And Ubiquitin Ligases (E3s)

STRUCTURAL AND FUNCTIONAL RELATIONSHIPS BETWEEN UBIQUITIN CONJUGATING ENZYMES (E2S) AND UBIQUITIN LIGASES (E3S)

Jenny (Hong) Hong

A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Medical Biophysics University of Toronto

©Copyright by Jenny Hong Hong 2013

Structural and Functional Relationships between Ubiquitin Conjugating Enzymes (E2s) and Ubiquitin Ligases (E3s)

Jenny Hong Hong

Master of Science

Graduate Department of Medical Biophysics

University of Toronto

2013 Abstract

The first part of the thesis describes a systematic function analysis that identified in vitro E2 partners for ten different HECT E3 ligase proteins. Using mass spectrometry, the linkage composition for the resulting autoubiquitylation products of a number of functional E2-HECT pairs was determined. HECT domains from different subfamilies catalyze the formation of very different types of Ub chains, largely independent of the E2 in the reaction.

The second part of the thesis describes the characterization of the RAD6-interactome. Using affinity purification coupled with mass spectrometry, I identified a novel RAD6-interacting E3 ligase, KCMF1, which binds to a different surface on RAD6 than the other RAD6-associated E3 ligases. KCMF1 also recruits additional proteins to RAD6, and this new complex points to novel

RAD6 functions. Interestingly, the RAD6A R11Q mutant polypeptide, found in X-linked mental retardation patients specifically loses the interaction with KCMF1, but not with other RAD6- associated E3 ligases.

Acknowledgments

During my time working on the project, I was fortunate to receive several contributions from many individuals. Without their assistance, I would not have been able to complete my thesis.

I am greatly indebted to my supervisor, Dr. Brian Raught, who has provided me with ample advice, guidance and encouragement in helping me to develop better research skills. Thank you for allowing me to present at various conferences. I particularly enjoyed the trip to San Diego. I had a very memorable learning experience at the lab.

I would like to thank my supervisory committee members, Dr. Cheryl Arrowsmith, Dr. Daniel Durocher, and Dr. Stephane Angers. Their advice and support have been instrumental to the development of my project. In particular, I would like to thank Dr. Arrowsmith for allowing me to use the resources at her lab to complete the structural component of my project. I have come to love using the LEX system. I would also like to thank the members of the Arrowsmith lab, especially Dr. Lilia Kaustov and Shili Duan, who have taken the time to teach me how to purify proteins and perform NMR experiments. I had a lot of fun hanging at the lab.

I would like to thank all the past and present members of the Raught lab, for keeping a friendly and collaborative workspace. I would particularly like to thank Etienne, for his constant humor in the lab and his input on my experiments. Thank you Michael for sharing your experiences and knowledge with me. It has been nice to have someone at the lab with whom I can speak Chinese to. I also would like to thank Yasmina, Tharan, Deborah, Megan, and Connie for all the fun that we had.

Finally, I would like to thank my family and friends, who have been with me through the tough and happy times. I would like to dedicate this thesis to my mother, Yumin, who have given up so much in order for me to study at Canada. I hope that she will be proud of who I am today and what I have accomplished so far. Hopefully, I will become the woman that she has always wanted me to be.

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Table of Contents

Chapter 1 ...... 1

1 Introduction ...... 2 1.1 The ubiquitin system ...... 2 1.1.1 E1 Ubiquitin Activating Enzyme ...... 5 1.1.2 E2 Conjugating Enzyme ...... 6 1.1.3 E3 Ubiquitin Ligases ...... 7 1.2 E2-E3 interactions and mechanisms of ubiquitylation ...... 10 1.2.1 E2-E3 interaction ...... 10 1.2.2 Selecting for the correct Ub linkage ...... 11 1.2.3 Mass spectrometry-based analysis of linkage types or ubiquitin products ...... 15 1.3 Rad6 and its E3 ligases ...... 16 1.3.1 Functional relationships between Rad6 and its E3 ligases ...... 17 1.3.2 Structural relationships between Rad6 and its E3 ligases ...... 20 1.3.3 Rad6 and X-linked Mental Retardation (XLMR) ...... 22 1.4 Affinity purification coupled to mass spectrometry and protein-protein interactions ..... 23 1.4.1 Analysis of protein interaction data ...... 24 1.5 Thesis rationale and outline ...... 26

