 12.2 Chaperones, ubiquitin–proteasome system and autophagy. Different layers of quality control machineries maintain cellular protein homeostasis. As a first layer, the chaperone Hsp70 as well as its cofactors (Hsp40) and nucleotide exchange factors (NEF) mediate protein folding, escort target proteins to their destination and support import of nascent polypeptide chains into organelles. The ubiquitin proteasome system (UPS, 26S Proteasome, consisting of its 19S regulatory particle and the 20S proteolytic core) represents the second layer of protein quality control. Substrate proteins targeted with the small modifier ubiquitin (E1, E2 and E3 enzymes mediate covalent attachment of ubiquitin to the substrate) are degraded by the 26S proteasome, whereby ubiquitin itself is recycled. The accessibility of defective proteins to the 26S proteasome is supported by the AAA-ATPase Cdc48, which extracts ubiquitinated membrane proteins. As a third layer, protein aggregates/inclusion, organelles or pathogens are targeted to autophagy. This requires the engulfment by an autophagosome, which expands around the substrate by the lipidation of the ubiquitin-like modifier Atg8 with phosphatidylethanolamine (PE). Subsequently, the autophagosome fuses with a cellular lysosome, where final degradation occurs. (NEF, nucleotide exchange factor; ATP, adenosine triphosphate; ADP, adenosine diphosphate; P, phosphate).

substrates, or target recognition, is mostly mediated by the E3 ligase enzymes. Ubiquitination can either result in the attachment of single ubiquitin moieties (mono-ubiquitination), or chains build on lysine residues within ubiquitin itself (poly-ubiquitination). The seven lysine residues of ubiquitin allow the formation of different types of ubiquitin-chains. Often, these serve as targets for different ubiquitinlinkage specific binding proteins, thereby dictating the downstream consequences of ubiquitination. Finally, ubiquitin is a reversible process, being cleaved by ubiquitin-specific peptidases called deubiquitinases (Komander and Rape, 2012). Ubiquitin chains formed via lysine 48 of ubiquitin

are the canonical signal targeting proteins for degradation by the 26S proteasome. This protease complex degrades proteins into short peptides by multiple proteolytic activities within its barrel shaped 20S core particle, being its 19S particle responsible for regulatory functions (Fig. 12.2). Role of Cdc48/p97/VCP The 26S proteasome is only able to degrade soluble proteins. Thus, substrates bound to larger structures, such as protein complexes, or embedded into membranes, need to be extracted before degradation, with the help of accessory factors (Fig. 12.2). The main component of the UPS exerting this function

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is the ATPase and ubiquitin-dedicated chaperone Cdc48 (p97/VCP in mammals). Cdc48 segregates proteins by an ATPase driven mechanism, thereby allowing proteolysis by the 26S proteasome. Cdc48 assembles with several ubiquitin-binding co-factors, which assist it in substrate recognition. Moreover, Cdc48 also directly binds ubiquitinligases and deubiquitinating proteins, thus acting as a general hub in ubiquitin related processes (van den Boom and Meyer, 2018). Autophagy In contrast to soluble proteins, larger structures are refractory to proteasomal turnover. Interestingly, proteasomes can be much smaller in size than protein aggregates. In fact, cryo-EM structures identified entire proteasomes incorporated into the structure of a sub-set of large protein aggregates (Guo et al., 2018). While being consistent with biomedical data suggesting up to 50% proteasome entanglement in neurons, these structures also possibly explain reduced proteasomal activity in neurodegeneration (Pontano Vaites and Harper, 2018). Protein aggregates can be targeted for degradation inside the lysosomes, organelles that are called vacuoles in yeast and plants (Khaminets et al., 2016). In this process, termed macroautophagy (hereafter autophagy), substrates are engulfed by double-membrane bound autophagosomes, which subsequently fuse with the vacuole/lysosome (Fig. 12.2). Vacuoles are acidic compartments containing promiscuous proteolytic enzymes that then deconstruct the engulfed substrates. Similarly to the UPS substrates, which are ear-marked by ubiquitin, the autophagosome membrane is ear-marked by the small ubiquitin-like modifier Atg8 (LC3 in mammals). Atg8 is a cytosolic protein that gets covalently conjugated to the lipid phosphatidylethanolamine at the autophagosomal membrane, on induction of autophagy. In addition to aggregated proteins, a broad range of substrates can be targeted for turnover by autophagy, including entire organelles or pathogens. These pathways of selective autophagy utilize specific autophagy receptors, which characteristically have a dual organization, consisting of a substrate recognition domain and an Atg8 interacting motif. Therefore, autophagy receptors promote the engulfment of their substrates by bridging the autophagosomal membrane

to the target substrates. For example, the selective turnover of mitochondria, or mitophagy, depends on the ubiquitination of a myriad of substrates at the mitochondrial outer membrane, which engage several autophagy receptors like Optineurin, NDP52 and p62 (Geisler et al., 2010; Narendra et al., 2010; Lazarou et al., 2015; Khaminets et al., 2016; McWilliams and Muqit, 2017). Moreover, similar to misfolded soluble substrates, modification by ubiquitin of protein aggregates targets them for degradation by selective autophagy, utilizing specific receptors containing ubiquitin-binding domains. For example, Hsp42 dependent aggregate formation has been shown to be required for the turnover of defective proteasome subunits by autophagy (Marshall et al., 2016). This common feature of the UPS and selective autophagy ensures efficient degradation of aberrant proteins once they have been tagged for degradation (Lu et al., 2017). Protein inclusions When efficient degradation fails, especially during acute stress or when the proteostasis network is perturbed, proteins are sequestered into inclusions, thereby minimizing their reactive surface compared to soluble proteins (Miller et al., 2015; Sontag et al., 2017). Such inclusions are often transient structures, which can either be resolved by disaggregating chaperones, or instead be degraded by selective autophagy, in case they persist in the cytoplasm. General features of mitochondria Mitochondrial functions Mitochondria are central organelles of all eukaryotic cells, functioning as energy-converting powerhouses, metabolic factories and signalling centres (McBride et al., 2006; Nunnari and Suomalainen, 2012). They are required for oxidative phosphorylation (OXPHOS), thus being known as the ATP powerhouse. In addition, mitochondria are key for many metabolic processes, like the synthesis of phospholipids (Silva Ramos et al., 2016; Tatsuta and Langer, 2017). Moreover, the assembly of iron– sulfur-clusters (essential enzymatic cofactors) starts within mitochondria, reason why these organelles are essential for cellular viability (Braymer and Lill, 2017; Cardenas-Rodriguez et al., 2018). Finally, mitochondria are active components of many

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signalling pathways, such as programmed cell death, ageing, cellular differentiation and organism development (Green et al., 2014; Kauppila et al., 2017; Noguchi and Kasahara, 2018; Pallafacchina et al., 2018; Paupe and Prudent, 2018; Zhang et al., 2018). Sub-compartmentalization of mitochondria Mitochondria are bound by two separate membranes, the outer mitochondrial membrane and the inner mitochondrial membrane ( Jakobs and Wurm, 2014; Schorr and van der Laan, 2018). The two compartments bound by these membranes are called intermembrane space and matrix. The inner membrane forms large invaginations, called cristae, harbouring the respiratory OXPHOS chain complexes. Moreover, the part of the inner membrane that lines parallel to the outer membrane is called inner boundary membrane. Finally, cristae and inner boundary membrane are connected at cristae junctions, and the outer membrane and the inner boundary membrane make close contacts, termed contact sites. Due to their high biosynthetic demands, mitochondria are extremely rich in proteins, many of which are assembled into large complexes, often embedded into the mitochondrial inner and outer membrane, respectively. Given that 99% of mitochondrial proteins are nuclearencoded, most of the organellar proteome needs to be post-translationally imported into the respective sub-compartment within mitochondria (Harbauer et al., 2014). Import of mitochondrial proteins Proteins targeted to mitochondria are mainly imported via two channels spanning both mitochondrial membranes, called TOM (translocase of the outer membrane) and TIMs (translocases of the inner membrane) (Wasilewski et al., 2017; Wiedemann and Pfanner, 2017; Pfanner et al., 2019). Together with their interaction partners, these two channels allow directing each protein to their final subcellular destination. Once inside mitochondria, imported proteins must assemble with those encoded by the mitochondrial DNA, which in humans are 13. Interestingly, translation of nuclear-encoded and mitochondrial-encoded OXPHOS components coordinately adapt to metabolic conditions stimulating respiratory growth

(Couvillion et al., 2016). Moreover, this response was shown to be unidirectionally controlled by cytosolic translation components. Mitochondrial protein quality control Owing to the dimensions of the protein transport pores, proteins cross the membranes in an unfolded state. In addition, the unfolded import-competent state must be protected from non-native interactions. Therefore, chaperones – ensuring proper folding and assembly into active proteins – but also proteases -allowing to eliminate faulty polypeptides- are of extreme importance for mitochondrial protein quality control (Rugarli and Langer, 2012; Voos, 2013; Voos et al., 2016) (Fig. 12.3). In addition, once proteins have reached their destination, compartment-specific mechanisms of protein quality control locally protect the proteome. Upon mitochondrial stress, several proteostasis networks have been nicely shown to operate at damaged mitochondria, for example to protect cells from death signals (D’Amico et al., 2017; Priesnitz and Becker, 2018). Chaperones guiding mitochondrial proteins To support efficient import of proteins targeted to mitochondria, these are kept in the cytoplasm in an unfolded state by the cytosolic Hsp70 and Hsp90 machineries (Deshaies et al., 1988; Young et al., 2003; Craig, 2018). In turn, mitochondrial Hsp70 (mtHsp70) – residing at the matrix site of the TIM complex – is crucial for the import and subsequent folding of proteins into mitochondria (Kang et al., 1990; Liu et al., 2001; Schulz et al., 2015). Protein folding inside the mitochondrial matrix largely also depends on the Hsp60-Hsp10 chaperonin, a member of the GroEL family of bacterial chaperones (Cheng et al., 1989; Reading et al., 1989). Importantly, the Hsp70 and Hsp60 systems are not only required for the folding of newly imported proteins, but also support refolding and prevent aggregation of unfolded proteins, which might occur upon proteotoxic stress (Kubo et al., 1999; Bender et al., 2011). Under conditions of severe protein folding stress, as for example acute heat stress, the abundance of unfolded proteins exceeds the capacity of mitochondrial chaperones to maintain these in a soluble state, resulting in protein aggregation. Such aggregates can be resolved

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Figure 12.3  Mitochondrial quality control systems. For each individual compartment, mitochondria possess their own quality control proteins required for maintenance of the mitochondrial proteome. Within the matrix, mitochondrial Hsp70 and its cofactors (Hsp10 Hsp60, Hsp100) maintain proper protein folding after import. Turnover of misfolded proteins is regulated via the proteases LON and ClpXP. Proteolytic turnover of inner membrane proteins is mediated by m-AAA at the matrix side and i-AAA on the intermembrane side. The i-AAA protease maintains also proteolytic control of outer membrane proteins like Om45 and Tom20. The protease Htra2 maintains proteolysis of misfolded and damaged proteins in the intermembrane space. At the outer membrane, the AAA-ATPase Msp1 mediates segregation of outer membrane proteins to the cytosol, which results in subsequent proteasomal turnover. Polypeptides are imported via the TOM (translocase of the outer membrane) and TIM (translocase of the inner membrane) channels. On import stress, polypeptides reaching the matrix are degraded by the Lon protease.

by mitochondrial Hsp100 chaperones, which are required for efficient recovery from acute heat stress (Schmitt et al., 1996) (Fig. 12.3). Mitochondrial proteolytic machinery Incorrect complex assembly or mis-targeting of membrane spanning proteins is especially prone to result in the accumulation of non-native proteins, which can undergo non-specific interactions, potentially detrimental for the cell. To remove misfolded proteins, mitochondria are equipped with a diversified set of proteases (Hamon et al., 2015; Quirós et al., 2015) (Fig. 12.3). The substrates targeted by mitochondrial proteases mainly depend on their sub-mitochondrial localization and structural properties. In general, mitochondria have ATP-dependent and independent proteases

in all its sub-compartments. The first belong to the AAA+ superfamily, characterized by their oligomerization into a beta-barrel structure, enclosing a chamber with ATP-dependent pulling activity. In addition to the complete turnover of their substrates, mitochondrial proteases are also required to process and thus mature pre-proteins, for example by removing the mitochondrial targeting sequence. The main protease degrading misfolded proteins in the mitochondrial matrix is Pim1/LON (Wagner et al., 1994). In addition, the matrix protease ClpXP has been implicated in degradation of misfolded proteins, but its function is not yet well understood (Haynes et al., 2007). Interestingly, a role of LonP1 and ClpP in regulating heteroplasmy was suggested (Latorre-Pellicer et

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al., 2016). Indeed, human mitochondrial DNA shows extensive sequence variability, suggested to impact on mitochondrial proteostasis, dependent on LonP1 and ClpP. These findings have clinical implications, in what regards mitochondrial replacement therapies, which should be considered when choosing the mitochondrial DNA donor (Latorre-Pellicer et al., 2016). The mitochondrial inner membrane harbours two AAA proteases, which are anchored to the inner membrane by transmembrane domains (Gerdes et al., 2012; Rugarli and Langer, 2012; Levytskyy et al., 2017; Patron et al., 2018). The m-AAA protease exposes its catalytic domain to the matrix, while the catalytic domain of the i-AAA protease faces the intermembrane space. These proteases degrade misfolded proteins of the inner membrane, being their substrate specificity mainly depending on the topology of the respective substrates (Leonhard et al., 2000; Almajan et al., 2012; Stiburek et al., 2012; Anand et al., 2014; Kondadi et al., 2014; König et al., 2016; Wai et al., 2016; Wang et al., 2016; Pareek et al., 2018; Sprenger et al., 2019). In the inner mitochondrial membrane space, misfolded and damaged proteins are degraded by the proteases Omi/HtrA2 and Atp23 (Osman et al., 2007; Clausen et al., 2011). At the outer membrane, proteins are surveilled by mitochondrial and cytoplasmic quality control machineries in parallel, as discussed later. The unfolded protein response – UPR Mitochondrial stress inhibits mitochondrial translation, but also impacts on nuclear expression. This was termed unfolded protein response (UPR) and depends on the transcription factor ATFS-1 (Nargund et al., 2012; Jovaisaite and Auwerx, 2015; Münch and Harper, 2016; Higuchi-Sanabria et al., 2018; Shpilka and Haynes, 2018). In intact mitochondria, ATFS-1 is imported into the matrix and degraded by the protease LON. However, upon import inhibition, ATFS-1 is diverted from the mitochondria to the nucleus. There, it up-regulates critical detoxifying genes, encoding proteins ensuring proper translation, folding and turnover at mitochondria, thus restoring mitochondrial homeostasis.

Mitochondrial roles in quality control of cytosolic components Non-native proteins are a general threat for the cellular proteome and their spatial sequestration – into aggregates, inclusions or organelles – is a common strategy to limit such effects (Sontag et al., 2017). Moreover, misfolded proteins are transported between organelles, as it has been shown for terminally misfolded cytosolic proteins, which are transported into the nucleus for degradation (Park et al., 2013). Interestingly, novel roles of mitochondria in coping with cytosolic or cytosolic-exposed proteins have recently emerged. Mitochondrial contributions to mitigate aggregation toxicity Asymmetric inheritance of protein aggregates It is known that protein aggregates that cannot be efficiently cleared by proteolytic systems are asymmetrically inherited during cell divisions, thereby ensuring that one cell deriving from such a division is free of damaged aggregated proteins (Aguilaniu et al., 2003; Shcheprova et al., 2008; Clay et al., 2014; Coelho et al., 2014; Hill et al., 2017; Saarikangas et al., 2017). Interestingly, an active role of mitochondria in restricting the mobility and thus inheritance of protein aggregates residing in the cytosol has been identified (Zhou et al., 2014). In addition, proteins aggregated inside the matrix were sequestered into specific deposits that were also retained in the mother cell (Bruderek et al., 2018). Finally, asymmetric inheritance depended on mitochondrial size and actively engaged the motor components involved in mitochondrial transport (Böckler et al., 2017). Consistent with a cellular mechanism providing for rejuvenated daughter cells, a filtering process that prevented feeble mitochondria from being inherited had equally been shown (Higuchi et al., 2013; Nyström, 2013). In contrast, however, under conditions of mild heat stress these damage-retention quality control mechanisms were inhibited. Instead, inheritance of toxic components to the daughter cell was promoted, which consequently increased longevity of the mother cell (Baldi et al., 2017).

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Import and turnover of cytosolic proteins in mitochondria Recently, it has been observed that mitochondria can also function as a place to dispose misfolded cytosolic proteins under stress conditions. As previously mentioned, initially it had been observed that cytosolic protein aggregates are tethered to mitochondria, which facilitates asymmetric inheritance, keeping daughter cells free of the damaged proteins (Zhou et al., 2014). Consistently, purification of such aggregates revealed a physical interaction with the mitochondrial import pore (Ruan et al., 2017). Moreover, it was observed that the clearance of cytosolic aggregates was supported by import of cytosolic proteins into the mitochondrial matrix, where these proteins were handled by the Lon mitochondrial protease Pim1 (Ruan et al., 2017). The mitochondrial import and clearance of cytosolic proteins was in particular observed on acute heat shock and inhibition of cytosolic Hsp70, suggesting that this pathway functions to buffer extensive cytosolic proteotoxic stress (Ruan et al., 2017). The presence of ubiquitinated proteins inside mitochondria was suggested, which could perhaps result from similar surveillances principles (Lehmann et al., 2016). However, this does not necessarily imply a functional role of ubiquitin inside mitochondria. Import of cytosolic aggregated proteins required the disaggregase Hsp104, probably through generating soluble proteins for mitochondrial import. Surprisingly, this process was independent of cytosolic Hsp70, which usually cooperates with Hsp104 (Ruan et al., 2017). In conclusion, borrowing mitochondrial proteolytic capacity seems to have beneficial effects for stress-release of cytosolic protein load. Nevertheless, it remains an open question to which quantitative extend mitochondrial import of cytosolic proteins contributes to cytosolic proteostasis. In addition, it is still unclear if and how the import of aberrant proteins affects mitochondrial proteostasis and which mechanisms might protect mitochondria. For example, acute heat stress will also affect mitochondrial proteins and the additional uptake of non-native cytosolic proteins can be expected to pose a major challenge for the mitochondrial proteostasis network.

Turnover of cytosolic-exposed proteins by mitochondrial proteases Transmembrane proteins residing at the outer membrane of mitochondria can, in principle, be degraded by outer membrane-embedded proteases, or be subject to membrane extraction to the cytoplasm or to mitochondria, for turnover. Interestingly, recent studies revealed a role of the i-AAA protease, or Yme1, for turnover of two outer membrane anchored proteins. Indeed, proteolysis of Tom22 and Om45 was independent of the proteasome pathway but instead depended on Yme1 (Wu et al., 2018). In addition, proteolysis required substrate dislocation by Yme1, after recognition of their inner-membrane-space domains by the Mgr1/Mgr3 complex. Mgr1/Mgr3 interact with Yme1, thus enhancing its catalytic activity (Dunn et al., 2008). These findings show a cross-membrane mechanism for proteolytic control at mitochondria. Quality control of mitochondrial proteins in the cytoplasm Defects in mitochondrial targeting and import of proteins result in mis-localization of mitochondrial precursor proteins to the cytosol. Multiple concerted responses operating at the mitochondrial surface are now shown, allowing these proteins to be degraded in the cytosol by the UPS. Rescue of mitochondrial import overload by cytosolic machineries The import machinery at the outer membrane has an upfront role in determining mitochondrial biogenesis. Indeed, it is now clear that the import process is highly regulated, both under physiological and pathophysiological conditions (Harbauer et al., 2014). Moreover, it plays critical roles in surveilling translocation quality and in signalling import stress (Wasilewski et al., 2017). Pre-import chaperones The classical cytosolic Hsp70 and Hsp90 chaperones, their co-factors Sti1 and Ydj1, and ubiquilins (chaperone-like factors), associate with mitochondrial pre-proteins and also physically interact with the outer membrane components of the mitochondrial import channel (Deshaies et al., 1988; Young

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et al., 2003; Hoseini et al., 2016; Zanphorlin et al., 2016; Jores et al., 2018; Opaliñski et al., 2018). Supporting these physical interaction evidences, a genetic synthetic growth defect was observed between TOM and STI1, which encode protein forming an important scaffold, by simultaneously binding to Hsp70 and Hsp90 (Hoseini et al., 2016). Moreover, among the TOM components, a prominent role of Tom70 in import control has been suggested (Backes et al., 2018; Hansen et al., 2018; Opaliñski et al., 2018). In fact, Tom70 has a tetratricopeptide repeat, a domain known to bind to Hsp90 (Zanphorlin et al., 2016). Consistently, chemical inhibition of the Hsp70/90 interaction with Tom70 reduced the mitochondrial association of protein aggregates (Pavlov et al., 2018).

function, they have been implicated in targeting mislocalized mitochondrial precursor proteins to the 26S proteasome for degradation (Itakura et al., 2016; Whiteley et al., 2017). In addition, by binding to mitochondria-targeted membrane proteins, ubiquilins prevent their aggregation, thus exerting a chaperone like function. At this step ubiquilin binding still allows correct targeting of the bound protein. However, prolonged binding will result in ubiquitination of the bound protein by ubiquitin-ligases recruited by the UBA domain, and subsequent targeting for degradation (Itakura et al., 2016). Thus, ubiquilins not only function in targeting already ubiquitinated substrates for turnover but themselves exert an important role in triage of mitochondrial proteins (Fig. 12.4).

Dual role of ubiquilins in control of protein import Ubiquilins were proposed to be involved at the earlier steps of mitochondrial protein biogenesis (Itakura et al., 2016; Whiteley et al., 2017). Ubiquilins are substrate receptors for proteasomal degradation, typically harbouring a ubiquitin-binding (UBA) domain for recognition of ubiquitinated cargo and a ubiquitin like domain, which is required for proteasomal targeting (Buchberger, 2002; Funakoshi et al., 2002). In line with their canonical

Recognition of J-proteins by mitochondrial receptors Recent findings shed additional light on the early steps of mitochondrial protein biogenesis, by identifying how – once translated – mitochondrial proteins are targeted intracellularly to the surface of the organelle (Hansen et al., 2018; Opaliñski et al., 2018). Djp1, a J-protein that localizes to the surface of the endoplasmic reticulum, was found to contribute to mitochondrial protein import, in cooperation with pre-protein receptors. Therefore,

Figure 12.4 Dual role of ubiquilins in control of mitochondrial pre-proteins. The ubiquitine-proteasome receptors Ubiquilin 1 and 2 (UBQLN) guide mitochondrial membrane proteins after translation to the import channel. However, on prolonged binding, UBQLNs recruit E1, E2 and E3 ligase enzymes, required for their ubiquitination, allowing subsequent turnover by the proteasome.

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a pathway termed ER-Surf has been proposed, in which the endoplasmic reticulum provide a surface to capture mitochondrial preproteins (Fig. 12.5). In addition, the recognition by J-proteins of mitochondrial translocase components seems to be broad but specific, because although Djp1

cooperates with Tom70, Tom22 recruits the J-protein Xdj1 (Opaliñski et al., 2018). Consistent with an important role of J-proteins, Ydj1 and Sis1 were found to mediate import of beta-barrel proteins to the mitochondrial outer membrane ( Jores et al., 2018).

Figure 12.5  ER-associated J-Proteins promote mitochondrial import. The J-protein Djp1, which localizes at the ER surface, contributes with the TOM channel at the mitochondrial outer membrane to import mitochondrial pre-proteins.

Figure 12.6  Cellular responses to mitochondrial import stress. Different quality control pathways are described maintaining cellular homeostasis on import overload at the mitochondrial surface. Protein import can be reduced due to intra- or extracellular stresses, causing the cytosolic accumulation of protein aggregates destined for the mitochondria (mPOS, mitochondrial precursor over-accumulation). The accumulation of unimported mitochondrial proteins can activate the UPRam (unfolded protein response activated by mis-targeting of proteins), which mediates the turnover of these proteins via the 26S proteasome. Similarly, import clogging at the TOM channel activates the MitoCPR pathway (mitochondrial compromised protein import response). Thereby, the AAA-ATPase Msp1 interacts with the TOM channel via Cis1 and mediates the extraction of clogging proteins to the cytosol, where they are targeted for proteasomal turnover.

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UPRam – unfolded protein response activated by mistargeting of proteins The global cellular responses caused by accumulation of mitochondrial precursor proteins in the cytosol were recently addressed, by provoking a defective protein import in the intermembrane space (Wrobel et al., 2015). Interestingly, mis-targeted mitochondrial proteins activated a concerted proteostatic response in the cytosol, whereby protein synthesis was inhibited and the proteasome was activated. Importantly, these responses were key in alleviated systemic pathology of the organelle and organismal death. In conclusion, UPRam allows buffering the consequences of physiological slowdown in mitochondrial protein import, thus promoting cellular survival under stress (Fig. 12.6). Consistently, it was previously shown that reduced mitochondrial-membrane potential induced aggregation in the cytosol (Erjavec et al., 2013). A role of faulty protein import and accumulation of unprocessed mitochondrial proteins in the cytosol was equally proposed. Such defects generated by mitochondrial dysfunctions could be compensated for by a boost in cytosolic protein quality control, thus maintaining viability despite chronic failures in mitochondrial function (Erjavec et al., 2013). Another study suggested that defects in protein import lead to increased levels of reactive oxygen species, which in turn affect protein synthesis by modification of cytosolic ribosomes (Livnat-Levanon et al., 2014). Interestingly, reducing cytosolic synthesis of mitochondrial proteins has even been shown to reduce mitochondrial degeneration, emphasizing its impact on mitochondrial integrity (Wang et al., 2008). mPOS – mitochondrial precursor overaccumulation stress Simultaneously to UPRam, a study addressed the global consequences of aberrant accumulation of mitochondrial precursors in the cytosol, triggered either by impairing protein import or by clinically relevant mitochondrial damage (Wang and Chen, 2015). Consistent with UPRam, a cytosolic proteostatic network could be observed (Fig. 12.6). In particular, ribosomal biogenesis was modulated, where cap-dependent and thus major translation was down-regulated, to suppress protein synthesis. Moreover, cap-independent

translation was up-regulated for a particular set of proteins that prevent ribosome assembly, thus reinforcing inhibition of general translation. Finally, this cytosolic network also suppressed cell death, confirming the physiological relevance of mPOS. MitoCPR – mitochondrial compromised protein import response An artificial precursor protein leading to clog of the protein import machineries revealed a role of the dislocase Msp1(yeast)/ATAD1(mammals) (Weidberg and Amon, 2018) (Fig. 12.6). Msp1 is a AAA-ATPase inserted at the outer membrane and facing the cytosol, assembling into a hexameric ring (Wohlever et al., 2017). An analysis of the genes transcriptionally correlated with import clogging allowed the identification of Cis1, which physically interacts with Msp1 but also with the Tom70 component of the outer membrane translocase. Clearance of the precursor proteins, which depended on Cis1, Tom70 and Msp1, also required the proteasome to degrade the non‑imported proteins. IPTP – Interplay between mitochondrial translation and cytosolic responses Apart from the responses just described, primarily induced at the surface of mitochondria, a proteostasis retrograde mechanism initiated in the matrix was also reported. The unfolded protein response (UPR) had already revealed that mitochondrial stress can inhibit translation at the mitochondria. Now, a crosstalk mechanism of mitochondrial translation accuracy impacting on cytoplasmic proteostasis was also proposed (Suhm et al., 2018). Mitochondrial translation is signalled by a novel interorganellar proteostasis transcription program (IPTP), impacting chronological lifespan. Hyperaccurate mitochondrial translation stimulated Hsp104-mediated refolding and proteolytic capacity of a proteasomal model substrate. This infers that decreased accuracy of mitochondrial translation impaired management of cytosolic protein aggregates, eliciting a general transcription stress response. It also shows that cytosolic proteostasis, nuclear stress signalling and mitochondrial translation are closely coordinated in determining cellular homeostasis and lifespan (Suhm et al., 2018).

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Turnover of mitochondrial proteins by the UPS In contrast to the nucleus, proteasomes are not present in mitochondria. Nevertheless, the UPS was shown to degrade some mitochondrial proteins after their insertion in the outer membrane, independently of the import surveillance mechanism described below. In the endoplasmic reticulum, ubiquitinated proteins are extracted to the cytoplasm by Cdc48/p97 to be degraded by the proteasome (Rape et al., 2001; Franz et al., 2014). This process is called ERAD (endoplasmic reticulum associated degradation). A similar mechanism was identified in mitochondria and named OMAD, for outer membrane associated degradation, in analogy to ERAD (Neutzner et al., 2007; Braun and Westermann, 2017). OMAD – extraction of cytosolicexposed proteins by the cytosolic dislocase Cdc48 As previously mentioned, Cdc48 assembles with a myriad of partners that assist the AAA protein in extracting proteins from complexes or membranes (Fig. 12.7). Cdc48 was suggested to extract the

yeast mitofusin Fzo1 from mitochondria under oxidative stress conditions (Heo et al., 2010; Esaki and Ogura, 2012). In absence of external stress, Cdc48 formed a complex with Doa1, Ufd1 and Npl4 to retrogradely translocate ubiquitinated membrane-anchored proteins to the cytoplasm, including tagged versions of Msp1 and Tom70 (Wu et al., 2016). Although tagged Fzo1 was also a MAD substrate (Wu et al., 2016), the endogenous protein is instead stabilized by endogenous Cdc48 (Simões et al., 2018), demonstrating limitations of working with tagged proteins, and thus accessing the real Cdc48 substrates. Nevertheless, the work from Wu et al. (2016) clearly highlights the importance of Cdc48 in quality control mechanisms at the outer membrane. In mammals, p97 was also required for the extraction and proteasomal turnover of outer membrane proteins, under damaging conditions, including mitofusins and Mcl-1 (Neutzner et al., 2007; Tanaka et al., 2010; Xu et al., 2011). Interestingly, Cdc48 performs opposing roles to MAD, by instead increasing the stability of the Fzo1 protein (Fig. 12.7). In fact, mitochondria form a dynamic network that is continuously remodelled by fusion and fission events. Fzo1, present at the

Figure 12.7 Dual roles of Cdc48 at the mitochondrial surface. The ubiquitin-specific AAA-ATPase Cdc48 maintains protein quality control at the mitochondrial outer membrane and regulates mitochondrial dynamics. On the one hand, Cdc48 acts as a cytosolic dislocase, which segregates ubiquitinated outer membrane proteins, allowing their degradation by the 26S proteasome. On the other hand, Cdc48 protects ubiquitination on Fzo1, and controls a cascade of deubiquitinases, like Ubp12, thus promoting the mitochondrial outer membrane fusion process.

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mitochondrial outer membrane, is required for mitochondrial fusion. In contrast to MAD, Fzo1 was shown to be protected from the UPS by Cdc48 (Simões et al., 2018). Instead, Cdc48 was required for the turnover of a deubiquitinating enzyme which inhibits Fzo1. In addition, Cdc48 served as a binding platform, allowing crosstalk regulation between deubiquitinases and thus promoting membrane merging and mitochondrial fusion. Extraction of cytosolic-exposed proteins by the mitochondrial dislocase Msp1 Similar to Cdc48/p97, Msp1/ATAD1 is a AAAATPase at the mitochondrial outer membrane that assembles into an hexameric ring, as previously mentioned (Schnell and Hebert, 2003). Therefore, Msp1/ATAD1 constitutes an alternative machinery to segregate substrates from the mitochondria. Indeed, recent findings showed that Msp1/ATAD1 participates in a local organelle surveillance pathway, to deal with proteins inappropriately inserted into mitochondria (Hegde, 2014; Okreglak and Walter, 2014; Opaliñski et al., 2014). Correct targeting of proteins to their respective organelles is a general challenge, given that the vast majority of organellar proteins are synthesized as precursors on cytosolic ribosomes and have to be transported to their intracellular destinations (Schnell and Hebert, 2003). This challenge is even bigger for tail-anchored proteins, i.e. with a single transmembrane segment at the C-terminus. Msp1 was shown to extract tail-anchored proteins mis-targeted from peroxisomes to mitochondria (Chen et al., 2014; Okreglak and Walter, 2014). Consistently, purified Msp1 drove ATP-dependent extraction of tailanchored proteins from the lipid bilayer (Wohlever et al., 2017). It is highly likely that the proteasome will degrade these mis-localized proteins, once extracted by Msp1, as it is the case in mitoCPR. In conclusion, the Msp1/ATAD1 protease ensures the fidelity of organelle specific-localization of tail anchor proteins. Moreover, as previously described, it also functions in pre-protein clearance during mitochondrial import stress. This highlights critical functions of an outer membrane dislocase in maintaining mitochondrial integrity. IMS proteins The proteasome was also shown to degrade proteins present at the inner-membrane space, after

their retrograde translocation back to the cytosol, mediated by Tom40 (Bragoszewski et al., 2015) This is consistent with a previously observed accumulation of mitochondrial inner membrane proteins, upon chemical inhibition of the proteasomal activity (Radke et al., 2008). Interestingly, under those conditions, the inner membrane space protease Omi/HtrA2 could degrade inner membrane proteins. However, it should be noted that mitochondrial proteases can also be sensitive to proteasomal inhibitors. Collectively, it is possible that faulty folding of inner membrane space proteins during import could lead to ubiquitination of their cytosolic exposed parts, providing access to the UPS. This is consistent with the observation that a fraction of newly synthesized intermembrane space precursors are degraded in the cytosol before reaching this subcompartment, event in the absence of stress-inducing conditions (Bragoszewski et al., 2013; Kowalski et al., 2018). In conclusion, the UPS plays a constitutive role for the biogenesis of intermembrane space proteins. Mitochondrial damage overload In addition to the previously described removal of individual proteins for proteasomal degradation, excessive damage can be repaired by eliminating whole mitochondrial fragments, by mitophagy, thus protecting the healthy mitochondria (Harper et al., 2018; Pickles et al., 2018). In addition, selected mitochondrial components can be eliminated from the mitochondrial network by mitochondria-derived vesicles or mitochondrial derived compartments (Sugiura et al., 2014; Hughes et al., 2016) (Fig. 12.8). Mitophagy The panoply of different mechanisms described, e.g. UPRmt, UPRam, POS, mitoCPR, allow coping with transient stress, still repairable at the level of the mal-functioning proteins. However, prolonged or acute stress conditions can no longer be reversed by these pathways, and mitochondria need to be eliminated by mitophagy, in order to restore cellular homeostasis. A prominent role of mitochondrial fission in allowing the detachment of damaged pieces from the entire network was recently proposed (Burman et al., 2017). One of the early steps and hallmarks in mitophagy is the general ubiquitination of outer

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Figure 12.8 Elimination of damaged mitochondria. Local mitochondrial defects cause the budding of mitochondrial-derived vesicles (MDVs), which can consist only of the outer membrane, or of outer and inner membrane. Such vesicles, forming on oxidative stress, segregate from the mitochondrial network and fuse subsequently with lysosomes, where final degradation takes place. However, acute mitochondrial defects initiate the clearance of mitochondria via mitophagy. Loss of the mitochondrial membrane potential causes the accumulation of the kinase Pink1 at the outer membrane. Pink1 recruits and phosphorylates Parkin, which activates the E3 ligase. In addition, Pink1 phosphorylates ubiquitin, which is required for Parkin activation as well. Activated Parkin ubiquitylates several mitochondrial outer membrane proteins, which serve as a platform for the recruitment of the ubiquitin-like modifier LC3 via different adaptor autophagic receptors. LC3 lipidation with PE (Phosphatidylethanolamine) allows autophagosome expansion, which engulfs the whole damaged mitochondria. Subsequent mitochondrial degradation takes place within the lysosome on fusion of autophagosomes with lysosomes.

membrane proteins. This is believed to recruit several autophagy receptors and therefore mark mitochondria that should be eliminated. The most famous components performing this task are the E3 ligase Parkin and the mitochondrial kinase Pink1 (Pickles et al., 2018). In healthy mitochondria, Pink1 is imported into mitochondria, processed during import and then released to the cytosol and degraded by the N-end rule. In contrast, upon loss of membrane potential, the kinase is arrested due to import failure, thus integrating in the outer membrane, and exposing the catalytic domain to the cytosol (Matsuda et al., 2010; Vives-Bauza et al., 2010). There, it phosphorylates Parkin at its ubiquitin-like domain, which changes its conformation and leads to enzymatic activation. Moreover, Pink1 phosphorylates serine 65 of ubiquitin chains assembled by Parkin at the outer membrane, further increasing Parkin recruitment to the mitochondria and activation, by feedforward loop mechanisms. These chains bind several receptors like Optineurin, NDP52 and p62, which then engage the autophagic machinery and culminates by releasing mitochondria into the lysosome for destruction (McWilliams and Muqit, 2017) (Geisler et al., 2010; Narendra et al., 2010; Lazarou et al., 2015; Khaminets et al., 2016). In addition to Parkin, other E3 ligases

stimulate mitophagy (Covill-Cooke et al., 2018). For example, the outer membrane RING ligase March5 induced mitophagy on hypoxic conditions, together with the autophagic receptor FUNDC1 (Chen et al., 2017). In addition, March5 has many additional physiological functions (Covill-Cooke et al., 2018). Finally, ubiquitin-independent mitophagy pathways have also been described as for example the role of the ATG8 receptor NIX in hypoxia (Khaminets et al., 2016). Mitochondrial-derived vesicles In contrast to mitophagy, where the whole organelle is degraded, mitochondria can also dispose content in the form of vesicles, called MDVs (mitochondria-derived vesicles), which transport proteins and lipids to other cellular organelles (Sugiura et al., 2014). Mitochondria can form different types of vesicles, with different cargoes and also with different cellular destinations, thus facilitating intracellular communication. In yeast, vesicles allowing selective degradation of a protein subset were also found, suggesting that budding of mitochondria could be a conserved mechanism (Hughes et al., 2016). Pink1 and Parkin are also involved in the formation of MDVs containing oxidized cargo and formed after oxidative stress.

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Consistent with their quality control roles, these MDVs were destined to the lysosomes, suggesting a similar role to mitophagy of Pink1 and Parkin, just more confined and not so extreme. Interestingly, however, a role of Pink1 and Parkin in repressing vesicle formation was also recently reported (Matheoud et al., 2016). Indeed, MDVs containing antigens were negatively regulated by Pink1 and Parkin. These MDVs were targeted to the cellular surface, to present the cargo on major histocompatibility (MHC) class I molecules, triggering an immune response. Consistently, Pink1 and Parkin also prevented the activation of an inflammatory response caused by excessive mutations in the mitochondrial DNA (Sliter et al., 2018). Finally, other roles of Parkin have been proposed, as for example in mitochondrial trafficking along neurons (Scarffe et al., 2014). Role of ribosomal quality control (RQC) in mitochondria Fidelity of protein synthesis is essential for mitochondrial function, since the vast majority of mitochondrial proteins are synthesized by cytosolic ribosomes. However, protein synthesis at ribosomes can go wrong, resulting in aberrant translation products, which are subjected to ribosomal quality control (RQC) (Brandman and Hegde, 2016) (Fig. 12.9). Failure in RQC of mitochondrial proteins results in defective proteins, which are still imported into mitochondria and interfere with mitochondrial proteostasis (Izawa et al., 2017). RQC monitors ribosomal activity and is

activated on stalling of translation. Potential causes for stalling are damaged or truncated mRNAs, particular mRNA sequences including poly(A)tails, excessive mRNA secondary structures and insufficient amounts of amino acids or tRNAs (Brandman and Hegde, 2016). As a consequence of stalling, ribosomes are disassembled and the potentially defective mRNA is degraded, as well as the nascent polypeptide chain. RQC The first step of RQC is the splitting of the ribosome, resulting in a 60S ribosomal subunit bound to the nascent polypeptide chain. This complex is then recognized by the RQC complex, which consists of Rqc1, Rqc2, the E3 ubiquitin ligase Ltn1 (Listerin in mammals) and Cdc48 with its co-factors Npl4 and Ufd1 (Brandman et al., 2012). The function of Ltn1 is to ubiquitylate the stalled polypeptide at the ribosomal exit site, resulting in proteasomal degradation (Bengtson and Joazeiro, 2010). This process requires the action of Cdc48 to extract the ubiquitinated nascent chain from the 60S ribosome (Defenouillère et al., 2013). In addition to ubiquitination, the stalled polypeptide can be further modified by addition of multiple alanine and threonine residues, at the C-terminal. The synthesis of this amino acid extension -termed CAT-tail (c-terminal Alanine Threonine tail) occurs independently of mRNA or 40S subunits. Instead, it depends on the recruitment of charged t-RNAs by Rqc2 (Shen et al., 2015). CAT-tails increase the probability of exposing lysines present

Figure 12.9  Ribosomal quality control. Translation of defective proteins requires the ribosomal quality control (RQC) machinery. On stalling of translation, ribosomes disassemble resulting in the 60S subunit bound to the nascent polypeptide chain. This recruits the RQC complex consisting of Rqc1, Rqc2 and Ltn1. Rqc1 recruits Cdc48 for the segregation of the polypeptide chain from the 60S subunit, which is subsequently degraded via the 26S proteasome. Rqc2 recruits charged tRNAs for the assembly of a CAT tail at the C-terminus of the nascent poly-peptide. The CAT tail increases exposure of lysine residues, which are ubiquitinated by the E3 ligase Ltn1, thus addressing the stalled polypeptide for proteasomal degradation. Additionally, the peptidyltRNA hydrolase Vms1 mediates the removal of the bound tRNA from the nascent chain, facilitating release from the ribosome.

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on the stalled chain to Ltn1. In turn, Rqc1 recruits Cdc48 to ribosomal subunits. When ubiquitination and degradation are compromised, the nascent polypeptides are still released from the ribosome and form CAT-tail dependent aggregates in the cytosol (Choe et al., 2016; Defenouillère et al., 2016; Yonashiro et al., 2016). Role of Vms1/ANKZF1 in RQC Another factor recently implicated in RQC is Vms1. Initially, Vms1 had been identified as a cofactor recruiting Cdc48 to mitochondria on stress (Heo et al., 2010). Despite being a cytosolic soluble protein, Vms1 translocates into mitochondria on oxidative stress, dependent on ergosterol peroxide, suggesting the presence of an oxidized sterol receptor at the outer membrane (Nielson et al., 2017). Further studies revealed that Vms1 binds to ribosomes and the RQC complex, suggesting a role of RQC in mitochondrial quality control and a general function of Vms1 in RQC (Izawa et al., 2017). Indeed, Vms1 and its human homologue ANKZF1 have then been identified as peptidyl-tRNA hydrolase, required to remove the bound tRNA from the nascent chain, thereby facilitating release from the ribosome (Zurita Rendón et al., 2018; Verma et al., 2018). Mitochondrial functions of Vms1/ ANKZF1 Vms1, despite being a general factor in RQC, appears to particularly affect mitochondrial proteostasis (Izawa et al., 2017). Indeed, a combined deletion of the genes encoding the proteins Vms1 and Ltn1 resulted in severe growth inhibition under respiratory conditions. This growth defect was entirely dependent on CAT-tail formation by Rqc2. Consistently, aggregation and sequestration were observed, mainly of mitochondrial proteins, highlighting the importance of RQC for mitochondrial integrity (Izawa et al., 2017). Conversely, overexpression of Vms1 reduced the Rqc2-dependent aggregation, by inhibiting Rqc2 binding to ribosomes. The strong impact of Vms1 on mitochondrial proteostasis might be explained by differences in the fate of cytosolic and mitochondrial proteins. Nascent mitochondrial proteins might still be partially imported, thanks on their N-terminal mitochondrial targeting sequence. This might reduce the efficiency of Ltn1-dependent

ubiquitination on ribosome stalling. Consequently, an increase in the requirement of Vms1 in RQC, to clear nascent mitochondrial polypeptides and prevent clogging of the import channel, is not surprising. Though initially identified as a Cdc48interacting protein (Heo et al., 2010), the function of Vms1 in RQC was reported to be independent of Cdc48 (Verma et al., 2018). In contrast, the general role of Vms1 in RQC was shown to depend on Cdc48 interaction (Izawa et al., 2017). Concluding remarks Increased proteotoxic burden is a hallmark of neurodegeneration. The main cellular strategies to cope with proteotoxic stress are to increase the levels of chaperones, activate proteolytic pathways and reduce protein synthesis. Recent events highlighted that these main strategies come in different flavours and involve crosstalk between different cellular compartments. Knowing that mitochondria joined this team considerably broadens our knowledge of the ubiquitin dependent and independent crossfunctional cellular stress response mechanisms. Hopefully these findings will get us closer to therapies for the myriad of diseases caused by insufficient handling of protein damage. Acknowledgements We are grateful to T. Tatsuta for critical reading of the manuscript. This work was supported by grants of the Deutsche Forschungsgemeinschaft (DFG; ES338/3-1, CRC 1218 TP A03; to M.E.-H.), of the Fritz-Thyssen foundation (10.15.1.018MN, to M.E.-H.), of the Center for Molecular Medicine Cologne (CMMC; CAP14, to M.E.-H.), was funded under the Institutional Strategy of the University of Cologne within the German Excellence Initiative (ZUK 81/1, to M.E.-H.) and benefited from funds of the Faculty of Mathematics and Natural Sciences, attributed to M.E.-H. References Aguilaniu, H., Gustafsson, L., Rigoulet, M., and Nyström, T. (2003). Asymmetric inheritance of oxidatively damaged proteins during cytokinesis. Science 299, 1751–1753. https://doi.org/10.1126/science.1080418. Almajan, E.R., Richter, R., Paeger, L., Martinelli, P., Barth, E., Decker, T., Larsson, N.G., Kloppenburg, P., Langer, T., and Rugarli, E.I. (2012). AFG3L2 supports mitochondrial protein synthesis and Purkinje cell

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Interplay of Ubiquitination and SUMOylation with miRNAs Yashika Agrawal1,2 and Manas Kumar Santra1*

13

1National Centre for Cell Science, Savitribai Phule Pune University Campus, Pune, Maharashtra,

India.

2Department of Biotechnology, Savitribai Phule Pune University, Ganeshkhind, Pune,

Maharashtra, India.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.13

Abstract A myriad of processes occurring in each individual cell govern the functions of tissues and the overall activity of multicellular organisms. Stringent and precise multilevel control of these cellular pathway components dictates the normal functioning of cells. This control, to maintain the homeostasis in the cells, is exercised at various levels including transcription, post-transcription, translation and post-translation. miRNAs have emerged as major players regulating the post-transcriptional events in the cells, whereas ubiquitination and sumoylation are among the major post-translational events. The following sections in this chapter will discuss miRNAs, namely their synthesis and functions, the processes of ubiquitination and sumoylation and how these different level modifications crosstalk with each other in order to regulate the normal and diseased states of individuals. The life as small RNAs Introduction The last decade has witnessed a burst of knowledge in the field of RNA biology with most significant advances occurring in the branch of small noncoding RNAs that regulate genes and genome. The discovery of these small non-coding RNAs and their involvement in almost all the major

biological processes including development, cell differentiation, cell proliferation, cell death, neuronal patterning, fat and glucose metabolism, immunity in animals and apical basal patterning and development of leaves and flowers in plants (He and Hannon, 2004) have provided a new avenue for understanding the various aspects of the complex life processes rendering these small non-coding RNAs as potential therapeutic targets. These small non-coding RNAs of ≈ 20–30 nucleotide length are classified into three main categories – short interfering RNAs (siRNAs), microRNAs (miRNAs), and piwi-interacting RNAs (piRNAs). miRNAs: biogenesis in animals and plants Though a few miRNAs are transcribed as monocistronic transcripts, most of the miRNAs in animals are clustered and are expressed as polycistronic primary transcripts (Lee et al., 2003; Cai et al., 2004). The miRNAs in animals are mainly transcribed by RNA Polymerase II to generate the primary transcripts (pri-miRNA), which contain the stem loop structures and are usually several kilobases long (Lee et al., 2004). These pri-miRNA are then processed in the nucleus by a microprocessor complex containing Drosha and Di George syndrome critical region in gene 8 (DGCR8) in humans and Drosha and Pasha in D. melanogaster and C. elegans. This processing result in ≈ 60–70

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nucleotide hairpin structures called precursor miRNAs (pre-miRNA) (Denli et al., 2004; Gregory et al., 2004; Han et al., 2004; Landthelar et al., 2004). The RAN-GTP dependent Exportin-5 (EXP5) then transports the pre-miRNAs to the cytosol where they are cleaved by an endonuclease RNA III enzyme Dicer, which functions in association with a trans-activation response RNA-binding protein (TRBP) and protein kinase, interferon-inducible double-stranded RNA-dependent activator (PRKRA, also known as PACT) to generate ≈ 22 nucleotide miRNA duplexes (Grishok et al., 2001; Hutvágner et al., 2001; Ketting et al., 2001; Knight and Bass, 2001; Chendrimada et al., 2005; Haase et al., 2005; Lee et al., 2006). The miRNA duplex incorporates into the Argonaute (Ago) family protein complexes forming the RNA-induced silencing complex (RISC) wherein one strand is degraded (known as passenger strand or miRNA*) and the other remains bound to Ago complex as mature miRNA (known as guide strand or miRNA) (Khvorova et al., 2003). Thereafter, Dicer with its interacting proteins associates with the Ago family proteins containing the mature miRNA to form the RISC loading complex (RLC) (Gregory et al., 2005; Maniataki and Mourelatos, 2005; MacRae et al., 2008). The miRNA then guides the complex to its target mRNA which is silenced either by translation repression or by degradation (Kim et al., 2009; Carthew and Sontheimer, 2009). Unlike animals, the plant miRNAs are mostly present in intergenic regions of the genome and are present as independent transcriptional units (Chen, 2008). Most of the plant pri-miRNAs have been shown to have 5’cap and poly-adenylated tails (Xie et al., 2005; Megraw et al., 2006). Furthermore, the processing of plant miRNAs is wholly dependent on Dicer like-1 (DCL-1) which is affirmed by the absence of homologues of Drosha and its cofactors (DGCR8 and Pasha) (Reinhart et al., 2002; Papp et al., 2003). Another distinguishing factor between plant and animal miRNAs is the methylation of plant miRNAs after Dicer processing, which is facilitated by a methyl transferase, Hua Enhancer (HEN1) (Chen, 2008). This methylation is essential for the activity of the mature miRNA complex. The miRNA/miRNA* duplex then associates with the Ago protein complex for its further activity.

miRNAs: functions in animals and plants The advancements in the identification and understanding of miRNAs have established them as essential gene regulatory units playing roles in various biological processes including development of an organism, its cellular differentiation, metabolism and death. Functions of some miRNAs in animals and plants are discussed here. The first two miRNAs discovered in C. elegans lin-4 and let-7 was shown to have roles in early and late stages of larval development respectively (Reinhart et al., 2000; Boehm and Slack, 2005). Another miRNA lsy-6 was found to be involved in the neuronal patterning ( Johnston and Hobert, 2003; Chang et al., 2004). Similarly, the over-expression of miRNA bantam in Drosophila melanogaster was suggested to induce growth and inhibit apoptosis (Nolo et al., 2006) and miR-14 was found to be involved in the fat metabolism and suppressed cell death (Xu et al., 2003). Furthermore, miR-430 family was found to be involved in the zygotic development and neurogenesis of zebrafish as well as in the early development of frogs (Giraldez et al., 2005; Watanabe et al., 2005). Roles of several miRNAs in different stages of mice development and cell differentiation have also been recognized; miR-196 has role in the limb development (Hornstein et al., 2005), miR-1 shows the activity in differentiation of cardiomyocytes (Zhao et al., 2005) and mir-181 causes an increase in B lymphocytes and regulates the mouse haematopoietic lineage differentiation (Chen et al., 2004). Additionally, fat metabolism in humans is found to be regulated by miR-143 via enhancing the levels of extracellular signal-regulated kinase-5 (ERK5) suggesting its role in adipocyte differentiation (Esau et al., 2004). The micoRNA-155 is seen to be involved in the innate immunity (Thai et al., 2007) and miR-375 in humans is indicated to be an inhibitor of glucose-stimulated insulin secretion (Poy et al., 2004). Presence of miR-375 in the pituitary glands of zebrafish embryos indicates its role in hormone secretions as well (Wienholds et al., 2005). miRNAs like miR-32 have exhibited role in antiviral defence mechanisms (Lecellier et al., 2005). Furthermore, the tumour suppressor or oncogenic roles of miRNAs in various cancers are now well established. miR-15a and miR-16-1 exert tumour suppressor activity in chronic lymphocyte

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leukaemia (Calin et al., 2002; Cimmino et al., 2005), miR-17–92 cluster exhibits oncogenic activity in B cell lymphomas via up-regulation of c-Myc (He et al., 2005), miR-93 and miR-106a promote malignancy and enhance migration and invasion of breast cancer (Manne et al., 2017) and miRs-372 and 373 induce tumorigenesis in human fibroblasts (Voorhoeve et al., 2006). Analogous to animals, miRNAs play essential part in maintaining the normal growth patterns in the plants as well. Studies conducted on maize established the role of miR-165/166 in abaxial polarity ( Juarez et al., 2004) and two other reports revealed the role of miR-167 in plant reproductive development (Ru et al., 2006; Wu et al., 2006). Some other miRNAs were found to be involved in plant hormonal pathways including miR-159 which is regulated by Gibberellic Acid and leads to late flowering in high amounts (Achard et al., 2004) and miR-164 which affects the Auxin signal transduction pathways and leaf patterning (Nikovics et al., 2006). In Arabidopsis, elevated levels of miR-172 and miR-319 govern the developmental floral defects like absence of petals, the transformation of sepals into carpels (Aukerman and Sakai, 2003), patchy leaf shapes and delayed the flowering times (Palatnik et al., 2003). The functions of the above discussed miRNAs and many others discovered till date emphasize their importance in the growth and development of living organisms, necessitating a thorough understanding of different aspects of their activities and regulation. Ubiquitination Ubiquitination is a biochemical pathway of covalent attachment of the ubiquitin (Ub) protein to other target proteins. Ub is an evolutionary conserved, ubiquitously present, small protein of 8.5 kDa consisting of 76 amino acids (Goldstein et al., 1975; Schlesinger and Goldstein 1975; Nakayama and Nakayama, 2006). It usually ligates to the lysine residue of its substrate proteins through an isopeptide linkage between the C-terminal glycine (glycine 76) of Ub and the amino group of lysine of the substrate protein (Yang et al., 2013). Ub contains seven lysine residues at K6, K11, K27, K29, K33, K48, and K63 through which the Ub chain is extended thereby determining the fate of the target proteins. Ubiquitination can be

mono-ubiquitination where single Ub binds on a single lysine of substrate protein, multi-monoubiquitination where attachment of ubiquitin molecules occurs on various lysines scattered over the substrate or polyubiquitination where ubiquitin chains (at least four Ub conjugated with each other) attach on one or several lysines of substrate protein. Monoubiquitination predominantly promotes protein trafficking whereas polyubiquitination promotes either protein trafficking (through K63) or protein degradation (through K11 and K48) (Verhelsta et al., 2011; Bielskienė et al., 2015). Ubiquitination of the substrate protein is a multistep reaction carried by three enzymes, the ubiquitin-activating enzyme (E1), ubiquitinconjugating enzyme (E2) and ubiquitin ligase (E3). E1 enzyme activates ubiquitin in an ATP-dependent manner, wherein E1 catalyses acyl-adenylation of C-terminus of Ub which then transfers to cysteine residue of E1. E1 then interacts with E2 and transfers the Ub to its active cysteine. The E3 enzyme then interacts with both the substrate protein and E2-Ub and transfers the Ub to a lysine residue of substrate protein (Pickart, 2001). Almost 600–1000 E3 ligases have been identified in humans as opposed to 38 E2 conjugating enzymes and only two E1 enzymes (UBA1, UBA6) (Deshaies and Joazeiro, 2009; Schulman and Harper, 2009). Based on their domain structure and biochemical features, the E3 ligases have mainly been classified into four families – HECT (Homology to E6-AP C-terminus), RING finger (Really Interesting New Gene), U box type and PHD type (Plant homeodomain) (Glickman and Ciechanover, 2002; Nakayama and Nakayama 2006). Of these, RINGfinger E3 ligase is the largest family accounting for almost 95% of E3 ligases (Deshaies and Joazeiro, 2009). In HECT E3 ligase, Ub of E2 binds to active cysteine of E3 and is then transferred to substrate; whereas in RING finger and U-box type, E2 bound Ub and E3 bound substrate are brought in close contact to facilitate the direct transfer of Ub to the substrate (Grande et al., 2012). After the ubiquitination and trafficking or degradation of substrates, the Ub moieties are recycled through the deubiquitination process catalysed by Deubiquitination enzymes (DUBs) or Ubiquitin specific proteases (USPs) (Yang et al., 2013).

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Sumoylation Sumoylation is also a biochemical process very similar to ubiquitination in which a small ubiquitin-like modifier (SUMO) protein moiety is enzymatically conjugated to the target proteins (Matunis et al., 1996; Mahajan et al., 1997). SUMO is a ≈ 12 kDa protein consisting of about 100 amino acids which is synthesized as an inactive precursor. It undergoes a cleavage at its C-terminus by a family of SENP (sentrin/SUMOspecific protease) enzymes which exposes its di-glycine motif for its conjugation with the lysine residues of the substrates (Gong et al., 1999; Kim and Baek, 2006; Cashman et al., 2014). Analogous to the ubiquitination pathway, SUMOylation also involves a cascade of three enzymes – The activating enzyme E1 that activates the SUMO proteins in an ATP-dependent manner by forming a heterodimer of SAE1 and SAE2 in mammals. In this process, a thioester bond is formed between the active-site cysteine residue of SAE2 and the C-terminal glycine residue of SUMO. The second is the conjugating enzyme E2 of which Ubc9 is the only known enzyme. The active SUMO protein is then passed to the active site cysteine of Ubc9 which can then directly bind to the consensus SUMOylation motif of the target proteins. The third is the ligase enzyme E3 which transfers the SUMO protein to the substrate via an isopeptide linkage between C-terminal carboxyl group of SUMO and the lysine residue of substrate. Though E2 can directly recognize and bind to the substrate, presence of E3 in facilitating this process has also been observed (Desterro et al., 1997; Johnson and Blobel 1997; Lee et al., 1998; Gong et al., 1999; Sampson et al., 2001). Though a consensus motif [Ψ KxD/E (where Ψ is a large hydrophobic residue)] for SUMOylation is recognized, it is not an essential requirement for the sumoylation of proteins. The SUMOylation also depends on the growth microenvironments and stress stimuli. The SUMOylation plays major role in myriad of cellular functions including gene regulation, cell development and differentiation and disease progression (Hannoun et al., 2010). The SUMOylated proteins are deSUMOylated after their functions by the SUMO specific protease (SENP) and the SUMO proteins are then recycled.

The post-transcriptional and post-translational interactions Though gene expression and protein functions are regulated at various levels in the cells; the post-transcriptional and the post-translational regulation imparts rapid response and higher sensitivity towards internal or external cellular changes. The mRNA-protein correlation is essential for the normal function and about 33.15% of total variation in this correlation is contributed by the post-transcriptional biological properties (Wu et al., 2008) necessitating the better understanding of these highly dynamic processes. Micro-RNAs and the ubiquitination pathway There exists an either direct or indirect interplay between miRNA and the ubiquitination regulatory pathways which defines the fate of the cell. miRNAs control many regulatory pathways, most prominent of which include the developmental and the oncogenic processes. The continued comprehensive study into the role of miRNAs in cancer has provided us with the understanding of oncomiRs and tumour suppressor miRNAs. A multilevel complex interaction between several miRNA families and the ubiquitination pathway exists in multifarious cancers which provide a significant insight into developing the anti-cancer therapies. Our previous work identifies an indirect interaction between miRNAs and the ubiquitination machinery and establishes their role in cancer cell invasion. Our work identified miR-93 and miR106a as oncomiRs that act by post-transcriptional inhibition of tumour suppressor FBXO31 resulting in inhibition of senescence and activation of Slug which promotes the cell migration and invasion. We also discovered a feedback mechanism where Slug transcriptionally up-regulates miR-93 and miR106a leading to continued inhibition of FBXO31 expression at the post transcriptional level. But this phenomenon in the cells differs during genotoxic stress conditions wherein the miRs 93 and 106a gets inhibited, resulting in elevated FBXO31 levels, which then post-translationally down-regulates Slug levels via K48 linked polyubiquitination and degradation, preventing the cancer cell invasion and migration (Manne et al., 2017).

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An indirect regulation exists between Ubiquitin proteasome system (UPS) and the oncomiR-424 wherein miR-424 suppresses the E3 ubiquitin ligase COP1, thus preventing the ubiquitin mediated proteasomal degradation of STAT3, a key substrate of COP1 (Dallavalle et al., 2016). Another study that portrays the complex interplay existing between the microRNAs and UPS was conducted on autosomal dominant polycystic kidney disease (ADPKD) by de Stephanis et al. (2018). They described the role of miR-501-5p in regulation of p53 levels and cellular apoptosis. They describe the phenomenon wherein miR-501-5p suppresses the expression of PTEN and TSC1 genes leading to the activation of mTOR kinase, which in turn increases the expression levels of E3 ligase MDM2 resulting in the proteasomal degradation of p53 and thus the inhibition of apoptosis. Hence, inhibiting the MDM2/ mTOR signalling would lead to restoration of p53 function. Ubiquitination regulating the microRNA processing Ubiquitination plays important role in the biogenesis and activity of miRNAs thereby modulating the post-transcriptional events by the post-translational modifications (PTMs). Drosha is an indispensable enzyme for the processing of pri-miRNA to premiRNA in the nucleus. The regulation of Drosha by PTMs represents the requirement of maintaining protein homeostasis in miRNA biogenesis. Acetylation at the N-terminus of Drosha is essential for its stability and the processing of pri-miRNA to premiRNA in the nucleus; but the lack of acetylation at Drosha leads to its ubiquitin mediated proteasomal degradation, which was evident on infection of gastric mucosa AGS cells with Helicobacter pylori. The infected AGS cells did not show any difference at the mRNA levels of Drosha but the protein levels were decreased owing to lesser acetylation and increased ubiquitination mediated degradation (Tang et al., 2013). Ubiquitin proteasome system maintains the required levels of majority of cellular proteins and degrades the damaged or redundant proteins. The Ago proteins are essential for the processing/activity of miRNAs which are also regulated by the UPS. The mouse homologue of lin-41 interacts with and ubiquitinates Ago2 protein thus negatively regulating miRNA pathway (Rybak et al., 2009). A similar

mechanism of interaction between miRNA pathway and the ubiquitin proteasome pathway exists in plants. F-box protein FBXW2, a component of Cullin-RING E3 ubiquitin ligase, has been shown to negatively regulate Ago1 in Arabidopsis thaliana (Earley et al., 2010). In addition to the miRNA processing enzymes, the miRNA functional complex is also controlled by the UPS. Trinucleotide repeat containing six (TNRC6) is a part of the RISC–miRNA complex that facilitates the suppression of target mRNA. An E3 ubiquitin ligase Tripartite motif 65 (TRIM65) proteasomally targets TNRC6 and thereby relieves the miRNA driven repression of target mRNA, thus regulating the activity of miRNA (Li et al., 2014). Micro-RNAs regulating the ubiquitination process Interplay of miRNA with ubiquitination machinery occurs at various levels and one of them is the direct interaction of miRNAs with F-box proteins to modulate the cellular activities. One such oncomiR is miR-223, which suppresses the E3 ubiquitin ligase FBXW7 to promote oesophageal squamous cell carcinoma (Kurashige et al., 2012). FBXW7 and FBXW11 are known to be involved in malignancy via the SCF-E3 ubiquitin ligase. It has been shown that miR-182 suppresses the activity of both FBXW7 and FBXW11 by binding to their 3′UTR thereby promoting the tumorigenesis of non-small cell lung cancer (Chang et al., 2018). Additionally, miR-27a post transcriptionally represses FBXW7 in the G2/M and early G1 phases of the cell cycle but relieves it during the G1/S phase boundary. This release allows FBXW7 to facilitate the proteasomal degradation of its target protein Cyclin E to promote the S-phase progression of cell cycle. Furthermore, the miR-27a represses the tumour suppressive activity of FBXW7 and thereby promotes the paediatric acute lymphoblastic leukaemia (ALL) (Lerner et al., 2011). It is an established fact that the activity of p53 strongly regulates the fate of cancer cells necessitating the continuous studies to identify new facets of this protein’s functions and regulation. The study by Yang et al. (2017) proposes that oncomiR-100 inhibits apoptosis in poorly differentiated gastric cancer cells by ubiquitin-mediated degradation of p53. The oncomiR-100 inhibits an E3 ubiquitin ligase RNF144B by binding to its 3′UTR, which

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is then unable to ubiquitinate and degrade pirh2 E3 ubiquitin ligase. The active pirh2 ubiquitinates and degrades p53 inhibiting the apoptosis of gastric cancer cells. The addition of antagomiR-100 reverses this process and promotes the ubiquitination mediated degradation of pirh2 E3 ligase resulting in active p53 and apoptosis of gastric cancer cells. Thus, this work provides a new mechanism of ubiquitin mediated regulation of p53 by miR-100 (Yang et al., 2017). A subset of tumour cells is known as the tumourinitiating cells (TICs) or cancer stem cells (CSCs) and have the enhanced self-renewal capacity and the ability to repopulate tumour after anticancer therapies. These CSCs have thus been strongly implicated in progression and metastasis of varied tumours including many subsets of breast cancers. A study by Guarnieri et al. proposes the role of miR-106b–25 cluster in regulation of these CSCs via modulation of NOTCH signalling pathway. miR-106b–25 cluster represses the E3 ubiquitin ligase neural precursor cell expressed developmentally down-regulated gene 4-like (NEDD4L) which prevents the degradation of NOTCH1 protein thereby promoting the stem cell properties of TICs (Guarnieri et al., 2018). ITCH is HECT type E3 ubiquitin ligase known to promote tumour progression and metastasis in multiple cancers by proteasomally targeting the large tumour suppressor (LATS) 1 and 2 kinases of the Hippo signalling pathway (Harvey and Tapon, 2007; Salah et al., 2011). A study in pancreatic cancer shows the role of miR-106b in post-transcriptional repression of ITCH, thus inhibiting metastatic progression (Luo et al., 2016). TNFα signalling pathway is essential in many biological processes including immune response, cell proliferation, differentiation and apoptosis. For the specific signal transduction, the balance between the cascade proteins and their expression is maintained by the ubiquitin proteasome system (UPS). In the Synovial Fibroblasts (SFs) of Rheumatoid Arthritis (RA) condition, miR-17 enhanced the Lys63-linked polyubiquitination of TNFα in activated RA-SFs to stabilize certain TNFα cascade proteins. Conversely, high level of Lys48-linked ubiquitination was observed for TRAF2, cIAP1 and cIAP2 in presence of miR-17 under similar conditions in RA-SFs indicating the

inhibition of downstream signalling cascade. Additionally, the levels of DUBs, USP2 and PSMD13 were suppressed and the binding of TRAF2 with the cIAP1/cIAP2 complex was also interfered by miR-17, thereby inhibiting the cascade (Ahmed et al., 2016). Disruption in UPS is known to be associated with many diseases including cancer (Mani and Gelmann, 2005) and the dysregulation of Cullin proteins in the Cullin ring finger E3 ligases (CRLs) has been shown to promote tumorigenesis (Chen et al., 1998; Chen et al., 2014; Sang et al., 2015). Osteosarcoma has been a leading cause of death in children and adolescents with a 5 year survival rate of 15%–30% (Ward et al., 2014). A study conducted on osteosarcoma demonstrated that CRL4B forms an E3 complex with DNA damage binding protein 1 (DDB1) and CUL4-associated factor 13 (DCAF13) (CRL4BDCAF13) which proteasomally degrades the tumour suppressor PTEN (phosphatase and tensin homologue deleted on chromosome 10). This study further established that miR-300 decreased the stability of CRL4BDCAF13 and inhibited the proteasomal degradation of PTEN (Chen et al., 2018). Another study by Zhou et al. (2018) reveals the negative effect of miR-192-5p on deubiquitinating enzyme USP1 in osteosarcoma. miR-192-5p inhibits the proliferation and cell migration of osteosarcoma by targeting USP1, thereby acting as a tumour suppressor. These studies provide new avenues for osteosarcoma therapy. miRNAs have been implicated in almost all biological processes and are believed to be critical during development in both the vertebrates and invertebrates (Brennecke et al., 2003; Johnson et al., 2005). miR-135a is seen to be highly expressed in zygotes of mouse but its levels decline thereafter. It was found that miR-135a is required for the first cell division of zygote and it exerts its function by modulating the function of E3 ubiquitin ligases. miR-135a suppresses the E3 ubiquitin ligase Seven in absentia homologue 1 (Siah1) in the mouse zygotes by binding to its 3′UTR. This results in the high expression levels of chemokinesin DNA binding protein (Kid) promoting chromosome compaction and proper cell division. Conversely, inhibition of miR-135a results in higher Siah1 levels, which then proteasomally degrades Kid

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leading to disruption in the first cleavage of zygote. This work establishes the crosstalk of miRNA and the ubiquitination pathway during a key developmental process in mouse (Pang et al., 2011). The E3 ubiquitin ligase Nedd4 (neural precursor cell expressed developmentally down-regulated protein 4) is involved in the actin dependent patterning of heart in Drosophila thus regulating the heart development in fly. miR1 directly binds to 3′UTR of Nedd4 and thereby modulates the expression levels of Nedd4 substrates in heart (Zhu et al., 2017). Micro-RNAs and the SUMOylation pathway As discussed earlier, the micro-RNA biogenesis and functions are closely linked with post-translational modifications where both influence the other to modulate their activity. This segment encompasses the available information interlinking the interactions between SUMOylation and miRNA pathways. SUMOylation regulating the microRNA processing Though it is normally accepted that SUMO modifications directly affect the substrate proteins yet a study proposes that SUMOylation can regulate a target without directly modifying it and can enhance the levels of certain proteins. This study demonstrates that SUMOylation can negatively regulate the expression levels of miR-34b/c through AKT and FOXO3a. SUMOylation at Lys476 of AKT enhances its kinase activity, which prevents the nuclear localization of FOXO3a thereby diminishing the levels of tumour suppressor miR-34b/c and resulting in increased levels of oncoprotein c-Myc (Li et al., 2018). Thus, this study suggests a paradigm in which SUMOylation affects the miRNA levels which in turn alters the levels of other target proteins. The general process of miRNA maturation has been discussed earlier. However, certain miRNA families like let-7 which contain short G-rich stretches in their terminal loop require an hnRNP K homology (KH)-type splicing regulatory protein (KHSRP) as an essential component of Drosha complex for miRNA processing. The study by Yuan et al. (2017) suggests the role of SUMOylation in

processing of TL-G-Rich miRNAs via KHSRP regulation. They demonstrate that modification of KHSPR by SUMO1 at Lys87 inhibits its interaction with the pri-miRNA/Drosha-DGCR8 complex resulting in the defects of pre-miRNA formation. The decrease of TL-G-Rich miRNAs like let-7 consequently leads to tumorigenesis (Yuan et al., 2017). Exosomes in the cytosol act as the storage compartments for RNAs including mRNAs, miRNAs and other non-coding RNAs. But the process of sorting these miRNAs into different exosomes is not yet clear. Villarroya-Beltri et al. (2013) identified that the RNA-binding protein heterogeneous nuclear ribonucleoprotein A2B1 (hnRNPA2B1) recognizes the EXOmotifs of specific miRNAs, binds to them and controls their loading into exosomes. The SUMOylation of hnRNPA2B1 is a prerequisite for its binding to the miRNAs thereby controlling the activity of miRNA functions (Villarroya-Beltri et al., 2013). SUMOylation also modulates the activity of enzymes involved in miRNA biogenesis and thus regulates the post-transcriptional events by the post-translational changes on the miRNA pathway proteins. A study in alveolar macrophages shows that cigarette smoke induces SUMOylation of RNA endonuclease enzyme Dicer which is essential for generation of mature miRNA from pre-miRNA. This SUMOylation reduces the activity of Dicer resulting in miRNA processing defect, thus altering the miRNA profile of cells (Gross et al., 2014). The gene silencing activity of small RNAs requires their association with the Ago family proteins. The SUMOylation of Ago2 at Lys402 is shown to be essential for its RNAi activity providing a new insight into the Ago2 mediated control of gene expression ( Josa-Prado et al., 2015). Conversely, another report by Sahin et al. (2014) suggests that SUMOylation of Ago2 at Lys402 by SUMO1 or SUMO2/3 antagonizes its stability and enhances the turnover of Ago2 protein. The role of miRNAs as post-transcriptional regulators is well established. However, increasing evidences suggest the direct functions of pri-/ pre-miRNAs in regulation of gene expression (Liu et al., 2008; Trujillo et al., 2010; Yue et al., 2011; Roy-Chaudhuri et al., 2014). The SUMOylation of DGCR8 at Lys707 by SUMO1 is shown to stabilize

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its association with pri-miRNA without altering its association with Drosha and the Microprocessor complex. This enhanced affinity of SUMOylated DGCR8 with pri-miRNA has been linked to the direct functions of pri-miRNA in recognition and repression of target mRNA and also with the observed DGCR8’s function in regulation of tumorigenesis and cell migration (Zhu et al., 2015). Furthermore, the miRNA biogenesis has been closely linked to SUMOylation with various factors involved in the processing pathway being SUMOylated for the formation of mature miRNA. The SUMOylation of TARBP2 at Lys52 has been implicated in its stabilization by preventing its Lys48 linked polyubiquitination and degradation. The SUMOylated TARBP2 recruits Ago2 to constitute the RLC and simultaneously promotes the loading of pre-miRNA into the RLC for its further processing into mature miRNA (Chen et al., 2015). Micro-RNAs regulating the SUMOylation process Neurobiologists worldwide have been working to understand and prevent the ever increasing incidences of ischaemic strokes. The ischaemic strokes are caused due to deprivation of oxygen to certain parts of brain leading to permanent brain damage. Lee et al. (2012) have discovered a link between two groups of miRNAs, ubiquitin like modifiers (ULMs) and neuroprotection in ischaemic conditions using hibernating torpor ground squirrels as model organisms. They identified that compared to the active animals, the miRNAs of miR-200 family (miR-200a, b, c/miR-141/miR-429) and the miR182 family (miR-182/miR-183/miR-96) were consistently repressed during the torpor phase of squirrels. Additionally, the expression of various ULMs including SUMOylation and their conjugation with target proteins was found to be increased (Lee et al., 2012). The same group later ascertained the negative role of miR-182 and miR-183 in the process of SUMO conjugation. They identified the small molecules that inhibit miRNAs 182 and 183 and enhance the global SUMOylation in cells and observed these effects to be cytoprotective during oxygen and glucose deprivation in neurons (Bernstock et al., 2016). In addition to the involvement of SUMO modifications in neuronal activity, their importance in the cardiac functions is also well recognized. Any

alteration or defect in Vascular Smooth Muscle Cells (VSMCs) results in a variety of diseases including atherosclerosis (Kawai-Kowase et al., 2009), hypertension (Wang et al., 2007), cancer (Coinu et al., 2006), vascular aneurysms and asthma (Satoh et al., 2009) rendering it essential to understand their normal regulation. miR-200c was found to inhibit the expression of Krϋppel-like transcription factor 4 (KLF4) and the SUMOconjugating enzyme Ubc9. Existence of a feedback loop was also discovered wherein high levels of Ubc9 SUMOylate KLF4 which then transcriptionally represses miR-200c (Zheng et al., 2015). This work depicts the regulation of SUMOylation process by a micro-RNA and alternately the regulation of miRNA by SUMOylated protein. Moreover, miR-146a is shown to negatively regulate SUMO1 by targeting its 3′UTR and preventing its association with cardiac sarcoplasmic reticulum calcium ATPase pump (SERCA2a). The SUMOylated form of SERCA2a is known to be beneficial for heart failure patients making miR-146a and SERCA2a gene SUMOylation a combined therapeutic target for patients with heart failure (Oh et al., 2018). The diverse role of miRNAs and SUMO modifications are yet again exhibited by their involvement in the conversion of white adipose tissues to brown adipose tissues which burn the normal fat aiding in weight loss. Koh et al. have defined the role of miR30a-5p in repression of E2 SUMO ligase Ubc9 in human adipocytes which prevents the SUMOylation of PR domain containing 16 (PRDM16) protein and leads to acquisition of brown fat features (Koh et al., 2016). The bacterial and viral pathogens have evolved sophisticated methods to control the host machineries and to ensure their survival and proliferation. A study by Verma et al. (2015) depicts the mechanism adopted by the intestinal pathogen Salmonella typhimurium to promote its infection and intracellular survival in the host cells. They demonstrate the up-regulation of miR-30c and miR-30e on S. typhimurium infection. These miRNAs repressed the expression levels of crucial SUMO pathway enzymes Ubc9 and PIAS1. This depletion results in the SUMO-conjugated proteome decrease and increases the pathogenic infection (Verma et al., 2015). Another study performed in Epstein–Barr virus (EBV) predicts several viral miRNAs that modulate SUMO-regulated functional networks

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to promote their replication and infection in host (Callegari et al., 2014). In yet another work, a specific mechanism of infection of EBV in the host cells was identified. The active viral infection was accompanied with an increase in the SUMO conjugated proteins and down-regulation of SUMO targeted ubiquitin ligase RNF4. The viral miRNA miR-BHRF1-1 post-transcriptionally inhibits RNF4 to prevent the ubiquitin mediated degradation of SUMOylated viral proteins. Thus, this study recognizes a new strategy of viral interference with the SUMOylation pathway and describes the role of miR-BHRF1-1 in viral replication (Li et al., 2017). All these studies attest to the existence of a complex and intricate interconnection of two different levels of regulation to maintain the cellular homeostasis. Both the post-transcriptional and the post-translational regulation of gene expression are essential to maintain the homeostasis and both these processes are individually regulated. Yet, their inter-regulation further enforces the importance of their precise functions and the need to ascertain their error-free roles. Though the regulation of miRNAs with SUMOylation and with ubiquitination is being explored; yet the complex inter-connections point to the presence of a threeway regulation between miRNAs, ubiquitination and SUMOylation. Thus, many more aspects of these inter-regulations need to be further explored to provide a better insight into the pathway modulations and to exploit them for therapeutic purposes. Acknowledgement Part of this work was financially supported by the Department of Biotechnology Grant (BT/ PR6690/GBD/27/475/2012) to M.K.S and partly by the National Centre for Cell Science, Department of Biotechnology, Ministry of Science and Technology, Government of India (to M.K.S.). Y. A. is a DBT senior research fellow. References

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The Role of Ubiquitination and SUMOylation in DNA Replication Tarek Abbas1,2,3*

14

1Department of Radiation Oncology, University of Virginia, Charlottesville, VA, USA.

2Department of Biochemistry and Molecular Genetics, University of Virginia, Charlottesville, VA,

USA.

3Center for Cell Signaling, University of Virginia, Charlottesville, VA, USA.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.14

Abstract DNA replication is a tightly regulated conserved process that ensures the faithful transmission of genetic material to define heritable phenotypic traits. Perturbations in this process result in genomic instability, mutagenesis, and diseases, including malignancy. Proteins involved in the initiation, progression, and termination of DNA replication are subject to a plethora of reversible post-translational modifications (PTMs) to provide a proper temporal and spatial control of replication. Among these, modifications involving the covalent attachment of the small protein ubiquitin or the small ubiquitin-like modifier (SUMO) to replication and replicationassociated proteins are particularly important for the proper regulation of DNA replication as well as for optimal cellular responses to replication stress. In this chapter, we describe how the ubiquitination and SUMOylation processes impact DNA replication in eukaryotes and highlight the consequences of deregulated signals emanating from these two versatile regulatory pathways on cellular activities.

Regulation of eukaryotic DNA replication Initiation of DNA replication Eukaryotic DNA replication is tightly regulated such that cells replicate their entire genome once and only once in a given cell cycle (Machida et al., 2005). For mammalian cells, this is no easy task since each proliferative somatic cell must efficiently replicate approximately 6 billion base pairs (in male cells) from roughly 250,000 replication origins scattered throughout the genome with each division cycle (Cadoret et al., 2008; SequeiraMendes et al., 2009; Karnani et al., 2010). With roughly 600 million new blood cells born in the bone marrow of an adult human (Doulatov et al., 2012), one cannot grasp the magnitude of the task the replication machinery has to accomplish. The core machinery of DNA replication is highly conserved in all living organisms, but eukaryotes diverge significantly in its regulation owing to the larger, more complex genomes (Kaguni, 2011). In bacteria (e.g. Escherichia coli), replication initiates at individual replication initiation sites or

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Figure 14.1  Regulation of replication initiation in eukaryotes. A model depicting the step-wise assembly of the pre-replication complex (Pre-RC) in late mitosis and during G1 phase of the cell cycle, followed by replisome assembly. The six-subunit ORC complex binds to origins of DNA replication in late M and early G1. This is followed by the recruitment of the replication licensing proteins CDC6 and CDT1, and the loading the of the MCM2–7 helicase (origin licensing). At the G1/S transition, the Dbf4-dependent kinase (DDK) and CDK enzymes promote the assembly of the replicative helicase, or the CMG complex, which is marked by the recruitment of the GINS complex (Sld5, Psf1, Psf2, Psf3), along with CDC45. MCM10 aids in this process by recruiting and stabilizing DNA polymerase α (POL α). Other proteins [e.g. Treslin (Sld3 in yeast), RecQL4 (Sld2 in yeast), and TopBP1] help in the replisome assembly (not shown). As DNA synthesis begins in S-phase, the unwound DNA is stabilized by the single-stranded DNA binding protein RPA, and DNA polymerases (POL ε and POL δ) initiate replication.

origin of chromosome replication (OriC) where two replication forks assemble and move in opposite direction at a rate of 1 Kb/sec/fork to replicate the entire 4.4 Mb circular chromosome within 30 minutes (Katayama, 2017). The AAA+ ATPase replication initiator protein DnaA, which is conserved in virtually all bacteria, recognizes and binds with high specificity to high density GATC repeat sequences (DnaA box) within these replicons, and both DNA binding and ATP

hydrolysing activities of DnaA are essential for replication initiation (Hansen and Atlung, 2018). Initiation of DNA replication in eukaryotes (Fig. 14.1) is similarly dependent on the binding of a DnaA-like six-subunit origin recognition complex (ORC) to replication origins in an ATP-dependent manner (Bell and Stillman, 1992; Bell and Dutta, 2002). ORCs from various eukaryotes exhibit a wide range of sequence-recognition specificities. For example, whereas ORC from budding yeast

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specifically recognizes 11-bp or 17-bp conserved sequences within the ≈ 400 autonomously replicating sequences (ARS) (Dhar et al., 2012), the fission yeast ORC recognizes AT stretches (but without sequence consensus) through the AThook motif present on the Orc4 subunit (Chuang and Kelly, 1999; Segurado et al., 2003; Dai et al., 2005; Hayashi et al., 2007). The six-subunit ORC complex from high eukaryotes binds DNA without sequences specificity (Vashee et al., 2003; Schaarschmidt et al., 2004), although replication initiates from genomic loci that are enriched for AT-rich sequences, dinucleotide repeats, asymmetrical purine-pyrimidine sequences, and matrix attachment region (MAR) sequences (Li and Stillman, 2012; Kumar and Remus, 2016). Additional epigenomic features, such as the DNA topology, transcription factors and regulatory elements, local chromatin environment as well as the replication initiation proteins CDT1 and CDC6 play a role for the selectivity of ORC to stably bind replication origins (Masai et al., 2010; Li and Stillman, 2012; Kumar and Remus, 2016). Replication initiation in high eukaryotes is also dependent on histone methylation. For example, recent studies demonstrate a critical role for histone H4 methylation at Lys-20 (H4K20) at replication origins in the nucleation of DNA replication (Tardat et al., 2010; Beck et al., 2012a). Mono-methylation of H4K20 (H4K20me1) is catalysed by the histone methyltransferase (HMT) SET8 (also known as PR-SET7), which deposits a single methyl group on Lys-20 of nucleosomal histone H4 (Nishioka et al., 2002; Xiao et al., 2005). When tethered to specific genomic loci, catalytically active, but not inactive, SET8 recruits pre-RC proteins on chromatin and replication initiates from these sites (Tardat et al., 2010). Mono-methylated Lys-20 of H4 is subject for further methylation [di- and tri-methylation (H4K20me2 and H4K20me3, respectively)] by the SUV4-20H1/H2 HMTs (Schotta et al., 2008). The conversion of H4K20me1 to H4K20me2/3 by SUV4-20H1/H2 likely plays an important role for SET8-dependent replication initiation, as the recruitment of ORC1 as well as the ORC-associated protein (ORCA) protein (both capable of binding H4K20me in vitro) to chromatin requires SUV4–20H1/H2 (Beck et al., 2012a).

Cell cycle regulation of replication initiation in eukaryotes Initiation of eukaryotic DNA replication is cell cycle regulated, requires the ordered assembly of several proteins at replication origins, and occurs in two distinct steps that are temporally separated within the cell cycle (Fig. 14.1). The first step involves the establishment of pre-replicative complexes (preRCs) through the sequential assembly of ORC, CDC6, and CDT1, followed by the loading of the six-subunit helicase MCM2–7 (minichromosome maintenance proteins, subunits 2–7) at origins of replication in late mitosis (M) and early G1 (first Gap) phase of the cell cycle. Once the MCM2–7 complexes are loaded onto replication origins (origin licensing), the ORC-CDC6-CDT1 pre-RC components are no longer required to initiate replication. In the second step, licensed origins are activated in S phase (DNA synthesis phase) to generate active replication forks (origin firing), and this requires the conversion of the inactive double hexameric MCM2–7 helicase to an active replicative helicase, the CMG complex, which is composed of MCM2–7, its cofactor CDC45, and the GINS complex (Gambus et al., 2006; Moyer et al., 2006; Pacek et al., 2006; Ilves et al., 2010; Kang et al., 2012). This conversion process, which is highly conserved from yeast to human, requires the activity of the Dbf4-dependent kinase (DDK) and the cyclin-dependent kinase (CDK). Both kinases are activated at G1/S transition, and their concerted activities promote the recruitment of several scaffolding proteins and DNA polymerase Polε to assemble the replisome. Studies in yeast have shown that while DDK phosphorylates multiple Mcm2–7 subunits to recruit the scaffolding protein Sld3 with its partners Sld7 and Cdc45, CDK phosphorylates the two other scaffolding subunits Sld2 and Sld3, thereby promoting their interaction with Dpb11 (TopBP1 in human) in cooperation with Polε and GINS (Gambus et al., 2006; Moyer et al., 2006; Pacek et al., 2006; Ilves et al., 2010; Muramatsu et al., 2010; Kumagai et al., 2010, 2011; Boos et al., 2011; Kang et al., 2012; Bruck and Kaplan, 2015, 2017; Fang et al., 2016). Replisome assembly also requires the action of multiple protein complexes involved in monitoring replication fork progression, in coordinating DNA synthesis with chromatin assembly, and in responding to

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genetic perturbations by generating checkpoint and damage signals (Leman and Noguchi, 2013). Progression and termination of DNA replication Origin firing in eukaryotes is temporally regulated with distinct early- and late-replicating genomic regions and exhibits significant flexibility that gives the cells control over situations that interfere with normal progression of replication forks (RenardGuillet et al., 2014). Activation of the CMG complex is tightly coupled to the activity of histone chaperones, nucleosome-remodelling complexes and chromatin-modifying enzymes (Groth, 2007, 2009; Jasencakova and Groth, 2010). These later factors facilitate nucleosomal disassembly ahead of the replication forks and reassembling nucleosomes with correct positioning following their passage. The DNA primase–POLα complex generates primers that will be extended by POLε (for continuous DNA synthesis of the leading strand) or POLδ (for the discontinuous replication of the lagging strand) (Bell and Dutta, 2002; Bell and Labib, 2016). Several other proteins are important for the maturation and ligation of the Okazaki fragments. In budding yeast, these include the flap endonuclease Rad27, the DNA helicase-nuclease Dna2, the Exo1 exonuclease and the DNA ligase Cdc9 (Bell and Labib, 2016). DNA topoisomerases relieve topological stresses created by the moving replication forks, and many proteins and protein complexes aid in removing other barriers to the progressing replication forks, such as tightly-bound non-histone proteins. Other proteins must be recruited to deal with difficult to replicate genomic sequences or with actively transcribing genomic templates. Progression of DNA replication is also tightly coordinated with the establishment of sister chromatid cohesion as well as with the activity of multiple proteins and protein complexes involved in the sensing and repair of DNA damage that may be encountered during DNA replication (Waters et al., 2009; Villa-Hernandez and Bermejo, 2018). Termination of DNA replication occurs at converging replication forks from neighboring origins of replication, although in some cases, termination occurs at chromosomal termination regions (TERs) defined by replication pausing elements contained within these TERs (Labib and Hodgson, 2007; Fachinetti et al., 2010). Genomic and mechanistic studies in budding

yeast identified 71 such regions, and further demonstrated that these TERs can influence fork progression and merging (Fachinetti et al., 2010). Replication across these TERs, which are characterized by the accumulation of X-shaped structures, can be facilitated by the Rrm3 DNA helicase, and the fusion of the converging forks at these sites is aided by DNA topoisomerase II (Topo II or Top2 in yeast), thus counteracting abnormal genomic transitions (Fachinetti et al., 2010). Termination of DNA replication is marked by the completion of local DNA synthesis, the decatenation of the two daughter strands by DNA topoisomerases and the final disassembly of the replisome (Dewar and Walter, 2017; Gambus, 2017). Ubiquitin-dependent regulation of DNA replication Overview of the ubiquitinproteasome system ATP-dependent and ubiquitin-mediated proteasomal degradation through the ubiquitinproteasome system (UPS) provides an efficient mean to regulate protein abundance and maintain homeostatic regulation of cellular physiology, and is involved in almost all cellular activities (Kornitzer and Ciechanover, 2000; Amir et al., 2001; Ciechanover and Schwartz, 2002; Glickman and Ciechanover, 2002; Hershko, 2005; Schwartz and Ciechanover, 2009). The process ensures the timely down-regulation of cellular proteins via the 26S proteasome, where roughly 80% of all intracellular proteins are digested into small peptides (Skaar et al., 2014). Proteasomal degradation is preceded by the covalent attachment of multiple copies of the highly conserved 76 amino-acid ubiquitin protein [linked together through Lys-48 (Lys-48 linkage) or Lys-11 (Lys-11 linkage)] to substrate proteins (Fig. 14.2). This occurs in a series of enzymatic reactions involving the activity of an E1 ubiquitinactivating enzyme, the transfer of the activated ubiquitin to an E2 ubiquitin-conjugating enzyme (UBC), and the selective transfer of ubiquitin to the substrate through the activity of an E3 ubiquitin ligase (Glickman and Ciechanover, 2002; Groll and Huber, 2003; Kornitzer and Ciechanover, 2000; Teixeira and Reed, 2013). Whereas Lys-48 and Lys-11- polyubiquitination signal proteolytic

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Figure 14.2  Regulation of protein ubiquitination. Protein ubiquitination involves the sequential activity of an E1 ubiquitin activating enzyme, an E2 ubiquitin conjugating enzyme and an E3 ubiquitin ligase (a cullin-based E3 ubiquitin ligase is shown as an example). The E3 ligase transfers the ubiquitin moiety (Ub) to the substrate through interaction with the E2-charged ubiquitin, forming a covalent isopeptide bond between the C-terminus of ubiquitin and a specific Lys residue on the substrate. Polyubiquitin chains (poly-Ub) can be formed by covalently conjugating the C-terminus of a ubiquitin moiety to one of seven Lys residues (e.g. Lys-48) or to the fist Met residue (M1) on another ubiquitin moiety. Polyubiquitination through Lys-48 (K48), and Lys-11 (K11) linkages directs the substrate to the 26S proteasome, where the substrate is proteolytically degraded into small peptides, with the ubiquitin moieties released and recycled. Other homotypic poly-ubiquitin chains [e.g. M1, Lys-63 (K63)], or the attachment of single ubiquitin moieties to individual (mono-ubiquitination) or multiple (multi-ubiquitination) Lys residues do not signal protein degradation and serves other distinct biological functions. A set of deubiquitinating enzymes (DUBs), which are highly specific cysteine proteases, can cleave the isopeptide bonds between the ubiquitin and ε-amino group of the substrate Lys or the Lys of the other ubiquitin moiety in a polyubiquitin chain. DUBs can also cleave the peptide bond between ubiquitin and the N-terminal methionine of another ubiquitin moiety.

degradation, other homotypic poly-ubiquitin chains involving ubiquitin conjugation through Lys-63 or Met-1, or the attachment of single ubiquitin moieties to individual (mono-ubiquitination) or multiple (multi-ubiquitination) Lys residues do not signal protein degradation, but play a role in various cellular process (Fig. 14.2). These include activities that impact protein–protein interaction, transcription factor activation, protein synthesis, and cellular response to DNA damage (Wang et al., 2001; Tokunaga et al., 2009; Yang et al., 2010; Behrends and Harper, 2011; Dantuma and Pfeiffer, 2016; Schwertman et al., 2016). E3 ubiquitin ligases are critical for conferring specificity for the substrates to be ubiquitinated

and, in some cases, for dictating the nature of substrate ubiquitination (Zheng and Shabek, 2017). Cullin-RING (Really Interesting New Gene) E3 ubiquitin Ligases (CRLs) represent the largest family of E3 ubiquitin ligases in mammals, promoting the polyubiquitin-mediated degradation of approximately 20% of total cellular proteins via the proteasome (Hotton and Callis, 2008; Deshaies and Joazeiro, 2009; Soucy et al., 2009; Duda et al., 2011; Hua and Vierstra, 2011; Lipkowitz and Weissman, 2011; Sarikas et al., 2011; Lydeard et al., 2013; Chen et al., 2015). Other E3 ubiquitin ligases including the HECT (Homologous to the E6-AP Carboxyl Terminus) domain containing E3 ubiquitin ligases are described in more details in

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recent excellent reviews (Li et al., 2008; Deshaies and Joazeiro, 2009; Skaar et al., 2014; Zheng and Shabek, 2017). CRLs are involved in many cellular processes, including DNA replication, cell cycle progression and cellular proliferation (Petroski and Deshaies, 2005; Bosu and Kipreos, 2008; Hotton and Callis, 2008). CRL family members include eight cullin proteins (cullin 1–3, 4A, 4B, 5, 7 and cullin 9) and a cullin-like protein ANAPC2 or APC2. The multi-subunit CRL1 E3 complex, better known as the SCF ligase (SKP1-Cullin1-FBox protein), is the prototype of this family of E3 ligases and is best known for its role in controlling cell cycle progression, proliferation, and differentiation (Nakayama and Nakayama, 2005; Maser et al., 2007; Welcker and Clurman, 2008; Huang et al., 2010; Duan et al., 2012; Lee and Diehl, 2014). The SCF ubiquitin ligase is built around the cullin 1 scaffold subunit, which binds the SKP1

(S-phase kinase-associated protein 1) adaptor protein through its N-terminal domain (Fig. 14.3). The SKP1 subunit bridges one of several substrate receptors with their cognate substrates to the cullin 1 subunit (Wang et al., 2014). The cullin 1 C-terminal domain, on the other hand, is essential for substrate polyubiquitination through its interaction with a small RING domain protein (RBX1 or RBX2), which is essential for the recruitment of the E2 UBCs. The substrate specificity of the SCF ligase complex is dictated by a family of substrate receptors, which are collectively called F-box proteins owing to their interaction with the SPK1 protein through conserved F-box motif (Skaar et al., 2014; Heo et al., 2016). Mammalian cells express at least 69 F-box proteins, and thus assemble a large number of distinct SCF ligases. Each F-box protein is capable of recognizing a subset of ubiquitination substrates, commonly through interaction with

Figure 14.3  Regulation and restraint of origin licensing via the UPS. A schematic illustrating the various steps involved in origin licensing through the cell early part of the cell cycle and their regulation via the UPS. Three E3 ubiquitin ligases [APC/CCDH1 (left) SCFSKP2 (centre), and CRL4CDT2 (right)] ensure the ordered but restricted assembly of the various pre-RC components in late M and early G1 phase of the cell cycle. APC/CCDC20 (not represented schematically) helps promote mitotic cyclin degradation and helps assembly of the AP ligase APC/ CCDH1. These E3 ligases are activated at distinct phases of the cell cycle (represented below). Distinct substrate receptors, CDC20 or CDH1, an F-box protein (SKP2), or a DCAF (CDT2) is critical for bridging the substrates for polyubiquitination by their cognate E3 ligases (APC/C, SCF, and CRL4 ligase, respectively). The CRL4CDT2 ligase recognizes its substrates only when they interact with chromatin-bound PCNA, and thus, only targets chromatin-bound proteins for degradation. Other substrates are targeted for ubiquitination only in their soluble form (see text for details). M: mitosis. G1/S/G2: First gap, DNA synthesis and second phases of the cell cycle, respectively. APC* multiple subunits that together function as adaptor proteins

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phosphorylated residues within small consensus ‘degron’ motifs in these substrates (Kipreos and Pagano, 2000; Cardozo and Pagano, 2004; Skaar et al., 2013; Wang et al., 2014; Heo et al., 2016). Ubiquitination is reversible and protein abundance is controlled by a set of deubiquitinases or DUBs (Fig. 14.2). DUBs play pivotal roles in the regulation of protein turnover, protein or enzymatic activation, protein–protein interaction, protein recycling, and cellular localization (Mukhopadhyay and Dasso, 2007; Komander et al., 2009; Reyes-Turcu et al., 2009; Hickey et al., 2012), and are increasingly recognized as attractive therapeutic targets for cancer therapy (Hoeller and Dikic, 2009; Crosas, 2014; D’Arcy et al., 2015; Pfoh et al., 2015; Lei et al., 2017; Harrigan et al., 2018). Biochemically, DUBs hydrolyse the isopeptide bonds between the ε amino group of Lys side chains of the target substrate and the C-terminal group of ubiquitin, or the peptide bond between the α amino group of the target protein and the C-terminus of ubiquitin (Wilkinson, 1997). Regulation of cell cycle control of replication via the UPS The SCFSKP2 E3 ubiquitin ligase (Fig. 14.3), composed of the core SCF complex and the substrate receptor SKP2 (S-phase kinase-associated protein 2), is one of the best characterized SCF ligases and best known for its role in promoting cell cycle progression through the activation of CDKs (Nakayama and Nakayama, 2005; Skaar et al., 2013). CDK activity controls replication initiation, and the SCFSKP2 ligase is critical for increasing CDK activity in G1 and in early S phase, by promoting the ubiquitination CDK inhibitors p21CIP1, p27KIP1, and p57KIP2 (Nakayama and Nakayama, 2005; Skaar et al., 2013). SCFSKP2 also promotes progression through G2 phase, primarily through its ability to promote the ubiquitin-dependent proteolysis of cyclin A. The degradation of cyclin A in late S-phase ensures the availability of sufficient CDK1 molecules to assemble cyclin B–CDK1 complexes essential for progression through G2. Progression through S phase also requires the availability of sufficient CDK2 molecules for assembly with cyclin A, and this is mediated, at least in part, through the activity of the SCFFBXW7 ligase, which utilizes FBXW7 as a substrate receptor to degrade CDK2-phosphorylated cyclin E following entry

into S phase (Clurman et al., 1996; Koepp et al., 2001). CDK activity must be kept low during mitosis and in early G1, and this is facilitated by the multisubunit APC/C (anaphase promoting complex/ cyclosome) ubiquitin ligase (Fig. 14.3), which promotes the ubiquitination and degradation of cyclin A and cyclin B (den Elzen and Pines, 2001). APC/C complex is the largest E3 ubiquitin ligase in mammals that is built around the APC2 cullinlike scaffold and utilizes the CDH1 (Hct1 in yeast) or CDC20 substrate receptors for recognizing and promoting the polyubiquitination (both Lys-48and Lys-11-linked ubiquitin conjugation) of key drivers of the cell cycle (Visintin et al., 1997; Zachariae and Nasmyth, 1999; Pines, 2006; van Leuken et al., 2008). The specificity of the APC/C ligases is based on the substrate receptors CDH1/CDC20 ability to recognize degron motifs (destruction D-boxes and KEN-boxes) within the targeted substrates (Pfleger and Kirschner, 2000; Pfleger et al., 2001). The APC/CCDC20 is activated in G2 and in early mitosis in a cyclin B–CDK1-dependent manner, and this is critical for the initial degradation of mitotic cyclins (cyclin A in prometaphase and cyclin B in metaphase) (Rahal and Amon, 2008). The SCFβTRCP1 ligase utilizing the substrate receptor β-transducin-repeat-containing protein 1 (βTRCP1) aids in activating APC/CCDC20 both by stimulating CDK1 activity through enhancing the ubiquitination and degradation of the CDK1 tyrosine kinase inhibitor Wee1, and by relieving inhibition of the APC/CCDC20 via promoting the degradation of the F-box protein and early mitotic inhibitor 1 (EMI1), which is an endogenous inhibitor of APC/C (Guardavaccaro et al., 2003; Watanabe et al., 2004). Cyclin B–CDK1 subsequently phosphorylates the APC3 and APC1 subunits of the APC/C ligase, thereby facilitating the docking of CDC20 onto the APC/C ligase and the assembly of the active ligase complex (Fujimitsu et al., 2016). Cyclin B–CDK1 additionally phosphorylates CDH1, resulting in conformational changes in CDH1 that preclude the assembly of an active APC/CCDH1 ligase. In late mitosis and through G1, CDC20 is exchanged for CDH1/Hct1 following the dephosphorylation and activation of CDH1 by the CDC14A phosphatase (Cdc14 in yeast), and the newly assembled APC/CCDH1/Hct1 ligase complex