Chapter 2 ...... 28

2 A human ubiquitin conjugation enzyme (E2)-HECT E3 ligase function screen ...... 29 2.1 Chapter overview ...... 29 2.2 Contributions ...... 29 2.3 Materials and Methods ...... 30 2.3.1 Plasmids ...... 30 2.3.2 Protein purification ...... 30 2.3.3 Autoubiquitylation and E2 loading assays ...... 30 2.3.4 Mass Spectrometry ...... 31 2.4 Results ...... 32 2.4.1 Construction of an E2 library ...... 32 2.4.2 An E2-HECT E3 functional screen ...... 34

2.4.3 Characterization of Ub chain linkages in E2 – HECT reactions ...... 39 2.5 Discussion ...... 40

Chapter 3 ...... 42

3 Identification of a novel RAD6-associated E3 ligase, KCMF1, and specific loss of KCMF1 binding by an X-linked mental retardation-associated mutant R11Q RAD6A protein ...... 43 3.1 Chapter overview ...... 43 3.1.1 Contributions ...... 44 3.2 Materials and Methods ...... 44 3.2.1 Expression Constructs ...... 44 3.2.2 Stable Cell lines ...... 45 3.2.3 Transfection and shRNA interference ...... 45 3.2.4 Affinity purification ...... 45 3.2.5 Mass Spectrometry ...... 46 3.2.6 Protein Expression and Purification ...... 47 3.2.7 In vitro RAD6A binding assay ...... 47 3.2.8 NMR sample preparation ...... 48 3.2.9 NMR Spectroscopy and Data Analysis ...... 48 3.2.10 Autoubiquitylation assay and mass spectrometry analysis ...... 49 3.2.11 Immunofluorescence and Image Acquisition and Processing ...... 49 3.2.12 Live-cell confocal microscopy ...... 50 3.2.13 qRT-PCR ...... 50 3.3 Results ...... 53 3.3.1 A mammalian RAD6 interactome ...... 53 3.3.2 A RAD6A XLMR mutant protein loses the interaction with the newly identified RAD6 interactors ...... 54 3.3.3 KCMF1 interact with all the newly identified RAD6 interactors ...... 56 3.3.4 KCMF1 directly binds to RAD6A in vitro ...... 57 3.3.5 The C-terminus of KCMF1 is necessary and sufficient to bind to RAD6A ...... 58 3.3.6 RAD6A binds to KCMF1 through its N-terminal helix and residues around its active site 60 3.3.7 KCMF1 inhibits RAD6A chain formation activity in vitro ...... 63

3.3.8 Intracellular localization of KCMF1 ...... 66 3.3.9 KCMF1 may localize to endoplasmic reticulum and lysosomes ...... 70 3.3.10 KCMF1 localizes to aggresomes in response to proteasome inhibition ...... 71 3.3.11 Loss of RAD6A in cells results in a lower number of KCMF1-positive aggresomes 75 3.4 Discussion ...... 76 3.4.1 A novel RAD6 associated E3 ligase, KCMF1 ...... 76 3.4.2 Functional relationships between RAD6 and KCMF1 ...... 78 3.5 Future Directions ...... 80