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maintains low cyclin B levels ( Jaspersen et al., 1999; Donzelli et al., 2002; Sullivan and Morgan, 2007; Robbins and Cross, 2010). APC/CCDH1/Hct1 activation is facilitated by APC/CCDC20, which mediates the release of the CDC14A from centrosomes (and yeast Cdc14 phosphatase from the nucleolus) through an unknown mechanism (Shirayama et al., 1999; Bembenek and Yu, 2001; Kaiser et al., 2002; Mocciaro et al., 2010; Chen et al., 2016). APC/CCDH1 ligase activity is critical for inactivating mitotic CDK and for exit from mitosis. This is accomplished via the APC/C CDH1/Hct1-dependent polyubiquitination and degradation of not only mitotic cyclins, but also of CDC20, thereby stabilizing the APC/CCDC20 ligase ubiquitination substrate and the CDK inhibitor p21 (or its homologue in yeast, Sic1) (Shirayama et al., 1999; Amador et al., 2007). APC/CCDH1 maintains low CDK activity through early G1 by promoting the degradation of the SKP2 subunit of the SCFSKP2 ligase (Bashir et al., 2004; Wei et al., 2004). This prevents the premature degradation of the CDK inhibitors p21 and p27, which can bind to and inhibit CDK2 in G1 (Abbas and Dutta, 2009). At the G1/S transition, the APC/CCDH1/Hct1 ligase is inactivated through the phosphorylation of the CDH1/Hct1 subunit by cyclin E-CDK2 (Cappell et al., 2016). Further inhibition of CDH1 (and CDC20) is mediated by EMI1, and this has been proposed to mark a ‘point of no return’ for entry into S-phase (Reimann et al., 2001; Cappell et al., 2016). Stabilization of mitotic cyclins is essential for the completion of DNA synthsis and for progression throguh G2 (Di Fiore and Pines, 2007). Ubiquitin-dependent restraint of origin licensing One of the most important features of regulating DNA replication in eukaryotes is the uncoupling of origin licensing, which takes place in late M and early G1, from origin firing in S-phase (Fig. 14.3). This ensures that replication initiates from individual origins of replication during S phase and is prevented from firing again until nuclear division is completed. The fluctuating CDK activity during the cell cycle, which is largely dependent on the ubiquitin-dependent proteolysis described above, is essential for this uncoupling process. The rising CDK activity in S phase is incompatible for origin licensing as many of the origin licensing

proteins are phosphorylation substrates for CDK. CDK-dependent phosphorylation of certain replication licensing proteins suppresses origin licensing, either because this triggers their ubiquitination and proteolytic degradation or results in their exclusion from the nucleoplasm (Blow and Dutta, 2005; Abbas and Dutta, 2017). For example, CDK-phosphorylated human ORC1 protein, the largest subunit of the ORC complex, undergoes ubiquitin-dependent proteolysis specifically in S phase cells, and this is mediated by the SCFSKP2 ubiquitin ligase (Méndez et al., 2002; Tatsumi et al., 2003). Unlike human ORC1, ORC1 from Drosophila undergoes ubiquitin-dependent proteolysis via the APC/CFZr/CDH1 E3 ligase as soon as cells exit mitosis and requires a domain in the N-terminus of Drosophila ORC1 that is non-conserved in human ORC1 (Araki et al., 2003, 2005; NarbonneReveau et al., 2008). The replication licensing protein CDC6 is also targeted for proteolysis through the UPS, and this ensures that replication occurs only once during each division cycle. Although yeast Cdc6, a factor essential for loading Mcm2–7 onto replication origins is ubiquitinated through the SCFCdc4 E3 ubiquitin ligase, mammalian CDC6 was previously shown to be shuttled outside the nucleus through the rising CDK activity at the G1/S transition, and this was sufficient to prevent origin relicensing (Aparicio et al., 1997; Tanaka et al., 1997; Saha et al., 1998; Fujita et al., 1999; Jiang et al., 1999; Petersen et al., 1999; Alexandrow and Hamlin, 2004). However, recent evidence suggests that mammalian CDC6 also undergoes ubiquitin-dependent proteolysis. Three E3 ubiquitin ligases are implicated in restricting the expression of mammalian CDC6 to late mitosis and early G1. In G1 cells, and in cells exiting the cell cycle into quiescence, mammalian CDC6 is ubiquitinated and degraded via the APC/ CCDH1 ligase, and this is dependent on an interaction between the substrate receptor CDH1 and CDC6 (Petersen et al., 2000). This ubiquitin-dependent degradation of CDC6 ensures that origin licensing is completed before cells transverse through G1 phase and is dependent on the D-box and KENbox motifs of CDC6, since a combination of point mutations of these motifs stabilizes CDC6 both in G1 and in quiescent cells. In cells entering the cell cycle from quiescence, and as cyclin E-CDK2 activity builds up, CDC6 is phosphorylated by cyclin

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E-CDK2, and this prevents CDC6 recognition by CDH1, and promotes origin licensing before entry into S phase (Mailand and Diffley, 2005). In S and in G2 phases of the cell cycle, mammalian CDC6 is ubiquitinated and degraded via the activity of the CRL4CDT2 and the SCFCyclinF, respectively, and both of these activities are essential for preventing origin relicensing and rereplication (Clijsters and Wolthuis, 2014; Walter et al., 2016). The role of CRL4CDT2 and the APC/C ubiquitin ligases in restraining origin licensing In addition to CDC6, CDT1 is another major ubiquitination and degradation substrate for the CRL4CDT2 ubiquitin ligase (Fig. 14.3). The multisubunit CRL4 ligase complexes share common features with SCF ligases but utilize a different set of substrate adaptors collectively known as DCAFs (DDB1 and Cullin 4 associated factors) (Angers et al., 2006; Higa and Zhang, 2007). DCAFs include at least 49 family members of WD motif-rich proteins that, similar to the F-box protein substrate receptors of the CRL1 ligases, recognize and recruit substrates for polyubiquitination by the CRL4 ligase (Angers et al., 2006; He et al., 2006; Higa et al., 2006; Jin et al., 2006). The core CRL4 complex is comprised of one of two paralogues, cullin 4A or cullin 4B, that binds DDB1 (DNA damage-specific protein-1) through its N-terminus (Fig. 14.3). DDB1 is an adaptor protein that is analogous to the SKP1 subunit in the SCF ligases, and functions to bridges one of the DCAFs to the cullin subunit. The C-terminus of the cullin 4 subunit binds to RBX1 or RBX2, which are required for the recruitment of E2 UBCs, necessary for polyubiquitination. The DCAF CDT2 assembles with CRL4 to form a rather unique E3 ubiquitin ligase that appears to recognize its substrates when they interact with the DNA polymerase δ processivity factor proliferating cell nuclear antigen (PCNA) through a specialized PCNA-interacting protein motif or PIP-box, and only when PCNA is loaded onto chromatin (Arias and Walter, 2006; Senga et al., 2006). This likely restricts the CRL4CDT2 activity to S and early G2 phases of the cell cycle as well as during the repair of certain DNA lesions that requires PCNA (e.g. nucleotide excision repair) (Higa et al., 2003; Abbas and Dutta, 2011; Havens and Walter, 2011; Abbas et al., 2013). The PIP-box contained within

CRL4CDT2 substrates, commonly referred to as the ‘PIP degron’, is a variant of the PIP-box motif that is commonly used by many proteins to interact with PCNA, and contains, in addition to the canonical sequence [Q-X-X-(I/L/M)-X-X-(F/Y)-(F/Y)], conserved Thr and Asp acid residues at positions 5 and 6 respectively, as well as a basic amino acid residue c-terminal of the PIP-box (at position +4), as well as a second basic amino acid at position +3 or +5 (or both) (Havens and Walter, 2009, 2011; Abbas et al., 2010; Michishita et al., 2011). The ability of CRL4CDT2 to prevent origin relicensing and rereplication was initially attributed to its ability to specifically target CDT1 for proteolysis during S phase (Arias and Walter, 2006; Jin et al., 2006; Nishitani et al., 2006; Senga et al., 2006). In fact, in various eukaryotes, with the exception of budding yeast where Cdt1 is exported to the cytoplasm along with the Mcm2–7 complex (Devault et al., 2002; Tanaka and Diffley, 2002), deficiency in cullin 4, DDB1 or in CDT2 induces rereplication and genomic instability reminiscent of that seen following CDT1 overexpression (Vaziri et al., 2003; Zhong et al., 2003; Jin et al., 2006; Lovejoy et al., 2006; Sansam et al., 2006; Tatsumi et al., 2006; Kim et al., 2008). Rereplication induced by CRL4CDT2 inactivation results in the accumulation of DSBs, presumably due to the accumulation of replication intermediates and replication fork stalling/collapse, and activates DNA damage checkpoints, both of which can be partially suppressed through the co-depletion of CDT1 (Zhu et al., 2004; Lovejoy et al., 2006; Zhu and Dutta, 2006). We now know that CRL4CDT2 prevents rereplication through promoting the polyubiquitination and degradation of multiple proteins involved in origin licensing during S and G2 (Fig. 14.3). These include not only CDC6 and CDT1, but also SET8 and p21, both of which bind PCNA through PIP degrons (Abbas et al., 2010; Abbas et al., 2008; Centore et al., 2010; Clijsters and Wolthuis, 2014; Jørgensen et al., 2011; Kim et al., 2008; Nishitani et al., 2008; Oda et al., 2010; Tardat et al., 2010). The Drosophila melanogaster E2f1 transcription factor is another ubiquitination substrate for CRL4CDT2 whose degradation in S-phase is critical for rereplication suppression and is dependent on the interaction between E2f1 and PCNA through a PIP degron that is absent in the human protein (Shibutani et al., 2008).

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The PCNA-dependent and CRL4CDT2 catalysed polyubiquitination and degradation of chromatinbound p21 in S phase is important for sustaining elevated CDK2 activity essential for S phase progression and for freeing PCNA from inhibitory p21 (Abbas and Dutta, 2009). Increased stability of p21 following CRL4CDT2 inhibition contributes to the rereplication phenotype observed in these cells, presumably because of inhibition of CDK activity, a condition compatible for origin licensing, but not likely to be sufficient to do so in the absence of stabilized CDT1 and SET8. This is evident by the fact that the expression of PCNA binding-deficient mutant of p21 (p21ΔPIP), which is resistant to CRL4CDT2-mediated polyubiquitination and degradation induces robust senescence but is only associated with minor rereplication (Kim et al., 2008; Benamar et al., 2016). This is contrary to the role of stabilized SET8 in rereplication induction in cells with inactivated CRL4CDT2, which is both necessary and sufficient to induce rereplication (Abbas et al., 2010; Tardat et al., 2010; Benamar et al., 2016). It is important to note that both p21 and CDT1 are also necessary for rereplication induction in cells expressing CRL4CDT2-resistant mutant SET8 protein (SET8 ΔPIP) (unpublished observations). Although the role of SET8 in promoting rereplication when stabilized in S phase is not entirely clear, it is likely to be dependent on its ability to monomethylate H4K20 and the subsequent conversion of this histone mark to H4K20me2/3 at replication origins (Abbas et al., 2010; Tardat et al., 2010 Beck et al., 2012a;). Unlike chromatin-bound CDT1, p21 and SET8, soluble forms of these proteins are targeted for ubiquitination and proteolysis both in late G1 and/ or S phase by other ubiquitin ligases, most notably, the SCFSKP2 ligase (Fig. 14.3). This E3 ligase targets CDT1 for ubiquitination and degradation following its phosphorylation at Thr-29 by cyclin A-CDK2 in late G1 and in S phase (Li et al., 2003; Liu et al., 2004; Takeda et al., 2005). Similarly, p21 is phosphorylated at Ser-130 by CDK2 and this promotes the degradation of soluble p21 at the G1/S transition and in S phase (Bornstein et al., 2003). Soluble SET8 was also suggested to be targeted for ubiquitination via the SCFSKP2 ligase in S phase, although it is not clear whether this requires SET8 phosphorylation (Yin et al., 2008; Oda et al., 2010). However, depletion or deletion of SKP2,

unlike CRL4CDT2 inactivation, does not induce rereplication, suggesting that even in the presence of increased soluble fractions of these proteins, the CRL4CDT2 is sufficient to prevent origin licensing by efficiently removing the chromatin-bound forms of these proteins. Although CDT1, p21 and SET8 are largely undetectable in late G1 and throughout most of S-phase, they reappear in late S phase and in G2 (Abbas et al., 2010). In the case of CDT1, this accumulation is critical for progression through G2, but this is not likely to be dependent on CDT1 ability to bind chromatin. This conclusion stems from the observation that CDT1 is phosphorylated by CDK1 in late S and early G2, and this prevents CDT1 from binding to chromatin, and that abolishing CDK1dependent phosphorylation of CDT1 inhibits cell cycle progression (Rizzardi et al., 2015). CDT1 is additionally phosphorylated by the stress-activated mitogen-activated protein kinases (MAPK) p38 and JNK and this too, precludes recognition by CRL4CDT2 (Chandrasekaran et al., 2011). A recent study suggested that CDT1 is also ubiquitinated and degraded in G2 cells via the SCFFBXO31 ubiquitin ligase, and that inactivation of this pathway results in minor rereplication ( Johansson et al., 2014). It is unclear from this study however, how the stabilized CDT1 in G2 cells with inactivated SCFFBXO31 gains access to chromatin in the presence of elevated CDK1 activity. The reaccumulation of SET8 in G2, similar to CDT1, is critical for cell cycle progression, and this is thought to be mediated through its ability to promote histone H4 methylation needed for chromatin condensation prior to entry into mitosis (Beck et al., 2012b; Jørgensen et al., 2013). Following the accumulation of methylated H4K20, and from prophase to early anaphase, cyclin B/ CDK1 phosphorylates SET8 on Ser-29, and this removes SET8 from chromatin, without inhibiting its methyltransferase activity (Wu et al., 2010). The dephosphorylation of SET8 in late M-phase by the CDC14 phosphatase primes the SET8 protein for proteolytic degradation via the APC/CCDH1 ligase (Wu et al., 2010). The importance of p21 reaccumulation in G2 on the other hand, is not clear, but may be important to restrict cyclin A-CDK2 activity. In addition to the mechanisms by which CDT1 is targeted for proteolysis in late G1 and in S phase, metazoans evolved another mechanism

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to suppress CDT1 activity through the expression of a small protein inhibitor of CDT1 called geminin (Wohlschlegel et al., 2000; Tada et al., 2001). Geminin is under the transcriptional control of E2F1, which transactivates dozen other genes essential for S-phase progression, including cyclin E (Wong et al., 2011). Geminin, however, undergoes ubiquitin-dependent degradation in late mitosis and early G1 via the APC/CCDH1 E3 ligase activity (McGarry and Kirschner, 1998). In late G1 and early S-phase, geminin is phosphorylated by cyclin E-CDK2, and this prevents its recognition by CDH1, stabilizing the protein, which directly binds CDT1 and sterically hinders its ability to recruit MCM2–7 complexes to replication origins (Tada, 2007; Caillat and Perrakis, 2012). At the same time, cyclin E-CDK2 phosphorylates CDH1, thereby inactivating APC/CCDH1 (Cappell et al., 2016). In addition, residual CDH1 is inhibited by EMI1, marking a ‘no return’ decision to enter S-phase (Cappell et al., 2016). EMI1 also binds CDC20 and inhibits the APC/CCDC20 ligase in S-phase, thereby stabilizing mitotic cyclins A and B, which are essential for the completion of DNA synthesis and G2

progression (Reimann et al., 2001; Di Fiore and Pines, 2007; Cappell et al., 2016). As mentioned above, in S phase, the SCFSKP2 ligase cooperates with CRL4CDT2 to promote the degradation of soluble and chromatin-bound CDT1, respectively. The former pathway is aided by cyclin A-CDK2, which phosphorylates CDT1 at Thr-29, and requires EMI1 for suppressing APC/CCDC20, which would otherwise ubiquitylate and degrade not only cyclin A, but also geminin. Whereas suppressing geminin initiates rereplication in certain cell types, it is insufficient to do so in some other cancer cell types or in non-malignant cells (Machida and Dutta, 2007; Zhu and Depamphilis, 2009; Benamar et al., 2016). Activation of S phase APC/C ligase on the other hand (e.g. by depleting EMI1), is sufficient to initiate rereplication in the majority of mammalian cells examined (Machida and Dutta, 2007; Benamar et al., 2016). Together, these findings highlight the importance of CRL4CDT2 and EMI1 for restraining origin licensing in S phase by preventing the accumulation of chromatin-bound and active CDT1, as well as other replication licensing proteins.

Figure 14.4  Posttranslational modification by SUMOylation. A schematic representing the various steps involved in the SUMOylation and deSUMOylation cycle. SUMO E1, E2 and E3 enzymes (mammalian representative of these enzymes is shown) promote the conjugation of SUMO to substrate proteins. DeSUMOylation is catalysed by SUMO-specific proteases [mammalian SENPs (Sentrin/SUMO-specific proteases) is shown as a representative example] and is involved both in SUMO maturation and in the removal of SUMO moieties from protein substrates.

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SUMO-dependent regulation of replication initiation Overview of the SUMOylation process Modification via the small ubiquitin-like molecule SUMO (Fig. 14.4) also plays important roles in the regulation of eukaryotic DNA replication as well as the regulation of multiple other cellular activities including DNA repair, transcription, nuclear transport, and protein quality control (Sarangi and Zhao, 2015; Jalal et al., 2017; Zilio et al., 2017). Similar to ubiquitination, sumoylation involves the covalent conjugation of SUMO or SUMO chains to the ε amino-group Lys residue of substrates, and requires the sequential action of E1 activating, E2 conjugating, and E3 ligase enzymes ( Johnson, 2004; Gareau and Lima, 2010; Lamoliatte et al., 2014), reminiscent of that involved in protein ubiquitination. SUMO, like ubiquitin, is usually conjugated to Lys side chains of substrate protein and can be conjugated at single Lys in the substrate proteins (mono-sumoylation), at multiple Lys residues of the substrate proteins (multi-sumoylation), or form various length chains at single Lys in the protein substrates (poly-sumoylation) (Fig. 14.4). Like ubiquitination, protein modification by sumoylation is reversible and is regulated by a set of SUMO-specific cysteine proteases (Mukhopadhyay and Dasso, 2007; Hickey et al., 2012). SUMO proteases deconjugate SUMO proteins using their isopeptidase activity, cleaving between the terminal Gly of SUMO and the substrate Lys (Hickey et al., 2012). The first described SUMO protease, the S. cerevisiae protein U1p1 (UBL-specific protease 1), exhibits distant similarity to certain viral proteases but is unrelated to any known deubiquitinating enzyme (Li and Hochstrasser, 1999). Mammalian cells express at least six SUMO-specific proteases, known as SENPs or Sentrin/SUMO-specific proteases (SENP1-SENP3 and SENP5-SENP7), that share significant sequence homology with U1p1, and can be broadly classified into three subfamilies based on their sequence homology, subcellular localization and substrate specificity (Mukhopadhyay and Dasso, 2007; Hickey et al., 2012). Three additional SUMO-specific proteases, DESI1 (deSUMOylating isopeptidase 1), DES12 and USPL1

(ubiquitin-specific protease-like) exist in mammalian cells and share only little sequence similarity to U1P or SENPs (Schulz et al., 2012; Shin et al., 2012). Some SUMO-specific proteases are also important for SUMO maturation, as they cleave the precursor or inactive form of SUMO at the c-terminus to expose two glycine residues. SUMO proteases play important roles in protein–protein interaction and in regulating cellular localization, and significant effort is dedicated for the development of pharmacological inhibitors of this class of proteases for therapeutic purposes (Mukhopadhyay and Dasso, 2007; Hickey et al., 2012; Kumar and Zhang, 2015; Bialik and Woźniak, 2017). Regulation of replication initiation proteins via SUMOylation As is the case for ubiquitination, modification of replication initiation proteins by sumoylation helps restrict origin licensing to late mitosis and early G1. Initial studies in budding yeast demonstrated that multiple subunits of the ORC complex undergo sumoylation, although the functional significance of these modifications is not entirely clear (Cremona et al., 2012). Studies of human ORC2 demonstrated that this subunit is sumoylated in G2/M. ORC2 sumoylation restricts the ORC complex to centromeric regions within the genome and enhances the demethylation of histone H3 lysine 4 (H3K4) in centromeric chromatin via the recruitment of the H3K4 demethylase KDM5A (Craig et al., 2003; Prasanth et al., 2004; Lee et al., 2012; Huang et al., 2016). Inhibition of ORC2 sumoylation results in rereplication, polyploidy and DNA damage at centromeric chromatin that correlate with the accumulation of H3K4 trimethylation (H3K4me3) in centromeric chromatin, reduced transcription from centromeric α-Satellites, and replication from decondensed pericentric heterochromatin (Huang et al., 2016). It remains to be seen whether the sumoylation of other ORC subunits or ORC2 from the other eukaryotes play a specialized role in the regulation of origin licensing as that seen for human ORC2 or exhibit similar, generalized, and, possibly, redundant function in preventing relicensing of replication origins from centromeric heterochromatin through promoting epigenetic changes.

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Similar to ORCs, multiple subunits of the MCM2–7 complex from various eukaryotes were found to be sumoylated (Golebiowski et al., 2009; Elrouby and Coupland, 2010; Cremona et al., 2012; Hendriks et al., 2014; Ma et al., 2014; Schimmel et al., 2014; Tammsalu et al., 2014; de Albuquerque et al., 2016; Wei and Zhao, 2016). MCM2–7 sumoylation appears to also negatively regulate replication initiation. This is supported by the finding that, in both man and yeast, the sumoylation of the six-subunit complex is detectable in G1, preceding DDK1-mediated phosphorylation of the MCM4 subunit, but is rapidly declined as cells enter S phase and remains undetectable until the G1 of the next cycle; the exception is with the yeast Mcm7 subunit, which persists throughout S phase and peaks with the completion of replication (Cremona et al., 2012; Schimmel et al., 2014; de Albuquerque et al., 2016; Wei and Zhao, 2016). Studies in yeast demonstrate that Mcm2–6 sumoylation increases its association with the PP1 phosphatase, thereby preventing premature phosphorylation of Mcm4, an essential step for CMG formation and origin firing (Davé et al., 2014; Hiraga et al., 2014; Mattarocci et al., 2014; Wei and Zhao, 2016). At the G1/S transition, and as cells enter S phase, the DDK kinase activity rises, and this, combined with Mcm2–6 desumoylation, potentially via the Ulp2 protease (de Albuquerque et al., 2016; Wei and Zhao, 2016), aid in Mcm4 phosphorylation, CMG activation, and origin firing (Wei and Zhao, 2016). A further evidence in support of a negative role for sumoylation in the regulation of replication initiation in eukaryotes is obtained from a study in Xenopus, where the expression of SUMO-specific proteases or a dominant-negative SUMO E2 was found to increase origin firing (Bonne-Andrea et al., 2013). Because the PPIDDK-mediated regulation of MCM2–7 activation is conserved across eukaryotic species (Wotton and Shore, 1997; Lee et al., 2003; Cho et al., 2006; Masai et al., 2006; Montagnoli et al., 2006; Tsuji et al., 2006; Cornacchia et al., 2012; Hayano et al., 2012; Yamazaki et al., 2012), these studies support the conclusion that negative regulation of MCM2–7 phosphorylation through sumoylation is an evolutionary conserved mechanism that regulates replication initiation in eukaryotes.

Ubiquitin and SUMO regulation of DNA synthesis Proteolytic and non-proteolytic roles for ubiquitin and SUMO at the replisome Emerging evidence support important roles for protein ubiquitination and sumoylation in the regulation of unperturbed DNA synthesis (Fig. 14.5), as well as in coordinating DNA synthesis with chromatin dynamics (Almouzni and Cedar, 2016; García-Rodríguez et al., 2016; Henikoff, 2016; Talbert and Henikoff, 2017). Proteomic analysis demonstrated that many of the components of the replisome were found to be ubiquitinated (Wagner et al., 2011). Although ubiquitination plays both proteolytic and non-proteolytic functions during DNA synthesis, sumoylation of replisome components almost invariably plays only nonproteolytic regulatory roles. The non-proteolytic regulatory functions of ubiquitin and SUMO are not always apparent, although in some cases their role is beginning to be appreciated. For example, the catalytic subunit of polymerase δ in the fission yeast Saccharomyces Pombe is stable despite undergoing ubiquitination in unperturbed S phase (Roseaulin et al., 2013). Pol2, the catalytic subunit of DNA polymerase ε, however, is ubiquitinated and degraded via the SCFPof3 ligase (Roseaulin et al., 2013). This implies that the synthesis of the leading strand requires a ‘fresh’ supply of DNA polymerase, whereas the synthesis of the discontinuous lagging strand does not (Roseaulin et al., 2013). In mammalian cells, both regulatory subunits of DNA polymerase δ (POL δ), p66 and p12, are ubiquitinated during S phase, and this modification appears to regulate protein–protein interactions either within the polymerase holoenzyme or with other replication factors (Liu and Warbrick, 2006). Interestingly, the suppression of fork progression in response to DNA damage is mediated, at least in part, through ubiquitin-dependent proteolysis of the p12 subunit via the CRL4CDT2 ligase, which requires the interaction of p12 with PCNA (Terai et al., 2013). This is only one of the several examples of the role of UPS in regulating DNA replication under replication stress or in response to DNA damage, which are described in greater details in several recent outstanding reviews (Sommers et al.,

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Figure 14.5 Ubiquitin and SUMO regulation of DNA synthesis. A schematic model of the replication fork during DNA synthesis in eukaryotic cells. The CMG replicative helicase (MCM2–7/GINS/CDC45) unwinds the duplex DNA ahead of the replication fork. Topoisomerase TOPO I is important for the relaxation of the positive supercoiling building ahead of the replication fork. TOPO II (not shown) can resolve the intertwining of the daughter DNA strands resulting from the fork rotation behind the replication fork. Single-stranded DNA (ssDNA) is coated by the ssDNA binding protein RPA (replication protein A). The replication factor C (RFC) loads PCNA and DNA polymerase ε (POL ε) to synthesize the leading strand (continuous replication). On the lagging strand, DNA polymerase α (POL α), which is stabilized by the Minichromosome maintenance protein 10 (MCM10), synthesizes short RNA/DNA primer. RFC subsequently displaces POL α, and polymerase δ (POL δ) synthesizes short DNA segments (Okazaki fragments). The flap structure-specific endonuclease FEN1 processes the 5′ ends of Okazaki fragments, and the DNA ligase I (LIG I) joins the DNA fragments (discontinuous replication). Many of these proteins (shown on right) are modified by ubiquitination and SUMOylation and this is important for the regulation of DNA synthesis (see text for details).

2015; García-Rodríguez et al., 2016; Renaudin et al., 2016). Another replisome protein that undergoes both proteolytic and non-proteolytic ubiquitination is the minichromosome maintenance protein 10 (MCM10). Mcm10 was first identified by Lawrence Dumas and colleagues in a screen for temperaturesensitive mutants for S phase progression defects in S. cerevisiae and denoted as dna43 (Dumas et al., 1982). MCM10 was subsequently identified (and the gene sequenced) in an independent study aimed at identifying replication initiation mutants that are defective in the maintenance of minichromosomes (Merchant et al., 1997). MCM10 is an essential DNA replication factor and is conserved in all eukaryotes but is absent in bacteria and archaea. The protein functions primarily as a scaffold protein with DNA binding properties but lacks enzymatic functions. Initial studies in fission yeast demonstrated that Mcm10/Cdc23 plays a role in

replication initiation through facilitating Cdc45 chromatin binding, an essential step in CMG activation (Gregan et al., 2003). Subsequent studies showed that Mcm10 facilitates the initial strand separation through its binding to origins through its Zink finger-dependent DNA binding activity (Kanke et al., 2012; van Deursen et al., 2012; Thu and Bielinsky, 2013). In budding yeast, Mcm10 appears to play an additional role in replication elongation through interacting with and stabilizing the catalytic subunit of DNA polymerase α (Pol1) (Ricke and Bielinsky, 2004). In G1 and in S phase, the budding yeast Mcm10 undergoes mono-ubiquitination at two Lys residues (diubiquitination) and this was shown to be essential for its interaction with PCNA and for cell growth (Das-Bradoo et al., 2006; Thu and Bielinsky, 2013). Similar to budding yeast, mammalian MCM10 interacts with and stabilizes the catalytic subunit of DNA POL α (p180) (Fig. 14.5), and this is important for efficient DNA

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synthesis (Chattopadhyay and Bielinsky, 2007; Zhu et al., 2007). Whether mammalian MCM10 undergoes di-ubiquitination, and if this regulates its ability to interact with PCNA or with other components of the replisome is not known. Mammalian MCM10 however, was shown undergo ubiquitindependent degradation both in unperturbed cells and following exposure to of cells to ultraviolet radiation (UV) (Kaur et al., 2012; Romani et al., 2015). Although MCM10 degradation following DNA damage may be important to halt DNA synthesis in the face the bulky DNA lesions induced by UV, the significance of its proteolytic degradation during unperturbed S phase remains to be determined. Systematic and proteome-wide proteomic studies demonstrate that many of the replisome proteins that are regulated via the UPS are also sumoylated (Cremona et al., 2012; Tammsalu et al., 2014; Bursomanno et al., 2015). Similar enrichment for poly-sumoylated proteins during DNA synthesis is also observed using an in vitro replication assay in Xenopus egg extract (BonneAndrea et al., 2013). Additional studies utilizing a method of isolating proteins on nascent DNA coupled with mass spectrometry (iPOND-MS) also demonstrate that chromatin isolated within the vicinity of the replisome is significantly enriched for sumoylated proteins (Lopez-Contreras et al., 2013; Dungrawala et al., 2015). These studies also demonstrated a relative depletion of ubiquitination events, suggesting an interaction between ubiquitination and sumoylation at the replisome. The identity of the E3 SUMO ligase responsible for protein sumoylation at the replisome is not known, but PIAS1 is a good candidate given its enrichment at these active replicating sites (Lecona et al., 2016). The USP7/HAUSP (Herpesvirusassociated ubiquitin-specific protease) DUB is another protein that is enriched at active DNA synthesis sites and may be responsible for the observed depletion of ubiquitinated proteins (Lecona et al., 2016). USP7 is a SUMO-DUB (SDUB), and is one of only two DUBs (the other is USP11) that have been shown to deubiquitylate sumoylated proteins (Hendriks et al., 2015; Lecona et al., 2016). Pharmacological inhibition of USP7 slows replication fork progression, inhibits new origin firing, and reverses the high-SUMO and low-ubiquitin chromatin environment observed

at or near the replisome (Bonne-Andrea et al., 2013; Lopez-Contreras et al., 2013; Lecona et al., 2016). How USP7 regulates new origin firing and replication progression is not entirely clear, but likely dependent on the stabilization of sumoylated replisome components that are essential for the replisome activity (Lecona et al., 2016; Wei and Zhao, 2016). This conclusion is substantiated by the reduced replication progression in SUMO E2 and E3 mutants as well as by the prolonged S phase progression seen in human cells with inactivated UBC9 SUMO-conjugating enzyme (Cremona et al., 2012; Schimmel et al., 2014; Hang et al., 2015). These studies, however, do not exclude the possibility that the accumulation of ubiquitinated proteins (or their ubiquitin-dependent degradation) upon USP7 inhibition may contribute to the inhibition of replication progression or new origin firing. One of the most notable examples of replisome proteins that is regulated by sumoylation is the budding yeast Pol2. Pol2 sumoylation is mediated by the Nse2/Mms21 SUMO ligase, and this sumoylation, as well as the sumoylation of Mcm6, is reduced not only in cells with mutations in Nse2, but also in cells deficient in Rtt107, a multi-functional scaffolding protein that plays multiple roles in replication progression (Hang et al., 2015). Although the main function of Pol2 sumoylation is not entirely clear, it is tempting to speculate that it may have important regulatory role for controlling DNA polymerase activity during replication fork progression. Significantly, the Nse2/Mms21 SUMO ligase, along with the Ubc9 SUMO-conjugating enzyme, also plays a role for the sumoylation of Smc5 and Smc6 subunits of the SMC (structural maintenance of chromosomes) SMC5/6 complex, and this is important for the repair of collapsed replication forks and for counteracting recombinogenic events at damaged replication forks (Ampatzidou et al., 2006; Branzei et al., 2006; Chen et al., 2009; Xue et al., 2014). Interestingly, Rtt107, which plays an important role in cellular response to replication stress to reduce replication-associated recombination, forms two additional and distinct complexes with the cullin 4 E3 ubiquitin ligase Rtt101Mms22 (Collins et al., 2007; Hang and Zhao, 2016; Xue et al., 2014), and with the Slx4 scaffolding protein (Hang and Zhao, 2016). The Rtt101Mms22 ubiquitin ligase ubiquitylates acetylated histone H3, and this facilitates nucleosome assembly during replication

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(Han et al., 2013). The Rtt107–Slx4 complex on the other hand, is critical for controlling recombination during DNA replication, particularly under conditions of replicative stress (Chin et al., 2006; Roberts et al., 2006; Ohouo et al., 2010). The Ataxia telangiectasia related protein kinase ATR and the checkpoint protein CHK1 play important roles in stabilizing stalled replication forks and for preventing their collapse into DSBs. In mammalian cells, the generation of DSBs following ATR inhibition is dependent on the SLX4 scaffold endonuclease, and requires the activity of the RNF4 E3 ubiquitin ligase that promotes the ubiquitin-dependent degradation of sumoylated proteins at stalled replication forks (Ragland et al., 2013). Interestingly, RNF4 also promotes the polyubiquitination of activated Fanconi anaemia proteins FANCD2 and FANCI following their ATR-dependent sumoylation by the SUMO E3 ligases PIAS1/PIAS4 at stalled replication forks (Gibbs-Seymour et al., 2015). Ubiquitinated FANCD2 and FANC1 are subsequently removed from the stalled replication sites through the activity of the DVC1-p97 segregase complex, and inactivation of FANCD2/FNACI sumoylation compromises cell survival in response to replication stress (Gibbs-Seymour et al., 2015). This example highlights the interplay between sumoylation and ubiquitination in the regulation of DNA replication at active replication sites both during normal replication and in response to replication stress. In addition to undergoing mono-ubiquitinated, the p66 subunit of the mammalian DNA POL δ is also mono-sumoylated by SUMO3, and this modification likely regulates protein–protein interaction or impacts the polymerase function (Liu and Warbrick, 2006). Other proteins involved in the synthesis of DNA lagging strand, such as the flap endonuclease 1 protein (FEN1), also undergoes sumoylation. FEN1 sumoylation in human cells is mediated by SUMO3 and begins in S phase and peaks in G2/M (Guo et al., 2012). FEN1 sumoylation promotes its ubiquitination and degradation via the PRP19 E3 ligase, which interacts with sumoylated FEN1 at least in part through its SIM (sumo-interacting motif) motif (Guo et al., 2012). Interestingly mutation of Ser-187 in FEN1 to Ala abrogates the phosphorylation at this site and precludes FEN1 sumoylation resulting in cell cycle delay and polyploidy (Guo et al., 2012).

In addition to the various components of the replicative helicases and polymerases, other components of the replisomes, including topoisomerases, DNA primase, the clamp loader RFC complex, as well as the nucleosome remodelling factor FACT were also found to be sumoylated (Golebiowski et al., 2009; Elrouby and Coupland, 2010; Cremona et al., 2012; Ma et al., 2014; Tammsalu et al., 2014). Among these, the sumoylation of DNA topoisomerase (TOP1) is best understood. The PIAS1 SUMO ligase was recently shown to sumoylate TOP1, and this is essential for reducing R-loop-mediated stalling of replication forks (Li et al., 2015). Biochemically, TOP1 sumoylation inhibits its catalytic activity, thereby reducing the nicking of DNA at transcriptionally active sites (Li et al., 2015). TOP1 sumoylation also enhances its binding to active RNA polymerase II, resulting the recruitment of splicing factors to suppress R-loop formation (Li et al., 2015). The role of sumoylation and/or ubiquitination in the regulation of other replisome components as well as other complexes involved in replication progression, such as components of the SMC complex (e.g. cohesin, condensin), is less understood, although emerging evidence support an important role for sumoylation in cohesion establishment (Rudra and Skibbens, 2013). PCNA: A central hub for ubiquitination and SUMOylation signalling One of the best examples for the role of ubiquitination and sumoylation in the regulation of DNA replication progression involves PCNA (Fig. 14.5). The homotrimeric DNA polymerase sliding-clamp coordinates the activity of many proteins involved in DNA replication, DNA repair and other chromatinrelated transactions (Choe and Moldovan, 2017; Ulrich and Takahashi, 2013). Although PCNA can be ubiquitinated at multiple Lys residues (McIntyre and Woodgate, 2015), only the mono-ubiquitination of PCNA at a conserved Lys residue (Lys-164 in human PCNA) is well understood. This particular modification is carried out by the Rad6-Rad18 E2-E3 ubiquitin conjugating enzyme/ligase and is one of the best understood posttranslational modifications of this protein. Such modification impacts the affinity of PCNA for different DNA polymerases, and is essential for error-prone translesion DNA synthesis (TLS) through the recruitment

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of translesion Y-family DNA polymerases [e.g. polymerase eta (POL-η)] to replication factories to bypass replication-stalling DNA lesions (Yang et al., 2013). This recruitment is dependent on the ubiquitin-binding domain of TLS polymerases, which has a high-affinity to mono-ubiquitinated PCNA (Bienko et al., 2005; Plosky et al., 2006). The CRL4CDT2 and RNF8 ubiquitin ligases are two other E3 ligases that can substitute for Rad18 in promoting PCNA mono-ubiquitination (Zhang et al., 2008; Terai et al., 2010). Although this posttranslational modification is significantly stimulated in cells exposed to bulky DNA lesions, such as those induced by UV (e.g. cyclobutane pyrimidine dimers), mono-ubiquitinated PCNA is detectable in normal proliferating cells in the absence of DNA damage, perhaps to aid in the replication of difficult to replicate DNA sequences or to cope with replication stress (Leach and Michael, 2005; Frampton et al., 2006; Terai et al., 2010). In S. cerevisiae, the heterodimeric E2 ubiquitin conjugating enzyme, Ubc13-Mms2, which is recruited to chromatin by the RING-finger protein Rad5, can convert the mono-ubiquitinated Lys on PCNA to Lys-63-linked polyubiquitin chain to participate in gap-filling damage tolerance (Prakash, 1981; Hoege et al., 2002; Torres-Ramos et al., 2002; Branzei et al., 2004; Haracska et al., 2004) and in template switching, an error-free pathway of DNA that utilizes the newly replicated sister chromatid as a template for replication (Hoege et al., 2002; Branzei et al., 2008, 2011; Choi et al., 2010; Hedglin and Benkovic, 2015). In mammals, this biochemical activity is carried out by the SNF2 histone linker plant homeodomain RING helicase (SHPRH) or by the helicase-like transcription factor (HLTF), and this was shown to suppress PCNA-dependent TLS and mutagenesis (Motegi et al., 2008; Unk et al., 2008). Interestingly, RAD18 itself can be mono-ubiquitinated, and this PTM inhibits its ability to mono-ubiquitylate PCNA and, the same time, suppresses its interaction with SHPRH or HLTF (Lin et al., 2011; Moldovan and D’Andrea, 2011; Zeman et al., 2014). Template switching is further facilitated by USP7, which deubiquitinates and stabilizes both HLTF and RAD18 through enhancing the interaction between the non-ubiquitinated RAD18 and HLTF (Qing et al., 2011; Zeman et al., 2014). Under replicative stress (e.g. following treatment with the alkylating agent

methyl methanosulfonate (MMS)), USP7 also deubiquitinates and stabilizes both RAD18 and POL-η, and this promotes TLS (Qian et al., 2015; Zlatanou et al., 2016). Under these conditions, the E3 ubiquitin ligase TNF receptor associated factor (TRAF)-interacting protein (TRIP) also facilitates TLS by promoting the Lys-63-polyubiquitination of POL-η, which is required for its focus formation at damage sites (Wallace et al., 2014). Several activities restrain TLS activity to reduce or prevent the mutagenic load caused by the lowfidelity polymerases. USP7 likely plays a role in this regulatory step by removing the mono-ubiquitin moiety on PCNA (Kashiwaba et al., 2015). The isopeptidase USP1, however, plays a more prominent role in deubiquitinating PCNA and in turning off TLS (Huang et al., 2006; Andersen et al., 2008). TLS is also restrained under conditions of increased DNA damage by UV irradiation, and this is mediated by USP10, which also deubiquitinates PCNA (Park et al., 2014). USP10-dependent PCNA deubiquitination requires the activity of EFP, an ISG15 E3 ligase, which ISGylate monoubiquitinated PCNA, thereby recruiting USP10 to deubiquitylate PCNA (Park et al., 2014). Following the release of POL-η, PCNA is de-ISGylated by UBP43, and engages the replicative DNA polymerases to resume normal replication, and inactivation of this pathway increases mutagenesis. In yeast, however, increased PCNA mono-ubiquitination, for example through inactivating the PCNA deubiquitinase Ubp10, does not increase mutagenesis, suggesting the existence of other mechanisms to suppress TLS (Gallego-Sanchez et al., 2012). The TLS polymerases themselves are subject to proteolytic degradation, and in the case of POL-η, this is mediated by MDM2 ( Jung et al., 2012). POL-η can also be mono-ubiquitinated by the PIRH2 E3 ligase, and this suppresses its interaction with mono-ubiquitinated PCNA ( Jung et al., 2010, 2011). Inactivation of TLS polymerases through ubiquitination is also conserved in yeast. For example, the S. cerevisiae homologue of POL-η, Rad30, as well as the Rev1 polymerase undergo proteolytic degradation, and for Rad30, this is mediated via the SCFUfo1 ubiquitin ligase (Waters and Walker, 2006; Skoneczna et al., 2007; Plachta et al., 2015). Sumoylation also plays important roles for regulating PCNA function. In fact, PCNA is sumoylated at the same Lys residue that is

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subject to mono-ubiquitination, suggesting that both sumoylation and ubiquitination of the same residue on PCNA is tightly regulated for optimal activity of this important protein. In yeast, PCNA sumoylation on Lys-164 (and to a lesser extent on Lys-127) is cell cycle regulated, preceding the entry of cells into S-phase, and is robustly induced by severe or lethal DNA damage (Hoege et al., 2002; Branzei, 2011; Hedglin and Benkovic, 2015). PCNA sumoylation, which is catalysed by the Ubc9 SUMO-conjugating enzyme, appears to interfere with PCNA-polymerase binding and with DNA repair, and is likely to be important for unloading PCNA during normal replication (Hoege et al., 2002; Branzei, 2011; Hedglin and Benkovic, 2015). Inactivation of UBC9 function in human cell lines prolongs S-phase, but it is unclear whether this is due to suppression of PCNA sumoylation (Schimmel et al., 2014). PCNA sumoylation is also important for the recruitment of the Srs2 helicase and anti-recombinase to suppress spontaneous and DNA damage-induced homologous recombination during S phase (Papouli et al., 2005; Pfander et al., 2005; Armstrong et al., 2012; García-Rodríguez et al., 2016; Zilio et al., 2017). Inactivation of PCNA

sumoylation was also shown to suppress post-replication repair associated with template switching (Branzei et al., 2008), and this likely due to interference between SUMO–PCNA interaction with Srs2 and/or with Rad18, Rad5 and ELg1 (an alternative subunit of the RFC clamp loader) (Pfander et al., 2005; Parnas et al., 2010). Sumoylation of mammalian PCNA is less abundant (Gali et al., 2012), reflecting the lower recombination activity in mammals. Regulation of replication termination by ubiquitin and SUMO How eukaryotic DNA replication is terminated is not entirely clear, but emerging evidence support important roles for ubiquitination in this process (Fig. 14.6). Studies in budding yeast and in Xenopus egg extracts show that the disassembly of the CMG complex is dependent on Lys-48-linked polyubiquitination of the MCM7 subunit of the MCM2–7 helicase (Maric et al., 2014; Moreno et al., 2014). Ubiquitinated MCM7 is recognized by the hexameric AAA+ adenosine triphosphatase (ATPase) and the segregase Cdc48/p97, which

Figure 14.6 Ubiquitin-dependent regulation of replication termination in eukaryotes. A model depicting the termination of eukaryotic DNA replication at converging replication forks, and its regulation by the ubiquitination of the Mcm7 Subunit of the Mcm2–7 helicase complex leading to the disassembly of the CMG complex (Mcm2– 7-GING-Cdc45). Mcm7 ubiquitination is promoted by the SCFDia2 E3 ubiquitin ligase in S. cerevisiae and by the CRL2Lrr1 E3 ubiquitin ligase in metazoans and is extracted through the activity of the p97 chaperon.

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on ATP hydrolysis promotes protein unfolding (Barthelme et al., 2014; Maric et al., 2014; Moreno et al., 2014). This triggers MCM7 translocation through the Cdc48/p97 ring, with the consequent disassembly of the hexameric MCM2–7 complex and replication termination (Bell, 2014; Lengronne and Pasero, 2014). Mcm7 polyubiquitination in S. cerevisiae is mediated by the SCF E3 ligase and the F-box protein Dia2, and inactivation of this pathway prevents CMG disassembly resulting in replication defects, although Mcm2 proteolysis is not required for Mcm2–7 disassembly and replication termination (Maric et al., 2014; Moreno et al., 2014; Morohashi et al., 2009). Polyubiquitination of the MCM7 in C. elegans and in Xenopus, is carried out by the replisome associated ubiquitin ligase CRL2Lrr1, which is similarly required for replication termination (Dewar et al., 2017; Sonneville et al., 2017). A role for protein sumoylation is replication termination is also beginning to emerge. In S. cerevisiae for example, the termination of DNA replication is associated with a specific reduction in Mcm7 sumoylation, which unlike the sumoylation of the other Mcm2–6 subunits, is concordant with the completion of DNA replication concurrent with increases in polyubiquitination of this subunit. (Wei and Zhao, 2016). It remains to be determined if the sumoylation of Mcm7 interferes with or is coordinated with the polyubiquitination of this subunit and with replication termination. As mentioned above, the Top2 DNA topoisomerase in budding yeast has been implicated in promoting replication across TERs, and this is important for the merging of the converging replication forks at these replication termination sites (Fachinetti et al., 2010). Top2 is also important for the decatenation of sister chromatids (Lee and Bachant, 2009). Interestingly, a subset of Topo II in various eukaryotes, including human TOPO II, is found to be sumoylated. In mitosis, the sumoylation of metazoan Topo II is essential for its recruitment to kinetochores, and interference with this sumoylation results in elevated frequency of segregation errors and aneuploidy (Lee and Bachant, 2009). A similar function for Topo II sumoylation in promoting replication termination is expected, but a concrete evidence for this prediction is yet to emerge.

Concluding remarks Significant advances in our understanding of the molecular and biochemical activities that function to control DNA replication have been made in the last few decades. The identification and characterization of the various PTMs of the many proteins that are associated with almost every step of DNA replication enriched our appreciation of the complexity underlying this highly conserved and important biological activity. In particular, the covalent attachment of ubiquitin and/or the ubiquitin-related protein SUMO on specific Lys residues on replication and replication-related proteins to form monomers and polymers of ubiquitin or SUMO chains ensures the timely and efficient temporal and spatial control of replication both during normal proliferation and in response to various perturbations. Modification of replication proteins by ubiquitin and SUMO involves both proteolytic and non-proteolytic functions that operate cooperatively through convoluted feedback mechanisms that, together with other PTMs, provide rich and complex networks of protein-protein communications to control both the fidelity and robustness of DNA replication. The execution of these modifications by a diverse and highly specific set of E2–E3 pairs of ubiquitin and SUMO conjugating enzymes and ligases, as well as their reversal by an equally diverse and specific set of ubiquitin- and SUMO-proteases, adds a readily apparent new layer of complexity that will require significant more research to fully understand and appreciate. While we know a great deal about the mechanisms involved in the ubiquitination and sumoylation of replication proteins and their impact on replication, proteome-wide studies indicate that many more replication and replication-related proteins are modified by these versatile moieties, both during normal replication and in response to cellular stresses, particularly those that cause replication stress. For these, the challenge is to understand the functional significance of these additional modifications and to identify the biochemical activities underlying their regulation. It is expected that new breakthroughs will come to be soon realized given the recent development of novel state-of-art biochemical protocols and assays (e.g. iPOND-MS and proximity labelling

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assays), gene-specific editing tools (e.g. CRISPR/ Cas and TALENs) as well as new genetic screening and functional assays. Lessons from past research, as outlined in this chapter, indicate that few family members of the ubiquitin and SUMO conjugating and deconjugating enzymes, such as the SCF, APC/C and CRL4 ubiquitin ligases, the USP7 deubiquitinase, the UBC9 SUMO conjugating enzyme as well as the SUMO ligase PIAS1, play key role in the regulation of the various aspects of eukaryotic DNA replication. These will likely to dominate the scene in future research in this area. Acknowledgement We apologize for authors whose primary work was not cited due to space limitations. Work in the author’s laboratory is supported by NIH grant R00 CA140774. References

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Roles of Ubiquitination and SUMOylation in DNA Damage Response

15

Siyuan Su1,2, Yanqiong Zhang1,2 and Pengda Liu1,2*

1Lineberger Comprehensive Cancer Center, The University of North Carolina at Chapel Hill,

Chapel Hill, NC, USA.

2Department of Biochemistry and Biophysics, The University of North Carolina at Chapel Hill,

Chapel Hill, NC, USA.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.15

Abstract Ubiquitin and ubiquitin-like modifiers, such as SUMO, exert distinct physiological functions by conjugating to protein substrates. Ubiquitination or SUMOylation of protein substrates determine the fate of modified proteins, including proteasomal degradation, cellular re-localization, alternations in binding partners and serving as a protein-binding platform, in a ubiquitin or SUMO linkage-dependent manner. DNA damage occurs constantly in living organisms but is also repaired by distinct tightly controlled mechanisms including homologous recombination, non-homologous end joining, inter-strand crosslink repair, nucleotide excision repair and base excision repair. On sensing damaged DNA, a ubiquitination/SUMOylation landscape is established to recruit DNA damage repair factors. Meanwhile, misloaded and mission-completed repair factors will be turned over by ubiquitin or SUMO modifications as well. These ubiquitination and SUMOylation events are tightly controlled by both E3 ubiquitin/SUMO ligases and deubiquitinases/deSUMOylases. In this chapter, we will summarize identified ubiquitin and SUMO-related modifications and their function in distinct DNA damage repair pathways, and provide evidence for responsible E3 ligases, deubiquitinases, SUMOylases and deSUMOylases in these processes. Given

that genome instability leads to human disorders including cancer, understanding detailed molecular mechanisms for ubiquitin and SUMO-related regulations in DNA damage response may provide novel insights into therapeutic modalities to treat human diseases associated with deregulated DNA damage response. Introduction DNA encodes for inheritable genetic information that is not only essential to exert normal cellular function but also indispensable to maintain the human society. Thus, DNA should be stable while versatile. Although certain genetic changes are permissible to drive evolution (usually at a low mutation rate), improper damaged DNA need to be repaired timely. With the development of technology, human beings are exposed to more DNA damaging cues nowadays such as wireless internet (Wi-Fi) (Akdag et al., 2016), ultraviolet (UV) radiation from sun exposure (Sinha and Häder, 2002) and even microwave ovens (Sagripanti et al., 1987) used on a daily basis. If the damaged DNA is detected and repaired to a level tolerated by cells, cells will survive and may develop neoplastic transformation; otherwise cells will die and be cleared. Damaged DNA is actively monitored by DNA

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damage sensors and repaired by DNA damage repair factors. Notably, in most prokaryotes such as bacteria, a SOS response is commonly triggered by DNA damage to repair damaged DNA and also contributes to anti-antibiotic features (Kreuzer, 2013). In this chapter, we will focus on DDR (DNA damage response) in eukaryotes given its close relationship to human physiology and pathology (Ciccia and Elledge, 2010). In response to genotoxic challenges, eukaryotes activate DNA damage checkpoints to suppress DNA replication, arrest cell cycle, stop proliferation and meanwhile activate signal transduction pathways to directly repair damaged DNA, or promote transcription of repair enzymes. Mechanisms sensing and repairing damaged DNA are conserved in eukaryotes. Factors inducing DNA damage can be divided into two categories: intrinsic factors and exogenous factors. The most frequent sources of intrinsic DNA damage are from inaccurate DNA replication, free radicals generated in vivo under oxidative stress or from normal biological processes including meiotic recombination and V(D)J recombination during antibody production (Hartlerode and Scully, 2009). Strong environmental cues including UV radiation, X-ray, gamma-ray and other chemical mutagens also cause various types of DNA damage, including DSBs (doublestrand breaks), SSBs (single-strand breaks), DNA base mutation, deletion, insertion, deamination, chemical modifications and formation of pyrimidine dimmers. Accordingly, distinct DNA damage responses are triggered. For example, UV-induced DNA crosslinking is resolved by NER (nucleotide excision repair) (Marteijn et al., 2014), unmatched, modified and damaged DNA bases are removed and refilled by the mismatch repair mechanism (Li, 2008), SSBs and DSBs are repaired by either HR (homologous recombination) (Li and Heyer, 2008) or NHEJ (non-homologous end joining) (Chang et al., 2017). Similar to prokaryotes, eukaryotes also utilize a SOS response in coordinating different repair pathway choices in responding to severe DNA damages (Fu et al., 2008). The eukaryotic DNA damage repair systems include DSB repair, inter-strand crosslink repair (ICLR), nucleotide excision repair (NER) and base excision repair (BER) (Hoeijmakers, 2001). Cells would need acute responses to repair damaged DNA-otherwise severe unrepaired DNA

damage leads to cell death (Nowsheen and Yang, 2012). The fastest reaction in cell is through biochemical reactions-indeed protein translational modifications have been observed and proven to play indispensable roles in this regard. For example, ATM, ATR or DNAPK controls phosphorylation of a large group of ‘SQ/TQ’ containing substrates including Chk1 and Chk2 (Chen and Poon, 2008), while as protein kinases themselves, Chk1 and Chk2 will further amplify the DNA damage signals by phosphorylating more substrates such as Cdc25A, p53, PML, Plk3 and many others (Bartek and Lukas, 2003). It is a kinase network or landscape that transduces the DNA damage signals in an acute and spatial-tempo dependent manner (Chen and Poon, 2008). In addition to extensively studied and well-characterized protein kinase cascades in DDR, ubiquitination and its close cousin, SUMOylation are other types of protein modifications that exert indispensable roles in both sensing and repairing damaged DNA (Brinkmann et al., 2015; Wang, Z. et al., 2017). In this chapter, we will summarize recent progress on ubiquitin and SUMO-related regulations on DDR, list all identified ubiquitination and SUMOylation events during DDR, further illustrate their physiological and pathological function and provide new insights into future research directions or therapeutic modalities targeting these identified ubiquitination or SUMOylation events. Overview of the ubiquitin signalling Ubiquitin is a 76 amino-acid protein highly conserved among eukaryotic species. Usually ubiquitin is considered as a modifier for proteins-attachment of ubiquitin moiety to a lysine residue on target proteins regulates important cellular processes including cellular trafficking, immune sensing, protein translation, metabolism, cell cycle and autophagy (Finley, 2009). Protein ubiquitination is a three-step enzymatic reaction requiring three types of enzymes, including E1 ubiquitin-activating enzyme, E2 ubiquitin-conjugating enzyme and E3 ubiquitin ligase. In mammals, there are one major E1, forty E2s and more than 600 E3s. E3 ubiquitin ligases are mainly divided into three families based on their structures and mechanisms of ubiquitin transfer, including RING (Really Interesting New Gene), HECT (Homologous to E6-AP Carboxyl

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Terminus) and RBR (RING-Between-RING) domain containing E3 ubiquitin ligase families (Zheng and Shabek, 2017). For RING and RBR families of E3 ligases, activated ubiquitin by E1 will be conjugated to E2, and it is the E2 enzyme directly transferring ubiquitin to substrates that are determined by E3 ligases. While for HECT domain containing E3 ligases, ubiquitin will be transiently transferred from E2 to E3 then transferred to substrates. In this process, E3 ubiquitin ligases determine the substrate specificity. Notably, each ubiquitin contains seven lysine residues. Addition of a ubiquitin to a prior ubiquitin molecule can be linked through each of seven lysine residues in ubiquitin, or through a head-totoe ligation, leading to formation of poly-ubiquitin chains in different linkages. According to the position of linked lysine residue, poly-ubiquitin chains can be linked through M1 (head-to-toe), K6, K11, K27, K29, K33, K48 and K63 linkages. The exact structures for poly-ubiquitin chains in a variety of linkages remain unclear, while some conformation for di-ubiquitin chains or shorter chains have been determined. K48-(Zhang, N. et al., 2009) or K11linked (Bremm et al., 2010) poly-ubiquitin chains adopt compact structures (Saeki, 2017) that fit well to the 26S proteasome recently determined by Cyro-EM (Dong et al., 2018). Thus, these two linkages are poised for protein degradation-an energy dependent process to destroy and recycle unwanted proteins. M1 and K63 (Weeks et al., 2009) linkages are in more labile structure with a great degree of flexibility (Kulathu and Komander, 2012; Sekiyama et al., 2012). These two types of ubiquitin chains usually serve as a binding platform for protein factors in various physiological conditions such as innate immunity (Xia et al., 2009) and cellular trafficking (Erpapazoglou et al., 2014). Recently, we found a protein modification independent function of K63-linked poly-ubiquitin chains in directly binding exposed naked DNA to facilitate DNA damage repair (Liu et al., 2018). K29 (Kristariyanto et al., 2015) and K33 chains adopt a zigzaging conformation (Michel et al., 2015). Notably, multiple linkages of poly-ubiquitin chains have been indicated to play critical roles in DDR, including K63, K6, K27 and others (Elia et al., 2015a). In addition, a given poly-ubiquitin chain can be either composed of one linkage (homotypic) or several different linkages to form chains with mixed

linkages or branched chains (heterotypic) (Meyer and Rape, 2014; Ohtake and Tsuchiya, 2017). Moreover, more than one poly-ubiquitin chain can be covalently attached to the same ubiquitin molecule on different lysine residues (Suryadinata et al., 2014). To make it more complicated, the ubiquitin molecule itself also undergoes various post-translational modification (PTM) events, including phosphorylation (Koyano et al., 2014) and acetylation (Ohtake et al., 2015), adding another layer of regulation on poly-ubiquitin chains. These distinct linkage composition and ubiquitin modifications on substrates create unique languages coding for distinct biological meanings, which have been referred to as ‘ubiquitin codes’ (Komander and Rape, 2012; Yau and Rape, 2016). Ubiquitination is a reversible protein modification and a result of a balance between adding and removing ubiquitin moieties. Various deubiquitinating hydrolases or deubiquitinases (DUB) have been identified as key enzymes for the removal of ubiquitin polypeptides from target proteins (Komander et al., 2009). To deal with the complicated ubiquitin system, mammalian cells develop DUBs that can be in large divided into seven families, including five families of cysteine proteases and one family of Zn-dependent metalloprotease (Komander et al., 2009). Specifically, cysteine proteases include USPs (ubiquitin specific proteases), OTUs (ovarian tumour proteases), UCHs (ubiquitin carboxyl-terminal hydrolases), Joshphin family of proteases and MINDYs (motif interacting with ubiquitin containing novel DUB family)(Abdul Rehman et al., 2016). The family of Zn-dependent metalloprotease consists JAMMs ( JAB1/MPN/ MOV34 metalloproteases), also termed as MPN+ family of DUBs (Clague et al., 2013). The role of DUBs in DDR is just began to be appreciated (Kee and Huang, 2016) and there is limited knowledge about whether and how these DUBs recognize ubiquitinated proteins in a linkage-specific manner, but the general impression is that compared with E3 ubiquitin ligases, DUBs are lacking certain substrate specificity-which means that a small number of DUBs may govern deubiquitination of a large spectrum of ubiquitinated substrates. Usually, USP DUBs directly bind substrates owing to the presence of protein interacting motifs (Faesen et al., 2011; Ye et al., 2009), while OTU DUBs exert certain ubiquitin linkage specificity, such as targeting

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the M1-linkage in LUBAC signalling (Keusekotten et al., 2013) and NF-kB signalling (Rivkin et al., 2013), or K63-linkage in mTOR signalling (Wang, B. et al., 2017) and non-canonical NF-kB signalling (Hu et al., 2013). How DUBs control DDR has been understudied. Thus, the ‘ubiquitin codes’ are produced by ‘ubiquitin writers’, most of the time are E3 ubiquitin ligases (Natarajan and Takeda, 2017) and removed by ‘ubiquitin erasers’ that are DUBs. Accordingly, different ‘ubiquitin code’ can be read and interpreted by various ‘ubiquitin readers’ that carry out distinct biological functions (Pinder et al., 2013). Making sure damaged DNA is repaired correctly and timely is key to maintain genome integrity, otherwise unrepaired DNA lesions may cause cells to die or accumulated DNA alternations may induce tumorigenesis (O’Connor, 2015). Thus, critical steps of DDR including the sensing of DNA damage, the recruitment of DNA damage repair factors and the repair of DNA lesions, are tightly controlled. Besides DNA repair, equally important is other cellular responses to DNA damage, such as cell cycle arrest and apoptotic cell death, if the DNA damage is very severe and unrepairable. Because of its importance to cell survival and function and its dynamic response to environment cues, DDR is tightly regulated by protein post-translational modifications. One indispensable mechanism to ensure accurate and efficient DDR is to utilize the ubiquitin signalling. Indeed, upon DNA damage, a ubiquitin landscape is quickly established to label the damage foci, recruit repair factors and regulate the entire repair process by multiple E3 ubiquitin ligases including RNF8, RNF168, BRCA1, BMI1, Ring1B, Rad18 and others (Messick and Greenberg, 2009). The efforts to investigate contribution of ubiquitin linkages start early. Initially, ectopically expressed K6 and K63-linked, but not K48-linked ubiquitin was enriched at sites of DNA damage (Sobhian et al., 2007). Following studies demonstrated that K48-linked ubiquitin chains also play a critical role in removing Ku80/Ku70 complexes to facilitate the progression of NHEJ. More recent non-biased large-scale studies examining endogenous ubiquitin linkages observed a dramatic accumulated K6- and K33-linked ubiquitin chains with DDR (Elia et al., 2015a). Thus, a variety of distinct linkages of ubiquitin chains may play important roles in guiding proper sensing and repair of damaged DNA

under different pathophysiological conditions. Given that sensing and repair of damaged DNA are complicated processes and there is no clear boundary between these two consecutive events, in the following section, we will summarize distinct ubiquitin events and their roles in DDR in a DNA damage repair mechanism dependent manner. Overview of the SUMO signalling In addition to ubiquitin, there are many ubiquitin like (UBL) molecules with similar sequence/structure composition but distinct function (Hu and Hochstrasser, 2016). Small Ubiquitin-like MOdifier (SUMO) is a highly conserved (approximately 12 kDa) protein produced as an immature precursor that needs to be cleaved by sentrin/SUMO-specific protease 1 (SENP1) prior to conjugation. SUMO has similar conjugation pathways as ubiquitin, but the process is carried out by SUMO-specific enzymes. First, E1-activating enzyme (the heterodimer SAE1/SAE2) charges C-terminal di-glycine residues of mature SUMO in an ATP-dependent manner. Then the activated SUMO is transferred to the E2 conjugating enzyme UBC9 via a thioester transfer step. Next, UBC9 directly conjugates the SUMO molecule to the lysine residues of substrate proteins through an isopeptide linkage, or with the assistance of SUMO E3 ligases. The E3-ligating enzymes improve conjugation by either recognizing target lysines or enhancing SUMO discharge from the E2 to the substrate. The most well characterized SUMO E3s are Protein Inhibitor of Activated STAT (PIAS 1–4) (Rytinki et al., 2009), with an SP-RING domain similar to the RING motif in many E3 ubiquitin ligases. The SUMO conjugation system has relatively fewer enzymatic components than the ubiquitin system. Compared with approximately 40 different E2-conjugating enzymes for ubiquitin, only one E2 (Ubc9) in SUMO system has been identified so far. Moreover, only a handful of SUMO E3 enzymes have been identified compared to around 600 E3 ubiquitin ligases. Notably, plants and metazoan have more enzyme isoforms of SUMO E3s compared with lower eukaryotes like in yeast. However, unlike only one unified ubiquitin molecule, there are more than one SUMO isoform in the SUMO system: SUMO1, SUMO2/3 and the

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recently described ones including SUMO4 (Baczyk et al., 2017) and SUMO5 (Liang et al., 2016). SUMO proteins have high similarities to the tertiary folding structure of ubiquitin while they share limited sequence identity (less than 20%) and have different surface charge distributions (Huang et al., 2004). SUMO1 was first identified as a human ubiquitin-like protein that interacts with RAD51/ RAD52 proteins (Shen et al., 1996), Promyelocytic leukaemia (PML) components (Boddy et al., 1996), and conjugates GTPase RanGAP1 to recruit it to nuclear pore complex protein RanBP2 (Matunis et al., 1996; Mahajan et al., 1997). SUMO2 and SUMO3 are nearly identical in sequence (97% identity, referred to as SUMO2/3) but distinct from SUMO1 (50% identity). SUMO4 is reported as a new IĸBα modifier (Guo et al., 2004) but another study showed SUMO4 cannot be processed to a mature form due to its unique proline-90 residue (Owerbach et al., 2005). Recently, SUMO5, previously reported as a pseudogene (Su and Li, 2002), could form novel poly-SUMO isoforms that regulate PML nuclear bodies (Liang et al., 2016). In addition, SUMOylation occurs most frequently (≈ 75%) at a lysine residue within a consensus sequence ‘ψKxE/D’ (where ψ represents a hydrophobic amino acid and x any amino acid) (Bernier-Villamor et al., 2002; Hendriks et al., 2017; Lamoliatte et al., 2017) but ubiquitination has little preference for lysine context. SUMOylation of different forms of SUMO modifiers can occur on the same or different substrates. Some proteins are preferentially modified by one type of SUMO isoform while others could be modified by different SUMO isoforms. SUMOylation can also be in the form of chains as polySUMO as in the ubiquitin system, and the chains are only generated on SUMO2/3 but not SUMO1(Sarge and Park-Sarge, 2009). SUMOylation is also a reversible process, similar to deubiquitination, but in which deSUMOylation involves the removal of SUMO terminal glycine from the lysine residues of the substrate protein by specific proteases (Nayak and Müller, 2014). Unlike the array of proteases in the ubiquitin system, the SUMO protease family has just been found to be limited. SUMO proteases can be divided into three classes, including (1) thiol proteases, (2) cysteine proteases and (3) a mammalian specific SUMO-specific protease USPL1(Nayak and Müller, 2014). SUMO thiol proteases include

six sentrin (SUMO)-specific proteases termed as SENPs including SENP-1, -2, -3, -5, -6 and -7 in mammals (Hickey et al., 2012). Notably, although SENP-8 was originally identified as a deSUMOylase, later it was proven that the true substrate for SENP-8 is another ubiquitin-like molecule Nedd8 (Gan-Erdene et al., 2003; Mendoza et al., 2003). SENP1–3 and SENP5 are related to the yeast deSUMOylase Ulp1, and SENP6 and 7 are close to yeast deSUMOylase Ulp2. These SENPs differ in SUMO maturation (C-terminal hydrolase) and isopeptide cleavage activity. Additionally, different SENPs have their preferences for different SUMO modifier isoforms. For example, both SENP1 and SENP2 can process SUMO1and SUMO2/3, while SENP3 and SENP5 are mainly involved in SUMO2/3 deSUMOylation. PolySUMO chains of SUMO2/3 are dissociated by SENP6 and SENP7 (Hickey et al., 2012). Notably, the SUMO cysteine proteases include Desi-1 and Desi-2 are only present in plants and metazoan (Nayak and Müller, 2014). Interestingly, SUMO conjugation can be achieved in both SUMO E3 ligase dependent and independent manners (Nayak and Müller, 2014). Similar to poly-ubiquitination, poly-SUMOylation chains can also serve as a binding platform for protein factors and to date there are some SUMOylation binding domains characterized, including a hydrophobic core sequence ([V/I]-x[V/I]-[V/I]) (Heerwagen et al., 1995) surrounded by negatively charged residues, or a protein motif composed of [I/V/L]-[D/E]-[I/V/L]-[D/E][I/V/L] (Ouyang et al., 2009) (Table 15.1). While on the other hand, there are more than 16 wellcharacterized ubiquitin binding domains (Grabbe and Dikic, 2009). Given that thousands of proteins have been identified to be modulated by this modification, it is not surprising that SUMOylation plays a broad spectrum of cellular functions in development, growth, metabolism, and DNA damage response (Nayak and Müller, 2014). Ubiquitin and SUMO signalling in HR DNA double-strand breaks are the most severe type of DNA damage, whose repair is governed by two major pathways: Homologous Recombination (HR) and Non-Homologous End-joining (NHEJ)(Lieber, 2010). The HR pathway requires

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Table 15.1  Comparison of Ub and SUMO conjugation system Components in conjugation system

Ubiquitination

SUMOylation

Conjugates

Ubiquitin

SUMO1, SUMO2/3 SUMO4, SUMO5

E1 activating enzymes

UBE1 (UBA1)

SAE1-SAE2 (UBA2)

E2 conjugating enzymes

~ 40 conjugation enzymes

UBE2I (UBC9)

E3 ligases

~ 600 E3 ligases

PIASs, RanBP2, Siz1#, CBX4

Conjugate removing enzymes

~ 100 DUBs

SENP1–3 and 5–7, DeSi1/2, USPL1

Conjugation sites

Little preference for lysine context

Frequent consensus sequence ‘ψKxE/D’

the presence of a homologous DNA sequence as the repair template; thus, it is mainly functional in S and G2 phases (Longhese et al., 2010). The NHEJ pathway, by its name, is an error-prone DNA damage repair pathway because it directly glues two broken DNA ends without caring about whether the repair products faithfully resemble their original DNA sequence. While on the other hand, as NHEJ does not require the presence of a nearby template chromatin to repair DNA lesions, it is more versatile for acute repair and to promote DNA evolution. Notably, NHEJ occurs during the entire cell cycle. Once DSBs occur, these damaged free DNA ends can be recognized by either the Mre11/ Rad50/Nbs1 (MRN) complex or the Ku70/Ku80 complex, leading to HR or NHEJ, respectively. The determining step for HR repair is DNA end resection, where broken double-strand DNA (dsDNA) will be resected into a long ssDNA (single-strand DNA) that intrudes into dsDNA to search for homologous sequence. DNA end resection is carried out by the MRN complex. MRN searches for free DNA ends-Rad50 binds dsDNA to allow perfusion of MRN complexes along DNA for this search and Mre11 carries out a nucleolytic reaction to exert two functions: (1) recruit Exo1 (Exonuclease 1) to initiate resection and (2) remove Ku70/ Ku80 from binding broken DNA ends to promote HR and suppress NHEJ. This process was recently confirmed by single molecule imaging (Myler et al., 2017). This process can be antagonized by BRCA1 binding to DNA (Paull et al., 2001), resulting in inhibition of the nucleolytic activities of MRN and suppression of HR. In addition, Exo1 protein stability is governed by the E3 ubiquitin ligase Cyclin F (Elia et al., 2015a). Upon MRN loading onto DNA, Nbs1 is poly-ubiquitinated by the E3 ubiquitin

ligase Skp2 in a K63-linkage dependent manner to recruit the kinase ATM to sites of damage, where ATM phosphorylates histone H2 at Ser139, forming γ-H2Ax foci. Notably, γ-H2Ax foci serve as red flags to earmark DNA damage sites (Fig. 15.1 and Table 15.2). The DNA damage signal can be further amplified by a way that MDC1 (Mediator of DNA damage checkpoint protein 1) binds and protects γ-H2Ax, bringing in another MRN complex through binding Nbs1 (Stewart et al., 2003; Lukas et al., 2004) and a second ATM kinase [Nbs1 binds ATM (Falck et al., 2005)] to phosphorylate MDC1 that is necessary to recruit a critical E3 ubiquitin ligase RNF8. MDC1 undergoes K48-linked ubiquitination as a protein turnover control with unknown E3 ligases (Shi et al., 2008), a process blocked by USP7 (Su et al., 2018). In addition to ubiquitination, MDC1 is also SUMOylated by PIAS4 to promote MDC1 protein turnover (Luo et al., 2012). RNF8 is a key Ring-finger E3 establishing and orchestrating a ubiquitin landscape on histones at sites of DNA damage by ubiquitinating H2A or H2Ax in a K63-linkage specific manner with the help of the E2 enzyme UBC13 (Kolas et al., 2007; Mailand et al., 2007). In addition, RNF8 also promotes ubiquitination of Nbs1 to facilitate the MRN complex formation and HR (Lu et al., 2012). The critical role of RNF8 in positively regulating DDR is evidenced by the observation that RNF8 deletion leads to cellular sensitivity to IR and arrested G2/M transition (Huen et al., 2007; Kim et al., 2007; Kolas et al., 2007; Mailand et al., 2007). Notably, the role of RNF8 in DDR is antagonized by DUBs such as USP11 (Yu, M. et al., 2016) and BRCC36 through specifically removing K63-linked ubiquitin chains RNF8/UBC13 produce. In addition to establishing

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Figure 15.1 Ubiquitin and SOMO modifications in HR.

K63- and K48-linked ubiquitin chains, RNF8 also produces K11-linked ubiquitin chains on unknown substrates to inhibit transcription, and this function of RNF8 is antagonized by the DUB cezanne (Paul and Wang, 2017). Once the initial ubiquitin signal is established by RNF8/UBC13, another E3 ubiquitin ligase RNF168 recognizes ubiquitinated/ SUMOylated H2A and further ubiquitinates H2A at K13-K15 residues to amplify the ubiquitin signalling (Doil et al., 2009; Panier et al., 2012). While other reports support the notion that although RNF168 functions depending on RNF8, RNF8 and RNF168 ubiquitinate non-histone proteins and histones, respectively, to establish the ubiquitination landscape on DNA damage (Mattiroli et al., 2012; Panier et al., 2012). RNF168 itself could be poly-ubiquitinated by the HECT type E3 ligases TRIP12 and UBR5, restricting the spreading of ubiquitinated γ-H2Ax and preventing genome-wide transcriptional suppression, which could be potentially detrimental to cells (Gudjonsson et al., 2012). In addition, RNF168 binds and ubiquitinates PML to trigger subsequent SUMO2 modification of PML that facilitates formation of PML nuclear bodies (Shire et al., 2016). While a viral E3 ubiquitin ligase ICP0 targets both RNF8 and RNF168 to negatively regulate their function (Lilley et al., 2010). In addition to RNF168, RNF8mediated ubiquitin signalling also recruits other E3 ubiquitin ligases including Rad18 (Huang et al., 2009) and HERC2 (Bekker-Jensen et al., 2010; Wu et al., 2010) to amplify ubiquitin signalling. Beyond RNF8, another E3 ubiquitin ligase CHFR also triggers the first wave of ubiquitination events at DSBs by at least ubiquitinating PARP1 (poly-ADP-ribose polymerase I) that may regulate ubiquitination and poly-ADP-ribosylation (Fig. 15.1 and Table 15.2). Nonetheless, established K63-linked ubiquitin chains by multiple E3 ubiquitin ligases mentioned above serve as a binding platform to recruit proper DNA damage repair factors, such as Rap80/ BRCA1 and 53BP1, which determines repair by HR or NHEJ. The UIM (ubiquitin interacting motif) in Rap80 binds K63-linked poly-ubiquitin chains and promotes the assembly of the Rap80/ ABRA1/BRCA1 complex (Kim et al., 2007; Sobhian et al., 2007; Wang, B. et al., 2007; Yan et al., 2007), which is essential for HR. On the other hand, RNF168 ubiquitinates 53BP1 through K63-linked poly-ubiquitination to promote 53BP1

Table 15.2  Summary of modified DDR members by ubiquitin and SUMO Repair pathway

Substrates

E3 ligase/linkage/function

DUB/linkage/function

DDR

H1

RNF8

DDR

H2A -K13/ K15

RNF1682,5,6, K63, recruits 53BP1, USP167, interacts with HERC2 RAP80, RAD18, RNF169 USP38–10, counteracts RNF168

SUMOylase/function

, K63, recruits RNF168

1–5

USP4411, counteracts RNF8/ RNF168-mediated histone ubiquitination BRCC3612, reverses H2A ubiquitination by RNF8/RNF168 POH1, negatively regulates 53BP1 accumulation DDR

H2A-K119/ K120

RING1B/BMI113–15, recruits DNA repair factors

DDR

H2A-K127/ K129

BRCA1-BARD116, K6, maintain chromatin in a transcriptionrepressive status

DDR

H2BK120

RNF20/RNF4017, monoubiquitination, promotes HR

DDR

BMI1

HR

Nbs1

CBX4, influences DDR Ubsignalling18 Skp219, K63, promotes ATM binding to Nbs1 and HR RNF820, facilitates MRN complex formation and HR

HR

MDC1

RNF421, K48, MDC1 degradation

HR

PARP1

CHFR23, K63, important for first wave of ubiquitination in HR

HR

RPA

RNF424, K48, promotes HR; RFWD325,26, mixed linkages, promotes HR PRP1927, unknown linkage, binds and ubiquitinates RPA-ssDNA to bring ATRIP to ATR activation

PIAS4, drives RNF4 interaction21,22

DeSUMOylase/function

RNF16828, triggers PML SUMO2 modification

HR

PML

HR

BRCA1

HR

BRCA2

HERC230, inhibits BRCA1 binding to BRCA2

HR

53BP1

RNF1683, K63, promotes 53BP1 recruitment to the site of DNA damage

HR

CtIP

promotes RNF13832, K63, CtIP accumulation and HR activation

PIAS1/4, increases BRCA1: BARD1 E3 ligase activity in vitro29

PIAS1 and PIAS4, promotes DSB repair31

BRCA1-BARD133, K63, maintain CtIP on chromatin HR

RNF168

TRIP12/UBR5, K48, removes RNF168 to prevent widespreading histone ubiquitination

HR

unknown

Rad18, interacts with Rad51c to promote HR 35

HR

911

Rad6-Rad1836,

HR

Exo1

SCF-CyclinF37, K48, degradation

HR

PALB2

KEAP139, blocks PALB2/ BRCA1 complex formation and suppresses HR

HR

Claspin

APC/Cdh140, K48, degradation β-TRCP40, K48, degradation

PIAS4, increases protein stability and promotes its transcription34

PIAS4, reduces its stability38

USP740, reverses β-TRCP mediated ubiquitination, stabilizes Claspin

HR

ERCC6

N/A

USP741

HR

Chk1

N/A

USP742, stabilizes Chk1

HR

Mdm2

N/A

USP743, stabilizes Mdm2

SENP6, promotes its hypoSUMOylation38

Table 15.2 Continued Repair pathway NHEJ

Substrates

E3 ligase/linkage/function

Ku80

RNF844,

DUB/linkage/function

SUMOylase/function

DeSUMOylase/function

required for RAD51 accumulation48

SENP6, promotes its hypoSUMOylation48

K48, degrades Ku80

RNF13845, K48, degrades Ku80 F-box proteins46, degrade Ku80 and promotes NHEJ RNF126, releases Ku70/80 for NHEJ to continue RNF144A47, K48, degrades DNAPK

NHEJ

DNAPK

NHEJ

RPA70/RPA1

NHEJ

XLF

β-TRCP49, K48, degrades phosphorylated XLF

NHEJ

XRCC4

Fbw750, K63, enhances the binding between XRCC4 and Ku70/80, promotes NHEJ repair

regulates localization51

MonoUb52, stablizes DNA ligase IV Template switching/ Translesion synthesis

PCNA

RAD1853, monoUb, facilitates TLS and stimulates the E3 activity of FANCL

USP154 USP755, suppresses induced PCNA monoUb

Rad5, K63, promotes template switching repair in yeast RNF856, K48, Plays a role in DNA Damage Tolerance (DDT)56

Template switching/ Translesion synthesis

KAP1

FA

FANCD2 and FANCL60,61, monoubiquitination, FANCI promotes BRCA1/2 pathway

auto-SUMO ligase57, DSBassociated transcriptional repression58

SENP7, promoting chromatin relaxation59

FA

FANCG

BRCC6, the inhibition of which K63Ub62, required for binding with Rap80–BRCA1 complex and improved HR increased HR efficiency

NER

DDB2

DDB163, K48, degrade DDB2

USP2464, degrades DDB2

NER

RNA polII

Rsp565, K63 or mixture of monoand poly-Ub, prerequisite step for degradation by Elong1-Cul3

Ubp265, trims K63 Ub chains on RNA PolII into mono-Ub for proofreading

Elong1-Cul365, K48, degrades RNA polII

Ubp366, reverses K48 Ub chains on RNA polII

NER

H2B

N/A

USP767, promotes base-excision repair

NER

XPC

UV-DDB268, enhances XPC binding with DNA

USP1169, increase XPC retention on the damaged DNA

RNF11170, triggers XPC release from damaged DNA sites, allow binding of other NER factors BER

MUYH, RNA Polβ

MULE71, K48, promotes degradation

BER

APE1

Mdm272 and UBR373, promotes degradation

BER

PNKP

Cul4A-DDB1-STRAP74, promotes degradation

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loading onto sites of damage (Bohgaki et al., 2013) and subsequent repair of damaged DNA through NHEJ. BRCA1 then facilitates Rad51 loading by complexing with BRCA2/PALB2 (Sy et al., 2009; Zhang, F. et al., 2009a,b) and Rad51 is indispensable to search for homologous DNA sequence for HRmediated DNA damage repair. Moreover, BRCA1 also promotes DNA end resection by recruiting the resection enzyme CtIP and excluding 53BP1 thus inhibiting NHEJ ( Jiang and Greenberg, 2015). Ubiquitination of PALB2 by the E3 ligase Keap1 has been observed to specifically block the BRCA complex formation, rather than targeting PALB2 for degradation, thus suppressing HR (Orthwein et al., 2015). The RING-type E3 ligase RNF138 has been shown to ubiquitinate CtIP, promoting its accumulation to the site of DNA damage, thereby activating HR repair. This ubiquitination occurs at a relatively early stage of DNA resection. On the other hand, CtIP could also be ubiquitinated by BRCA1BARD1 E3 ligase, which serves to maintain CtIP on the chromosome after DNA damage. Another DNA repair protein under regulation of ubiquitination is RPA, which binds naked ssDNA after DNA resection. Both RNF4 (Galanty et al., 2012) and RFWD3 (Elia et al., 2015b) bind and ubiquitinate RPA, promoting the removal of RPA from DNA damage sites and suppressing HR repair, while PRP19 (Maréchal et al., 2014) ubiquitinates RPA and brings along ATRIP, which in turn activates ATR kinase and promotes HR pathway. Moreover, RFWD3 ubiquitinates RPA to promote replication fork restart and increase HR efficiency at stalled replication forks during DNA replication (Elia et al., 2015b). RPA also undergoes SUMOylation by unknown SUMOylase(s), which promotes RPA binding to Rad51 (Dou et al., 2010) to facilitate HR. In addition, SUMOylation of ATRIP has also been observed to facilitate ATRIP interaction with ATR, while the identities of the SUMOylase(s) remains unknown (Wu et al., 2014) (Fig. 15.1 and Table 15.2). Intriguingly, BRCA1 itself functions as a E3 ligase by complexing with BARD1 and multiple substrates have been identified in DDR including but not limited to H2A, H2AX, RNA polII, TFIIE, NPM1, CtIP, tubulin, ER-α and claspin (Wu et al., 2008; Densham and Morris, 2017). BRCA1 undergoes SUMOylation by PIAS1/4 and SUMO

conjugation promotes BRCA1 E3 ligase activity in vitro (Morris et al., 2009). The E3 ligase HERC2 negatively regulates BRCA2 protein stability by attaching K48-linked ubiquitin chains and BARD1 binding to BRCA2 protects BRCA2 from HERC2dependent degradation (Wu et al., 2010). The APC/Cdh1 E3 ligase negatively regulates DDR by targeting Claspin for K48-linked ubiquitination and degradation (Bassermann et al., 2008; Gao et al., 2009; Oakes et al., 2014). In addition, Claspin is also targeted by anther E3 ligase β-TRCP for degradation, where USP7 specifically antagonizes β-TRCP but not Cdh1-mediated Claspin proteolysis (Faustrup et al., 2009). Notably, FANCG undergoes K63-linked ubiquitination to facilitate its association with BRCA1/Rap80 to promote HR for resolving DNA crosslinks, a process that is antagonized by the DUB named BRCC36(Zhu et al., 2015) (Fig. 15.1 and Table 15.2). In addition to well-established ATM/MDC1/ RNF8 signalling in response to DSBs, the BAL1/ BBAP E3 ligase complex has been observed to be able to sense and transduce DNA damage signals independent of the ATM/MDC1/RNF8 signalling that is associated with PARP1 activation and BRCA1 recruitment (Yan et al., 2013). Notably, deSUMOylation by SENP7 of KAP1 (KRAB-associated protein 1) relaxes chromatin structure to promote HR (Garvin et al., 2013), while SUMOylation of Tyrosyl-DNA phosphodiesterase 1 (TDP1) promotes TDP1 enrichment on damage sites although the identity of the SUMOylase(s) is elusive (Hudson et al., 2012) (Fig. 15.1 and Table 15.2). Ubiquitin and SUMO signalling in NHEJ The NHEJ repair pathway starts with binding of damaged DNA by the Ku70/80 heterodimers through the Ring-like structure, enabling the recruitment of DNA repair factors functioning in NHEJ, including DNAPK, XLF, PAXX, XRCC4, DNA ligase IV, Artemis and DNA polymerases μ and λ (Lieber, 2010). Initially, Ku70/Ku80 needs to be loaded efficiently to ensure timely repair of damaged DNA, but during NHEJ repair Ku70/ Ku80 rings need to be efficiently and timely removed. This is partially achieved by either

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RNF8 (Feng and Chen, 2012) or RNF138 (Ismail et al., 2015)-mediated K48-linked ubiquitination of Ku80 to remove Ku80/Ku70 complexes from DSBs to allow NHEJ to occur. On the other hand, the APC (Anaphase Promoting Complex) catalyses K48-linked ubiquitination of RNF8 to antagonize the negative regulation of Ku80 by RNF8, facilitating NHEJ (Ma et al., 2018). In addition to single subunit Ring figure E3 ligases including RNF8 and RNF168, a group of F-box E3 ligases including Fbxl12, β-TRCP, Fbh1, Fbxl19, Fbxo24, Fbxo28 and Kdm2b have been observed to target Ku80 for ubiquitination and degradation, therefore facilitating NHEJ (Postow and Funabiki, 2013). RNF126 ubiquitinates and degrades Ku80 to release Ku70/Ku80 from damaged DNA to complete NHEJ. Deficiency in RNF126 leads to extended NHEJ process (Ishida et al., 2017). In addition to proteasomal degradation, Ku80/ Ku70 can also be removed by VCP/p97-which is important for Ku70/Ku80 extraction from DSBs on K48-linked ubiquitination in a Ufd1/Npf4 dependent manner, therefore suppressing NHEJ and facilitating HR (van den Boom et al., 2016). Interestingly, Ku70 has been observed to display a DUB activity towards stabilizing the proapoptotic protein Bax, thus exerting roles in cell apoptosis in addition to DNA damage (Rathaus et al., 2009). In yeast, Yku70 is SUMOylated by yeast SUMOylases including Mms21 and Siz1/2, and SUMO conjugation promotes Yku70 association with DNA (Hang et al., 2014) (Fig. 15.2 and Table 15.2). The E3 ubiquitin ligase RNF144A targets cytosolic DNAPK for K48-linked ubiquitination and degradation to promote DNA damage-induced cellular apoptotic response (Ho et al., 2014). DNAPK recruits DNA damage repair factors to the site of lesions, including Artemis that trims the DNA ends with overhangs, and DNA ligase IV, which ligates blunt-ended DNA. In addition, DNAPK also phosphorylates and recruits XRCC4, PAXX, XLF to complex with DNA ligase IV to form a ligase complex with optimal activity for NHEJ. Notably, an important factor in this complex, XLF, undergoes Akt-mediated phosphorylation that triggers its association and degradation by the E3 ubiquitin ligase β-TRCP in a K48 linkage dependent manner (Gan et al., 2015; Liu et al., 2015) to suppress NHEJ.

Figure 15.2 Ubiquitin and SOMO modifications in NHEJ.

XRCC4 undergoes K63-linked ubiquitination by Fbw7 to facilitate its association with Ku complexes, thus enhancing NHEJ (Zhang et al., 2016). In addition, mono-ubiquitination of XRCC4 was also observed with unknown E3 ligase(s) to stabilize DNA ligase IV (Foster et al., 2006). Moreover, SUMOylation of XRCC4 by PIAS retains XRCC4 in cytoplasm, thus impairing NHEJ (Yurchenko et al., 2006) (Fig. 15.2 and Table 15.2). Whether and how DNA ligase IV is subjected to ubiquitinmediated regulation remains unknown.

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Ubiquitin and SUMO signalling in inter-strand crosslink repair (ICLR) Inter-strand crosslinks can be induced by exposure to alkylating agents, platinum and psoralens in environment and by clinical treatments and they are toxic to cells given that they strongly prevent transcription and replication due to the inability of the dsDNA for proper separation and is strongly associated with a human disorder called fanconi anaemia (FA) (Deans and West, 2011). Thus, many members repairing ICLs are named after this disorder and the repair pathway to resolve ICLR is also called FA pathway that mainly relies on HR but with distinct sets of nucleases and other DNA processing enzymes. Mechanistically, ICLs are recognized by FANCM that recruits subsequent associated proteins such as FANCL, FANCG and others to form a core FA complex, where FANCL exerts an E3 ligase activity to mono-ubiquitinate FANCI and FANCD2 (Longerich et al., 2009; Miles et al., 2015). These mono-ubiquitin events serve as a binding platform for Pol V and FAN1, respectively, that will activate ATR. On the other hand, the core FA complex also recruits BTR (the Bloom’s syndrome complex) and FANCJ to facilitate BRCA1-mediated HR repair. Although no poly-ubiquitination event has been reported on the FA pathway, mono-ubiquitination of FANCI and FANCD2 serves as a signalling antenna for FA repair progression. Rad18 has been observed to be critical for FANCI and FANCD2 monoubiquitination in its E3 ligase dependent manner (Williams et al., 2011), however, whether Rad18 directly ubiquitinates FANCI and FANCD2 warrants further investigation. In addition, biallelic mutations of the RFWD3 E3 ubiquitin ligase lead to FA, supporting its critical role in FA while with the exact substrate(s) for RFWD3 in FA remain unknown (Knies et al., 2017). Moreover, the E3 ubiquitin ligase Fbw7 targets the key FA pathway member FAAP20 for ubiquitination and degradation in a GSK3 phosphorylation dependent manner to clear FAAP20 on completion of FA repair (Wang et al., 2016). In addition to ubiquitination, FANCI also undergoes SUMOylation by PIAS1/4 and this modification promotes FANCI protein degradation to terminate FA signalling (Gibbs-Seymour et al., 2015) (Fig. 15.3 and Table 15.2).

Figure 15.3 Ubiquitin and SOMO modifications in FA.

Ubiquitin and SUMO signalling in nucleotide excision repair (NER) UV irradiation from sunlight or clinical applications trigger the formation of double thymidine dimmers, a type of DNA lesions that will be resolved by NER. NER can be divided into global genome nucleotide excision repair (GG-NER) (Yu, S. et al., 2016) and

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transcription coupled nucleotide excision repair (TC-NER) (Pani and Nudler, 2017). XPC is the sensor for both NERs by complexing with Rad23B and CETN2 to label damaged DNA to initiate NER. XPC undergoes UV-DDB2-mediated ubiquitination to enhance its binding to DNA (Sugasawa et al., 2005), as well as SUMOylation in a DDB2 and XPA-dependent manner to prevent XPC proteasome degradation (Wang et al., 2005; Wang, Q.E. et al., 2007). XPC is stabilized by SUMOylation via unknown SUMOylase(s) in this process (Wang, Q.E. et al., 2007). After NER initiation, RNF111 medaited ubiquitination of prior ubiquitinated or SUMOylated XPC facilitates the release of XPC from damage sites to allow binding of NER factors such as XPG and XPF (van Cuijk et al., 2015). In addition, USP11 deubiquitinates XPC to extend its retention on damaged DNA, thus enhancing NER (Shah et al., 2017). Consistent with this observation, reduced USP11 expression was observed in human skin cancer patients, highlighting its role as a tumour suppressor in promoting NER (Shah et al., 2017). In addition, USP24 deubiquitinates and stabilizes DDB2 that promotes XPC ubiquitination and NER. The Flap endonuclease 1 (FIN1) that exerts endonuclease activity in NER is SUMOylated by unknown SUMOylase(s) and this SUMOylation event promotes FIN1 degradation to suppress NER (Guo et al., 2012). In TC-NER, the RNA PolII/CSB (ATPase) complex is indispensable to fill in the DNA gaps and VCP/p97 promotes their proteolytic clearance (He et al., 2016, 2017), while USP7 together with UVSSA, deubiquitinates RNA PolII and CSB to stabilize these proteins (Higa et al., 2016), both of which are essential for TC-NER. In addition, SUMOylation of C-terminus of CSB by unknown SUMOylase(s) has been observed to facilitate CSB’s function in NER (Sin et al., 2016). Notably, NER also induces H2A ubiquitination in a manner depending on the MRN/MDC1/RNF8 signalling (Marteijn et al., 2009). XPF/ERCC1 is an essential downstream factor of both GG-NER and TC-NER serving as a damage repair nuclease complex. USP45 specifically deubiquitinates XRCC1 to promote its translocation to damage sites (Perez-Oliva et al., 2015) while the identity of E3 ligase(s) responsible for XRCC1 ubiquitination remains elusive (Table 15.2). In yeast, Rad1 endonuclease cleaves ssDNAs to facilitate NER. Rad1 is SUMOylated by yeast

SUMOylases Siz1/2 to release Rad1 from binding ssDNA (Sarangi et al., 2014b). In addition, the nuclease complex scaffolding protein Saw1 is SUMOylated by Siz1/2 as well to attenuate Rad1 binding while meantime promotes Slx4 interaction to tone down NER (Sarangi et al., 2014a). The yeast Topoisomerase II (Top2) is SUMOylated by Siz1/2 to promote Top2 centromeric localization to facilitate damage repair (Bachant et al., 2002; Takahashi et al., 2006; Takahashi and Strunnikov, 2008). Siz1/2 SUMOylase also SUMOylates the DNA ligase scaffolding protein Lif1, which leads to release of Lif1 from binding DNA (Vigasova et al., 2013), and the DNA recombination mediator, Rad52, to reduce Rad52 binding to Ufd1 and DNA (Sacher et al., 2006; Torres-Rosell et al., 2007; Altmannova et al., 2010; Bergink et al., 2013), to terminate repair (Table 15.2). Ubiquitin and SUMO signalling in base excision repair (BER) BER repairs damaged DNA bases in a highly coordinated order with a rapid speed. Recognition of the damaged bases is carried out by DNA glycosylases such as Msh2, Mlh1 and MutYH. Upon excision of the damaged DNA bases by AP endonucleases (such as APE1), the gaps will be filled by PNKP and XRCC1/DNA ligase III. Ubiquitination mediated protein stability control of BER components was firstly observed in early 2000s (HernandezPigeon et al., 2004). Soon afterwards, MutYH (Dorn et al., 2014) and RNA Polβ (Parsons et al., 2009) levels were found to be negatively regulated by the E3 ligase Mule. In addition, both Mdm2 (Busso et al., 2009) and UBR3 (Meisenberg et al., 2012) target APE1 for proteasomal degradation to restrain APE1 expression and activity in BER. PNKP is recognized and degraded by a E3 ligase complex composed of Cul4A–DDB1–STRAP, a process that can be antagonized by ATM-mediated phosphorylation of PNKP (Parsons et al., 2012). As a PARP-dependent E3 ligase, RNF146 ubiquitinates XRCC1 and DNA ligase III to facilitate BER (Kang et al., 2011; Zhou et al., 2011). Moreover, the E3 ligase CHIP was observed to govern the protein turnover of a handful of BER members, including XRCC1, OGG1 and RNA Polβ (Parsons et al., 2008). In addition, Cullin 1 and Cullin 4-based E3 ligases have also been implicated in degrading BER

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components UNG and SMUG1 induced by Vpr (Schröfelbauer et al., 2005). Interestingly, Rad7 and San1 E3 ligases target variants or mutated, but not WT-Msh2 for proteasomal degradation, suggesting that in addition to control of normal BER process, certain ubiquitin signalling may also govern aberrant protein turnovers for BER members under pathophysiological conditions. In yeast, SUMOylation of the DNA glycosylase TDG attenuates TDG binding to DNA to negatively regulate BER (Hardeland et al., 2002; Steinacher and Schär, 2005; Baba et al., 2005, 2006; Smet-Nocca et al., 2011) (Fig. 15.4 and Table 15.2). Discussion and future perspectives Genome stability is essential for normal cell physiology such as development, metabolism, proliferation in individuals, and also indispensable to faithfully pass genetic information to next generation. While certain flexibility is also allowed to gain advantages to adapt to environment or for evolution for better survival. In this chapter, we focus on DNA damage repair regulations in individuals rather than across different generations. The tight while tempo and spatial control of genome stability is achieved by a delicate DNA damage sensing, initiating, repair and termination system, mechanisms of which are conserved evolutionarily from yeast to human. Although distinct types of DNA damages are repaired by a variety of mechanisms, all key components in these repair pathways are controlled at their cellular levels – both protein abundance and enzyme activity. Although DNA damage induced transcriptional regulation of certain genes is also present (Elkon et al., 2005; Alvarez-Fernandez et al., 2010), as an acute response, protein post-translational regulations play a more important role. In addition to protein phosphorylation that can amplify signals quickly towards a large-scale, protein ubiquitination and SUMOylation provide a powerful approach to properly earmark unnecessary proteins for degradation, alter protein cellular localization, and more importantly, provide a platform for protein binding to recruit necessary DNA damage repair factors. This is partially achieved by the uniqueness of the ubiquitin code. The ubiquitin code is composed of types of ubiquitin modifications

Figure 15.4 Ubiquitin and SOMO modifications in BER.

(mono-ubiquitination or polyubiquitination), versatile ubiquitin linkages that are gaining more and more attention due to their underappreciated physiological functions (Swatek and Komander, 2016), composition of polyubiquitin chains (homogenous or branched chains), post-translational modifications of a single ubiquitin molecule and complexity in ubiquitination accepting sites on substrates. Different combinations of these ubiquitin codes provide distinct biological meanings that can be interpreted by different ubiquitin code reader proteins. Compared with ubiquitin, less is known about SUMO, and it remains to be determined whether SUMO molecules are also undergoing posttranslational modifications and whether branched SUMO chains are present. In addition to ubiquitin or SUMO molecules, cellular levels, cellular location and activities of ubiquitin or SUMO enzymes are also tightly controlled to ensure proper repair of damaged DNA. If not, unfaithful repair of damaged DNA, delayed repair and insufficient repair will lead to genome instability. Genome instability has been linked and shown as the cause for a variety of human disorders, including Xeroderma pigmentosum, Cockayne

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syndrome, Fanconi anaemia, Bloom syndrome, Ataxia telangiectasia, Hutchinson–Gilford Progeria syndrome, other rare genetic diseases and cancer (Watanabe and Maekawa, 2013). These diseases are resulted from DNA nucleotide changes, nucleotide insertion, deletion, translocation and changes or exchanges at chromosomal levels. Thus, it is not surprising that dysregulation of key ubiquitin E3 ligases, deubiquitinases, SUMOylases and deSUMOylases are observed in cancer. For example, cancer patient derived Fbw7 mutations occur in its substrate binding region, leading to inability for cancerous mutated Fbw7 to target its physiological substrates for degradation. Thus, aberrantly accumulated Fbw7 substrates [FAAP20 in ICLR (Wang et al., 2016)] lead to improper ICLR facilitating tumorigenesis. Another example is the E3 ubiquitin ligase SPOP. Mutations in substrates binding regions of SPOP in cancer similarly impair DDR by disrupting normal substrate degradation process, while the exact identity of the SPOP substrates in DDR remains unknown (Boysen et al., 2015). Compared with ubiquitin system, whether and how SUMO modifying enzymes (including both SUMOylases and deSUMOylases) contribute to human diseases are just began to be appreciated and further thorough investigations are warranted. Although E3 ubiquitin ligases usually do not display enzymatic activity but rather facilitate ubiquitin transfer from E2 enzymes to substrates, inhibitors targeting E3/substrate interactions have been developed such as Skp2 inhibitors (Wu, L. et al., 2012; Chan et al., 2013). These inhibitors demonstrate potential in treating cancer, however, they have not been applied in DNA damage studies. Similarly, only a few DUB inhibitors have been developed with promises in cancer therapy (Kategaya et al., 2017; Turnbull et al., 2017), although their function in DDR is understudied. Advances of detailed molecular understanding of the ubiquitin and SUMO-mediated regulatory signalling events will pave the foundation to identify new ubiquitin and SUMO modifying enzymes as potential drug targets to alter or correct defective DDR to treat human genetic diseases caused by genome instability. References

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The Role of Ubiquitination and SUMOylation in Telomere Biology Michal Zalzman1,2,3,4*, W. Alex Meltzer1, Benjamin A. Portney1, Robert A. Brown1 and Aditi Gupta1

16

1Department of Biochemistry and Molecular Biology, University of Maryland School of

Medicine, Baltimore, MD, USA.

2Department of Otorhinolaryngology-Head and Neck Surgery, University of Maryland School of

Medicine, Baltimore, MD, USA. Marlene and Stewart Greenbaum Cancer Center, University of Maryland School of Medicine, Baltimore, MD, USA. 4 The Center for Stem Cell Biology and Regenerative Medicine, University of Maryland School of Medicine, Baltimore, MD, USA. 3

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.16

Abstract Telomeres are a unique structure of DNA repeats covered by proteins at the ends of the chromosomes that protect the coding regions of the genome and function as a biological clock. They require a tight regulation of the factors covering and protecting their structure, as they are shortened with each cell division to limit the ability of cells to replicate uncontrollably. Additionally, they protect the chromosome ends from DNA damage responses and thereby, prevent genomic instability. Telomere dysfunction can lead to chromosomal abnormalities and cancer. Therefore, dysregulation of any of the factors that regulate the integrity of the telomeres will have implications to chromosomal stability, replicative lifespan and may lead to cell transformation. This chapter will cover the main factors participating in the normal function of the telomeres and how these are regulated by the ubiquitin and SUMO systems. Accumulating evidence indicate that the ubiquitin and SUMO pathways are significant regulators of the shelterin complex and other chromatin modifiers, which are important for telomere structure integrity. Furthermore, the crosstalk between these two pathways has been

reported in telomeric DNA repair. A better understanding of the factors contributing to telomere biology, and how they are regulated, is important for the design of new strategies for cancer therapies and regenerative medicine. Telomere structure and function Telomeres are DNA structures covered by proteins at the ends of the chromosomes that serve several key biological functions. Primarily, they function as a ‘biological clock’ that regulates the replicative lifespan of cells, as well as protect the integrity of the ends of the chromosomes from nucleolytic digestion (Vaziri et al., 1994; Vaziri and Benchimol, 1996; Karlseder et al., 1999; O’Sullivan and Karlseder, 2010). The telomeres consist of several components that cooperate to mediate telomere function: A DNA repeat sequence (TTAGGG in mammals), the Shelterin complex, a group of proteins that function to cover and compact the repeat sequences and interacting RNA components. Telomere length varies between organisms ranging from a few hundred base pairs in yeast, to tens of kilobase pairs in mammals. In humans, the length of

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The Hayflick limit and the end replication problem The telomere’s ability to act as a biological clock serves as a powerful mechanism for tumour suppression. Primary cells can potentially divide a limited amount of times before reaching a state where they can no longer replicate (Hayflick and Moorhead, 1961). This replication limit, called the Hayflick limit, was first described by Dr Leonard Hayflick in 1961. In this work, he and Dr Paul Moorhead demonstrated that unlike cancer cells, primary cells age in culture and eventually die, disproving a longstanding theory that all cell lines in culture are immortal (Hayflick and Moorhead, 1961). In the next decade, the connection between cellular ageing and telomeres was hypothesized to be a result of the ‘end replication problem’, which is a by-product of the linear nature of eukaryotic DNA. DNA polymerase can only synthesize DNA in the 5′–3′ direction to generate new DNA. Replication of the leading 5′–3′ DNA strand allows DNA polymerase to generate a complete complimentary strand. Conversely, the 3′–5′ lagging DNA strand, requires the activity of the enzyme Primase which adds RNA primers for the creation of 100–200 base pair sized DNA fragments, called the Okazaki fragments. This allows DNA polymerase to synthesize DNA in the 5′- 3′ direction. Following replication, the RNA primers are removed and the Okazaki fragments are ligated together. However, Primase cannot add RNA primers at the end of the lagging strand. This issue leads to an incomplete replication, creating a 75–300 nucleotide long overhang of the

Embryonic stem cells Telomere Length

the telomere can range anywhere from 5 kb–20 kb (Samassekou et al., 2010). Telomeres play a critical role in cellular replicative lifespan and protect the coding regions of the genome. Therefore, dysfunctions or disruptions of those nucleoprotein structures at the end of the chromosomes can present as serious pathologies. These telomere syndromes span several different disease areas including blood, lung and liver disease, bone marrow failure, age related disease, and cancer. Collectively, telomere erosion has a strong correlation with several different organ specific diseases. Understanding the mechanisms that regulate telomere length will help guide diagnosis, prevention and treatment.

Cancer cells

senescence

crisis

Number of Cell Doublings

Figure 16.1 Telomere length decreases as cells age. In culture, every cell division results in loss of telomeric DNA. The rate of telomere shortening varies between cell types and continues until the telomeres reach a point when they are short enough to induce a signal to enter into senescence, i.e. growth arrest. Further telomere erosion and in vivo clearance by the immune system leads to apoptosis or culture crisis.

3′ telomeric end (Makarov et al., 1997; McElligott and Wellinger, 1997; Chai et al., 2006). To protect this overhang, proteins are recruited for the further processing needed to properly create a protected DNA structure called a T-loop, resulting in overall resection of the end of the chromosome. Thus, as cells divide and DNA is replicated, the telomeres gradually shorten (Fig. 16.1). At a certain predetermined length, which varies between cell types, the telomeres become critically short and signal for the cell to cease further replication, inciting cellular senescence or death. Telomeres and protecting the genome The ability of cells to detect and repair DNA damage is a powerful mechanism for cellular maintenance and cancer prevention. One such trigger for this response is via the detection of exposed linear DNA (Chapman et al., 2012). Several cellular responses to detection of broken DNA exist, including repair of fragments, cell cycle arrest, or, if the damage is severe, apoptotic cell death ( Jackson and Bartek, 2009). These mechanisms have evolved to prevent aberrant genomic instability resulting in mutations that can lead to cancer. Paradoxically, because mammalian DNA is linear, and due to the 3′ overhang, telomeres can be recognized as DNA breaks by the DNA repair machinery and damage

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response agents, unless properly shielded. Therefore, to protect the telomeres, the repeat sequences fold into themselves to create a ‘T-loop’ structure to avoid recognition double-stranded DNA damage. The repeat sequence is masked by a network of proteins called the shelterin complex (Fig. 16.2).

DNA and the telomeric end, successfully blocking any DNA repair response (Fig. 16.2). The shelterin complex To properly form and conserve the T-loop structure, a number of proteins are required, collectively called the shelterin complex (Fig. 16.2). The shelterin complex is comprised of six proteins: TRF1 (telomere repeat binding factor 1) (Zhong et al., 1992; Chong et al., 1995), TRF2 (telomere repeat binding factor 2) (Bilaud et al., 1997), POT1 (protection of telomeres 1) (Baumann and Cech, 2001), TIN2 (TRF1-interacting nuclear protein 2) (Kim et al., 1999), RAP1 (repressor and activator protein 1) (Li et al., 2000), and TPP1 (POT1-and TIN2interacting protein) (Houghtaling et al., 2004). The shelterin complex protects the telomeres in multiple ways. First, through facilitating the T-loop formation by promoting the strand invasion of the 3′ DNA overhangs. This prevents the detection of the exposed single-stranded DNA, and therefore blocks the detection as DNA break (Griffith et al., 1999). Both TRF2 and TRF1 are able to remodel artificial telomeres in vitro to create the T-loop structures (Bianchi et al., 1997; Griffith et al., 1999; Stansel et al., 2001). Further, TIN2 acts to enhance the TRF1 mediated T-loop formation (Kim et al., 2003). Secondly, the shelterin complex interacts with and inhibits the DNA damage response

T-loops A displacement loop or a D-loop, is a DNA structure in which a double-strand of DNA is additionally occupied by a third single-stranded DNA based on base complementarity. In the context of the telomere, this structure is referred to as a T-loop and is created through the 3′ strand invasion of the G-rich overhang, created during DNA replication (Greider, 1999; Griffith et al., 1999; Murti and Prescott, 1999; de Bruin et al., 2000; Muñoz-Jordán et al., 2001). The displacement occurs at a distant place from the end of the telomere, creating a large duplex lariat structure (de Lange, 2005). Evidence for these structures first arose from work done in vitro, demonstrating that artificially generated telomeres form large loops only in the presence of a 3′ overhang (Greider, 1999). This looped structure shelters the exposed G-rich overhang, caused by the end replication problem, and in the process protects the C-rich shortened end of the telomere. This triplex nucleic acid structure allows DNA repair proteins to distinguish between breaks in the

TTAGGGTTAGGG AATCCCAATCCCAATCCCAATCCC-3’

50-300 nt 3’ overhang

9-15 kb repeats

TIN2 TRF1

TIN2

TRF2 RAP1

TRF1

TIN2

TIN2

TRF2

TRF1

RAP1

TRF2

TRF1

TRF2

POT1

3’

RAP1

RAP1

RAP1 RAP1

TRF2

TRF2

TTAGGGTTAGGG AATCCCAATCCC

POT1

POT1

RAP1 TRF2

POT1

Figure 16.2 Telomere structure. Telomeres are structures at the ends of the chromosomes that contain 6-nucleotide repeat sequences (top). Incomplete replication of the lagging strand results in a G-Rich 3′ overhang (middle). The repeat sequences are coated with a cluster of proteins called the shelterin complex. The shelterin complex helps loop the G-rich overhang into the double-strand telomere sequence, creating a T-loop (bottom).

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pathway (Karlseder et al., 2004). A deficiency of any of the shelterin complex components leads to telomere deprotection, genomic instability and potentially to cellular senescence. When TRF2 is down-regulated, the ATM kinase pathway is activated, leading to cell cycle arrest (Karlseder et al., 1999). DNA damage signalling is also seen in the absence of TIN2 or POT1 (Kim et al., 2004; Hockemeyer et al., 2005). Lastly, the shelterin complex is thought to inhibit the telomere maintenance enzyme, telomerase, which can add telomere repeats lost during cell replication (Loayza and de Lange, 2003; Liu et al., 2004; Kelleher et al., 2005; Lei et al., 2005). Protein turnover Concentrations and spatial gradients of proteins must be able to rapidly respond to extracellular cues and cell status (Korolchuk et al., 2010). Even subtle protein imbalances can drastically impact important cellular processes. Therefore, the regulation of protein degradation and turnover play an important role in the cell-life cycle (Ciechanover, 2005). Protein levels in the cells are at a constant state of turnover. Continuous synthesis of proteins and degradation is required for steady state protein levels and cellular homeostasis (Reinstein and Ciechanover, 2006). Hence, protein turnover plays an important role in regulating cellular fitness (Ciechanover, 2005). The balance is maintained through three major systems regulating the maintenance of proper protein folding and native conformation. The first is the chaperone system, which includes stress-induced heat shock proteins (HSPs) involved in protein folding. The second is the ubiquitin-proteasome system (UPS), controlling the degradation and clearance of misfolded proteins. Finally, the autophagy system, responsible for the recycling and degradation of long lived, structural proteins and organelles. (Eskelinen and Saftig, 2009; Yang and Klionsky, 2010). Protein turnover, or degradation, influences a variety of basic cellular functions. Another primary role of protein degradation is to serve as an intracellular quality control system through elimination of misfolded or damaged proteins. Accumulations of misfolded proteins can create non-physiological interactions with other proteins that are particularly harmful to the cell. Proteins can be damaged

in multiple ways, including genetic mutation, misfolding in the ER, translational errors, toxic factors from the environment, or intracellular toxic agents resulting from ageing or disease. Damaged proteins must either be quickly repaired or eliminated in order to prevent further harm to the cell (Goldberg, 2003). More recently, regulated protein degradation has been shown to control complex cellular processes including metabolism, cell cycle, transcription, signal transduction and apoptosis (Ciechanover, 2005; Chen and Sun, 2009). Owing to the amount of vital cellular functions influenced by protein turnover, it is not surprising that dysfunction in protein degradation has been implicated in multiple diseases (Reinstein and Ciechanover, 2006). Aberrant protein stabilization or accelerated degradation of proteins changes their steady-state levels, precipitating disease. Neurological disorders such as Parkinson’s disease are highly linked to protein turnover dysfunction. In many cases, aggregates of disease specific proteins are accumulated and cannot be degraded (Ciechanover and Brundin, 2003; Tanaka et al., 2004). Disruptions in protein turnover have also been identified in cancer, where stabilization of oncogenes and destabilization of tumour suppressors contribute to malignancy (Ohta and Fukuda, 2004). Proteins have developed specialized functions and are therefore degraded at widely different rates. The standard measurement is also known as protein half-life (Zhou, 2004; Hinkson and Elias, 2011). Protein half-lives can range from just a few minutes to hours, in the case of transcription factors and regulatory proteins, to up to multiple days for structural proteins (Goldberg, 2003). In order to control the various cellular functions regulated by protein turnover, cells must precisely control protein half-lives. The function of the ubiquitinproteasome system and the process of SUMOylation The Ubiquitin-Proteasome System (UPS) is a highly regulated apparatus responsible for intracellular protein turnover and degradation (Hershko and Ciechanover, 1998; Nandi et al., 2006). The UPS is a selective process orchestrated by a series of ubiquitin ligase enzymes specific to the pathway. The role of the UPS has been demonstrated in the

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Figure 16.3 The ubiquitin proteasome system. (A) Illustration of the ubiquitination process. (B) Polyubiquitinated protein is degraded by the proteasome.

turnover of up to 90% of all cellular proteins (Huang and Figueiredo-Pereira, 2010). As the name Ubiquitin suggests, the UPS is involved in the regulation of a wide array of biological processes including antigen presentation, DNA repair, protein trafficking, epigenetic regulation, and the cell cycle (Nandi et al., 2006; Al-Hakim et al., 2010). The proteasome has become a drug target, as fluctuations in proteasomal activity and defects in function have been linked to a variety of diseases (Dahlmann, 2007; Bedford et al., 2011). The ubiquitin gene encodes for a small, 76 amino acid protein. Ubiquitin is covalently conjugated to lysine residues of substrate proteins through an isopeptide linkage in a post-translational process called ubiquitination (Pickart and Eddins, 2004). Ubiquitination occurs through an enzymatic cascade in three steps: activation, conjugation, and ligation. Each step is facilitated by a distinct ubiquitin enzyme (Fig. 16.3A,B). The initial step, activation of the ubiquitin molecule by an ATP dependent E1 ubiquitin enzyme, produces a ubiquitin-adenylate intermediate. Next, the activated ubiquitin is transferred to the active site of an E2 ubiquitin enzyme via a trans(thio)esterification reaction (Ye and Rape, 2009). In the final ligation

step, E3 ubiquitin enzymes facilitate the transfer, either directly (HECT domain E3s) or indirectly (RING domain E3s), of the ubiquitin molecule from the E2 enzyme to a lysine residue in the substrate protein. Successive rounds of the ubiquitination process result in ubiquitin chains (Callis, 2014). The specificity of ubiquitination increases with each step, as only two genes are responsible for E1 ubiquitin enzymes and only 35 genes for E2 enzymes. However, there are well over six hundred E3 ubiquitin ligases that recognize target substrates, thereby conferring specificity to the UPS (Ardley and Robinson, 2005). E3 ubiquitin ligases fall into two main structural families that differ in how ubiquitin is transferred from E2 enzyme to substrate: the HECT (Homologous to the E6-AP Carboxyl Terminus) domain and the RING domain (Really Interesting New Gene) ligases (Buetow and Huang, 2016). HECT domain E3 ligases contain a catalytic cysteine residue that accepts the ubiquitin molecule from the E2 ligase, forming a thioester intermediate. It is the HECT domain E3 ligase that then directly transfers the ubiquitin to the substrate (Morreale and Walden, 2016). Approximately 30 HECT domain E3 ligases have been identified. Alternatively, ubiquitination with RING domain E3 ligases is facilitated by E2 ligases. The RING E3 ligase simply acts as a scaffold between the E2 ligase and substrate (Ozkan et al., 2005). RING domain E3 ligases are the predominant family in mammals, with over 300 enzymes identified. The Small Ubiquitin-like Modifier (SUMO) system regulates protein function by covalently attaching and detaching small protein chains that are analogous to ubiquitin. The enzymatic cascade of SUMOylation is similar to that involved in ubiquitination. Like the ubiquitin system, SUMOylation is involved in multiple cellular processes, such as regulation of transcription, apoptosis, transport to the nucleus, protein stability, cellular stress response and cell cycle progression [1]. The last four amino acids of the C-terminus of SUMO are cleaved off allowing the formation of an isopeptide bond between the C-terminal glycine residue and an acceptor lysine of the target protein. There are four SUMO proteins known as SUMO1–4. Similar to ubiquitination, SUMOylation is regulated by E1 activating enzymes, E2 conjugation enzymes, E3 SUMO ligases and SUMO specific proteases

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which are involved in the removal of SUMO conjugates. There are a number of diseases associated with SUMOylated proteins, such as Parkinson disease (PD) and Alzheimer’s disease. Histone SUMOylation was first identified in 2003. Histone 4 can be modified by SUMO through the HDAC. However, unlike ubiquitin, SUMOylation does not directly mark protein for proteasomal degradation, but often works to induce the function or shift the localization of the modified protein in the cell. Telomere regulation in cancer Cancer is considered a disease state in which dysfunctional cells replicate at a high rate and lose their ability to interact properly with their environment, resulting in abnormal growth and invasion into nearby and distant tissues. In 2016, an estimate of 1.6 million new cases of cancer were diagnosed, in the U.S. with roughly 600,000 deaths from the disease, making it the second leading cause of death ( Jemal et al., 2017). Cancer is thought to develop in a multistep process of sequential rounds of genetic mutations that convert a normal cell into a malignant cell ( Jonkers, 2012). However, in addition to mutation accumulation, a second crucial event must occur in order to ensure unlimited cell replication. Otherwise, a mutated cell will age and cease to divide before it can be detected as a tumour. This event must lead to the evasion or the reversal of telomeres shortening and physiological ageing during cell division (Sugimoto et al., 2004; Smith et al., 2016). Telomere dysfunction is further linked to cancer, as patients with telomere related diseases have an increased risk for developing cancer (de la Fuente and Dokal, 2007; Alter et al., 2009; Diaz de Leon et al., 2010; Alder et al., 2011). This is likely cause by shortened telomeres and faulty repair mechanisms that trigger chromosomal fusions and increase genomic instability. Telomerase Telomerase is a ribonucleoprotein complex that consists of an RNA component (TERC) and a reverse transcriptase (TERT). TERC provides the RNA template required for the reverse transcription activity of the enzyme TERT. Together with additional complex components, they add repeats to shortened telomeres. Telomerase activity is

most robust during embryonic development, and then persists in much lower levels in certain human adult tissues (Broccoli et al., 1995; Counter et al., 1995; Härle-Bachor and Boukamp, 1996; Schieker et al., 2004). Telomerase activity and the resulting telomere elongation can lead to the bypass of replicative senescence and to cell immortalization (Garbe et al., 2014; Smith et al., 2016). Expectedly, mutations in telomerase complex components are commonly found in telomere syndromes such as dyskeratosis congenita and aplastic anaemia (de la Fuente and Dokal, 2007; Savage and Alter, 2009; Diaz de Leon et al., 2010; Dokal, 2011; Nelson and Bertuch, 2012). The ability to maintain telomere length in cancer cells is traditionally attributed to the enzyme telomerase (Kunická et al., 2008). Telomerase is expressed in 85% of all cancers. The remaining telomerase negative cancers must activate other mechanisms to maintain telomere integrity which are collectively called: Alternative Lengthening of Telomeres (ALT) mechanisms (Bryan et al., 1997; Cesare and Reddel, 2010). While it has been shown that ALT can still function in the presence of telomerase overexpression, it is generally assumed that these mechanisms act in a mutually exclusive manner (Cerone et al., 2001; Grobelny et al., 2001; Perrem et al., 2001). In fact, inhibition of telomerase by a drug can ultimately lead to resistance through activation of ALT (Hu et al., 2012; Hu et al., 2016). Alternative lengthening of telomeres (ALT) The less understood mechanism of telomere maintenance in cancer is alternative lengthening of telomeres (ALT). The defining characteristic of the canonical ALT mechanism is its independence from telomerase activity. ALT dependent cancer cells also have other distinguishing characteristics, including extrachromosomal circular telomeric DNA (c-circles) (Cesare and Griffith, 2004; Henson et al., 2009), telomeric DNA associated with promyelocytic leukaemia (PML) bodies (Yeager et al., 1999), heterogeneous telomere lengths across different chromosomes, and increased telomere recombination events (Bailey et al., 2004). Unlike telomerase, that uses an RNA template, cancers that present with canonical ALT are thought to use telomeric DNA as a template for extension. The

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template can be a sister chromatid strand (sister chromatid exchange), extrachromosomal circular telomeric DNA, or a telomeric sequence from a separate chromosome (homologous recombination) (Cesare and Reddel, 2010; Yu et al., 2014). Therefore, proteins involved in homologous recombination (HR) were shown to be required for successful telomere maintenance in ALT (Zhong et al., 2007). Ubiquitination-mediated regulation of the shelterin The Shelterin complex caps the telomeres and acts as a protective layer covering the telomeric DNA at end of the chromosomes. Shelterin is composed of multiple proteins that have been shown to be regulated by the ubiquitin-proteasome system. The Ubiquitin-mediated degradation of the Shelterin component TRF1 is the first example and is facilitated by three E3 ligases: RLIM, FBX4 and β-TRCP1 (Lee et al., 2006; Her and Chung, 2009; Wang et al., 2013). RLIM targets telomere DNAbound TRF1 for proteasomal degradation (Her and Chung, 2009). Conversely, FBX4 binds to the N-terminal region of the dimerization domain of unbound TRF1 and targets it for degradation (Lee et al., 2006). Consequently, when either of these enzymes is depleted, TRF1 levels are stabilized, causing telomerase inhibition and leading to a decrease in telomere length and to impaired cell growth. Furthermore, TRF1 levels are also indirectly and independently regulated by the factors U2AF65 (Kim and Chung, 2014), TIN2 (Ye and de Lange, 2004), and the F-box protein β-TRCP1 (Wang et al., 2013), which were shown to positively regulate of TRF1 by acting as competitive inhibitors to FBX4, and physically preventing its interaction with TRF1 and subsequent ubiquitinmediated degradation. TIN2 itself acts as part of the shelterin complex, therefore, further regulation of TIN2 turnover is achieved by the ubiquitin system as the interaction with the E3 ligase SIAH2 sends it to proteasomal degradation (Bhanot and Smith, 2012). The turnover of another important shelterin subunit, TRF2, has also been shown to be regulated by ubiquitination. With either normal cell replication or telomere dysfunction, telomere shortening leads to reduced levels of TRF2 binding and as a

result, to a loss of TRF2 mediated telomere protection. Consequently, a cascade of events is triggered in the cell. First, the ATM kinase is activated which in turn phosphorylates the tumour suppressor p53. Then, the activated p53 triggers replicative senescence. Additionally, as a feedback loop, p53 induces the transcription of the E3-ubiquitin ligase SIAH1, which targets TRF2 for degradation. Finally, this cascade is further amplified, as a positive feedback loop, with increased p53 activation leading to further increased SIAH1 levels and TRF2 ubiquitination (Fujita et al., 2010). Another shelterin subunit regulated by ubiquitin-mediated degradation is TPP1. Although the E3 ubiquitin ligases for TPP1 are still unknown, it has been shown that inhibition of the proteasome system leads to stabilization of TPP1 protein levels. The human TPP1 levels are shown to be further regulated by interaction with the deubiquitinating enzyme USP7 which removes ubiquitin chains from it (Zemp and Lingner, 2014). In mice, the E3 ligase RNF8, ubiquitinates TPP1 and is required for its stabilization at telomeres (Rai et al., 2011). However, the role of ubiquitination of human TPP1 still remains to be discovered, as it has not been demonstrated to affect its function, or its interaction with other shelterin components such as TIN2, POT1 or its interaction with telomerase (Zemp and Lingner, 2014). SUMOylation-mediated regulation of the shelterin Crosstalk between SUMOylation and ubiquitination were also to contribute to regulation and turnover of TRF2. The E3 SUMO ligase PIAS1 interacts with and was shown to SUMOylate TRF2. This allows the interaction of the SUMO-targeted ubiquitin ligase RNF4 which in turn ubiquitinates TRF2 and send it to degradation by the proteasome and by that contributes to TRF2 turnover without affecting telomere integrity (Her et al., 2015). Finally, SUMOylation was also shown to tag Rap1 in the budding yeast, Saccharomyces cerevisiae. The SUMO-targeted ubiquitin ligase Uls1 binds SUMOylated Rap1, ubiquitinates it and send it to proteasome mediated degradation. Loss of Uls1 results in accumulation of poly-SUMOylated Rap1 and leads to telomere fusions. Elimination of Rap1 SUMOylation sites in Uls1-depleted cells prevents

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telomere fusion suggesting that poly-SUMOylated Rap1 is non-functional in telomere protection from NHEJ (Lescasse et al., 2013). In mammals, the shelterin complex component RAP1 forms a dimer with TRF2 to protect the telomeres from inappropriate processing by the homologous recombination pathway and from rapid telomere resection, which would otherwise result in telomere loss and fusions in both mouse and human cells (Rai et al., 2016). An additional role for SUMOylation in telomere length regulation through shelterin modulation has been shown in fission yeast (Miyagawa et al., 2014). SUMOylation of Tpz1, the fission yeast homologue of TPP1, is required to maintain telomere length. The mechanism was elucidated whereby SUMOylated Tpz1 recruits Stn-Ten1 to the telomere, which in turn inhibits the binding of telomerase and telomere replication. Mutation of the Lysine 242 on Tpz1 prevents its SUMOylation, prevents Stn-Ten1 recruitment, and results in abnormally long telomeres. These findings have yet to be demonstrated in mammalian systems, however, the CST complex (including STN and TEN1) are well conserved in humans and inhibit telomerase activity (Chen et al., 2012) suggesting the mechanism may be similar. In summary, the binding of shelterin complex subunits to the telomeric DNA is essential for telomere integrity and genome stability, but it relies heavily on the proper regulation of the Shelterin by the ubiquitin and SUMO systems. Telomere chromatin regulation by ubiquitin Post translation modification of histones tails such as methylation and acetylation are a powerful means for chromatin structure modulation. In yeast (Saccharomyces cerevisiae), the E2 Ubiquitin ligase Rad6 (Ubc2) was shown to regulate histone H3 methylation at lysine 4 (H3K4-me) through addition of one molecule of ubiquitin to histone H2B (mono-ubiquitination of H2B) at Lys 123. Furthermore, a mutation abolishing the Lysine 123 in H2B leads to telomere transcription and the expression of the long non-coding RNA TERRA (Sun and Allis, 2002). Another important modulator of telomere chromatin state by histone H2A ubiquitination is

mediated through the E3 ligase RNF8. RNF8 is DNA-damage-responsive protein that mediates histone ubiquitination signalling and plays a critical role in the cellular response to genotoxic stress and DNA damage repair (Huen et al., 2007). However, it was further shown to affect telomere stability and facilitate telomere fusion by ubiquitination of histone H2A and H2AX. Consistent with the critical effect of RNF8 on uncapped telomeres, loss of RNF8, as well as of the E3 ligase RNF168, reduces telomere-associated genome instability. These data suggest that H2A mono-ubiquitination may enhance cancer development by facilitating telomere fusion and dysfunction (Peuscher and Jacobs, 2011) and highlight mono-ubiquitination in the maintenance of telomere integrity. Poly-ubiquitination has also been shown to be important for telomere biology. The early embryonic gene Zscan4 (Zinc finger and SCAN domain containing 4) promotes genomic stability and telomere homeostasis in mouse embryonic stem (ES) cells (Zalzman et al., 2010). Zscan4 is transiently expressed (Zalzman et al., 2010), with protein level bursts associated with chromatin remodelling (Amano et al., 2013; Akiyama et al., 2015) and nuclear reprogramming during the generation of induced pluripotent stem (iPS) cells (Hirata et al., 2012; Jiang et al., 2013; Park et al., 2015). The human ZSCAN4 has been shown to interact with shelterin complex components (Lee and Gollahon, 2014, 2015), and has been suggested to play a role in cancer (Zalzman et al., 2010; Lee and Gollahon, 2014; Portney et al., 2018). Given the important role of ZSCAN4 and its transient expression in the cell (Falco et al., 2007; Zalzman et al., 2010), maintaining the delicate balance between its protein synthesis and degradation is critical for stem cell and potentially cancer cell function. Therefore, stringent regulation of the levels of ZSCAN4 is required to effectively control its function. Indeed, ZSCAN4 protein degradation was shown to be regulated by the ubiquitin-proteasome system (Portney et al., 2018). The E3 ubiquitin ligase RNF20 negatively regulates ZSCAN4 protein by poly-ubiquitination. Further, RNF20 depletion does not affect ZSCAN4 RNA transcription, yet it leads to the accumulation and stabilization of ZSCAN4 protein, suggesting it as a negative regulator of ZSCAN4 protein stability. Due to the important role of ZSCAN4 in the

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generation of iPS cells, these data have important implications for role of ubiquitination in the regulation of telomere and genomic stability (Portney et al., 2018). Discussion and perspectives The function of the telomeres as a biological clock requires a tight regulation of the factors covering and protecting their structure, in order to limit the ability of cells to replicate uncontrollably. Telomeres also protect the chromosome ends from activating DNA damage responses and thereby, prevent chromosomal fusions and genomic instability. Telomere dysfunction leads to increased chromosomal abnormalities and cancer development (Feldser et al., 2003; Blasco, 2005; Gilley et al., 2005). Consequently, a dysregulation of any of the factors that regulate telomere structure integrity and length will cause major implications to chromosomal integrity, cellular lifespan and cancer transformation. A better understanding of the process and factor controlling telomere processing is important for the development of new strategies for cancer therapies and regenerative medicine. Compelling evidence suggest that the ubiquitin and SUMO pathways are important regulators of both the shelterin and telomere chromatin structures. Moreover, these posttranslational modifications contribute to the cellular response to damaged telomeres. Further research is needed to promote our understanding of the effect of these modifications on telomere regulation and function and their significance to human health. Likewise, additional studies are needed to determine the underlying mechanisms, by which Ubiquitination and SUMOylation regulate telomere factors and protect the genome. The crosstalk between the two pathways has been demonstrated in both genomic and telomeric DNA repair. Further research will elucidate and the modifications unique to telomere maintenance and repair, which may allow to device new strategies for targeting the telomeres without triggering unwanted mechanisms for genomic DNA repair. References

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Role of Ubiquitin and SUMO in Intracellular Trafficking Maria Sundvall1,2,3*

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1Institute of Biomedicine, University of Turku, Turku, Finland.

2Western Cancer Centre FICAN West, Turku University Hospital, Turku, Finland. 3Department of Oncology and Radiotherapy, University of Turku, Turku, Finland.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.17

Abstract Precise location of proteins at a given time within a cell is essential to convey specific signals and result in a relevant functional outcome. Small ubiquitin-like modifications, such as ubiquitin and SUMO, represent a delicate and diverse way to transiently regulate intracellular trafficking events of existing proteins in cells. Trafficking of multiple proteins is controlled reversibly by ubiquitin and/or SUMO directly or indirectly via regulation of transport machinery components. Regulation is dynamic and multilayered, involving active crosstalk and interdependence between post-translational modifications. However, in most cases regulation appears very complex, and the mechanistic details regarding how ubiquitin and SUMO control protein location in cells are not yet fully understood. Moreover, most of the findings still lack in vivo evidence in multicellular organisms. Posttranslational modifications in regulation of cellular processes General principles of ubiquitination and SUMOylation Posttranslational modification (PTM) of proteins is a powerful, fast and often transient way to control the fate of existing proteins and cell

behaviour. In addition to e.g. chemical groups, such as phosphate groups, proteins can be modified by small polypeptides, such as ubiquitin and SUMO. Ubiquitin and SUMO share a similar three-dimensional structure and are principally covalently linked at lysine (K) residues of substrate proteins by a similar conjugation pathway (Hershko et al., 1998; Hay, 2013). Unlike ubiquitin, SUMO is preferentially attached at SUMO consensus sites ΨKxE in substrates Ψ = hydrophobic residue with high preference for I or V, x = any amino acid) under steady state conditions, but under stress conditions in particular more nonconsensus sites are SUMOylated (Hendriks et al., 2016). Mammals express ubiquitously SUMO1, SUMO2 and SUMO3, whereas SUMO4 and SUMO5 are only expressed in some tissues and their functional role is unclear (Pichler et al., 2017). SUMO2 and SUMO3 are nearly identical and resemble approximately 50% to SUMO1 (Geiss-Friedlander et al., 2007). Precursors of ubiquitin and SUMO are processed to mature forms prior to conjugation and activated by an E1 enzyme in a reaction depending on ATP. Subsequently ubiquitin and SUMO are transferred to an E2 enzyme, and E3 ligase alone or with E2 ligates them to substrates by an isopeptide bond (Hershko et al., 1998; Hay, 2013). Modification is reversible and specific proteases can cleave ubiquitin and SUMO from substrates (Williamson et al., 2013; Kunz et al.,

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2018). Proteins can be tagged by a single ubiquitin or SUMO either at one or at multiple lysines. In addition, modifications can exist as chains. Ubiquitin contains seven internal lysine residues (K6, K11, K27, K29, K33, K48, K63) that are involved in the formation of ubiquitin-ubiquitin polymers, known as polyubiquitin chains (Pickart et al., 2004; Heride et al., 2014). In addition, the linkage between the amino-terminal amino group of methionine on a ubiquitin can be conjugated with a target protein and the carboxy-terminal carboxy group of the incoming ubiquitin for linear chains (Walczak et al., 2012). SUMOs can also form polymeric chains through internal lysine residues (Geiss-Friedlander et al., 2007). Monoubiquitination and different chain types determine the fate of the modified protein (Piper et al., 2014). For example, K48 ubiquitin chains are considered classical signals for proteasomal degradation and K63 ubiquitin chains are linked to trafficking and DNA damage response (Pickart et al., 2004). Recently basic principles of the field have been challenged by e.g. discoveries of mixed polyubiquitin chains and ubiquitination of non-lysine residues (Piper et al., 2014). Less is known regarding the consequences of attachment of either single or many SUMO moieties. SUMO chains are at least implicated in recruitment of SUMO-targeted ubiquitin ligases (LallemandBreitenbach et al., 2008). Components, regulation and function of ubiquitin and SUMO machinery Multiple enzymes involved in ubiquitin conjugation have been recognized. The human ubiquitin machinery comprises a network including two ubiquitin E1 enzymes, approximately 40 ubiquitin E2s, and more than 600 E3 ubiquitin ligases in the human genome (Heride et al., 2014). Three major types of E3 ligases are really interesting new gene (RING) type E3s comprising most of human E3s, homologous to E6-AP carboxyl terminus (HECT) and RING-between-RING (RBR) E3s ligases (Deshaies et al., 2009; Rotin et al., 2009; Wenzel et al., 2012). Ubiquitin is cleaved by approximately 100 deubiquitinating enzymes (DUBs) (Williamson et al., 2013; Heride et al., 2014). Intriguingly, only one heterodimeric

E1 (Sae1/Aos1–Sae2/Uba2) and E2 (Ubc9) are known to be involved in SUMO conjugation (Hay, 2013). Three classes of SUMO E3s have been widely accepted and characterized including SP-RING Siz/PIAS ligases, RanBP2 and ZNF451 ligases (Geiss-Friedlander et al., 2007; Rytinki et al., 2009; Cappadocia et al., 2015). Both E2 and E3 can select substrates for SUMOylation, and spatial and temporal regulation of co-localization appears integral for substrate selection (Pichler et al., 2017). Cysteine proteases of the sentrinspecific protease (SENP) family members reverse SUMO conjugation in mammals (Kunz et al., 2018). Moreover, desumoylating isopeptides 1 and 2 and ubiquitin-specific protease-like 1 can deSUMOylate proteins (Shin et al., 2012; Schulz et al., 2012). Whereas ubiquitin machinery is widely expressed within a cell, components of SUMO conjugation pathway mainly localize at the nuclear pores and nucleus. Thus, most of SUMOylated substrates are nuclear proteins, although SUMO modified proteins outside of nucleus exist (GeissFriedlander et al., 2007). Ubiquitination is regulated by extracellular stimuli including growth factors and cytokines, stress and cell cycle changes (Pickart et al., 2004; Heride et al., 2014). Different types of stress stimuli such as heat shock, hypoxia, reactive oxygen species, DNA damage and proteotoxic stress regulate the activity of SUMOylation machinery (Hietakangas et al., 2003; Shao et al., 2004; Bossis et al., 2006; Galanty et al., 2009; Morris et al., 2009; Seifert et al., 2015). Both ubiquitin and SUMO modifications can alter the function, activity, location and stability of their targets. Ubiquitin and SUMO are recognized by either ubiquitin-binding domains (UBD) or sumo-interacting domains (SIM), respectively, and serve as platforms for non-covalent protein–protein interactions (Seet et al., 2006). These domains have been identified in hundreds of proteins. PTMs are also involved in regulation of conjugation specificity and activation (Pichler et al., 2017). For example, SUMO-targeted ubiquitin ligase (STUbL) RNF4 contains multiple SIMs and a RING-domain to bind SUMOylated proteins and an E2 ubiquitinconjugation enzyme (Sun et al., 2007; Tatham et al., 2008).

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Ubiquitin and SUMO as signals regulating membrane trafficking and endocytosis General principles of membrane protein trafficking and endocytosis in cells Internalization and endocytosis of cell surface proteins including different receptors often occurs via the clathrin-dependent endocytic pathway. Cell surface receptors are clustered to pits coated with clathrin that pinch off of the membrane forming vesicles and early endosomes. From early endosomes cargo can be directed back to plasma membrane via recycling endosomes or destined to lysosomal degradation via late endosomes and multivesicular bodies (Mellman and Yarden, 2013). Also, other types of endocytic routes exist, including cholesterol-rich membrane structures, such as lipid rafts and caveolae (Barbieri et al., 2016). Several adaptor proteins and PTMs control these processes (Piper et al., 2014). Membrane protein trafficking and endocytosis system are tightly connected to protein homeostasis. Ubiquitin and SUMO in receptor internalization at the cell surface and in endocytic compartments Initial studies in yeast suggested that ubiquitin can function as a sorting signal regulating the internalization and endosomal targeting of cell surface receptors (Kölling et al., 1994; Hicke et al. 1996, Terrell et al., 1998). After that several studies in different systems and organisms have confirmed that ubiquitin is important regulator of endocytosis and its most critical functional role is likely at the sorting endosomes (Mellman and Yarden, 2013). Endocytosis of human receptor tyrosine kinases (RTKs) is suggested to be regulated by multimonoubiquitination and K63-polyubiquitination, and there is some controversy regarding the significance of specific ubiquitination type due to methodological challenges to address this complex regulatory system (Haglund et al., 2003; McCullough et al., 2004; Huang et al., 2006; Sundvall et al., 2008; Huang et al., 2013). During endocytosis the ubiquitin moieties of cargo are recognized by different endocytic adaptors and regulators via UBDs, such as Eps15,

epsin and endosomal sortin complex required for transport (ESCRT) (Piper et al., 2014). Ubiquitin is also indirectly involved in control of endocytosis as components of endocytic machinery are actively regulated by ubiquitination (Piper et al., 2014). SUMOylation has been implicated in the regulation of endocytic processes, although when compared to ubiquitin the evidence is less extensive, and more work is needed to make general conclusions. Nevertheless, endocytosis of kainate receptor GluR6 is regulated by SUMOylation and non-SUMOylated mutant of GluR6, GluR6K886R, is endocytosis-impaired due to unknown mechanisms (Martin et al., 2007). SUMOylation of Smoothened (Smo) promotes its localization at the cell surface (Ma et al., 2016). Intriguingly, mechanistically SUMO interferes with efficient Smo ubiquitination by recruiting deubiquitinase UBPY/ USP8 in a SIM-dependent manner (Ma et al., 2016). SUMOylation regulates also cell surface expression and activity of VEGFR2 receptor tyrosine kinase (Zhou et al., 2018). VEGFR2–SUMO1 fusion protein but not SUMOylation defective mutant VEGFR2 accumulated at the Golgi suggesting that mechanistically SUMO regulates exocytosis of VEGFR (Zhou et al., 2018). SUMOylation can also indirectly regulate endocytosis. Components of endocytic machinery are modified and regulated by SUMO, such as CIN85 (Tossidou et al., 2012) and arrestin (Wyatt et al., 2011). Interestingly, also dynamin interacts with several members of the SUMOylation machinery (Mishra et al., 2004). Moreover, SUMO can be important for membrane binding of proteins. SUMOylation of PTEN at lysine 266 within CBR3 loop fosters binding of PTEN to plasma membrane via electrostatic interactions (Huang et al., 2012). Ubiquitin and SUMO as signals regulating nucleocytoplasmic shuttling and subnuclear targeting General principles of nuclear import, subnuclear targeting and export Passage of proteins in and out of the nucleus through nuclear pore complexes is tightly regulated.

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In principle, many of the nuclear proteins contain nuclear localization signal (NLS) and nuclear export signal (NES) that facilitate trafficking via associations with karyopherins including importins and exportins (CRM1) together with Ran-GTP, respectively (Stewart M, 2007). Ubiquitin and SUMO in nuclear targeting and trapping Molecular mechanism by which ubiquitin machinery controls protein functions are very complex but, nevertheless, some evidence exists to suggest the role for ubiquitin as a signal controlling nucleocytoplasmic trafficking. Ubiquitination of p53 contributes to nuclear export, and although the regulation of p53 ubiquitination and trafficking has turned out to be very complicated and likely dependent of conditions, attachment of monoubiquitin in particular is suggested to play a role in trafficking (Lohrum et al., 2001; Li et al., 2003). Moreover, similar type of regulation has been suggested for another tumour suppressor protein, PTEN and NF-kB essential modulator NEMO (Huang et al., 2003; Trotman et al., 2007). SUMO was initially discovered as a modifier of RanGap1 targeting it to nuclear pore complex (Matunis et al., 1996, Mahajan et al., 1997). Later SUMO has been implicated in both increasing and decreasing the nuclear accumulation of some proteins. Mechanistically, covalent linkage of SUMO may directly block protein interactions relevant for transport or generate SIM-mediated interaction platform facilitating or interfering with transport. SUMOylation is suggested to regulate nuclear import of full-length IGF receptor (Sehat et al., 2010). The levels of SUMOylation defective mutant IGFR are similar at the cell surface compared with wild type receptor, but the mutant receptor cannot translocate to nucleus unlike wild type (Sehat et al., 2010). Specifically IGFR is suggested to interact with RanBP2 at nuclear pores and that RanBP2 acts as an SUMO E3 ligase for IGFR (Pancham et al., 2015). SUMOylation may also increase the stability of IGFR (Pancham et al., 2015). Trafficking of SUMO machinery components is also under the control of SUMO. SUMOylation of Sae2 in the c-terminus within functional NLS efficiently increases nuclear accumulation (Truong et al., 2012).

SUMO has been suggested to regulate nuclear export of some proteins, including transcriptional repressor TEL, Kruppel-like transcription factor (KLF2), Serine hydroxylmethyltransferase 1 (SHMT1), PTEN, p53, ErbB4 and Notch (Wood et al., 2003; Du et al., 2008, Anderson et al., 2009; Bassi et al., 2013, Santiago et al., 2013, Knittle et al., 2017; Antila et al., 2018). SUMOylation adjacent to NES of KLF5 interferes with its interaction with nuclear export receptor CRM1 resulting in the inhibition of efficient export and increased accumulation in the nucleus (Du et al., 2008). ErbB4 RTK undergoes regulated intramembrane proteolysis (RIP) releasing an intracellular domain (ICD) that can translocate to nucleus and regulate transcription. PIAS3 catalysed SUMOylation within NES of ErbB4 increases the nuclear accumulation of a tyrosine phosphorylated ICD by altering the interaction with CRM1 (Sundvall et al., 2012; Knittle et al., 2017). SUMOylation deficient mutant of ErbB4 accumulates less in nucleus and cannot convey efficiently nuclear signalling (Knittle et al., 2017). Another receptor undergoing RIP, Notch, is also SUMOylated in the nucleus and SUMOylation increases nuclear accumulation of the ICD, but mechanisms resulting in accumulation remain to be elucidated (Antila et al., 2018). SUMOylation of SHMT1 increases nuclear accumulation and mutation of the SUMO motif prevents translocation to the nucleus due to unknown mechanisms (Anderson DD et al., 2009). Interestingly, same sites are also ubiquitinated with K63 polyubiquitin chains increasing stability in the nucleus (Anderson et al., 2012). SUMOylation of PTEN (at K254) is suggested to regulate the efficient nuclear accumulation and subsequently DNA damage response. When compared to wild type, SUMOylation deficient mutant of PTEN localizes less into the nucleus and cannot efficiently regulate homologous recombination (Bassi et al., 2013). Nucleocytoplasmic distribution of SUMOylation-deficient transcriptional repressor TEL also changes compared to wild type (Wood et al., 2003). However, the direct mechanism how SUMO regulates subcellular localization of PTEN or TEL is not clear (Wood et al., 2003; Bassi et al., 2013). On the contrary, SUMOylation of p53 stimulates its nuclear export by increasing the

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disassembly of p53 from the CRM1 in the cytosol (Santiago et al. 2013). Moreover, SUMOylation regulates nuclear export and intranuclear distribution of adenovirus E1B-55K protein (Kindsmüller et al., 2007). Altogether nuclear export and SUMOylation appear to be closely connected due to CRM1-mediated interaction of export complexes with SUMO E3 ligase RanBP2 (Ritterhoff et al., 2015) and SUMOylation also regulates nuclear transport via covalent modification of transport machinery (Rothenbusch et al., 2012). Regulation of subnuclear targeting by ubiquitin and SUMO Ubiquitin and SUMO system has been indicated in targeting proteins into certain subnuclear structures. Subnuclear targeting can direct proteins into locations essential for their functions, trap protein in locations where they are not available to regulate e.g. transcription or alter their susceptibility to regulators of stability. Both ubiquitin and SUMO are implicated in correct targeting of DNA repair factors to the sites of DNA damage (Ulrich, 2014). For example, BRCA1 targeting to double-strand breaks is regulated by K63-linked ubiquitination and SUMO and the process involves RNF4 (Galanty et al., 2009; Morris et al., 2009; Guzzo et al., 2012). PML bodies are subnuclear structures involved in regulation of transcription and host a lot of transcription factors and their regulators (Zhong et al., 2000a). PML is strongly SUMOylated and SUMOylation regulates the integrity of PML bodies and stability of PML as SUMOylation deficient mutant PML does not form nuclear bodies when expressed in PML null cells (Ishov et al., 1999; Müller et al., 1998; Zhong et al., 2000a,b). Several nuclear proteins are hosted in PML bodies and SIM-mediated interactions are thought to be important for assembly (Shen et al., 2006). SUMO E3 ligases and deSUMOlases have specific nuclear localizations and regulate substrate localization, but regulation is often not SUMO-dependent (Sachdev et al., 2001; Kotaja et al., 2002; Hietakangas et al., 2003). On the other hand, SUMO is suggested to be important in the regulation of subnuclear localization of Nuclear Factor of Activated T-cells (NFAT1) (Terui et al., 2004) or nucleolar localization of Proline-, glutamic acid- and leucine-rich protein 1 (PELP1) involved in ribosome biogenesis

and regulation of transcription (Finkbeiner et al., 2011). The genetic evidence in vivo and the significance of ubiquitin and SUMO mediated regulation of trafficking in human diseases Genetic studies using targeted gene disruption in mice suggest that ubiquitin and SUMO pathways are essential, but a lot of redundancy is evident with many of the pathway components. For example, SUMO1 and SUMO3 knockout mice are viable, but SUMO2 knockout mice die during embryogenesis as well as Ubc9 knockout is lethal in mice (Nacerddine et al., 2005; Evdokimov et al., 2008; Zhang et al., 2008; Wang et al., 2014). Unfortunately, very little is known regarding genetic models of modification-deficient mutants and their phenotypes in vivo. Deregulation of ubiquitin conjugation machinery and altered protein ubiquitination has been reported in diseases such as neurodegenerative diseases and cancer (Popovic et al., 2014). SUMOylation seems to be a general protective mechanism against the damage caused by stresses such as low oxygen and nutrient deprivation and may also protect neurons after stress and support the growth of cancer cells. Mutations targeting ubiquitin conjugation machinery, such as E3 ligases, and somatic mutations altering ubiquitin ligase binding and subsequent trafficking of targets, including deregulated endocytosis of RTKs, in cancer have been reported (Mellman and Yarden, 2013). SUMOylation seems to be up-regulated in cancer due to e.g. overexpression of the pathway components, such as Ubc9 and some E3 ligases (Seeler et al., 2017). Interestingly, it has been reported that the SUMOylation site is essential for leukaemic transformation mediated by PML-RARalpha in acute promyelocytic leukaemia (Zhu et al., 2005). Moreover, a germline variant of melanoma lineage-specific microphthalmia-associated transcription factor (MITF), MITF-E318K, increases predisposition to sporadic melanoma and renal cell carcinoma. This mutation disrupts SUMOylation site of MITF and results in increased transcriptional activity promoting tumourigenic properties in experimental models, but no changes

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in trafficking were observed (Bertolotto et al., 2011; Yokoyama et al., 2011). However, despite extensive sequencing efforts, the evidence is so far lacking for ubiquitin or SUMO binding sites to be systematically targeted by somatic mutations in cancers. Development and research efforts using drugs that specifically inhibit the conjugation pathway activity and components will help to understand the role of SUMO and non-proteolytic functions of Ubiquitin in different contexts. Conclusions Emerging evidence strongly implicates both ubiquitination and SUMOylation in the regulation of intracellular trafficking of several proteins at multiple levels. However, these regulation systems are highly dynamic, complex and complicated with several overlapping compensatory mechanisms and the possibility of targeting same residues with different PTMs, making it challenging to design conclusive studies. In most cases the precise convincing mechanistic insights on how this regulation of trafficking happens remain to be elucidated. In addition, the field is still lacking evidence supporting many general findings in vivo. Nevertheless, conceptually ubiquitin and SUMO have already emerged as powerful and potent regulators of essential cellular processes, including protein trafficking to control temporal and special protein functions, and these PTMs are definitely an important focus for future research. Acknowledgements The laboratory of the author has been supported by grants from the Academy of Finland, Sigrid Juselius Foundation, Finnish Medical Foundation Duodecim, Turku University Foundation, Paulo Foundation and Instrumentarium Science Foundation. The author has been supported by Pfizer, Novartis, Celgene, MSD (conference participation costs not related to the topic of this manuscript) and Roche (conference participation costs and consultant fee not related to the topic of this manuscript). Dr Johanna Ahlskog and Dr Anna Knittle are acknowledged for valuable comments regarding the manuscript.

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Roles of Ubiquitination and SUMOylation in the Regulation of Angiogenesis

18

Andrea Rabellino1*, Cristina Andreani2 and Pier Paolo Scaglioni2

1QIMR Berghofer Medical Research Institute, Brisbane City, Queensland, Australia.

2Department of Internal Medicine, Hematology and Oncology; University of Cincinnati,

Cincinnati, OH, USA.

*Correspondence: [email protected] https://doi.org/10.21775/9781912530120.18

Abstract The generation of new blood vessels from the existing vasculature is a dynamic and complex mechanism known as angiogenesis. Angiogenesis occurs during the entire lifespan of vertebrates and participates in many physiological processes. Furthermore, angiogenesis is also actively involved in many human diseases and disorders, including cancer, obesity and infections. Several inter-connected molecular pathways regulate angiogenesis, and post-translational modifications, such as phosphorylation, ubiquitination and SUMOylation, tightly regulate these mechanisms and play a key role in the control of the process. Here, we describe in detail the roles of ubiquitination and SUMOylation in the regulation of angiogenesis. Introduction The growth of new blood vessels from the existing vasculature is a process known as angiogenesis (Carmeliet, 2003; Ucuzian et al., 2010). In vertebrates, angiogenesis occurs across the entire lifespan and participates in multiple physiological processes, such as pregnancy, embryonic development and wound healing. Moreover, many diseases can promote de novo angiogenesis, a process also known as pathological angiogenesis or neoangiogenesis. In this regard, a well-known example

is tumorigenesis-induced angiogenesis, during which hypoxic and starved cancer cells activate the molecular pathways involved in the formation of novel blood vessels, in order to supply nutrients and oxygen required for the tumour growth. Additionally, more than 70 different disorders have been associated to de novo angiogenesis including obesity, bacterial infections and AIDS (Carmeliet, 2003). At the molecular level, angiogenesis relays on several pathways that cooperate in order to regulate in a precise spatial and temporal order the process. In this context, post-translational modifications (PTMs) play a central role in the regulation of these events, influencing the activation and stability of many growth factors, membrane receptors and downstream signalling effector molecules. Here, we will focus on the role of ubiquitination and SUMOylation in the regulation of angiogenesis. Molecular basics of angiogenesis Blood vessels arise from endothelial precursor cells, in a process known as vasculogenesis. Further stabilization of the new blood vessels, including their expansive growth and the formation of collateral bridges is known as angiogenesis (Carmeliet, 2003). During angiogenesis, a dynamic

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and complex crosstalk occurs between endothelial cells and the extracellular matrix in a tightly regulated manner in order to promote endothelial cells proliferation and differentiation, cytoskeletal reorganization and cell migration, and the formation of novel vessels (Carmeliet, 2003; Huang and Bao, 2004; Muñoz-Chápuli et al., 2004; Ucuzian et al., 2010). Endothelial cells, fibroblasts, platelets, inflammatory cells and cancer cells (Ucuzian et al., 2010) can all act as sources of angiogenic factors. Key pro-angiogenic factors are the Vascular Endothelial Growth Factors (VEGF1–5) and their receptors (VEGFR1, VEGFR2 and VEGFR3), the Placental Growth Factors (PlGFs), the Fibroblast Growth Factors (FGF1 and FGF2) and FGF receptors (FGFR1–4), the Transforming Growth Factor (TGF-β) family, the Tumour Necrosis Factor (TNF-α), the family of the Angiopoietins (ANG1 and ANG2) and the TIE-1 and -2 receptors, Ephrins and Leptins (Carmeliet, 2003; Huang and Bao, 2004; Ucuzian et al., 2010). On the other hand, anti-angiogenic factors include the Thrombospondins (TSP1–4 and TSP-5/COMP), Angiostatins and Endostatins (Huang and Bao, 2004). Moreover, other players may differentially contribute to the control of angiogenesis, like the Matrix Metalloproteinases (MMPs), Integrins, and the extracellular matrix (ECM) (Kessenbrock et al., 2010). These factors activate several downstream signalling pathways. For example, VEGF, and similarly FGF, mainly activate the ERK/MAPK pathway (Larsson et al., 1999; Cross et al., 2000; Wu et al., 2000), leading to the transcription of master genes involved in cell proliferation, such as MYC, ELK-1, FOS, etc. (Muñoz-Chápuli et al., 2004). On the other end, VEGF can also act independently of the ERK/MAPK cascade by activating other pathways such as the STAT signalling (Muñoz-Chápuli et al., 2004). Interestingly, additional stimuli can cooperate with angiogenic factors. Accordingly, Nitric Oxide (NO) is able to potentiate the VEGFdependent activation of the angiogenic pathways (Donnini and Ziche, 2002). VEGF also directly controls the migration of endothelial cells during angiogenesis activating the RHO GTPases RHO and RAC, which are required for cell motility and the formation of focal adhesions (Soga et al., 2001a,b). Moreover, other factors, such as the Protein Kinase C (PKC) (Yamamura et al., 1996), or the receptor NOTCH (Hellström et al., 2007),

can regulate cell migration in response to VEGF. On the other hand, anti-angiogenic factors such as the Endostatins are potent inhibitors of endothelial cells migration counteracting the formation of focal adhesions (Dixelius et al., 2003; Eriksson et al., 2003). Typically, during endothelial cell migration, cell proliferation is enhanced, while apoptosis is repressed. Generally, the apoptotic signals that regulate angiogenesis and the fate of endothelial cells are mediated by TNF-α and TGF-β signalling (Polunovsky et al., 1994; Choi and Ballermann, 1995). During angiogenesis, however, apoptotic pathways are inhibited by the crosstalk between Integrins, VEGF and FGF cascades that converge toward the activation of the AKT pathway (Gerber et al., 1998). Other signalling pathways involved in angiogenesis include WNT signalling (Dufourcq et al., 2002), and the pathways activated by cytokines, such as Pleiotrophin and Midkine (Stoica et al., 2001, 2002), and oestrogens (Hyder et al., 1996). Because hypoxia is an important factor in angiogenesis, also Hypoxia-Inducible Factors (HIFs) play a fundamental role in neo-angiogenesis during tumour development (Pugh and Ratcliffe, 2003). HIF is a basic helix–loop–helix heterodimeric transcription factor that under hypoxic condition binds to hypoxic response elements (HREs) of the DNA inducing the transcription of a series of hypoxia-related genes, many of which are involved in angiogenesis (Semenza, 2000; Wenger, 2002). For example, VEGF transcription is directly upregulated by HIF activity in hypoxic conditions (Levy et al., 1998; Pugh and Ratcliffe, 2003; Zhang et al., 2012; De Francesco et al., 2013). Accordingly, Hif-1α knock out mice show abnormal vascular development and embryonic lethality (Maltepe et al., 1997; Kotch et al., 1999). PTMs in angiogenesis PTMs are a series of covalent modifications that occur following protein synthesis, and regulate protein activity, turnover and/or localization. The most common PTMs include phosphorylation, acetylation, glycosylation, ubiquitination and SUMOylation. Every PTM is strictly regulated by several molecular mechanisms and feedback loops. Interestingly, every single PTM described so far participates in the regulation and control

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of angiogenesis (Rahimi and Costello, 2015). In this chapter, we will focus on ubiquitination and SUMOylation, and will describe how these PTMs work and impact angiogenesis. Ubiquitination and SUMOylation in angiogenesis Ubiquitination and SUMOylation are PTMs that regulate the activity and fate of a plethora of proteins (Clague et al., 2015; Hendriks et al., 2015). Both ubiquitination and SUMOylation consist of the covalent binding of a small protein modifier (ubiquitin, Ub hereafter, or Small Ubiquitin-like Modifier, SUMO hereafter) to one or multiple lysine (K) residues of a target protein. Both processes require three consecutive steps (Fig. 18.1), sequentially catalysed by E1, E2 and E3 ligases (Swatek and Komander, 2016; Rabellino et al., 2017). While for ubiquitination, a variety of E1–3 ligases are known, for SUMOylation, only one E1 (SAE1/2) and one E2 (UBC9) are known, and only few classes of E3 ligases have been described

Figure 18.1 Ubiquitination and SUMOylation are reversible PTMs occurring through an E1, E2 and E3 enzymatic cascade. Ubiquitination and SUMOylation consist in the binding of either Ub or SUMO (Ub/S in the figure) modifiers to a final target protein. This process occurs trough a sequential enzymatic cascade involving E1, E2 and E3 ligases. The last step of the reaction is usually facilitated by an E3 ligase that promotes the interaction between the E2 ligase and the target protein to be modified. Both ubiquitination and SUMOylation are reversible processes: specific de-ubiquitinase and de-SUMOylase enzymes (DUBs) remove Ub/SUMO from the target protein.

so far, including RanBP2, PC2, TOPORS and the PIAS family (Rabellino et al., 2017). Both ubiquitination and SUMOylation start with the attachment of a single Ub or SUMO to the target protein: these mono-ubiquitination/ SUMOylation events have several repercussions on the fate of the target. Moreover, both Ub and SUMO often form complex branched chains, and the complexity and/or the length of the chains will determine the fate of the modified-target. For example, Ub contains seven K residues that can be ubiquitinated, thus participating to the formation of complex and branched Ub chains (Kim et al., 2011; Wagner et al., 2011). Owing to the presence of multiple SUMO paralogs, the SUMOylation machinery is more complex than ubiquitination. In vertebrates, five different SUMO genes exist and they encode for 5 different SUMO proteins (SUMO1–5): SUMO1, SUMO2 and SUMO3 are ubiquitously expressed, while SUMO4 and SUMO5 are tissue specific and not well characterized yet (Guo et al., 2004; Liang et al., 2016). In particular, SUMO2 and SUMO3 are 97% alike, however, they share only 50% homology with SUMO1 (Saitoh and Hinchey, 2000). Moreover, SUMO1 cannot be SUMOylated due to the lack of an internal acceptor K. Therefore, SUMO1 is not able to form SUMO chains and it is considered a SUMO-chain terminator (Matic et al., 2008). Both ubiquitination and SUMOylation are reversible modifications, and specific de-ubiquitination and de-SUMOylation enzymes (DUBs) are able to cleave Ub and SUMO from a modified protein (Wing, 2003; Yeh, 2009) (Fig. 18.1). Although they share a very similar enzymatic cascade, ubiquitination and SUMOylation play different roles in several cellular processes. The main function of ubiquitination is to target proteins for their proteasome-dependent degradation (Swatek and Komander, 2016). However, depending on the size and the level of complexity of the Ub chain(s), the outcome can be different: some evidences indicate that multiple short- or branched-chains are more prone to induce protein degradation, while a single chain or a mono-ubiquitination tags can have a major role in intracellular signalling. Ubiquitination has been linked to DNA damage repair, transcriptional regulation, autophagy, activation of kinases and signalling, and regulation of the endosomal compartments during their internalization

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( Johnson, 2002; Sun and Chen, 2004; Grumati and Dikic, 2018). Similarly, SUMOylation has been associated to many important cellular functions, such as nuclear trafficking, DNA transcription, DNA damage repair, regulation of the cell cycle, and innate immunity (Flotho and Melchior, 2013). Interestingly, ubiquitination and SUMOylation often cooperate. This is the case of DNA damage repair, where ubiquitination and SUMOylation tightly control the activity of the DNA damage repair machinery (Galanty et al., 2009; Morris et al., 2009). Alternatively, ubiquitination and SUMOylation cooperate to induce protein degradation, as in the case of PML and its oncogenic counterpart PML-RARA, where the SUMOylated PML is degraded after the specific ubiquitination of its SUMO chain (Lallemand-Breitenbach et al., 2008; Rabellino et al., 2012; Rabellino and Scaglioni, 2013). Finally, ubiquitination and SUMOylation can counteract each other’s function, as for MYC, where its SUMOylation inhibits the interaction with the ubiquitination machinery (Rabellino et al., 2016). SUMO-1 and the regulation of endothelial cells SUMO proteins are evolutionary conserved across the whole eukaryotic kingdom and play important role in every aspect of cell physiology, including angiogenesis (Flotho and Melchior, 2013). It has been shown that SUMO1 expression in porcine aortic endothelial cells (PAECs) promotes cell proliferation, cell migration, and resistance to apoptosis, in a SUMO1-dose-dependent manner. Importantly, expression of SUMO1 improves the ability of the endothelial cells to form tubes and branching points, underlying its role in angiogenesis. Accordingly, similar observations were also obtained by studying the SUMO1 knock in mouse model, which exhibits a higher neo-vasculogenesis capacity than the control counterpart (Yang et al., 2013). Taken together, these data indicate that SUMO1 is directly involved in the regulation of endothelial cells during angiogenesis. It is worth noting that it has been established that SUMO2 and SUMO3 can compensate for SUMO1 functions (Evdokimov et al., 2008). Based on these observations it will be interesting to determine whether SUMO2/SUMO3 can compensate for the role of

SUMO1 in angiogenesis or whether SUMO1 is indispensable for this process. The regulation of VEGFR by ubiquitination and SUMOylation One of the most important factors involved in angiogenesis is VEGF and its associated receptors, VEGFRs. Particularly, VEGFR2 is a major key player in both physiological and pathological angiogenesis and it is massively regulated by PTMs, including phosphorylation, ubiquitination and SUMOylation. In particular, the binding of VEGF to VEGFR2 causes the activation of the receptor through multiple phosphorylation events, followed by its ubiquitination and internalization via clathrin-mediated/endosomal structures (Duval et al., 2003; Ewan et al., 2006; Bruns et al., 2010). It has been shown that VEGFR2 ubiquitination is required of its internalization, and once internalized, the receptor can be degraded through the lysosomes or can be recycled back to the plasma membrane (Bruns et al., 2010; Jopling et al., 2014). Interestingly, it has been recently reported that VEGFR2 can be ubiquitinated and degraded also in a VEGF-independent manner: in this case, the E1 ubiquitin-activating enzyme UBA1 controls the basal levels of VEGFR2 as well as its activity (Smith et al., 2017). These findings suggest that ubiquitination can independently regulate the availability of the VEGFR2 receptor during angiogenesis. Finally, the balance between the ubiquitinated and de-ubiquitinated status of VEGFR has also very important repercussions on endothelial cells during angiogenesis. Lately, it has been demonstrated the de-ubiquitinating enzyme USP8 plays a central role in the regulation of this balance. Accordingly, USP8 modulates the trafficking of VEGFR2 through the endosome and lysosome compartments regulating the degradation of the receptor (Smith et al., 2016). Based on the studies summarized here, it is clear that ubiquitination plays a major role in the regulation of VEGFR signalling and trafficking in angiogenesis. Interestingly, a study using a knock out mouse model of the de-SUMOylase SENP1 has described that also SUMOylation regulates the intracellular trafficking of VEGFR (Zhou et al., 2018). In particular, it has been demonstrated that SENP1 protein levels increase in

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vascular endothelial cells in response to ischaemia. Further analyses have shown that SENP1 knock down in endothelial cells leads to an increase of the SUMOylation levels of VEGFR2, and to an impaired VEGFR2-dependent angiogenic signalling. Specifically, the SUMOylation of residue K1270 in VEGFR2 causes the receptor to accumulate in the Golgi compartment reducing its localization on the cell membrane (Zhou et al., 2018). Accordingly, analyses performed in diabetic mouse models, indicated that SENP1 expression was drastically reduced leading to an increase of VEGFR2 SUMOylation and inhibition of its signalling. All together, these data indicate that SUMOylation inhibits VEGFR2-dependent angiogenesis (Fig. 18.2), suggesting that the balance between the SUMOylated and non-SUMOylated VEGFR2 dictates its activation during angiogenesis (Zhou et al., 2018). Interestingly, SUMOylation can also indirectly control VEGFR by regulating its gene expression. In this context, it has been reported that the master regulator of lymphangiogenesis PROX1 induces VEGFR expression in a SUMO-dependent manner (Pan et al., 2009). Based on these evidences, we conclude that during angiogenesis, SUMOylation can positively control the activity of VEGFR by regulating its spatial localization and/or its gene transcription. Further analyses are needed to identify the SUMO E3 ligases that control these processes. The regulation of NOTCH during angiogenesis by ubiquitination and SUMOylation NOTCH proteins (NOTCH1–4) are transmembrane receptors that operate in many cell types and at various stages during development. After the binding of one of their ligands, NOTCH undergoes a catalytic cleavage that releases its intracellular domain. At this point, the NOTCH intracellular domain (NOTCH-ICD) translocates into the nucleus where it forms an active transcriptional complex by interacting with CSL/RBP-J and MAML (Siebel and Lendahl, 2017) (Fig. 18.2). Extensive analyses of the NOTCH signalling have underlined its pivotal role in development and angiogenesis. Accordingly, NOTCH signalling regulates the transcription of a series of genes

involved in angiogenesis, including VEGFR and Ephrins (Siekmann and Lawson, 2007; Kofler et al., 2011). These observations have been also validated in the Notch1–4 knock out mouse models, which exhibit severe defects in angiogenesis and vascular remodelling (Krebs et al., 2000). Several PTMs regulates NOTCH signalling, including ubiquitination and SUMOylation. In particular, different ubiquitin E3 ligases have been associated to its degradation. However, it is not clear whether these ubiquitination processes directly impact or not on the angiogenic role of NOTCH (Lai, 2002). So far, the only ubiquitin E3 ligase that has been linked to the angiogenic activity of NOTCH is FBW7 (Tsunematsu et al., 2004) (Fig. 18.2). Accordingly, it has been shown that the Fbw7 knock out mouse model is embryonically lethal, and embryos die at early stage with massive abnormalities in the vascular development. Particularly, Fbw7 knock out embryos show an impaired vascular remodelling in the yolk sac and brain, with the ablation of major veins formation. Molecular analyses revealed that this phenotype is caused by Notch4 accumulation in the embryos. The accumulation of Notch4 results in turn in the over-expression of Hey1, a transcriptional repressor directly regulated by Notch4 and involved in vascular development and angiogenesis. Taken together, these data highlight the role of the ubiquitin ligase FBW7 in the positive regulation of angiogenesis, by directly regulating the Notch4-Hey1 signalling pathway (Tsunematsu et al., 2004). Recently, it has also been shown that SUMOylation regulates angiogenesis by modulating NOTCH activity. For instance, in endothelial cells, the binding of the ligand DLL4 to NOTCH1 leads to VEGF transcriptional repression and to the inhibition of the VEGF signalling pathway (Fig. 18.2). This process impairs the angiogenic potential of endothelial cells (Benedito et al., 2009). Both in vitro and in vivo evidence has shown that inactivation of the de-SUMOylase SENP1 reduces cell motility, spheroid sprouting and capillary formation. This phenotype was associated to an increase of NOTCH1 activity, linking the function of SENP1 to NOTCH1 during angiogenesis. Noteworthy, the C-terminal domain of NOTCH-ICD contains several SUMO-binding motifs, and biochemical analyses confirmed that NOTCH-ICD

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Figure 18.2  Ubiquitination and SUMOylation during angiogenesis. (1) Pro-angiogenic stimuli, such as a hypoxic environment, stimulate the expression of HIF-1 that in turn promotes the transcription of pro-angiogenic genes including VEGF. The level of HIF-1 depends on its ubiquitination/SUMOylation status: in normoxic or nonangiogenic conditions, the ubiquitin ligase VHL ubiquitinates HIF-1α directing it to proteasomal degradation. Moreover, SUMOylation PIASy-dependent leads to proteasome-dependent degradation of HIF-1α. On the other hand, both de-ubiquitination by ubiquitin-specific proteases (i.e. USP20) and de-SUMOylation by SENP1 are necessary to sustain HIF-1α stability and activity during angiogenesis. (2) HIF-1 leads to the transcription of genes encoding for pro-angiogenic factors such as VEGF, FGF, TGF-β, ANG1–2, and Ephedrines. These proangiogenic factors bind to the corresponding receptors exposed on the vascular endothelial cells (VEGFR2, FGFR, TIE1–2, EPHs). (3) The angiogenic factors gradient induces the migration of specialized endothelial cells (tip cells) that will begin the sprouting of new vessels. De-SUMOylation of VEGFR2 by SENP1 is required for angiogenesis, while SUMOylation of VEGFR2 promotes its degradation. However, the specific SUMO E3 ligase of VEGFR2 is still unknown. VEGFR, TIE1–2 and EPHs, are directed to degradation by c-CBL-dependent ubiquitination in non-angiogenic conditions. Under normoxic conditions also VEGF is targeted for degradation by VHL-dependent ubiquitination. (4) Endothelial progenitors differentiate into proliferative stalk cells that build up the main body of the new vessels. (5) To stop the process of sprouting and tube formation, VEGF induces the tip cells to secrete DLL4 ligand that will bind to NOTCH receptor on stalk cells. Activation of NOTCH, and its cleavage into NOTCH-ICD followed by its translocation into the nucleus, leads to VEGFR2 transcriptional repression thereby suppressing endothelial proliferation. SUMOylation of NOTCH1 is required for its cleavage into NOTCH-ICD contributing to the anti-angiogenic activity of NOTCH. While SENP1 is responsible for the deSUMOylation of NOTCH, its specific SUMO E3 ligase has not been identified yet. Ubiquitination of NOTCH by FBW7 causes its inhibition and degradation.

is indeed SUMOylated on three residues (K2049, K2150 and K2252). Moreover, it has been shown that in endothelial cells, NOTCH-ICD exists predominately in its SUMOylated form and that SUMOylation of NOTCH1 is necessary for the cleavage and the formation of NOTCH-ICD upon DLL4 activation. Furthermore, SUMOylation

increases NOTCH-ICD transcriptional activity and half-life, potentiating its anti-angiogenic signal. These data indicate that SUMOylation is a fundamental step for the positive regulation of NOTCH1 during angiogenesis. According to this hypothesis, SENP1 interacts with NOTCH1 and regulates its level of SUMOylation, modulating

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its anti-angiogenic activity (Zhu et al., 2017). Interestingly, the SUMO E3 ligase involved in the SUMOylation of NOTCH1 has not been identified yet and further analyses are needed in order to address this topic. Noteworthy, SENP1 activity has also been directly correlated to erythropoiesis, where SENP1dependent de-SUMOylation of GATA1 is required during embryonic erythropoiesis (Yu et al., 2010). The VEGFR and NOTCH converging angiogenic signalling is regulated by ubiquitination Recent studies have demonstrated that the homeostasis between pro-angiogenic and anti-angiogenic signalling in endothelial cells is maintained by the balance between VEGFR and NOTCH signalling (Hellström et al., 2007; Lobov et al., 2007; Suchting et al., 2007; Benedito et al., 2009; Sakaue et al., 2017). In order to identify proteins involved in VEGFR activation that can also impact NOTCH signalling, human umbilical vein endothelial cells (HUVECs), a well-established cellular model used to study angiogenesis, were intensively screened in order to identify proteins that are up-regulated upon VEGFR activation but that can also impact NOTCH signalling. The findings revealed that when HUVEC cells are stimulated with VEGF, the zinc finger protein BAZF is up-regulated, leading to the induction of filopodia, cell elongation and the formation of a cellular network typical of angiogenesis. Accordingly, BAZF also negatively controls NOTCH signalling pathway, promoting VEGFdependent angiogenesis. Mechanistically, BAZF interacts with the NOTCH signalling factor CBF1 in a VEGFR-dependent way. Indeed, BAZF binding suppresses the transcriptional activity of CBF1 by releasing it from the promoters of the target genes. In addition, BAZF induces the ubiquitination of CBF1, targeting it for cytoplasmic translocation and proteasomal degradation. Further analyses showed that BAZF mediates the formation of a complex between CBF1 and the ubiquitin E3 ligase CUL3, with this effect being triggered by VEGFR activation. This finding indicates that VEGFdependent angiogenesis induces CUL3-dependent ubiquitination and degradation of CBF1 using BAZF as mediator. Accordingly, it has been shown that Bazf knock out mice suffer from angiogenic defects, up-regulation of the Notch signalling

during development, and impaired wound healing during adulthood (Ohnuki et al., 2012). Taken together, these data demonstrate that the ubiquitination machinery is able to regulate simultaneously pro- and anti-angiogenic factors in order to guarantee a fine-tuning of a complex mechanism such as angiogenesis. Hypoxia-induced angiogenesis Hypoxia-induced angiogenesis is a well-established hallmark of cancer (Hanahan and Weinberg, 2011). Accordingly, HIF-1, the master regulator of hypoxia, is up-regulated in several human cancers, and it associates with poor prognosis (Semenza, 2012). Interestingly, also HIF-1 is massively regulated by PTMs including ubiquitination and SUMOylation. HIF-1 is a heterodimeric protein composed by the HIF-1α and HIF-1β subunits. While HIF-1β is constitutively expressed, HIF-1α is tightly regulated by oxygen availability. Under hypoxic conditions, HIF-1α translocates from the cytosol to the nucleus where it interacts with HIF-1β promoting the transcription of hypoxic genes, including VEGFR (Eguchi et al., 1997). It has been established that, in normoxic conditions, HIF-1α expression is usually kept at undetectable levels. Accordingly, oxygen induces HIF-1α poly-ubiquitination and degradation by the ubiquitin E3 ligase complex PHD/ VHL/VBC (Masoud and Li, 2015) (Fig. 18.2). Even though other pathways contribute to regulate HIF-1α stability (for example by regulating its mRNA levels or its translation), HIF-1α ubiquitindependent degradation represents the major control mechanism. The mechanism of HIF-1α regulation by the ubiquitination machinery has been extensively elucidated. Briefly, in normoxic conditions, the proline residues P402 and P564 of HIF-1α are hydroxylated by the dioxygenases PHD1–3 in an oxygen-dependent way (Epstein et al., 2001; Ivan et al., 2001). In turn, this PTM activates the ubiquitination of HIF-1α by VHL, leading to its proteasome-dependent degradation (Maxwell et al., 1999; Ohh et al., 2000; Tanimoto et al., 2000). Furthermore, HIF-1α levels can be regulated by mechanisms independent from the classic PHD/VHL machinery. For example, the chaperone protein HSP90 protects HIF-1α from degradation. It has been shown that RACK1 mediates the dissociation of HSP90 from HIF-1α, which is in turn recognized by the ubiquitin ligase Elongin-B/C and

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degraded (Liu et al., 2007). Alternatively, the kinase PLK3 regulates HIF-1α levels during hypoxia by phosphorylating the serine residues S576 and S657 thereby inducing the degradation of HIF-1α (Xu et al., 2010a). Additionally, the transcription factor TAp3 is able to directly interact with HIF1α, promoting the recruitment of the ubiquitin ligase MDM2, followed by its poly-ubiquitination and degradation in an oxygen-dependent manner (Amelio et al., 2015). Since ubiquitination is largely involved in the control of HIF-1α levels, de-ubiquitinating enzymes play an equally important role in maintaining the physiological level of HIF-1α. In this context, the de-ubiquitinating enzyme USP20 is able to interact with HIF-1α and to regulate the transcription of downstream genes of HIF-1α such as VEGF (Li et al., 2005) (Fig. 18.2). Similarly, other de-ubiquitinating enzymes such as USP8 and UCHL1 were shown to modulate HIF-1α levels and stability (Troilo et al., 2014; Goto et al., 2015). Owing to the major role that HIF-1 plays in tumour-induced angiogenesis, the development of drugs able to promote its degradation has gained a lot of interest. In this scenario, the small molecules SCH66336 and Apigenin disrupt the interaction of HSP90 with HIF-1α, therefore inducing HIF-1α degradation (Osada et al., 2004; Han et al., 2005; Melstrom et al., 2011). Moreover, other small molecules were found able to activate or increase the activity of the ubiquitin ligase complex. This is the case of the small molecule LW6, which increases the expression of VHL with a mechanism that has not been clarified yet (Lee et al., 2010). Similar to ubiquitination, also SUMOylation has been suggested to regulate HIF-1α levels, however, it is not clear whether SUMOylation increases or decreases HIF-1α stability. HIF-1α SUMOylation on the residues K391 and K477 was described for the first time in 2004, when it was proposed that the binding of SUMO1 to HIF-1α promotes its stabilization and transcriptional activity (Bae et al., 2004). Similarly, the protein RSUME can SUMOylate HIF-1α, increasing its stability. RSUME is upregulated on hypoxic stress and promotes SUMO conjugation by interacting with UBC9 (CarbiaNagashima et al., 2007). Furthermore, the SUMO E3 ligase CBX4 increases hypoxia-induced VEGF expression and angiogenesis by SUMOylating HIF-1α on the residues K391 and K477, increasing

its transcriptional activity. These results were also corroborated by the observation that CBX4 expression positively correlates with the level of VEGF expression, angiogenesis and over-all survival in hepatocellular carcinoma patients (Li et al., 2014). Despite these observations, other results indicated that the binding of SUMO1–3 to HIF-1α negatively regulates its transcriptional activity without altering its half-life (Berta et al., 2007). A different and complex scenario has been reported about the effects of SENP1-dependent de-SUMOylation of HIF-1α. It has been shown that the de-SUMOylation of HIF-1α by SENP1 inhibits the interaction between HIF-1α and VHL, resulting in the stabilization of HIF-1α, therefore suggesting that HIF-1α SUMOylation promotes its degradation (Cheng et al., 2007). These data were confirmed by Senp1 knock out mice, which showed a lower induction of HIF-1 signalling (Xu et al., 2010b). Similarly, SENP1 stabilizes HIF-1α levels and downstream signalling during myocardial ischaemia/reperfusion injury (Gu et al., 2014). Taken together, these results indicate that de-SUMOylation plays a pivotal role in maintaining HIF-1α levels during angiogenesis and suggesting that SUMOylation might directly signal for the ubiquitination/degradation of HIF-1α. However, this evidence contradicts the hypothesis that SUMOylation is required for maintaining the stability of HIF-1α, and additional work is needed to solve these inconsistencies. Whether PIAS family members contribute to regulate HIF-1 activity is also controversial. PIAS proteins (PIAS1–3 and PIASy) are SUMO E3 ligases involved in the regulation of several cellular functions, including angiogenesis, and they have been also associated to human malignancies (Rabellino et al., 2017). It has been reported that in hypoxic condition, PIASy interacts with HIF-1α triggering its SUMOylation thereby promoting its degradation (Kang et al., 2010) (Fig. 18.2). Opposite results have been reported regarding the interaction of PIAS3 with HIF-1α. It has been shown that PIAS3 positively regulates HIF-1α transcriptional activity, however, this function is independent of the SUMO E3 ligase activity of PIAS3 (Nakagawa et al., 2016). Taken together, these controversial observations suggest that different PIAS family members might have different roles in HIF-1α regulation and activity. These

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controversies need to be addressed in more detail in the future. Role of the SUMO E3 ligase PIAS1 in angiogenesis The PIAS SUMO E3 ligases have been directly associated to angiogenesis independently of their ability to regulate HIF-1. In particular, the role of PIAS1 in angiogenesis has been recently characterized using the Pias1 knock out mice model (Constanzo et al., 2016). It has been demonstrated that ablation of Pias1 in mice is embryonically lethal due to major defects in the vascular plexus of the yolk sac and thus in angiogenesis and erythropoiesis. Accordingly, Pias1 null mice embryos showed a significant reduction in blood vessel size and branching, which correlates with a low expression of the endothelial activation markers Angp2 and Vcam-1 (Constanzo et al., 2016). Interestingly, Vegfr levels were up-regulated in the yolk sac of Pias1 null mice suggesting a compensatory mechanism required for the activation of angiogenesis. These data were confirmed by in vitro experiments performed in HUVEC cells. Accordingly, expression of PIAS1 in this endothelial cellular model induces the expression of angiogenic markers and the down-regulation of anti-angiogenic genes, while PIAS1 knock down reduces the ability of HUVECs to form de novo tubes and branching structures (Constanzo et al., 2016). These data underline the role of PIAS1 in regulating angiogenesis during embryogenesis, however it has not been described whether its function relies on its SUMO E3 ligase activity. This issue was investigated by a recent work in which PIAS1 was described as the SUMO E3 ligase of MYC (Rabellino et al., 2016). The transcription factor MYC is a master transcription regulator involved in several cellular functions, including angiogenesis, and it is causally implicated in several human malignancies (Baudino et al., 2002; Dang, 2012). It has been described that the PIAS1dependent SUMOylation of MYC increases its half-life and its transcriptional activity. Accordingly, analyses of Pias1 null mice recapitulate the characteristics of the Myc null mouse model, showing a developmentally delayed and hypoplastic yolk sac, lacking the characteristic microvillar structures of the vascular plexus (Rabellino et al., 2016). Taken together, these data strongly suggest that PIAS1 plays a fundamental role in angiogenesis, and this

activity is likely due to its SUMO E3 ligase activity. Further studies will shed more light on the role of this SUMO ligase. PML in angiogenesis The promyelocytic leukaemia gene PML was described for the first time as product of the chromosomal translocation t(15;17)(q24;q21) in acute promyelocytic leukaemia (APL) (Piazza et al., 2001). Soon, it became clear that PML is involved in the positive regulation of several tumour suppressive functions and other cellular processes, including angiogenesis (Salomoni and Pandolfi, 2002; Rabellino and Scaglioni, 2013). PML is massively regulated by several PTMs, including ubiquitination and SUMOylation, which influence its activity, functions and regulation, including the formation of the functional units of PML, known as PML Nuclear Bodies (PML-NBs) (Bernardi and Pandolfi, 2007; Rabellino and Scaglioni, 2013). It has been shown that PML negatively controls angiogenesis through the inhibition of HIF-1a translation by repressing mTOR activity (Bernardi et al., 2006). These findings elegantly described a new role of PML in controlling angiogenesis, however, whether PML ubiquitination or SUMOylation take part of this process is not clear. In most recent years, however, a new layer of complexity regarding how PML regulates the mTOR/HIF-1α pathway has been added. PML degradation is tightly regulated by a series of PTMs, including phosphorylation, SUMOylation and ubiquitination that occur in a precise spatial and temporal order (Scaglioni et al., 2006; Yuan et al., 2011; Rabellino and Scaglioni, 2013). It has been shown that under hypoxia conditions, the ubiquitin E3 ligase CUL3 substrate KLHL20 co-operates with CDK1/2 and with the isomerase PIN1 in order to induce PML ubiquitination and degradation in a HIF-1 dependent way. In this scenario, it has been also demonstrated that the KLHL20-dependent PML degradation promotes neo-angiogenesis (Yuan et al., 2011), pointing toward anti-angiogenic properties of PML. Furthermore, it has been shown that this mechanism is counteracted by SCP phosphatases, which de-phosphorylate PML blocking the KLHL20-dependent degradation, which in turn will inhibit HIF-1 signalling in a mTOR-dependent way (Lin et al., 2014). Interestingly, it has also been shown that PML-NBs are the site of the interaction

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between CUL3 and CBF1 during the regulation of the VEGF-dependent NOTCH signalling (Ohnuki et al., 2012), suggesting that PML and PML-NBs might regulate angiogenesis through several pathways and mechanisms. Despite the fact that SUMOylation has not been directly implicated in this process, based on the data available to date, we speculate that SUMOylation might be critical for the correct outcome of the process. Finally, the role of PML in the inhibition of angiogenesis has been also demonstrated by its role in the positive-regulation of the anti-angiogenic factor Interferon-α (INF-α). Degradation of PML massively reduces the angiostatic effects of INF-α. Interestingly, INF-α stimulation leads to the induction of PML, which in turn activates STAT1 and STAT2 anti-angiogenic activity by promoting STAT1/2 ISGylation (Hsu et al., 2017), an ubiquitin-like modification which functions and regulation are still largely unknown (VillarroyaBeltri et al., 2017). The regulation of TIE2 and FGFR by c-CBL ubiquitination The TIE2 receptor belongs to the family of the tyrosine kinase receptor (RTK) and is predominantly expressed on the surface of endothelial and hematopoietic cells (Dumont et al., 1992). The binding of TIE2 to its ligand ANG1 activates a downstream signalling cascade that positively regulates angiogenesis ( Jones et al., 2001). Accordingly, Tie2 null mice die at early embryonic stage due to the lack of the formation of the capillary plexus and severe heart defects (Dumont et al., 1994). It has been shown that ubiquitination regulates the turnover of TIE2 in a ligand-specific fashion. Indeed, the binding of ANG1 to TIE2 is sufficient to induce the activation of the receptor and its subsequent ubiquitination by the Ub E3 ligase c-CBL (Wehrle et al., 2009). FGFR is another RTK that plays an essential role in angiogenesis (Yang et al., 2015). Similar to TIE2, ubiquitination regulates the turnover of FGFR and modulates its downstream signalling. Also, in this case, the ubiquitin E3 ligase involved in the ubiquitination of FGFR is c-CBL (Wong et al., 2002; Haugsten et al., 2008) (Fig. 18.2). While these data indicate that ubiquitination of TIE2 and FGFR is necessary to regulate them during angiogenesis, it is not clear whether

SUMOylation may modulate the activity and the turnover of these receptors. Ubiquitination and de-ubiquitination of the WNT signalling in angiogenesis The WNT signalling pathways control a wide spectrum of cellular functions, including cell proliferation and migration. WNT pathways can be classified in canonical/β-catenin-dependent and non-canonical pathways, and they both regulate and control angiogenesis. Accordingly, both in vitro and in vivo studies demonstrated that WNT and its Frizzled receptors regulate the migration of endothelial cells during angiogenesis (Zerlin et al., 2008). In the canonical pathway, WNT up-regulates the level of cytosolic β-catenin by inhibiting its ubiquitin-dependent degradation (Li et al., 1999). The WNT-dependent accumulation of β-catenin promotes the nuclear translocation of β-catenin where it activates the transcription of genes involved in cell growth regulation and pro-angiogenic genes, such as VEGF and IL-8 (Tetsu and McCormick, 1999; You et al., 2002; MacDonald et al., 2009). Hence, the regulation of the β-catenin is critical, and ubiquitination plays a central role. It has been shown that c-CBL induces the ubiquitination of nuclear β-catenin thereby promoting its degradation, therefore negatively regulating angiogenesis. Interestingly, the re-localization of c-CBL from the cytoplasm to the nucleus it is induced by WNT (Chitalia et al., 2013), suggesting the activation of a feedback mechanism that controls this pathway. The re-localization of c-CBL is induced by the WNT-dependent phosphorylation of c-CBL on the tyrosine Y731, which promotes c-CBL dimerization, binding to the β-catenin and the nuclear re-localization (Shivanna et al., 2015). Another ubiquitination-dependent regulation of the WNT signalling during angiogenesis has been described for the de-ubiquitinase Gumby. Noteworthy, the Gumby mouse mutants show severe angiogenic impairment during embryogenesis (Rivkin et al., 2013). Accordingly, Gumby knock out embryos die at early stage due to the insufficient development of the branching of the vascular system. It was previously reported that Gumby interacts with DVL2, which also plays an important role in WNT signalling (Rual et al., 2005). Further analyses performed in both in vitro and in vivo setting, indicated that Gumby negatively

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regulates WNT activity in endothelial cells, compromising their angiogenic potential (Rivkin et al., 2013). Studies reported that SUMOylation of the co-repressors TBL1 and TBLR1 led to the activation of the WNT signalling in a β-catenin-dependent manner. Conversely, the SENP-1-dependent de-SUMOylation of TBL1 and TBLR1 inhibits this mechanism (Choi et al., 2011). Similarly, it has been demonstrated that the SUMO E3 ligase PIASy SUMOylates the WNT downstream effector TCF4, enhancing the β-catenin-dependent transcriptional activity of TCF4 (Yamamoto et al., 2003). These findings suggest that SUMOylation may play a role in the regulation of the WNT signalling, however, its role in this context has not been elucidated yet. Ephrins regulation during angiogenesis The binding of the membrane ligand Ephrins to their receptors initiates a series of downstream signalling that regulate the fate of endothelial cells during angiogenesis. Ephrins receptors are RTKs subdivided in two subclasses, EPHA and EPHB, activated by the ligands EphrinA and EphrinB, respectively. In vertebrates, a total of ten EPHA and six EPHB are expressed, and several in vitro and in vivo studies have underlined the role of EPH receptors and their ligands in the regulation of angiogenesis (Pasquale, 2005). Because Ephrin ligands are anchored to the cell membrane, the interaction with their receptors requires a cellto-cell interaction. Once the activated receptor induces downstream signalling cascade, the stimulus is extinguished through a process that includes the receptor internalization and its degradation via ubiquitination (Litterst et al., 2007). Similar to other RTKs involved in angiogenesis, it has been shown that c-CBL is the ubiquitin E3 ligase responsible for EPHB receptor ubiquitination (Fasen et al., 2008) (Fig. 18.2). To date, however, there are no evidences that SUMOylation is involved in the regulation of Ephrins. The role of extracellular Ub in the regulation of angiogenesis Extracellular Ub regulates immune responses during inflammation, organ injuries and fibrosis, and elevated plasma levels of Ub correlate with several human pathologies (Asseman et al.,

1994; Takagi et al., 1999; Majetschak et al., 2005; Sujashvili, 2016). It has been also shown that the extracellular Ub promotes angiogenesis. Accordingly, using cardiac micro-vascular endothelial cells (CMECs), which is the major cell type involved in cardiac angiogenesis, it has been demonstrated that extracellular Ub promotes the expression of VEGFR, thereby triggering cytoskeletal rearrangement, cell migration and tube formation (Steagall et al., 2014). These observations raise several questions regarding the molecular mechanisms by which extracellular Ub activates angiogenesis. Such aspects should be of potential interest in view of future therapeutic applications of this discovery. Conclusions and remarks Ubiquitination and SUMOylation are PTMs that play fundamental roles in every aspect of human physiology. Here we have summarized their major roles in angiogenesis known so far. Because of the extreme significance of angiogenesis in tumour development and in other human diseases, both ubiquitination and SUMOylation might represent valuable candidate targets for the generation of new, more effective drugs for the treatment of these pathologies. In particular, even though SUMOylation has being known for more than two decades (GeissFriedlander and Melchior, 2007), it is still a fairly unknown process, and its involvement in angiogenesis regulation remains largely uncharacterized. More efforts should be made in order to shed light on this important PTM and its contribution to angiogenesis. References

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The Role of SUMOylation and Ubiquitination in Brain Ischaemia: Critical Concepts and Clinical Implications

19

Joshua D. Bernstock1,2*†, Daniel G. Ye3†, Dagoberto Estevez4, Gustavo Chagoya4, Ya-Chao Wang5, Florian Gessler6, John M. Hallenbeck2 and Wei Yang5*

1Department of Neurosurgery, Brigham and Women’s Hospital, Boston, MA, USA.

2Stroke Branch, National Institutes of Health (NIH), National Institute of Neurological Disorders

and Stroke (NINDS), Bethesda, MD, USA.

3Medical Scientist Training Program (MSTP), Baylor University, Houston, TX, USA.

4Department of Neurosurgery, University of Alabama at Birmingham, Birmingham, AL, USA. 5Department of Anesthesiology, Duke University Medical Center, Durham, NC, USA.

6Department of Neurosurgery, Goethe University Frankfurt, Frankfurt am Main, Germany.

*Correspondence: [email protected] and [email protected] †Both authors contributed equally to this work

https://doi.org/10.21775/9781912530120.19

Abstract Brain ischaemia is a severe form of metabolic stress that activates a cascade of pathological events involving many signalling pathways. Modulation of these pathways is largely mediated by post-translational modifications (PTMs). Indeed, PTMs can rapidly modify pre-existing proteins by attaching chemical or polypeptide moieties to selected amino acid residues, altering their functions, stability, subcellular localizations, or interactions with other proteins. Subsequently, related signalling pathways can be substantially affected. Thus, PTMs are widely deployed by cells as an adaptive strategy at the front line to efficiently cope with internal and external stresses. Many types of PTMs have been identified, including phosphorylation, O-GlcNAcylation, small ubiquitin-like modifier (SUMO) modification (SUMOylation), and ubiquitination. All these PTMs have been studied in brain ischaemia to some extent. In particular,

a large body of evidence has demonstrated that both global SUMOylation and ubiquitination are massively activated after brain ischaemia, and this activation may play a critical role in defining the fate and function of cells in the post-ischaemic brain. The goal of this chapter will be to summarize the current findings on SUMOylation and ubiquitination in brain ischaemia and discuss their clinical implications. SUMOylation in brain ischaemia/ stroke SUMOylation SUMOylation is a post-translational modification in which a member of the SUMO family of proteins is conjugated to lysine residues in target proteins. SUMOylation was first implicated in nuclear function pathways (Matunis et al., 1996;

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Mahajan et al., 1997). However, it has since been implicated with numerous extranuclear substrates (Martin et al., 2007). Its vital role to cell function has become evident in knockdown and knockout models of SUMO or components of this pathway, which prove fatal in eukaryotic cells (Hayashi et al., 2002; Wang et al., 2014). As a result, SUMO genes are highly conserved and widely found in protozoa, metazoa, plants, and fungi. Four isoforms have been described in mammalian cells and are designated SUMO1 to SUMO4 (Geiss-Friedlander and Melchior, 2007; Bernstock et al., 2018a). Of these four isoforms, SUMO1–3 are the best characterized in the literature. SUMO proteins are members of the ubiquitin-like proteins family, and function in an enzymatic pathway analogous to the ubiquitination pathway. In fact, despite their low homology (18% with ubiquitin and   75  kDa) were found to be reduced by 7 months, just before the age of amyloid plaque onset (Lee et al., 2014). Interestingly, β-amyloid appears to impair activitydependent SUMOylation in the brain. When Ubc9 is up-regulated, allowing for increased SUMOylation in the system, β-amyloid-induced deficits to long-term potentiation are rescued (Lee et al., 2014). A wealth of data from human, animal, and cell studies all indicate that a decrease in high molecular weight SUMO2/3 is Alzheimer’s disease-specific. This decrease is likely to be detrimental, considering that increasing SUMOylation rescues diseased long-term potentiation (Lee et al., 2014). It is interesting to speculate what the role of decreased high molecular weight SUMO2/3 proteins could mean in the diseased system. Likely, this is an indication of decreased polySUMOylation. Recall that Ubc9, the only E2 enzyme available in the mammalian SUMOylation pathway, is inhibited in some familial and sporadic cases of Alzheimer’s disease (Ahn et al., 2009; Lee et al., 2013). This certainly would reduce SUMOylation of all forms, including polySUMOylation. However, in some models of disease Ubc9 is increased, but high molecular weight SUMO2/3 is still decreased (Lee et al., 2014). There must be more occurring in Alzheimer’s disease than just inhibition of polySUMO-chain formation. Indeed, various labs have found that polySUMOylation promotes subsequent ubiquitination and degradation (Lallemand-Breitenbach et al., 2008; Mullen and Brill, 2008; Tatham et al., 2008). Therefore, it is plausible that long polySUMO-chains are created in early stages of disease, as a means of targeting potentially toxic proteins to the proteasome, and thus early stage polySUMOylated proteins have been degraded by late stages of disease.

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Figure 25.7 β-amyloid structure and APP processing. (A) Monomeric structure of β-amyloid 1–42, (B) pentameric structure of β-amyloid 1–42, and (C) 12-mer structure of β-amyloid 1–42. Structures 1IYT, 2BEG, and 2MXU were created using the Protein Data Bank NGL Viewer. (D) Toxic β-amyloid is produced via APP cleavage. Amyloidogenic processing involves β–secretase cleavage, followed by γ–secretase cleavage. Nonamyloidogenic processing involves α–secretase cleavage followed by γ–secretase cleavage.

β-amyloid One of the amyloids present in Alzheimer’s disease is β-amyloid, which is the primary component of senile plaques (Fig. 25.7A,B,C; Crescenzi et al., 2003; Lührs et al., 2005; Xiao et al., 2015; Rose et al., 2018). While β-amyloid is the toxic amyloid and Amyloid Precursor Protein (APP) is the functional protein from which its cleaved, data suggests that APP is differentially SUMOylated in Alzheimer’s disease. The different post-translational modifications of APP lead to a change in cleavage pattern of the protein, affecting the downstream production of β-amyloid (Zhang and Sarge, 2008). APP is a type 1 transmembrane glycoprotein. Although the function of APP is currently unknown, it is believed to play a role in formation of the neuromuscular junction, and synaptic transmission, ion channel function (O’Brien et al., 2011). APP is predominantly cleaved through the non-amyloidogenic pathway in healthy brains; the protein is first cleaved by α-secretase (also called

ADAM) (Lammich et al., 1999) and then cleaved by γ-secretases (including the proteins presenilin, PEN2, APH1, niscatrin) (Xia et al., 2000). This process results in products which are thought to be neuroprotective and neurotrophic, and to prevent β-amyloid formation (Pearson and Peers, 2006). However, APP can also be processed naturally through the amyloidogenic pathway and it is the up-regulation of this processing that increases β-amyloid in AD. β-secretase first cleaves APP (Greenfield et al., 1999; Xia et al., 2000; Xu et al., 2009) followed by γ-secretase (Xia et al., 2000). β-amyloid 1–40 and β-amyloid 1–42 are the dominant products produced via amyloidogenic APP processing, but it is thought that other fragments may be produced through proteolytic degrading enzymes (Kaminsky et al., 2010) (Fig. 25.7D; adapted from Linchtenthaler, 2012). APP is SUMOylated in vitro by SUMO1 and SUMO2 at lysines 587 and 595, which are near the site of β-secretase cleavage (Nistico et al., 2014). Hippocampal neurons from Alzheimer’s

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disease patients also immunolabelled robustly with SUMO3 (Li et al., 2003). Indeed, mutating lysine at 587 or 595 to arginine produced an APP that could not be SUMOylated (Zhang and Sarge, 2008). These K587R and K595R APP mutants exhibited increased levels of β-amyloid aggregates, while overexpression of SUMO E2 enzyme decreased levels of β-amyloid (Zhang and Sarge, 2008). This finding lead researchers to speculate that SUMOylation of APP could act as a protective mechanisms against amyloidogenic processing of APP. Further studies involving the up-regulation of Ubc9 supported this hypothesis, with the resulting decrease in β-amyloid aggregate levels (Zhang and Sarge, 2008). APP SUMOylation is convoluted. For example, knockdown of SUMO2 decreased aggregate species but did not appear to have any effect on APP processing (Li et al., 2003), suggesting that perhaps there is an indirect role for SUMO2 in APP processing and β-amyloid production. The authors suggest that this may be through driving α secretase, as opposed to β secretase, cleavage of APP (Li et al., 2003). Indeed, the α secretase cleavage products had increased SUMO2-modifications (Li et al., 2003). Surprisingly, knockdown of SUMO1 or SUMO2/3 did not affect the levels of APP or β-amyloid (Dorval et al., 2007). Dorval and colleagues may have identified the critical link; overexpression of SUMO3 protein up-regulated β-secretase protein levels, likely providing the mechanism for increased β-amyloid production previously observed in the link between SUMO and β-amyloid (Dorval et al., 2007). This hypothesis is strengthened by the findings of Zhang and Sarge (2008), who found that overexpression of APP and SUMO3 lead to an increase in β-amyloid production (Zhang and Sarge, 2008). Yun and colleagues found similar results with SUMO1, detecting a direct link between SUMO1 depletion and decreased β-amyloid 1–40 levels (Yun et al., 2013). Although the extent of APP SUMOylation is unknown, polySUMOylation has been postulated. Tatham et al. (2001) immunoprecipitated APP from mouse brain and discovered that the protein was SUMOylated by both SUMO1 and SUMO2/3 (Gocke et al., 2005). While the extent of endogenous SUMOylation of APP is undetermined, research has provided insight into the importance of poly- versus monoSUMOylation of APP with

regards to β-amyloid production. HEK293T cells were transfected with APP and a SUMO3 variant which could not produce SUMO chains, but could still mono-SUMOylate (Dorval et al., 2007). MonoSUMOylation of APP resulted in an increase in β-amyloid generation, leaving the researchers to speculate that polySUMOylation may negatively regulate β-amyloid production (Dorval et al., 2007). Ubiquitination of β-amyloid has been studied using a transgenic mouse model of Alzheimer’s disease. In this study, the APPswe/PS1 mouse model was crossed to a UBB+1 mouse. The UBB+1 mutant inhibits the ubiquitin-proteasome system, and so this bigenic model allows for the study of the ubiquitin-proteasome system on β-amyloid. Looking at various ages, researchers found a significant decrease in β-amyloid deposition and in soluble β-amyloid (1–42) (van Tijn et al., 2012). The reduction in amyloid deposition was transient, only lasting until 9 months of age (van Tijn et al., 2012). Intriguingly, the animals still expressed astrogliosis and were as functionally impaired as APPswe/PS1 age-matched controls (van Tijn et al., 2012). Although complex, it appears that β-amyloid production is influenced by APP SUMOylation. Reduction of SUMO1, SUMO2, or SUMO3 leads to decreased aggregated β-amyloid (Li et al., 2003; Dorval et al., 2007; Yun et al., 2013). Furthermore, polySUMOylation may play a role in negatively regulating β-amyloid production (Dorval et al., 2007). However, it is unclear if simply an increase in available SUMO lead to the increase in peptide as opposed to the disruption of polySUMOylation. Ubiquitination of APP is less understood than SUMOylation. When the UPS is inhibited, Alzheimer’s disease-models display less soluble and insoluble β-amyloid (van Tijn et al., 2014). However, the possibilities of off-target effects are great, and the question still exists of direct APP ubiquitination. Tau The second amyloid present in Alzheimer’s disease is tau, the primary component of neurofibrillary tangles and the neuropathological hallmark of tauopathies. Functionally, tau helps to stabilize microtubules and plays an important role in axon development (Mietelska-Porowska et al., 2014). In Alzheimer’s disease and other tauopathies, tau

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becomes hyperphosphorylated. The hyperphosphorylated tau is prone to amyloid fibrillization, leading to neurofibrillary tangle formation, and to disruption of its native function in microtubule stabilization (Mietelska-Porowska et al., 2014) (Fig. 25.8; Fitzpatrick et al., 2017; Rose et al., 2018). While phosphorylation remains the best-characterized post-translational modification of tau, tau also undergoes SUMOylation and ubiquitination (Dorval and Fraser, 2006; Morris et al., 2015). Tau ubiquitination was identified by mass spectrometry of paired helical filaments isolated from Alzheimer’s disease brain (Morishima-Kawashima et al., 1993; Cripps et al., 2006) and from mouse brain tissue (Morris et al., 2015). This ubiquitination leads to downstream proteasome degradation of tau (David et al., 2002), although, tau can also be degraded through the ubiquitin-independent proteasome system (Shimura et al., 2004). Tau can also be SUMOylated. Specifically, tau is SUMOylated at lysine 340, which is within the fourth microtubule binding repeat domain (Luo et al., 2014) (Fig. 25.8B; adapted from Fitzpatrick et al., 2017). Therefore, it is hypothesized that on tau release from microtubules, the K340 residue is accessible to SUMOylation enzymes (Luo et al., 2014). Indeed, this hypothesis was tested by applying colchicine to induce microtubule depolymerization and increase the available tau pool. The available tau exhibited a significant increase in SUMOylation (Luo et al., 2014). When SUMO and human tau are overexpressed in cells, high molecular weight bands of SUMOylated tau species increase (Dorval et al., 2007; Luo et al., 2014). This increase is robust for SUMO1 and was observed for

SUMO2 and SUMO3 as well (Dorval et al., 2007; Luo et al., 2014). Furthermore, in vitro studies indicate that SUMOylation of tau elevates levels of tau phosphorylation (Neddens et al., 2018), leading to an increase in cytotoxicity. When overexpression of both tau and SUMO1 occur in HEK293T cells, tau undergoes a significant increase in phosphorylation at Thr205, Ser214, Thr231, Ser262, Ser396, and Ser404 (Yu et al., 2009). To confirm linkage of tau SUMOylation and phosphorylation, researchers inhibited protein SUMOylation using ginkgolic acid and observed reduced phosphorylation of tau (Yu et al., 2009). Another group’s findings further strengthen the link between tau SUMOylation and phosphorylation; when a SUMO-conjugation deficient tau mutant (K340R) is overexpressed, both SUMOylation and phosphorylation are markedly decreased on tau (Luo et al., 2014). One report suggests that there is a direct link between the SUMOylation/ubiquitination of tau and its state of solubility. When tau is overexpressed in HEK293T cells, SUMOylation of tau is increased while ubiquitination of tau is decreased. This coincided with an increase in tau aggregation and a decrease in tau degradation (Luo et al., 2014). Studies also indicate an important linkage between tau acetylation and ubiquitination, the importance of which is only just starting to be understood. Ubiquitination sites are often in competition for acetylation, as identified by mass spectrometry (Morris et al., 2015). Tau methylation also appears to compete for two lysine residues, increasing the complexity of modification crosstalk (Morris et al., 2015). When lysine 311 and 281 are mutated so as not to be ubiquitinated, acetylated, or methylated,

Figure 25.8  Tau structure. (A) Fibrillar tau of microtubule binding regions 3 and 4 and (B) schematic of primary structure of tau. Structure created from the Protein Data Bank NGL Viewer from 5O3T.

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microtubule binding and protein aggregation decreases, suggesting an important conformational regulatory mechanism at these sites (Morris et al., 2015). As previously mentioned, neurofibrillary tangles label robustly for ubiquitin (Drummond et al., 2018). Indeed, tau is known to be degraded by the proteasome through both ubiquitin-independent and ubiquitin-dependent processes (David et al., 2002; Cardozo et al., 2002). Data suggest that both neurofibrillary tangles and soluble tau species can be ubiquitin-labelled (David et al., 2002; Cardozo et al., 2002; Drummond et al., 2018). A particularly interesting study found a direct link between cytosolic Ubiquitin-C-terminal Hydrolase L1 (UCHL-1) and a naturally-occurring truncated variant of tau, which is cleaved at aspartic acid 421 by caspases (Corsetti et al., 2015). UCHL-1, which functions as a modulator of ubiquitin homeostasis and controls remodelling of synapses, was found to interact non-physiologically with the truncated tau fragment, contributing to the early synaptotoxicity observed in Alzheimer’s disease (Corsetti et al., 2015). In the case of tau, ubiquitination and SUMOylation appear to work in contrast. This hypothesis comes from the findings that neurofibrillary tangles probe highly positively for ubiquitin, but not for SUMO1 (David et al., 2002; Cardozo et al., 2002; Drummond et al., 2018), and cell studies that observed a link between increased tau aggregation with increased SUMOylation and decreased tau degradation with increased ubiquitination (David et al., 2002; Cardozo et al., 2002). As stated previously for general inclusion body formation, ubiquitination of the substrate drives the toxic protein to degradation. However, if toxic proteins overwhelm the system, ubiquitinated proteins may be left in inclusion bodies as a final attempt to regulate pathogenicity. This may be the case for tau and neurofibrillary tangle formation (David et al., 2002; Cardozo et al., 2002; Drummond et al., 2018). However, recent crosstalk and mutagenesis studies indicate that there may be much more complex signalling pathways at work (Morris et al., 2015). An equally confounding question is the role of SUMOylation in tau pathogenicity. Unbound tau is more likely to be SUMOylated, and this SUMOylated tau leads to an increase in

hyperphosphorylation of the protein and protein aggregation (Köpke et al., 1993; Mandelkow et al., 1994). Perhaps SUMOylation of tau is the first step on the road to fibrillization. Indeed, SUMO-conjugation could provide the critical conformational change needed to create the cross-β structure in tau. However, this hypothesis has yet to be tested. Another plausible consideration is that SUMOconjugation allows for exposure of numerous phosphorylation sites that would have otherwise been masked or helps to recruit kinases to tau for increased phosphorylation. Polyglutamine disorders Polyglutamine disorders involve the expansion of a toxic CAG stretch in disease-specific genes. CAG expansion in polyglutamine disorders is familial, with variable CAG stretches depending on genetic inheritance. On translation, the CAG stretch ultimately becomes a polyglutamine, or polyQ, stretch in the effected protein and undergoes a toxic gainof-function (Watase et al., 2002; Yoo et al., 2003). PolyQ rich proteins are prone to aggregation and can result in neuronal inclusions, as hallmarked in polyglutamine disorders (Zoghbi and Orr, 2000; Gatchel and Zoghbi, 2005). These polyglutamine pathologies cause neuronal death in specific brain regions, including the basal ganglia, cerebellum, brainstem, and spinal motor nuclei (Ross et al., 2002). Aggregation of polyQ proteins is thought to overwhelm the UPS and to compromise essential cellular functions of the machinery (Mayer et al., 1989). Ubiquitination reduces polyQ toxicity, likely by promoting degradation of toxic polyQ proteins. Indeed, when UPS functioning is accelerated, polyQ toxicity lessens (Verhoeft et al., 2002; Michalik and Van Broeckhoven, 2004). This hypothesis is further strengthened by the finding that ubiquitin ligase mutations enhance polyQ toxicity (Saudou et al., 1998; Cummings and Zoghbi, 2000; Fernandez-Funez et al., 2000) and overexpression of the E3 ubiquitin ligase, Parkin, reduces polyQ aggregation and suppresses toxicity (Tsai et al., 2003). However, there is a debate within the field whether polyQ proteins are degraded by the UPS (Venkatraman et al., 2004; Pratt and Rechsteiner, 2008; Juenemann et al., 2013). Some studies suggest that polyQ proteins can be degraded if in a soluble state (Verhoef et al., 2002;

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Kaytor et al., 2004; Michalik and Van Broeckhoven, 2004; Juenemann et al., 2013; Tsvetkov et al., 2013). Others indicate that the proteasome cannot degrade polyQ proteins (Dyer and McMurray, 2001; Jana et al., 2001; Holmberg et al., 2004; Venkatraman et al., 2004). Indeed, in various mouse models of polyQ diseases, there is a lack of UPS impairment with an increase of polyQ proteins (Bence et al., 2001; Jana et al., 2001; Bennett et al., 2005). Instead, work in animal models suggests that polyubiquitin chains on polyQ proteins causes a blockage of the UPS during degradation (Maynard et al., 2009). It is worth noting, however, that a simple blockage of the UPS may not be the only disruption to the UPS in polyQ disease; instead, proteasomal degradation and polyQ protein interaction is likely much more complicated. Beyond the general effect on the UPS, all major polyglutamine disorders have high immunostaining of neuronal inclusions with SUMO (Ueda et al., 2002; Dorval and Fraser, 2007). Thus, SUMOylation and ubiquitination likely play an important role in polyglutamine diseases. Huntington’s disease The neuropathological hallmark of Huntington’s disease is Huntingtin (Htt) with various expansions in the polyQ region of the N-terminal domain (MacDonald et al., 1993). A minimal polyQ length, around 34–45 repeats, must be exceeded for Huntingtin (Htt) to undergo a conformational change to a cross β sheet rich amyloid-like state (Poirier et al., 2002; Steffan et al., 2004). Functionally, Htt is neuroprotective and enhances production of neurotrophic factors (Cattaneo et al., 2009). In the disease state, Htt aggregates and causes neuronal death. It is still unknown whether the pathogenesis of Huntington’s disease is due to a gain- or lossof-function of Htt (Bence et al., 2001; Cattaneo et al., 2009). Both full-length and fragment Htt (Httex1p97QP) have been found in Htt inclusions, and both contain the polyQ N-terminal domain (Kachman et al., 1996; Davies et al., 1997). These inclusions correlate with increased survival in neurons expressing polyQ Htt (Arrasate et al., 2004), providing strong evidence that proteinaceous inclusions provide a ‘dumping ground’ for toxic, misfolded polyQ proteins. Both full-length and fragment Htt are regulated by post-translational modifications, including

ubiquitination (Kalchman et al., 1996; Davies et al., 1997) and SUMOylation (Pennuto et al., 2009; Ehrnhoefer et al., 2011; Zheng and Diamond, 2012). Indeed, when Huntington’s disease striatum samples were measured for levels of SUMOylation, the insoluble fraction of proteins had much higher SUMO2- and SUMO1-conjugation than agematched controls (O’Rourke et al., 2013). Huntingtin protein is SUMOylated on lysine residues in the N-terminus (Steffan et al., 2004). Increased SUMOylation at these residues correlates with increased polyQ-derived toxicity (MacDonald et al., 1993). If SUMOylation sites are disrupted or E3 ligases are down-regulated, Htt aggregation decreases (Steffan et al., 2004). SUMO2 seems to play a particularly important role in modifying Htt, as a dose-dependent increase in Htt aggregation is observed (Lee and Goldberg, 1998). Since both full-length Htt and fragment Htt share these lysine residues, it is unsurprising that the Htt fragment can also be SUMO1- and SUMO2-conjugated (O’Rourke et al., 2013). SUMOylation of the Htt fragment appears to stabilize the peptide (Steffan et al., 2004). Furthermore, SUMO1-conjugation to the Htt fragment increases Htt accumulation and toxicity, but decreases aggregation of the polyQ protein (Andersen, 2006). SUMOylation and ubiquitination appear to compete for modification at lysines 6 and 9. While SUMOylation promotes Htt aggregation, ubiquitination at these same sites promotes solubility (MacDonald et al., 1993). Furthermore, overexpression of ubiquilin, a ubiquitin-binding shuttle factor involved in shuttling polyubiquitinated proteins to the proteasome, has a neuroprotective effect in a mouse model of Huntingtin’s disease (El Ayadi et al., 2012). Generally, it appears that SUMOylation of Htt leads to increased aggregation and ubiquitination of Htt leads to increased solubility. As the lysine residues are consistent across full-length and fragmented Htt, both toxic species have similar PTM modifications. An important distinction seems to arise in the SUMO protein being conjugated to Htt. Whereas SUMO1 is found in Htt-rich inclusion bodies and drives the fragment Htt to be neurotoxic, it likely does not influence Htt aggregation (Andersen, 2006; O’Rourke et al., 2013; Kunadt et al., 2015). Instead, SUMO2 appears to be the key driving force behind Htt aggregation (Kunadt et al.,

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2015). These findings question if monoSUMOylation and polySUMOylation of Htt drive different functions. While both are neurotoxic, perhaps polySUMOylation is an attempt at cellular protection. Indeed, there is a debate about whether polyQ proteins can be degraded by the proteosome. If possible, research suggests that only soluble polyQ proteins are able to be degraded through these mechanisms (Verhoef et al., 2002; Kaytor et al., 2004; Michalik and Van Broeckhoven, 2004; Juenemann et al., 2013; Tsvetkov et al., 2013). Thus, the cell would need to control toxic polyQ production in some way. We discussed previously that polySUMOylation drives substrates to the UPS, and that the UPS leave non-degradable proteins in inclusion bodies. Thus, it is plausible that polySUMOylation drives inclusion body formation. This theory becomes even more interesting when ubiquitination, monoSUMOylation, and polySUMOylation are considered together. If the hypothesis stands that polySUMOylation controls non-degradable polyQ proteins by driving inclusion body formation, then where does ubiquitination fit? Likely, soluble polyQ proteins are able to be ubiquitinated and degraded via the UPS. If this is the case, then insoluble polyQ proteins are likely to outcompete ubiquitin and instead be SUMOylated. This then allows the available lysine residues to be mono- or poly-SUMOylated. Dentatorubral–pallidoluysian atrophy Dentatorubral–pallidoluysian atrophy (DRPLA) is a neuropathological disorder, characterized by dementia, epilepsy, and disrupted movement. DRPLA arises from a CAG expansion in the gene which encodes atrophin-1 (Yazawa et al., 1995; Schilling et al., 1999), the function of which is currently unknown. However, SUMO1-conjugation of polyQ atrophin-1 increases nuclear inclusion body formation and cell death (Terashima et al., 2002). Indeed, when a conjugation-null SUMO1 mutant is overexpressed with polyQ atrophin-1, inclusion formation is decreased (Terashima et al., 2002). Currently, the data are too sparse to hypothesize the role of SUMOylation and ubiquitination on atrophin-1. Spinal and bulbar muscular atrophy Spinal and bulbar muscular atrophy (SBMA) occurs when a CAG repeat occurs in the gene

which encodes Androgen Receptor, a transcription factor that is stimulated by the androgen hormone (Katsuno et al., 2006). SBMA is an X-linked neurodegenerative disorder which manifests as progressive muscle weakness due to motor neuron degeneration in the brain stem and spinal cord. PolyQ Androgen Receptor can be SUMOylated and ubiquitinated on the same lysine residue (Poukka et al., 2000). Functionally, SUMOylation of Androgen Receptor suppresses its transcriptional activity (Takahashi-Fujigasaki et al., 2001; Nishida and Yasuda, 2002). When SBMA is modelled in Drosophila, inhibition of SUMOylation and ubiquitination of Androgen Receptor increases nuclear and cytosolic inclusion formation and degeneration (Chan et al., 2002). Indeed, when SUMO3 is overexpressed in cells, Androgen Receptors exhibit decreased aggregation and increased solubility (Mukherjee et al., 2009). Unlike Htt, SUMOylation and ubiquitination of androgen receptor appear to work in concert with each other. Both SUMO- and ubiquitin-conjugation to androgen receptor keep the protein soluble and monomeric (Mukherjee et al., 2009). Spinocerebellar ataxia types I, III, and VII Spinocerebellar ataxia (SCA) is a neurodegenerative disorder characterized by degeneration of Purkinje cells in the cerebellum. The disorder arises from a CAG expansion in the genes which encode the ataxin family of proteins. This family of proteins bind DNA and are involved in various nuclear functions. SCA-I occurs when a polyQ stretch forms in the ataxin-1 protein. Ataxin-1 can be SUMOylated on at least five different lysine residues and when the polyQ region stretches in SCA-I, SUMOylation of ataxin-1 is decreased (Riley et al., 2005). Silencing of SUMO2/3 using siRNA raises levels of ataxin-1 (Guo et al., 2014), and overexpression of SUMO1 reduces SUMO 2/3 conjugation, ubiquitination, and degradation of the protein (Guo et al., 2014). Inhibiting the formation of SUMO chains by overexpressing a chain-deficient SUMO2 KR mutant decreases the SUMO 2/3 conjugated high molecular weight ataxin-1 species, reduces ubiquitination, and blunts degradation of ataxin-1 (Guo et al., 2014). When SUMO1 or E2 enzyme Ubc9 is overexpressed, polyQ ataxin-1 aggregation increases (Ryu et al., 2010). Conversely, genetically inhibiting

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ubiquitination enhances polyQ-mediated neurodegeneration in a mouse model of SCA-1 (Cummings et al., 1999) and mutating mouse Usp14, which encodes a deubiquitinating enzyme, leads to ataxia (Chernova et al., 2003). Blocking SUMO 2/3 with an siRNA reduces ubiquitination of ataxin-1, suggesting that the SUMO- and ubiquitination systems are working together (Guo et al., 2014). Machado-Joseph Disease (MJD), or SCAIII, arises from the aggregation of Ataxin-3. Functionally, ataxin-3 deubiquitinates proteins by disassembling lysine 48- and lysine 63-linked polyubiquitin chains (Winborn et al., 2008). Ataxin-3 interacts directly with the ubiquitin-selective chaperone valosin-containing protein (VCP), which is critical for appropriate proteasomal degradation (Watts et al., 2004). CAG expansion in the gene produces the most common form of autosomal dominant SCA. Indeed, overexpression of polyQ ataxin-3 compromises the ability of the ubiquitin proteasome system to function appropriately (Burnett et al., 2003), whereas expression of ubiquitin ligase Parkin reduces polyQ ataxin-3 toxicity (Tsai et al., 2003; Morishima et al., 2008). Overexpression of the ubiquitin ligase C terminus of Hsc-70 interacting protein (CHIP) also delays the age at onset of the disease phenotype (Al-Ramahi et al., 2006). Like Ataxin-1, Ataxin-3 can be SUMOylated. Ataxin-3 has two SUMOylation sites, lysine 166 and lysine 356 (Zhou et al., 2013; Almeida et al., 2015). SUMO1-conjugation to Ataxin-3 results in protein stability, leading to increased toxicity and apoptosis (Zhou et al., 2013). However, Hsp70 chaperone function is able to rescue SUMO-induced polyQMJD degeneration (Besnault-Mascard et al., 2005; Shirakura et al., 2005; Hayashi et al., 2006). SCA-VII arises from a polyQ stretch of ataxin-7. Ataxin-7 can be SUMO1- or SUMO2-conjugated at lysine 257 ( Janer et al., 2010). Both SUMO1and SUMO2- conjugated ataxin-7 are found in inclusion bodies; however, SUMO1-conjugation decreases ataxin-7 aggregate formation ( Janer et al., 2010). Mutagenesis of K257 increases levels of SDS-insoluble Ataxin-7 aggregates and increases the amount of caspase-3 positive cells (Ryu et al., 2010). SCA-I and SCA-III are similar, in that SUMO1conjugation promotes protein insolubility and

neurotoxicity. Thus far, we have considered SUMO1-conjugation as representing monoSUMOylation or multiSUMOylation, as opposed to chained polySUMOylation. If this is true, then the role of SUMO2/3 in SCA-I disease reduction is likely due to polySUMOylation of the protein. Indeed, polySUMOylation and ubiquitination can target substrates to the UPS. In SCA-I, data suggests SUMO2/3- or ubiquitin-conjugation inhibits disease progression, possibly via UPS-driven degradation. Contradictorily, SCA-VII forms aggregate to a lesser extent when conjugated with SUMO1. It is odd that proteins within the same family would have such drastically different functions from the same modification. Ultimately, we are unable to hypothesize why this may be. SCA-VII is less studied than SCA-I and SCA-III, and so the data are much more limited. Rare disorders Neuronal intranuclear inclusion disorder Neuronal intranuclear inclusion disorder (NIID) is a rare neurodegenerative disease that affects both the central and peripheral nervous system. Characterized by the presence of neuronal intranuclear inclusions, the symptoms include ataxia and movement disorders which eventually develop into dementia (Sung et al., 1980). While most cases of NIID are sporadic, some familial cases do exist as autosomal dominant disease (Kimber et al., 1998; Zannolli et al., 2002). The neuronal intranuclear inclusions which hallmark this disease stain strongly for SUMO and ubiquitin reactivity across all forms of the disease, including familial, juvenile, and sporadic (Pountney et al., 2003; McFadden et al., 2005; TakahashiFujigasaki et al., 2006). On further investigation, multiple SUMO substrates have been localized to inclusions from diseased patient tissues. For example, localization studies using sporadic and familial NIID tissues suggest that the SUMO substrates histone deacetylase 4 (HDAC4) and promyelocytic leukaemia (PML) are components of the neuronal intranuclear inclusions (Takahashi-Fujigasaki et al., 2006). Indeed, the highly SUMOylated RanGAP1 has also been localized to neuronal intranuclear inclusions in familial NIID tissue (TakahashiFujigaskai et al., 2006).

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There appears to be an interesting overlap of NIID nuclear inclusions with aggregation-prone proteins of other neurodegenerative diseases. For example, ataxin-3, ataxin-1, and multiple polyglutamine disorder-related proteins are commonly found within nuclear inclusions (Lieberman et al., 1998, 1999; Zoghbi and Orr, 2000). Since there is no defining toxic protein identified with NIID, these inclusions likely arise from a general misregulation of SUMO- and ubiquitination. Thus, it is unsurprising that other aggregation-prone proteins appear in NIID inclusions with various levels of SUMO- and ubiquitination. Prions While many human prion diseases exist, this section focuses on the priogenic protein PrP which is found in kuru, Creutzfeldt-Jakob disease, fatal familial insomnia, and Gerstmann–Straussler– Scheinker syndrome. Genetic mutations have been found in the PrP gene of the familial forms of these diseases. The first genetic linkage to prion disease was found in GSS, which contains a P102L mutation in the PrP gene (Chernoff et al., 1995). Since this finding, 40 more mutations have been found in the PrP gene across all human prion protein diseases (Mead, 2006; van der Kamp et al., 2009). These genetic mutations have been linked to the priogenic conversion of PrP to pathogenic PrPsc, causing loss of prion protein function, increased toxicity, and increased transmissibility. Like most other proteinaceous inclusions, PrP aggregates are ubiquitinated (Piccardo et al., 2014) and there exists a correlation between elevated levels of ubiquitin-conjugated protein and reduced proteasomal function (Kang et al., 2004; McKinnon et al., 2016). Ubiquitination of prion aggregates is believed to occur once PrP has converted to the priogenic PrPsc form (Kang et al., 2004). This is linked to the ubiquitin-proteasome system, as identified by expression of the dominant negative mutant of USP14, a deubiquitinating enzyme that controls trimming of polyubiquitin chains and regulates the proteasomal process. When the dominant negative USP14 is expressed in a prion disease-model system, pathogenic prion protein was reduced (Homma et al., 2015). Indeed, expression of wild type USP14 increased pathogenic prion protein (Homma et al., 2015).

While the ubiquitin proteasome system appears to be involved in providing a ‘dumping ground’ for most toxic amyloid and amyloid-like proteins, prions appear to affect the function of the UPS. Wild-type prion protein is degraded in a ubiquitindependent manner (Yedidia et al., 2001). However, mutant prion protein oligomers directly bind to and inhibit the activity of the proteasome (Kristiansen et al., 2007; Deriziotis et al., 2011). The SUMO pathway is also involved in prion diseases. Specifically, in Creutzfeldt-Jakob disease, SUMO2/3 protein levels are decreased in patients (Karu et al., 2014). The toxic PrP protein is also believed to be a SUMO target due to its involvement in various SUMOylation-dependent pathways; however, SUMOylation of PrPc has yet to be determined. Perhaps, as mentioned in previous sections, polySUMOylation is negatively impacted in prion diseases. This would also lead to a decrease in PrPsc targeting to the proteasome, which is already negatively impacted in prion diseases. Functional amyloid-like proteins. Mammalian Cytoplasmic polyadenylation element binding protein 3 Cytoplasmic polyadenylation element binding protein 3 (CPEB3) is an RNA-binding protein that undergoes stimulus-dependent conformational and functional change. In its basal state, CPEB3 is soluble and functions as a translation inhibitor (Fioriti et al., 2015). On neuronal stimulation, CPEB3 converts to an insoluble translation promoter (Fioriti et al., 2015). The insoluble, translation promoter form of CPEB3 is necessary for long-term memory maintenance (Fioriti et al., 2015). From studies conducted under denaturing conditions, CPEB3 appears to form amyloid-like fibrils (Stephan et al., 2015) and when expressed in yeast, exhibits prionlike transmissibility (Si et al., 2010). Work from our lab has shed insight into cellular regulation of CPEB3 structure and function. CPEB3 is SUMOylated in the basal state and deSUMOylated when functioning to promote translation (Drisaldi et al., 2015). There is a cyclical quality to CPEB3 deSUMOylation and SUMO

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protein production; SUMO2 mRNA appears to be a CPEB3 target (Drisaldi et al., 2015). Interestingly, recent work from our lab has further validated the hypothesis that SUMOylation renders CPEB3 soluble and functionally inhibitory. SUMOylated CPEB3 is retained in the membrane-less organelle P body, the function of which is to degrade mRNAs (Ford et al., 2019). However, on chemical long-term potentiation (cLTP) stimulation, CPEB3 exits the P Body and enters the polysome for mRNA translation (Ford et al., 2019). In vitro, SUMOylated and RNA-bound CPEB3 undergoes phase separation and forms P body-like droplets (Ford et al., 2019). Beyond SUMOylation, ubiquitination also plays an important role in CPEB3 structure and function regulation. On synaptic activation, levels of the E3 ubiquitin ligase Neuralized (Neur1) increase (Pavlopoulos et al., 2011). This increase in Neurl1 is necessary for healthy memory function, by facilitating synthesis of proteins involved in synaptic plasticity and synaptic remodelling (Pavlopoulos et al., 2011). The effect of Neurl1 on protein synthesis is tied to ubiquitination of CPEB3; when ubiquitinated by Neurl1, CPEB3 undergoes a functional conversion to promote translation of target mRNAs (Pavlopoulos et al., 2011). Our lab is excited to continue the investigation into ubiquitination and SUMOylation of CPEB3, and currently believe that a SUMO-ubiquitin switch may be involved in altering translation functionality. La La is an RNA-binding protein which binds and traffics various axon-bound mRNAs (Wolin and Cedervall, 2002). The successful binding and trafficking of mRNAs to La is critical for growth-cone guidance, axonal regeneration, and synaptic plasticity (Wolin and Cedervall, 2002). SUMOylation of La directly influences its role in these neuronal functions (van Niekerk et al., 2007) and disrupted SUMOylation of La has been discovered in various cancer cell lines. La depletion and mistrafficking appears to impair cell proliferation in cancer; however, the exact mechanism is still unclear (Kota et al., 2018). Both SUMO-1 and SUMO-2/3 can modify La in an overexpression system, although endogenous

SUMOylation is still unknown (van Niekerk et al., 2007). Non-SUMOylated La binds kinesin and La SUMOylated at lysine 41 binds dynein; thus, SUMOylation plays an important role in axonal trafficking (van Niekerk et al., 2007). Specifically, non-SUMOylated La moves anterograde, while SUMOylated La moves both anterograde and retrograde along the axon (van Niekerk et al., 2007). Beyond trafficking, SUMOylation also enhances mRNA binding to La (Kota et al., 2016). Indeed, when mutated to lack SUMOylation sites, mutant La is incapable of binding target mRNAs (Kota et al., 2016). Mitochondrial anti-viral signalling protein and RIG-1 Mitochondrial anti-viral signalling protein (MAVS) is a component of an anti-viral immune response. In the signalling cascade, MAVS is downstream of receptor RIG-1 and functions to activate kinases that ultimately activate the transcription factors NF-κB and IRF3 (McWhirter et al., 2005). On activation by lysine 63-polyubiquitinated RIG-1, MAVS undergoes a gain-of-function conformation change to a fibrillar state (Hou et al., 2011). These MAVS fibrils consist of three-stranded helixes (Xu et al., 2014) as opposed to the classic cross β sheet of true amyloids. The polyubiquitination of RIG-1 plays the critical role of exposing the CARD domain of the protein, allowing for downstream interaction with MAVS, fibrillization of MAVS, and further activation of the signalling cascade (Hou et al., 2011). The same E3 that ubiquitinates RIG-1, TRIM25, also ubiquitinates MAVS at lysine 7 and 10 (Castanier et al., 2012). The RING finger protein 5 (MARCH5) ubiquitinates filamentous MAVS, targeting the protein to proteasomal degradation (Yoo et al., 2015). MAVS is an interesting example of how E3 ubiquitination targets the same protein in different conformations, driving various downstream effects. For example, TRIM25 ubiquitination of MAVS is likely the E3 involved in CARD domain exposure and downstream fibril formation. Whereas MARCH5 specifically targets filamentous MAVS for degradation. It is still unknown if TRIM25 and MARCH5 ubiquitinate different residues and if there is a difference in monoubiquitination or polyubiquitination.

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Yeast PSI+ PSI+ is the prionoid form of Sup35, a yeast protein subunit of a translation termination factor (Stansfield et al., 1996). PSI+ causes read through of stop codons in translating mRNAs. Although generally non-toxic to the cell, overproduction of the PSI+ conformation will lead to a decrease in health from loss-of-function. Heat shock proteins (HSPs) are chaperone proteins involved in the clearance of misfolded proteins and play a pivotal role in regulation of yeast prion-like proteins. Indeed, HSP104 can eliminate PSI+ when overproduced in yeast (Allen et al., 2006). HSP104 functions with the UPS, and when ubiquitin or critical elements of the UPS are knocked out in yeast, Sup35 converts to PSI+ at a higher frequency (Allen et al., 2006; Tank and True, 2009). Similarly, inhibiting autophagy promotes PSI+ production in yeast (Speldewinde et al., 2015). This effect may not involve direct ubiquitination of Sup35/PSI+, as ubiquitin-conjugated Sup35 was not found by multiple labs (Allen et al., 2006; Tank and True, 2009). Instead, the actinassociated protein Lsb2 is ubiquitinated and drives PSI+ formation (Chernova et al., 2017). PIN+ PIN+ is the prionoid form of Rnq1, a yeast polyQ protein of unknown function. PIN1+, sometimes referred to as RNQ+, facilitates de novo formation of other yeast prionoids (Stein and True, 2011). Unlike Sup35, Rnq1 can be ubiquitinated endogenously (Allen et al., 2006). As with other yeast prion-like proteins, HSP104 is involved in eliminating misfolded PIN+ and ubiquitin ligases protect the cell from prion-like proteins produced by PIN1+ (Theodoraki et al., 2012; Yang et al., 2014). HSP104 functions alongside the UPS, and when critical elements of the UPS are knocked out in yeast, Rnq1 converts to PIN1+ and forms amyloid-like fibrils (Allen et al., 2006; Yang et al., 2014). Conclusion SUMOylation and ubiquitination are common post-translational modifications which regulate protein function, interaction, trafficking, and

structure. Often, SUMO and ubiquitin conjugation destines the substrate to the UPS for degradation. However, as discussed throughout this chapter, these moieties are multi-functional. The role of SUMOylation and ubiquitination in protein regulation is particularly interesting when amyloid and amyloid-like proteins are considered. A significant amount of energy is used in the conversion from a soluble to an amyloid fibrillar state, making the fibrillar state an unlikely resting state for the protein. Thus, it is intriguing as to how amyloids are produced. Post-translational modifications, in part, can trigger critical steps in the fibrillization process of amyloidogenic proteins. This chapter investigates the role of SUMOylation and ubiquitination of amyloids in disease and health (Table 25.1). SUMOylation and ubiquitination of α-synuclein, the amyloid involved in Parkinson’s disease and other synucleinopathies, are likely independent processes. Generally, monoubiquitination of α-synuclein promotes aggregation (Nonaka et al., 2005; Lee et al., 2008) while monoSUMOylation promotes solubility (Krumova et al., 2011) and polyubiquitination drives protein degradation via the UPS (Lee et al., 2009). We then consider SOD1, TDP-43, and FUS amyloids involved in ALS. First, SOD1 has an interesting distinction between physiological and pathological SUMOylation patterns. Physiological SOD1 can be SUMO1-conjugated, whereas pathological SOD1 can be SUMOylated by SUMO1, SUMO2, or SUMO3. All SUMOylation appears to drive aggregation, but perhaps the increased capacity for SUMOylation of pathological SOD1 increases its likelihood of aggregation in a correlative manner. Second, TDP-43 pathogenicity is also influenced by SUMOylation. It is hypothesized that deSUMOylation may drive mutant TDP-43 to the cytosol where it is neurotoxic (Seyfried et al., 2010). Finally, although less well studied than SOD1 and TDP-43, toxic FUS is ubiquitinated in ALS. This chapter then considers Alzheimer’s disease and the amyloid components β-amyloid and tau. Overall, those suffering from Alzheimer’s disease show a marked decrease in high molecular weight (>75 kDa) SUMO2/3. We speculate, throughout this chapter, that changes in SUMO2/3 are likely

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Table 25.1  SUMOylation and ubiquitination of toxic amyloids. Summary of SUMOylation and ubiquitination of all toxic amyloids listed in this chapter Disease

Substrate

SUMOlyation of inclusion bodies

Parkinson’s disease

a synuclein

Ubiquitination of inclusion bodies

Effect of ubiquitin on neurodegeneration

SUMO1, SUMO2 Unknown

Yes

Decreased

Parkin

SUMO1

Unknown

Yes

Unknown

DJ-1

SUMO1

Unknown

Unknown

Unknown

Multiple systems atrophy

a synuclein

SUMO1

Unknown

Yes

Unknown

Dementia with Lewy bodies

a synuclein

SUMO1

Unknown

Yes

Unknown

SUMO1, SUMO2/3

Unknown

Unknown

Unknown

TDP-43

SUMO2/3

Unknown

Yes

Unknown

FUS

Unknown

Unknown

Yes

Unknown

Neuronal intranuclear inclusion disorder

Various

SUMO1

Unknown

Yes

Unknown

Ischaemia

Various

SUMO1, SUMO2/3

Decreased

Unknown

Unknown

Alzheimer’s disease

Amyloid b/ APP

SUMO3

Reduced

Unknown

Unknown

tau

SUMO3

Increased

Yes

Unknown

Huntington’s disease

Huntingtin

SUMO1, SUMO2 Increased

Yes

Decreased

Dentatorubral Pallidoluysian atrophy

Atrophin 1

SUMO1

Unknown

Unknown

Spinal and bulbar muscular atrophy

Androgen receptor

SUMO1, SUMO3 Unknown

Yes

Unknown

Spinocerebellar ataxia

Ataxin

SUMO1

Increased

Yes

Decreased

Prion diseases

PrP

Unknown

Unknown

Yes

Decreased

Amyotrophic lateral SOD1 sclerosis

Effect of SUMO on neurodegeneration

Increased

alterations in the extent of polySUMOylation of the substrate. Therefore, we believe that polySUMOylation is decreased in Alzheimer’s disease. Although speculative, the cause for decreased polySUMOylation is likely to come from its role in the UPS. PolySUMOylation can act as a marker for proteasomal degradation, functioning with downstream ubiquitination. In the case of Alzheimer’s disease, we believe that high molecular weight SUMO2/3 is decreased because of polySUMO-targeting to the proteosome. Likely, polySUMOylation is an early stage phenomena, and decreases in late stage disease due to decreased proteasome integrity. Indeed, the decrease in high molecular weight SUMO2/3 is observed in early stages of Alzheimer’s disease, and later stages of disease are marked by UPS dysfunction.

We further explore the role of SUMOylation and ubiquitination in Alzheimer’s disease by considering the two amyloids involved, β-amyloid and tau. It appears that β-amyloid production is influenced by APP SUMOylation. In the case of tau, aggregation correlates with increased SUMOylation and degradation correlates with increased ubiquitination (Luo et al., 2014). As stated previously for general inclusion body formation, ubiquitination of the substrate drives the toxic protein to degradation. However, if toxic proteins overwhelm the system, ubiquitinated proteins may be left in inclusion bodies as a final attempt to regulate pathogenicity. We further hypothesize about the role of SUMOylation on tau fibrillization. We offer that SUMOylation of unbound tau may be the critical structural point that converts tau to an amyloid,

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allowing for downstream hyperphosphorylation and fibrillization. SUMOylation and ubiquitination of polyglutamine diseases are then considered. First, we investigate Huntington’s disease amyloid, Htt. Generally, SUMOylation of Htt leads to increased aggregation and ubiquitination of Htt leads to increased solubility. An important distinction arises when the SUMO protein being conjugated to Htt differs. SUMO2 appears to be the key driving force behind Htt aggregation. Both monoSUMOylation and polySUMOylation are neurotoxic, but we hypothesize that polySUMOylation plays a protective role. Research suggests that only soluble polyQ proteins are able to be degraded through proteasomal mechanisms (Verhoef et al., 2002; Kaytor et al., 2004; Michalik and Van Broeckhoven, 2004; Juenemann et al., 2013). However, the cell still needs to control toxic polyQ production in some way. Likely, polySUMOylation drives Htt into inclusion bodies. Let’s assume that polySUMOylation controls non-degradable polyQ proteins by driving inclusion body formation. We hypothesize that soluble polyQ proteins are ubiquitinated and degraded via the UPS. Insoluble polyQ proteins are incapable of being degraded via the UPS. Therefore, insoluble polyQ proteins are instead SUMOylated. This then allows the available lysine residues to be mono- or poly-SUMOylated. Not all polyQ proteins behave similarly to Htt. Both SUMO- and ubiquitin-conjugation to androgen receptor keep the protein soluble and monomeric (Mukherjee et al., 2009) in DRPLA. Furthermore, in SCA, SUMO1-conjugation promotes protein insolubility and neurotoxicity. In SCA-I, data suggests SUMO2/3- or ubiquitinconjugation inhibits disease progression, possibly via UPS-driven degradation. This again strengthens our hypothesis that polySUMOylation allows some amyloidogenic proteins to work via the UPS towards degradation or inclusion body formation. Our penultimate investigation into toxic amyloids considers NIID. NIID lacks a specific protein which drives pathology; instead, many of the polyQ proteins previously discussed are within the inclusions which hallmark the disease. These inclusions likely arise from a general misregulation of SUMOand ubiquitination. Finally, we consider prion diseases and the roles SUMOylation and ubiquitination play in

regulating the amyloidogenic protein. We speculate that polySUMOylation is negatively impacted in prion diseases. This would also lead to a decrease in PrPsc targeting to the proteasome, which is already severely impaired. After considering the wealth of knowledge on toxic amyloids, we investigate the more recent literature on functional amyloids in mammals and yeast. RNA-binding protein CPEB3 is likely regulated by a SUMO-ubiquitin switch, driving translation functionality. The data indicates that SUMOylation of CPEB3 produces an inhibitory function, and ubiquitination of the protein promotes translation activity. Another RNA-binding protein, La, also switches functions based on if the protein is SUMOylated or deSUMOylated. Finally, MAVS is an excellent example of functional ubiquitination. Ubiquitination of MAVS allows for the conformational change to a fibrillar structure, with downstream functional effects. Finally, we end our investigation with yeast functional amyloids. Thus far, yeast have been the most prevalent model organism used in functional amyloid studies. From this work, numerous yeast prion-like proteins have been discovered. Here, we consider PSI+ and PIN1+. There is a wealth of information suggesting that the UPS is disrupted when either of these yeast prionoids are expressed. Overall, we can draw numerous conclusions. First, and most importantly, there is no generic function for SUMOylation or ubiquitination of amyloids. Each protein is distinct and will be impacted by PTMs differently; this is true even within protein families, as identified in the SCA section. Second, the data suggests that inclusion bodies act as a ‘dumping ground’ in disease. Often, the UPS is unable to perform its function or is overwhelmed with misfolded proteins in amyloid diseases. The inclusion bodies offer a final attempt to compartmentalize toxic species. Third, we hypothesize that polySUMOylation plays an important role in many of the above mentioned diseases. Likely, polySUMOylation is a second method in which the cell can target toxic species for degradation or inclusion body formation. The functional amyloid section provides important examples of the complexity of PTMs, and the resulting downstream effects. One outstanding difficulty in the field is deciphering which SUMOylation and ubiquitination patterns are part of protein function, and which

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patterns are part of cellular stress resulting from pathology. Furthermore, PTMs are highly dynamic, altering as the cell requires. Complexity is increased when crosstalk is considered. We briefly mentioned the influence of tau SUMOylation on downstream hyperphosphorylation. Many more examples of PTM crosstalk are known for these proteins but were not covered. In order to fully understand the role SUMOylation, ubiquitination, and other PTMs play on amyloids in health and disease, then state of health, time of quantification, and extent of crosstalk all must be understood. Although a large task, quantitative proteomics allows for a deeper understanding of the role of PTMs on amyloid function. Soon, more methodological advancements will allow us to probe deeper and ask more complex questions. We are excited to see the field progress our understanding of cellular regulation of amyloids in health and disease, and to gain insight into the roles PTMs play in the cellular regulation of amyloids in the brain. Acknowledgements We would like to thank Joe Rayman and Pauline Henick for their comments on content and editing. References Aguzzi, A., Heikenwalder, M., and Polymenidou, M. (2007). Insights into prion strains and neurotoxicity. Nat. Rev. Mol. Cell Biol. 8, 552–561. Allen, C., Büttner, S., Aragon, A.D., Thomas, J.A., Meirelles, O., Jaetao, J.E., Benn, D., Ruby, S.W., Veenhuis, M., Madeo, F., et al. (2006). Isolation of quiescent and nonquiescent cells from yeast stationary-phase cultures. J. Cell Biol. 174, 89–100. Almeida, B., Abreu, I.A., Matos, C.A., Fraga, J.S., Fernandes, S., Macedo, M.G., Gutiérrez-Gallego, R., Pereira, P.J., Carvalho, A.L., and Macedo-Ribeiro, S. (2015). SUMOylation of the brain-predominant Ataxin-3 isoform modulates its interaction with p97. Biochim. Biophys. Acta 1852, 1950–1959. https://doi. org/10.1016/j.bbadis.2015.06.010. Al-Ramahi, I., Lam, Y.C., Chen, H.K., de Gouyon, B., Zhang, M., Pérez, A.M., Branco, J., de Haro, M., Patterson, C., Zoghbi, H.Y., et al. (2006). CHIP protects from the neurotoxicity of expanded and wild-type ataxin-1 and promotes their ubiquitination and degradation. J. Biol. Chem. 281, 26714–26724. Andersen, P.M. (2006). Amyotrophic lateral sclerosis associated with mutations in the CuZn superoxide dismutase gene. Curr. Neurol. Neurosci. Rep. 6, 37–46. Arai, T., Hasegawa, M., Akiyama, H., Ikeda, K., Nonaka, T., Mori, H., Mann, D., Tsuchiya, K., Yoshida, M., Hashizume, Y., et al. (2006). TDP-43 is a component of ubiquitin-positive tau-negative inclusions in

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Keeping Up with the Pathogens: The Role of SUMOylation in Plant Immunity Rebecca Morrell and Ari Sadanandom*

26

Department of Biosciences, Durham University, Durham, UK. *Correspondence: [email protected] https://doi.org/10.21775/9781912530120.26

Abstract Owing to the changing, challenging pressures the plant pathogens can exert on hosts, plants require mechanisms to quickly sense and respond to them. Post-translational modifications (PTMs) provide a molecular level of control that can rapidly alter the stability, interaction and localization of proteins. SUMO is being increasingly implicated as a critical modifier affecting plant susceptibility at all stages of pathogen disease progression. This review highlights how in pathogen-associated molecular patterns (PAMP)-triggered immunity (PTI) on pathogen detection a PAMP receptor is SUMOylated to enable downstream pathogen defence genes to be activated. In effector-triggered susceptibility (ETS) pathogens exploit the plants endogenous SUMO system to aid disease progression, injecting SUMO proteases into the plant cells. Finally, in effector-triggered immunity (ETI), many mutants in the SUMO system show increased disease resistance due to elevated levels of salicylic acid and the consequential downstream signalling of pathogen defence genes. The research presented aims to highlight the critical role SUMO plays in plant immunity, directly by SUMOylating critical pathogen defence molecules and also indirectly by affecting important hormone signalling pathways involved in pathogen defence. Introduction The world population chiefly relies on plants to sustain itself, requiring crops not only for food but

also for the production of biofuels, medicine and for material products such as clothing. However, crop security is increasingly under threat due to climate change; this is also resulting in pests and diseases changing their geographical ranges. Furthermore, the changing climate is also causing abiotic pressures to crops as well. In addition, an increasing population is demanding greater output of agricultural systems, as by 2050 the population is predicted to reach 9.7 billion people (UN, 2012). The FAO in 2009, estimated that to feed a population of 9.1 billion people in 2050, food production will have to increase by 70% to feed the additional 2.3 billion people. The pressure on the agricultural sector to boost productivity will be challenged by using scarce natural resources more efficiently and adapting to climate change. During the second half of the 20th century crop losses were substantially reduced due to the development of disease resistant varieties (FAO, 1991). This was achievable due to the identification of a wide variety of resistance (R) genes present in a diverse variety of wild crop relatives “genetic crop resources” which was bred into commercial varieties. These R gene, from wild relatives, generally could be crossed into agriculturally important varieties through breeding, not requiring genetic modification and hence the ensuing legislature. However, crop losses due to plant diseases are increasing due to pathogens developing resistance. There are many examples in today’s agricultural system of crops at severe risk due to plant pathogens, which are becoming more globally spread

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Immunity Plants have physical barriers to diseases including a waxy cuticle and plant cell wall. If these barriers become invaded the plant has to rely on a series of signalling cascades. Each cell must contain innate immunity to pathogens as unlike mammals, plants do not have mobile defender cells or an immune specific cells ( Jones and Dangl, 2006). The plant immune system response includes initial, low-level pathogen-triggered immunity (PTI) or microbetriggered immunity (MTI) and high-level, specific effector-triggered immunity (ETI) ( Jones and Dangl, 2006; Dodds and Rathjen, 2010). As has been reviewed in Jones and Dangl (2006) and is summarized in Fig. 26.1 as a ‘zig-zag’ model of plant immunity starts with phase 1 of disease progression. Transmembrane pattern recognition receptors (PRRs) detect MAMPs/PAMPs (microbial- or pathogen-associated molecular patterns) such as flagellin and PTI is triggered. However, the disease progression moves to phase 2 if the pathogen is capable of overcoming phase 1 and deploying effectors, resulting in ETS (effector triggered susceptibility). Typically, 15–30 pathogen effectors are injected into plant cells, by the type III secretion system. They function to diminish structure in the plant cell, aid pathogen dispersal or interrupt PTI or ETI. In phase 3 ETI is triggered if the plant has R (resistance) genes for the invading pathogen that are capable of detecting effector

due to global trade. One such example is the Cavendish banana cultivar, the most commonly consumed banana. This banana variety is at risk of extinction, due to low genetic diversity, as all the species are clones and possess no natural resistance to tropical race 4 (TR4) Fusarium oxysporum f. sp. cubense (Foc). TR4 is difficult to control and prevent and current predictions estimate the disease will cause industry losses exceeding $138 million per year, despite a slow rate of spread (Cook et al., 2015). Crops which receive extensive immune research include wheat and rice, two major crops, that provide a large number of the world calories. Pathogens of these crops cause severe problems as they are consumed so widely. Current concerning pathogens to these crops include wheat stem rust, fusarium head blight (FHB) and rice blast. Wheat stem rust is currently threatening to recur in Europe, after being eradicated in the mid-to-late twentieth century (Lewis et al., 2018), FHB has been spreading in North America to the Pacific Northwest (Marshall et al., 2012) and if farmers in the Mid-South America eradicated rice blast it would provide farmers with 69.34 million dollars annually and increase the rice supply to feed an additional one million consumers (Nalley et al., 2016). To help tackle these global problems the plants natural immunity needs to be studied to be fully utilised.

Pathogen virulence

Pathogen

P

P PRR E

E R

E

Host immunity Evolutionary time Figure 26.1  The zig-zag model of plant immunity. Immune activation by receptor molecules PRR of pathogen PAMPs/MAMPs triggers PTI. Pathogens suppress the plant immunity with effector molecules resulting in ETS. Host R genes encode receptors that detect and respond to the effectors triggering ETI. PRR = pattern recognition receptor, E = effector, R = resistance gene. The evolutionary time axis (black) represents the evolutionary time pressure on both pathogen and host to evolve new molecules to evade the immunity or pathogenicity respectively.

Role of SUMOylation in Plant Immunity |  491

molecules. The immune response is either triggered directly, commonly by the plants NB-LRR (nucleotide binding-leucine rich repeat) proteins which recognise an effector or indirectly by plants detecting host proteins that have been altered by effectors. When ETI is triggered it results in a series of signalling cascades and disease resistance at the site of infection, usually culminating in hypersensitive cell death response (HR). The HR leads to programmed cell death which results in restricted biotrophic pathogen growth at the site of infection (Dangl et al., 2013). The final phase, phase 4 results in, through natural selection, pathogens changing or losing the effector molecules to evade ETI. Naturally, through natural selection, plants are also evolving new proteins to detect the changing/new effector molecules. The signalling cascades involved in plant immunity have been well researched. Post-translational modifications (PTMs) of proteins involved in the signalling cascade have determined phosphorylation and ubiquitination as important PTMs to regulate immunity (Casey et al., 2017). A large number of phosphopeptides, phosphorylated after pathogen elicitor treatment, have been identified, indicating the important role of phosphorylation in immunity (Casey et al., 2017). In particular MAPK and CPDK protein kinases initiate phosphorylation cascades, that phosphorylates transcription factors, controlling defence gene expression and the detection of MAMPs/PAMPs (van den Burg and Takken, 2010a). Ubiquitination also plays an important role in plant immunity. The ubiquitin specificity is conferred by E3 ligases. One such E3 ligase, CRL3 (Cullin3 Ubiquitin-Like) has been implicated in immunity by ubiquitinating NPR1 (Nonexpresser of PR genes 1), a transcriptional co-activator that in the presence of salicylic acid (SA) becomes active, activating SA- responsive gene promoters and expression of pathogen defence genes. SA is an important signalling hormone in plant immunity, triggering expression of many plant immune signalling cascades. CRL3 ligase functions to maintain SA-responsive immune gene expression, preventing constant production of immune responsive genes and autoimmunity. However, paradoxically, promoters of systemic acquired response (SAR) phosphorylate NPR1, facilitating its interaction CRL3 ligase. Spoel et al., (2009) demonstrated

a dual role for the CRL3 ligase as both an activator and repressor of plant immunity (Spoel et al., 2009). The ubiquitin system has been less studied in crop varieties, however the wheat ubiquitin E2 enzyme TaU4 has also been shown to be involved in pathogen defence. Virus induced silencing of TaU4 results in enhanced disease resistance to Zymoseptoria tritici, which causes Septoria Leaf Blotch (Millyard et al., 2016). Small ubiquitin-like modifier (SUMO) A relatively new post-translational modification that has been implicated in plant immunity is SUMO (small ubiquitin-like modifier), a small polypeptide of approximately 100 amino acids, that is capable of post-translationally modifying proteins through the formation of a covalent bond to proteins (SUMOylation). This commonly occurs during a stress response. Exposing plants to abiotic stresses including heat shocking and high salt concentration results in an accumulation of SUMOylated proteins (Kurepa et al., 2003; Lois et al., 2003; Conti et al., 2008). SUMO acts as a quick response mechanism to modify the behaviour of the proteins. SUMOylation of target proteins can result in changes to the SUMOylated protein stability, localization and interaction with other proteins (Wilkinson et al., 2010). There have been eight SUMO genes identified in Arabidopsis thaliana, with four encoding SUMO protein; SUMO1, SUMO2, SUMO3 and SUMO5 (Hammoudi et al., 2016). SUMO1 and 2 have functional redundancy as single sumo1-1 or sumo2-1 mutants are phenotypically similar to WT (wild type), whereas sumo1-1 sumo2-1 double mutants are embryonically lethal (Saracco et al., 2007). SUMOylation tends to occur at a conserved motif in plant proteins, ψ-K-V-D/E (ψ denotes a hydrophobic residue), with the covalent bond forming between C-terminal glycine of SUMO and an amine side chain of the lysine (K) (Kurepa et al., 2003). The conjugation of SUMO to target proteins requires a number of enzymes to catalyse the reaction including activation, conjugation and ligation enzymes (Fig. 26.2). Firstly, a SUMO protease is initially required to cleave the C-terminal extension of an immature SUMO resulting in mature SUMO. Using ATP SUMO E1, activating

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enzyme (composed of SAE1a, SAE1b and SAE2) forms a thioester bond to the now activated SUMO. The E2 enzyme (SUMO conjugating enzyme 1, SCE1) then transfers the newly activated SUMO to the E3 ligation enzyme that catalyses the formation of a covalent bond between the SUMO and target protein. However, the E2 enzyme can also directly facilitate the SUMOylation of target proteins in the absence of E3. There are two E3 ligases in Arabidopsis, HIGHPLOIDY2 (HPY2) and SAP and Miz1 (SIZ1) (Ishida et al., 2012), compared to the 1400 genes encoding E3 ligases in ubiquitin (Kraft et al., 2005) the importance of the SUMO E3 ligases has been speculated (Yates et al., 2016; Garrido et al., 2018; Verma et al., 2018). SUMOylation is a dynamic process as it is responding to stress, thus is also reversible. SUMO is cleaved from target proteins with a SUMO protease. The cleavage of SUMO between the terminal glycine and the target protein recycles free SUMO maintaining the equilibrium of SUMO for signalling. Until very recently there were eight identified SUMO proteases in A. thaliana namely; OTS1, OTS2, ESD4, ELS1, ELS2, FUG1, SPF1 and SPF2 (Castro et al., 2018). These cysteine proteases were identified due to their high sequence similarity to

yeast ULP1 (Ubiquitin like specific protease 1) and ULP2 proteases and are characterized by a conserved H–D–C catalytic triad (Kurepa et al., 2003). Orosa et al. (2018) identified eight new SUMO proteases including Desi 3A, based on their sequence similarity to human Desi 1, the TAIR accessions are given in Table 26.1 and the phylogenetic relationship in Fig. 26.3. The addition of these extra proteases further supports the hypothesis that the specificity of the SUMO system in plants may be conferred by deSUMOylation (Yates et al., 2016; Garrido et al., 2018; Verma et al., 2018). In addition, there have not currently been identified any proteases capable of cleaving AtSUMO5 (Colby et al., 2006), highlighting the requirement for further research in the field. Proteins can also form non-covalent interactions with SUMO through a SIM site (SUMO-interacting motif). These motifs are short hydrophobic peptide sequences with clusters of valine, isoleucine and leucine followed by clusters of negatively charged acidic or phosphorylated amino acids (Minty et al., 2000). SUMOylation of a target protein can result in new protein–protein interactions if the corresponding interacting protein has a SIM site (Wilkinson et al., 2010).

Substrate

Substrate

E3

Protease S

Protease S

S

S

E2

S

E1 +ATP

E2

E1

Figure 26.2 The SUMO cycle of activation, conjugation and deconjugation. Initially an immature SUMO is processed by a SUMO protease to form a mature SUMO. This free SUMO is activated by an E1 activating enzyme, with ATP, transferred to an E2 conjugation enzyme, which, sometimes aided by an E3 ligase enzyme is covalently attached to a substrate target protein via a lysine amino acid in the target protein. The SUMO can then be removed from the target protein by a SUMO protease. Both the target protein and SUMO can undergo cycles of conjugation and deconjugation in response to stresses.

Role of SUMOylation in Plant Immunity |  493

Table 26.1  The existing nomenclature for the identified Arabidopsis thaliana SUMO proteases. It incorporates the newly identified Desi proteases. It has been suggested by Castro et al. (2018) that in the future the SUMO proteases may be spelled out with a prefix of the species followed by increasing numbering. Castro et al. (2018) gave the example for tomato Class II ULPs may be named SlOTS1, SlOTS2, and so on SUMO protease

Tair accession

Cited by

ESD4 (Early in Short Days 4)

At4g15880

Castro et al. (2018)

ELS1 (ESD4-Like SUMO Protease 1)

At3g06910

Castro et al. (2018)

ELS2 (ESD4-Like SUMO Protease 1)

At4g00690

Castro et al. (2018)

OTS1 (Overly Tolerant to Salt 1)

At1g60220

Castro et al. (2018)

OTS2 (Overly Tolerant to Salt 2)

At1g10570

Castro et al. (2018)

FUG1 (Fourth ULP Gene Class 1)

At3g48480

Castro et al. (2018)

SPF1 (SUMO Protease Related to Fertility 1)

At1g09730

Castro et al. (2018)

SPF2 (SUMO Protease Related to Fertility 2)

At4g33620

Castro et al. (2018)

Desi3a (DeSUMOylating Isopeptidase 3A)

At1g47740

Orosa et al. (2018)

Desi1 (DeSUMOylating Isopeptidase 1)

At3g07090

Orosa et al. (2018)

Desi2B (DeSUMOylating Isopeptidase 2B)

At4g25680

Orosa et al. (2018)

Desi2A (DeSUMOylating Isopeptidase 2A)

At4g25660

Orosa et al. (2018)

Desi4A (DeSUMOylating Isopeptidase 4A)

At4g17486

Orosa et al. (2018)

Desi4B (DeSUMOylating Isopeptidase 4B)

At5g47310

Orosa et al. (2018)

Desi3C(DeSUMOylating Isopeptidase 3C)

At5g25170

Orosa et al. (2018)

Desi3B (DeSUMOylating Isopeptidase 3B)

At2g25190

Orosa et al. (2018)

Figure 26.3  A phylogenetic tree of the identified SUMO proteases. The molecular phylogenetic analysis was performed using ClustalX Bootstrap Neighbour-Joining, trees were visualized using MEGA7.

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Phase 1-PTI The first step of plant immunity that results in PTI, is the recognition of MAMPs/PAMPs by PRRs. SUMO has been implicated as an important PTM that occurs to FLS2 (Flagellin Sensitive 2), a PRR that detects bacterial flagellin. On detection of flagellin, FLS2 complexes with BAK1 (BRI1 Associated Receptor Kinase) as a co-receptor, phosphorylating BIK1 (Botrytis Induced Kinase 1) resulting in its dissociation. The now free BIK1 activates downstream signalling components including Mitogen-Activated Protein Kinases (MAPKs) and Respiratory burst Oxidase Homologue Protein D (RbohD). These proteins then trigger immune signalling such as reactive oxidative species (ROS) bursts which attempt to stop pathogen development. Orosa et al. (2018) identified a SUMO site in FLS2, when the lysine, which SUMO covalently attaches too, was mutated to arginine (FLS2K/R) the FLS2K/R protein can no longer be SUMOylated. FLS2 was found to be SUMOylated in the presence of flagellin, which aided the release of BIK1. On infection with virulent bacterial pathogen Pseudomonas syringae pv. tomato (Pst) the non-SUMOylatable FLS2K/R was found to be more susceptible to the infection. SUMO is required for flagellin dependent release of BIK1 which results in downstream signalling, the FLS2K/R mutant is less capable of producing oxidative bursts and inducing MPK6/3 protein production due to less BIK1 being released from FLS2. SUMO protease Desi3A was identified in the paper as the SUMO protease that de-SUMOylates FLS2. The importance of SUMO proteases in conferring specificity in the SUMO system and in particular in its role in defence was highlighted. The desia3a-1 mutants show enhanced flagellin susceptibility, high ROS burst, increased protein production of MPK3/6 and resistance to bacterial infection of Pst. This is due to FLS2 being highly SUMOylated resulting in more free BIK1 capable of activating its downstream signalling components. So far only one PRR has been identified to be SUMOylated with a role in pathogen defence, but it is likely there are many more targets. Phase 2-ETS In phase 2 of pathogen disease progression, if the pathogen is able to overcome PTI, it uses the type

III secretion system to insert effectors into the plant cells, encoded by Avr genes. The role of SUMO in plant immunity had been speculated for a long time as plant pathogens including Xanthomonas campestris, Ralstonia solanacearum, Pseudomonas syringae, Erwinia pyrifoliae and Rhizobium spp. inject effectors that have ULP1 SUMO protease homology (Orth et al., 1999, 2000; Deslandes et al., 2003; Hotson et al., 2003; Hotson and Mudgett, 2004; Roden et al., 2004; Bartetzko et al., 2009; Kim et al., 2013). Several effectors with SUMO proteolytic activity have been identified. Indeed, X. campestris pv. vesicatoria (X.c.v.) effector AvrBsT has been shown to deSUMOylate its host target protein and disrupt the protein product, preventing the hypersensitive response in tobacco. This is particularly noteworthy due to the bacteria, which causes bacterial spot, not possessing a sumoylation/de-sumoylation system (Orth et al., 2000). Another type III effector protein from Xcv, XopD, was also found to encode an active SUMO protease (Hotson et al., 2003). The XopD gene has even closer gene homology to ULP1 possessing extensive sequence similarity with ULP1 catalytic domain. The protein is translocated to the plant cell nucleus and subnuclear foci on infection and mimics an endogenous plant SUMO isopeptidase with SUMO-conjugated proteins as its substrates (Hotson et al., 2003). Finally, a shorter version of XopD, lacking the N-terminal domain, XopDXcc8004, has also been identified as a type III effector from Xvc acting as a SUMO protease (Tan et al., 2015). XopDXcc8004 expression in Arabidopsis was capable of deSUMOylating host protein HFR1 this elicited host defence response genes solely dependent on its SUMO protease activity; in transgenic plants harbouring XopDXcc8004 (C355A) no elicitation was noticed (Tan et al., 2015). Finally, it has been suggested that necrotrophic fungi Botrytis cinerea and Plectosphaerella cucumerina may also inject effectors that target the hosts SUMOylation machinery. Transgenic Arabidopsis, containing mutations in SUMO E1 activating enzyme SAE2, which disrupt the interactions between SUMO E1 and E2 enzymes, prevents SUMO conjugation in planta. These plants were shown to be more susceptible to the necrotrophic fungi. Early after pathogen infection the hosts SUMO conjugation was post-transcriptionally down-regulated, suggesting the fungal

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pathogen was targeting the SUMOylation machinery (Castaño-Miquel et al., 2017). Furthermore, it has been recently demonstrated by Srivastava et al. (2018) that necrotrophic pathogen infection of Botrytis cinerea promotes the degradation of SUMO protease OTS1 (Overly Tolerant to Salt 1). The SUMO proteases (ots1ots2-1) double mutant was shown to be more susceptible to the necrotrophic pathogen as the degradation of the OTS1 protease results in JAZ ( Jazmonate Zim) SUMOylation and jasmonic acid ( JA) inhibition (Srivastava et al., 2018). In the ots1ots2-1 double mutant, more SUMOylated and non-SUMOylated JAZ6 accumulates ( JA repressor), yet there is no change in levels of endogenous JA. This is due to SUMOylated JAZ6 being more stable and inhibiting the activity of COI1 (Coronatine Insensitive 1). COI1 is a JA receptor and on perception of JA COI1 binds to JAZ6 promoting its degradation. Whilst both JAZ6 WT and the non-SUMOylatable JAZ6, JAZ62KR bind COI1 at equal levels, the presence of SUMO in JAZ6 WT, inhibits the COI1 receptor, preventing COI1 promoting degradation of JAZ6, due to COI1 having a SIM site. OTS1 and 2 regulate the levels of SUMOylated JAZ6, with SUMOylated JAZ6 accumulating in the ots1ots2-1 mutant. The increased stability of the JA repressor in the ots1ots2-1 mutant inhibits JA signalling with the JA signal not being transmitted down the signalling pathway despite equal levels of JA. Necrotrophic infection of Arabidopsis thaliana promotes degradation of SUMO proteases OTS1 and OTS2 which increases the abundance of SUMOylated JAZ6 and inhibits JA signalling promoting disease progression for the pathogen (Srivastava et al., 2018). Phase 3-ETI Plant ETI is established when a plant protein encoded by an R gene interacts with a pathogen effector protein. These intracellular receptors generally follow a ‘gene for gene’ model (Flor 1971) whereby one R gene codes for a receptor for one Avr gene produced by the pathogen. Currently there have not been any direct examples of NB-LRR proteins being SUMOylated. However, Gou et al., 2017 when examining SUMO E3 ligase siz-1 mutant suggested the phenotype is dependent on NB-LRR protein SNC1 (Suppressor of NPR1 constitutive). It has been demonstrated

SNC1 is SUMOylated in planta. In the siz-1 mutant SNC1 is activated/over-accumulated. Overexpressing an F-box protein, CPR1 (Constitutive Expressor of PR Genes 1), that degrades SNC1, in the siz-1 mutant background was able to restore the siz-1 mutant phenotype of disease resistance. The study suggests that SIZ1 E3 ligase may have a role in levels of SNC1 protein, mediated by the SUMOylation status caused by SIZ1. The activation of NB-LRR genes activates downstream signalling cascades resulting in salicylic acid release, as a local and systemic signal for resistance against biotrophs, as part of SAR (system acquired resistance). It has been well documented that Arabidopsis mutants defective in salicylic acid biosynthesis or responsiveness have impaired immunity defence and defective systemic acquired resistance (SAR) and are therefore more susceptible to pathogens. It had been reported that mutation or overexpression of components of the SUMO pathway have a role of salicylic acid biosynthesis and therefore plant immunity (Lee et al., 2007; Kim, 2009; van den Burg et al., 2010; Villajuana-Bonequi et al., 2014; Bailey et al., 2016). The siz1-1 SUMO E3 ligase mutant, characterised by Lee et al. (2007), has elevated accumulation of salicylic acid causing constitutive SAR. The high basal levels of SA results in genes involved in pathogen defence being constitutively expressed, resulting in increased resistance to bacterial pathogen Pst DC3000. The up-regulation of pathogen defence genes included PAD4 (Phytoalexin-Deficient 4), (SID2) Salicylic Acid Induction Deficient 2, (PR1) Pathogenesis related 1 and (EDS1) Enhanced Disease Susceptibility 1. The siz1-1 immune phenotype was reversed when bacterially derived salicylate hydroxylase (NahG) gene was overexpressed. The NahG gene hydrolyses SA, demonstrating how the increased pathogen resistance was due to increased SA levels (Lee et al., 2007). A SUMO protease that has been implicated in salicylic acid levels is ESD4-1 (Early in short days 4), esd4-1 has a similar phenotype to siz1-1 ( Jin et al., 2008), of early flowering under short day conditions (Villajuana-Bonequi et al., 2014). The increase in SA in esd4-1 protease mutant has been shown to be due to biosynthesis gene ICS1 (Isochorismate synthase 1), as when this ICS1 gene (sid2) is mutated in the esd4-1 background the esd4-1 phenotype is reversed. Whilst the esd4-1

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protease mutant has not been phenotyped for pathogen susceptibility it can be hypothesized that the mutant would have enhanced resistance as mutants which have enhanced levels of SA such as siz1-1 have enhanced resistance to pathogen. In addition, esd4-1 has increased expression of PR1, a pathogen defence gene (Villajuana-Bonequi et al., 2014). Another two reported SUMO proteases that have been implicated in immune response is OTS1 (overly tolerant to salt 1) and its homologue OTS2, both proteases are required to be mutated due to functional redundancy. The double mutation is more resistant to Pst DC3000 and as was shown with the other proteases, the ots1ots2-1 double knockout has an increased expression of pathogen defence genes such as PR1 and PR2. The ots1ots2-1 double mutants also have up-regulated SA biosynthesis genes including high expression of ICS1, resulting in increased levels of SA, causing more SA signalling. OTS1 and OTS2 are degraded by high levels of SA and overexpression of OTS1 results in reduced levels of ICS1. Hence, Bailey et al. (2016) hypothesized a feedback mechanism where high SA levels degrade OTS1/2, when OTS1/2 are more abundant they lower SA levels by reducing ICS1 expression. This feedback loop may be controlled by a transcription factor which when deSUMOylated by OTS1/2 in low SA levels (i.e. when the SUMO proteases are stable) are unable to promote expression of ICS1. However, on OTS1/2 degradation in high SA levels the SUMOylated transcription factor is capable of promoting ICS1 expression, this transcription factor is yet to be identified. Three SUMO isoforms seem to have a role in plant immunity. Saleh et al. (2015) demonstrated how, in Arabidopsis SUMO3 is covalently attached to NPR1, an important protein activated in pathogen defence. SUMO3 alters the protein interactions of NPR1 promoting interaction with TGA3 (TGA1A-RELATED GENE 3), a transcription factor promoting expression of pathogen defence genes and blocking interaction with PR1/2 gene repressor, WRKY70 (WRKY DNA-BINDING PROTEIN 70). SUMO3 expression has also been shown to be induced by SA and bacterial PAMP flagellin22 (van den Burg et al., 2010a). Sumo3-1 knockout mutant does not display a defence phenotype, however SUMO3 overexpression had activated plant defences and increased resistance to Pst (van den Burg et al., 2010a).

SUMO1 and 2 double knockdown mutant is required due to the functional redundancy of the SUMO proteins and the embryo lethality of a true sumo1sumo2-1 knockout mutant. The knockdown mutant (sum1-1amiR-sum2) has increased activation of SA-dependent defence genes, in addition to overexpression of WT or conjugation deficient SUMO mutants (van den Burg et al., 2010b). The activated defence genes meant the mutants were more resistant to Pst (van den Burg et al., 2010b). The differing phenotypes of SUMO1, 2 and 3 mutants suggest that SUMO1 and 2 may be working upstream of SA activation, preventing unnecessary activation of SA defence, whereas SUMO3 may be functioning downstream of the SA signalling promoting plant defences. Finally, SUMO E2 conjugating enzyme SCE1 has been implicated in plant immunity. Silencing SCE1 using VIGS (virus-induced gene-silencing) in Solanum peruvianum resulted in higher susceptibility to Gram-positive bacterial pathogen Clavibacter michiganensis subsp. Michiganensis (Esparza-Araiza et al., 2015). Phase 4 As outlined by Jones and Dangl (2006) the fourth and final phase of pathogen infection is evolution of new effector proteins that can evade ETI. When an R gene protein product is capable of resisting an effector molecule the R gene spreads in frequency throughout the host population. This creates a selection pressure on the pathogen effector molecules, ineffective effectors are removed and mutations and selection pressure on the effector molecules results in new effector molecules. The new effector molecules have no corresponding host resistance genes, resulting in susceptible plants. This results in a virulent pathogen and due to few plants having the corresponding R gene it results in a lot of disease susceptible plants. This places a selection pressure on host R genes, when a new R gene is formed it will quickly spread through the population resulting in resistant plants and the cycle will continue again. As SUMO is a PTM it is not yet clear if SUMO would play a role an important role in evolution. However as has already been highlighted some pathogen effector molecules have SUMO protease activity (Orth et al., 2000; Hotson et al., 2003) so it

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could be hypothesized that through evolution their target proteins to deSUMOylate may change. Future work Numerous other proteins that play an important role in plant immune defence have been identified as SUMO targets. These proteins were identified by purifying Arabidopsis thaliana SUMO-conjugated proteins after the Arabidopsis was subjected to a general stress and identifying the proteins using tandem mass spectrometry. The proteins identified included transcriptional repressor of defence-related genes HDA19 (histone deacytalase19), transcriptional corepressors like TPL (Topless), stress-responsive transcription factors, including five different WRKYs and vital regulator of basal immunity EIN3 (ethylene insensitive 3) (Miller et al., 2010). The effect of SUMO on these proteins still need to be determined, highlighting the amount of research still required in the field to fully understand the role of SUMO in plant immunity. A novel angle to study the role of SUMO and pathogens is to determine if viruses impact on the SUMO system. It has already been reported in human cells that pathogenic viruses target human cascades. This has been reviewed by Boggio and Chiocca (2006), the review highlights how SUMO appears to facilitate viral infection suggesting the SUMOylation machinery may be targeted by antivirals. Two of the main uses viruses have of SUMO system is SUMOylating viral transcription factors and SUMOylating viral regulatory proteins to localize near or inside the nuclear membrane (Boggio and Chiocca, 2006). The interaction of plant viruses and host SUMOylation system has not yet been studied but may yield new ways to manage plant viruses. As has been highlighted by de Vega et al. (2018) SUMOylation may provide a role in priming, along with other PTMs. It was hypothesized due to the siz1-1 mutant and ots1ots2-1 double mutant showing constitutive SAR and resistance against Pst (Lee et al., 2007; Bailey et al., 2016). Priming of plants involves three stages an initial prepriming stimulus or ‘naïve’ phase, a postpriming stimulus or ‘primed phase’ resulting in transcriptional, post‐translational, metabolic, physiological and epigenetic reprogramming. The third and final phase is the

‘post primed’ state typified by an amplified sensitization or perception of immune inducing signals (de Vega et al., 2018). As primed plants have an increased number of ‘inactive’ immune receptors these seem likely be controlled by PTMs (de Vega et al., 2018). However, very little research has been carried out in the field, examining if SUMO plays a role in priming, but it could potentially yield novel methods of controlling pathogen development. Finally, a lot of immunity research on SUMO, focuses on Arabidopsis thaliana, more research into crops is required to help develop better methods of control of plant pathogens. Firstly, characterization of the world crops SUMO systems needs to be undertaken, then studies into the interactions of the SUMO system and immunity can be undertaken. A challenge, however in using the SUMO system in crops is it may require genetic manipulation to be efficient against pathogens, which currently means those crops may not be commercially viable in many countries. Conclusions As has been shown in the knockout data for three of the SUMO proteases (OTS1, OTS2 and ESD4), these three protease mutants show elevated pathogen resistance due to increased levels of SA and therefore pathogen resistance genes. In these mutants, it has been shown there are elevated levels of SUMOylated proteins (Conti et al., 2008; Villajuana-Bonequi et al., 2014; Srivastava et al., 2018), it has been suggested transcription factors that suppresses the expression of early defence genes may be SUMOylated (van den Berg et al., 2010). Furthermore, specific examples have been given of the direct effects of SUMOylation on an important pathogen detection protein FLS2 on pathogen resistance, providing a direct model of SUMO and immunity (Orosa et al., 2018). Lastly the evidence that different pathogens actively infect plant cells with SUMO proteases, to aid its disease progression (Orth et al., 2000; Hotson et al, 2003) demonstrate the critical role SUMO plays in immunity. SUMO has been shown to directly impact on pathogens due to SUMOylating important proteins involved in immunity and indirectly by affecting hormone pathways that affect immunity. It is unsurprising that SUMO plays as important role in plant immunity as a large-scale proteomics

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analysis of Arabidopsis thaliana proteins that are SUMOylated identified numerous nuclear proteins involved in transcription, RNA metabolism, chromatin remodelling and DNA repair (Miller et al., 2010). This highlights SUMO as an important PTM deployed by the plant when a pathogen infection is occurring (Verma et al., 2018). It is worth noting, however, that despite SUMO playing a critical role in plant immunity, it has a pivotal role in all biological processes and is not just limited to immunity, hence its ability to both directly and indirectly and both positively and negatively impact on disease progression. The SUMO system is a highly suitable PTM to be employed on pathogen detection due to is ability to conjugate proteins, modify their behaviour and deconjugate rapidly in a dynamic system. This system is capable of rapidly responding to a changing landscape of pathogens and switching, quickly, plant development between growth and defence. References Bailey, M., Srivastava, A., Conti, L., Nelis, S., Zhang, C., Florance, H., Love, A., Milner, J., Napier, R., Grant, M., et al. (2016). Stability of small ubiquitin-like modifier (SUMO) proteases OVERLY TOLERANT TO SALT1 and -2 modulates salicylic acid signalling and SUMO1/2 conjugation in Arabidopsis thaliana. J. Exp. Bot. 67, 353–363. https://doi.org/10.1093/jxb/erv468. Bartetzko, V., Sonnewald, S., Vogel, F., Hartner, K., Stadler, R., Hammes, U.Z., and Börnke, F. (2009). The Xanthomonas campestris pv. vesicatoria type III effector protein XopJ inhibits protein secretion: evidence for interference with cell wall-associated defense responses. Mol. Plant Microbe Interact. 22, 655–664. https://doi. org/10.1094/MPMI-22-6-0655. Boggio, R., and Chiocca, S. (2006). Viruses and sumoylation: recent highlights. Curr. Opin. Microbiol. 9, 430–436. Casey, M., Srivastava, M., and Sadanandom, A. (2017). Posttranslational modifications in plant disease resistance (Wiley Online Library). https://doi. org/10.1002/9780470015902.a0023736. Castaño-Miquel, L., Mas, A., Teixeira, I., Seguí, J., Perearnau, A., Thampi, B.N., Schapire, A.L., Rodrigo, N., La Verde, G., Manrique, S., et al. (2017). SUMOylation inhibition mediated by disruption of SUMO E1-E2 interactions confers plant susceptibility to necrotrophic fungal pathogens. Mol. Plant 10, 709–720. Castro, P.H., Bachmair, A., Bejarano, E.R., Coupland, G., Lois, M.L., Sadanandom, A., van den Burg, H.A., Vierstra, R.D., and Azevedo, H. (2018). Revised nomenclature and functional overview of the ULP gene family of plant deSUMOylating proteases. J. Exp. Bot. 69, 4505–4509. https://doi.org/10.1093/jxb/ery301. Colby, T., Matthäi, A., Boeckelmann, A., and Stuible, H.P. (2006). SUMO-conjugating and SUMO-deconjugating enzymes from Arabidopsis. Plant Physiol. 142, 318–332.

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Index

A Acquired immunity  390, 391, 395, 402, 403 Activity-based probes (ABPs)  15, 20–26, 28, 114 Affinity tags  73–75, 88 Ageing  8, 167, 198, 290, 292, 294, 357, 358, 363, 364, 368–370, 372, 462 Aggrephagy  354, 355 AIDs  43, 313, 417–420, 425, 440, 442, 443 AIM-like receptors (ALRs)  379, 383 Alternative lengthening of telomeres (ALT)  294, 295 Alzheimer’s disease  294, 332, 338, 357, 457, 462–465, 467, 473, 474 Amyloid  332, 338, 357, 368, 369, 453–476 Amyotrophic lateral sclerosis (ALS)  460–462, 473 Angiogenesis  167, 313–323 APOBEC3  421, 423, 431, 434, 436–438, 441 ASC protein  380–385 Autophagy  15, 41, 42, 168, 194, 197, 207, 264, 292, 315, 349–358, 367, 370, 383, 399, 454, 473 Auxiliary (Aux) mediated ligation  18

B Base excision repair (BER)  95, 263, 264, 277, 278 BCA2  434, 435, 442, 443 B cells  43, 390, 402, 403 Biotin strategies  101–115

C Caenorhabditis elegans  217, 218, 249, 363–372 Cancer  15, 17, 26–28, 35, 39–43, 46, 51, 56, 59, 60, 166, 167, 170, 171, 173, 176, 177, 180–184, 218–222, 224, 237, 241, 263, 277, 279, 289, 290, 292, 294, 296, 297, 307, 308, 313, 314, 319, 357, 358, 372, 379, 383, 417, 418, 434, 472 Cell cycle regulation  233, 234 Chaperones  170, 193–199, 201, 202, 209, 234, 365 see also Heat shock proteins CRM1  174, 306, 307 Cytoplasmic polyadenylation element binding protein 3 (CPEB3)  471, 472, 475 Cytosolic DNA receptor  399, 400

D Dentatorubral-pallidoluysian atrophy (DRPLA)  469

Deubiquitinases (DUBs)  5, 6, 16–23, 25–28, 104, 105, 112–115, 171–173, 219, 222, 237, 245, 265, 266, 268, 269, 274, 275, 279, 304, 315, 337, 353, 365, 380, 382, 383, 398, 403, 454, 471 Dicer  218, 223 DNA damage  9, 10, 15, 39, 42, 55, 59, 95, 101, 104, 108, 141, 167, 169, 172, 174, 181, 183, 222, 234, 235, 239, 242, 243, 245, 247, 248, 263–279, 289–292, 296, 297, 304, 306, 307, 315, 316, 332, 337 DNA primase  234, 246, 290 DNA repair  10, 38–41, 59, 72, 123, 163, 167, 169, 242, 246, 248, 266, 274, 289–291, 293, 297, 307, 366, 368, 426, 427, 438, 454, 498 DNA replication  45, 51, 59, 114, 231–250, 264, 274, 290, 291, 393 Drosha  128, 217, 218, 221, 223, 224

E Effector-triggered immunity (ETI)  489–491, 495, 496 Effector-triggered susceptibility (ETS)  489, 490, 494, 495 Endocytosis  15, 72, 305, 307, 402, 454 Ephrins  314, 317, 323 Epstein-Barr virus  40, 42, 43, 224, 225, 392, 393

F FACS  79, 88, 111, 124–127, 130, 131 FGFR  314, 322 Fluorescence protein reconstitution  123–129, 130, 131 FRET  26, 27 FUS protein  460–462, 473

H Hayflick limit  290 HDAC  39, 42, 54, 294, 354, 355, 430, 470 Heat shock proteins (HSP)  53, 56, 170, 195, 197–199, 201, 202, 204, 292, 319, 320, 470, 473 see also Chaperones HECT  8, 15, 16, 23, 25–28, 106, 136, 148, 169, 177, 181, 219, 222, 235, 264, 265, 269, 293, 304, 337, 363, 364, 368, 427, 434 Hepatitis C virus  40, 41, 392 Homologous recombination (HR)  55, 59, 248, 263, 264, 267–274, 295, 296, 306 Human immunodeficiency virus (HIV)  391, 392, 417–443

502  | Index

Human papillomavirus (HPV)  40–42, 45, 167, 384, 394, 402 Human T-cell leukemia virus  45 Huntington’s disease  332, 338, 357, 468, 469, 475 Hypoxia inducible factor (HIF)  56, 60, 314, 319–321, 357, 369

I IL-1β  379, 380, 384, 385, 398 IMS proteins  206 Inclusion bodies  457, 458, 460–462, 467–470, 474, 475 Inflammasome  379–385, 398, 399 Innate immunity  218, 265, 316, 370–372, 389–391, 395–402, 441, 490 Interferon  4, 40, 43, 218, 322, 389, 390, 393–398, 400–402, 431–433, 436, 454 Inter-strand crosslink repair (ICLR)  263, 264, 276 Intrinsic immunity  390–395 IPTP 204 Ischaemia  317, 320, 331–342 ISG15  4, 17, 22, 26, 77, 81, 82, 86, 91, 103, 106, 247, 397, 454

J J-proteins  195, 202, 203

K Kaposi’s sarcoma-associated herpesvirus  40, 43, 44, 393, 418

L La  332, 472, 475 Lewy bodies  458–460 LUBAC  266, 383, 397, 398 Lysosomes  193, 194, 197, 207, 208, 316, 349, 351, 353, 354, 385, 441

M MARCH proteins  207, 382, 396, 397, 402, 403, 431, 433, 434, 472 Mass spectrometry (MS)  27, 71–92, 95–97, 103, 105–107, 109, 111, 115, 124, 128, 129, 135, 136, 143, 147, 148, 152, 154, 156, 157, 159, 184, 245, 249, 340, 341, 462, 466, 497 MaxQuant  85, 157–159 MDM2  17, 28, 166–178, 182, 183, 221, 247, 277, 320, 403, 428, 434 Merkel cell polyomavirus  40, 45 microRNAs (miRNAs)  43, 45, 217–225, 333, 334, 461 Minichromosome maintenance proteins (MCMs)  233, 238, 239, 241, 243–245, 248, 249 Mitochondria  57, 59, 103, 107, 112, 124, 128, 182, 193–209, 355, 356, 366, 370, 397, 460, 472 Mitochondrial anti-viral signaling protein (MAVS)  397, 398, 472, 475 Mitochondrial derived vesicles  207, 208 MitoCPR  204, 206 Mitophagy  194, 197, 206–208, 355, 356, 369 mPOS 204 Multiple systems atrophy (MSA)  460 Mx proteins  394, 395, 431

N Native chemical ligation (NCL)  18, 23, 26 NDSM  37, 53 see also SUMO consensus motif NEDD  17, 20, 22, 26, 37, 62, 72, 77, 81, 82, 91, 103, 106, 108, 177, 181, 183, 222, 223, 267, 339, 365, 370, 434, 454 Neuronal intranuclear inclusion disorder (NIID)  470, 471, 475 NOD-like receptors (NLRs)  379–385, 395, 398, 399 Non-homologous end joining (NHEJ)  263, 264, 266–269, 274, 275, 296 NOTCH  222, 306, 314, 317–319, 322 Nuclear export sequence (NES)  57, 174, 306 Nuclear localization signal (NLS)  7, 57, 148, 306, 392, 427 Nucleocytoplasmic shuttling  7, 305–307, 401 Nucleotide excision repair (NER)  239, 263, 264, 276, 277

O OMAD  205, 206 Origin recognition complex (ORC)  232, 233, 238, 242, 243

P p53  17, 28, 41–44, 139, 163–185, 221, 222, 264, 295, 306, 385 p63  163–166, 174–184 p73  163–166, 178, 180–184 PAMP-triggered immunity (PTI)  489, 490, 494 Parkinson’s disease  27, 107, 292, 332, 338, 357, 457, 459, 460, 473 Pathogen–associated molecular pattern (PAMP)  379–381, 390, 395, 489–491, 494, 496 Pattern recognition receptor (PRR)  379–381, 390, 395, 398, 400, 490, 494 PCNA  10, 139, 239, 240, 243, 244, 246–248 PDSM  37, 53, 72 see also SUMO consensus motif Pexophagy  351, 356 PIAS  8, 39, 41–43, 53–55, 98, 138–140, 143, 144, 173, 174, 183, 224, 245, 246, 250, 266, 268, 274–276, 295, 304, 306, 315, 320, 321, 323, 366, 371, 393, 397, 401, 403 PIN+  473, 475 PML  6, 8–10, 36, 38–40, 44, 45, 52, 54, 56–58, 72, 81, 98, 109, 142, 175, 181, 184, 264, 267, 269, 294, 307, 316, 321, 322, 371, 392–394, 470 Polyglutamine disorders  467, 468, 471, 475 Prions  456, 457, 471, 473, 475 Proteaphagy 356 Protein microarray  135–144 PSI+  473, 475

R Rev protein  435 Ribophagy 355 Ribosomal quality control (RQC)  112, 194, 208, 209 Rig-like receptor (RIG)  395, 397, 398, 472 RING  8, 10, 15, 16, 23, 25, 28, 38, 54, 55, 112, 148, 167, 169, 171, 175, 178, 182, 184, 207, 219, 221, 222, 235, 236, 247, 264–266, 268, 274, 275, 293, 304, 337, 356, 365, 382, 391, 397, 427, 431–434, 456, 472

Index |  503

RNA-induced silencing complex (RISC)  218, 221 RNF4  10, 38, 43, 45, 55, 72, 81, 112, 151, 157, 178, 225, 246, 274, 295, 304, 307 see also SUMO-targeted ubiquitin ligases ROS signaling  367, 368, 370, 371, 494

S SAE1/2  7, 36, 40, 52, 139, 179, 220, 266, 304, 306, 315, 332, 335, 455, 463, 492, 494 SENP (Sentrin proteases)  5–7, 22, 36, 37, 39, 40, 44, 51–53, 55–63, 84, 97, 114, 123, 173, 176, 220, 242, 266, 267, 274, 304, 316–320, 323, 332, 333, 336, 389, 399, 400, 403, 442, 456, 463 SENP inhibitors  51, 55, 60–63, 336 Sequential peptide strategy  83, 88, 89 Sequential protein strategy  87–89 Shelterin  289, 291, 292, 295–297 SMT  51, 57–59, 108, 455 see also SENP SOD1  457, 460, 461, 473 Solid phase peptide synthesis (SPPS)  18–20 Spinal and bulbar muscular atrophy (SBMA)  469 Spinocerebellar ataxia (SCA)  469, 470, 475 STAT  53, 221, 266, 314, 322, 390, 400–402, 430, 431, 435, 436, 438, 442 Stress response  38, 51, 55, 72, 108, 183, 193, 204, 209, 293, 333–335, 363–372, 491 Stroke see Ischaemia SUMO αK-GG strategy  76, 81, 82, 86 SUMO αK-NQTGG strategy  82–84, 88 SUMO consensus motif  8, 9, 37, 38, 53–54, 72, 128, 140, 141, 144, 172, 176, 179, 184, 220, 267, 303, 332, 363, 455, 459, 461 see also NDSM and PDSM SUMO-interaction motifs (SIMs)  9, 10, 38, 43, 44, 54, 55, 58, 72, 98, 128, 143, 144, 175, 178, 246, 304–307, 365, 367, 371, 372, 391–394, 427, 433, 492, 495 SUMO KO strategy  79–81, 86 SUMO Lys-C+Asp-N strategy  85, 86 SUMO-targeted ubiquitin ligases (STUbls)  10, 38, 39, 44, 45, 55, 58, 72, 178, 184, 295, 304, 365, 393 SUMO WaLP strategy  84, 85, 96, 97 Synucleinopathies  458–460, 473

T Tau  338, 357, 457, 462, 465–467, 473, 474, 476, 491

T cells  307, 390, 398, 402, 403, 418, 419, 422, 430, 435, 438, 439, 441 TDP-43  460–462, 473 Telomere  39, 167, 289–297 TIE2 322 T-loops  290, 291 Toll-like receptor (TLR)  9, 353, 381, 383, 384, 395–399 TRIM proteins  174, 221, 334, 381–383, 391, 392, 395–400, 421, 431–434, 436, 441, 472 TULIP 147–159

U Ubc9  7, 8, 36, 37, 40–45, 52–54, 58, 173, 177–179, 183, 220, 224, 245, 248, 250, 266, 304, 307, 315, 320, 332–335, 357, 363, 366, 367, 371, 392, 430, 455, 463, 465, 469 Ubiquitin code  9, 15, 103, 104, 265, 266, 278, 337 Ubiquitin-interacting motif (UIM)  9, 269 Ubiquitin proteasome system (UPS)  25, 177, 193–209, 221, 222, 234, 237–238, 243, 245, 292–296, 337–340, 349–354, 356–358, 364, 365, 367, 369–372, 436, 454, 458–461, 465, 467–471, 473–475 Ubiquitin remnant strategy  76–78, 84, 86, 88 Ubiquitin-specific proteases (USPs)  5, 6, 16, 17, 22, 26, 28, 54, 97, 112, 114, 172, 219, 222, 242, 245, 247, 250, 265, 267, 268, 274, 277, 295, 304, 305, 316, 320, 337, 341, 353, 382, 397–400, 403, 470, 471 UbiSite strategy  77, 78, 86, 90, 91, 106 Ulp  6, 37, 53, 55–58, 242, 243, 267, 366, 370, 492, 494 see also SENP Unfolded protein response (UPR)  200, 204, 371

V Vascular endothelial growth factors (VEGFR)  128, 305, 314, 316, 317, 319, 321, 323 Vif protein  421, 423, 434, 436–439, 441 Vpr protein  278, 421, 423, 424, 427, 428, 436, 438, 439 Vpu protein  421, 423, 428, 436, 438–441

W WNT  58, 112, 176, 314, 322, 323

X Xenophagy  354, 355

Y Yeast two-hybrid  43, 124, 130, 135, 143, 177

SUMOylation and Ubiquitination Current and Emerging Concepts

Most proteins undergo post-translational modifications that alter physical and chemical properties, folding, conformation distribution, stability, activity and function. Ubiquitin and SUMOs are related small proteins that are members of the large ubiquitin superfamily of post-translational modifiers. Written by highly respected leaders in their fields under the expert guidance of the editor, this volume covers the principles of ubiquitination and SUMOylation, presents detailed reviews of current and emerging concepts and highlights new advances in all areas of SUMOylation and ubiquitination. Topics of note include the ubiquitin superfamily, the ubiquitin toolbox, onco viral exploitation of the SUMO system, small molecule modulators of desumoylation, mass spectrometry, global proteomic profiling of SUMO and ubiquitin, biotin-based approaches, genetic screening, SUMOylation networks in humans, targets for ubiquitin ligases, regulation of p53, protein homeostasis, miRNAs, DNA replication, DNA damage response, telomere biology, intracellular trafficking, regulation of angiogenesis, brain ischemia, autophagy, assembly and activity, antiviral defence, HIV infection, amyloid and amyloid-like proteins and plant immunity. This comprehensive and up-to-date book is the definitive reference volume on all aspects of SUMOylation and ubiquitination and is an essential acquisition for anyone involved in this area of biology.

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