References ...... 82

Supplementary Information ...... 98

List of Tables

Table 2.1 Construction of a human E2 library…………………………………………………...34

Table 3.1 Primers used for cloning………………………………………………………………52

Table 3.2 A RAD6 interactome………………………………………………………………….54

Table 3.3 RAD6A R11Q mutation disrupts protein-protein interactions………………………..55

Table 3.4 Interactome of KCMF1………………………………………………………………..57

Table 3.5 KCMF1’s C-terminus interacts with RAD6A in vivo………………………………....60

Supplementary Table 1. Raw data showing aggresome-counting……………………………....98

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List of Figures

Figure 1.1 The ubiquitylation cascade...... 3

Figure 1.2 The ubiquitin code...... 5

Figure 1.3 Mechanisms of linkage-specific ubiquitin chain assembly...... 12

Figure 1.4 Biological functions of Rad6...... 18

Figure 1.5 ProHits Analyst module-Comparison page...... 25

Figure 2.1 Ub loading assays...... 35

Figure 2.2 Autoubiquitylation reactions...... 38

Figure 2.3 Ub linkage analysis...... 40

Figure 3.1 The KCMF1 C-terminus can bind to RAD6A ...... 58

Figure 3.2 KCMF1 interacting domains ...... 60

Figure 3.3 The interaction of KCMF1 CT with RAD6A...... 62

Figure 3.4 A comparison of the RAD6 binding sites for Rad18, Ubr1 and KCMF1...... 63

Figure 3.5 RAD6A autoubiquitylation reactions...... 64

Figure 3.6 Intracellular localization of KCMF1...... 67

Figure 3.7 Intracellular localization of the RAD6A and RAD6A R11Q proteins...... 68

Figure 3.8 KCMF1 colocalizes with RAD6A in the cytosol...... 68

Figure 3.9 KCMF1 proteins are in both the cytoplasmic and microsomal cellular fractions...... 69

Figure 3.10 KCMF1 localizes to ER and lysosomes...... 71

Figure 3.11 Proteasomal inhibition induces KCMF1 to form perinuclear aggregates...... 73

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Figure 3.12 ABHD10 localizes to mitochondria...... 75

Figure 3.13 Loss of RAD6A expression leads to decreasing amount of KCMF1-positive aggregates...... 76

Figure 3.14 A comparison of the RAD6 binding sites for ubiquitin and KCMF1...... 79

Supplementary Figure 1. The anti-KCMF1 antibody is specific to KCMF1 proteins…………...98

List of Abbreviations aa Amino acid

ABHD10 α/β hydrolase domain containing 10 protein

AcMPAG acyl glucuronide

AP-MS Affinity purification coupled to mass spectrometry

APC/C Anaphase Promoting Complex/cyclosome

ARF1 ADP-ribosylation factor 1

ATP adenosine‐5’‐triphosphate

Bard1 BRCA1 associated RING domain 1

BRCA1 Breast cancer type 1 susceptibility protein

BRR Basic rich region

CFTR Cystic fibrosis transmembrane conductance regulator

CID Collision-induced dissociation

COMPASS Complex Proteins associated with Set1

CSP Chemical Shift Perturbations

CT C-terminus of KCMF1

CTD C-terminus domain of Pol II

DAPI 4',6-diamidino-2-phenylindole

DSB Double stranded break

E. coli Escherichia coli

E6AP E6-associated protein

ECL Enhanced chemoluminescence

EDTA ethylenediaminetetraacetic acid

ENaC Epithelial sodium channel

ER endoplasmic reticulum

ESFT Ewing’s sarcoma family of tumors

FGF Fibroblast growth factor

FL Full length

GAPDH Glyceraldehyde 3-phosphate dehydrogenase

GFP Green fluorescent protein

GPM Global Proteome Machine

GST Glutathione S‐transferase

H. sapiens Homo sapiens

H2B Histone 2B

HECT Homologous to the E6-AP carboxyl terminus

HEK Human Embryonic Kidney

HEPES 4‐(2‐hydroxyethyl)‐1‐piperazineethanesulfonic acid

HERC Homologous to E6 associated protein carboxy-terminus and RCC1 domain protein

HERC4 HECT domain and RCC1-like domain-containing protein 4

HR Homologous recombination

HRP Horseradish peroxide

HSQC Heteronuclear Single Quantum Coherence

HUWE1 HECT, UBA and WWE domain containing 1 protein

IF Immunofluorescence

IPTG isopropyl-1-thio-D-galactopyranoside

ITCH Itchy homology E3 ubiquitin protein ligase

KCMF1 Potassium channel modulatory factor 1

LC-MS/MS Liquid chromatography-tandem mass spectrometry

LTQ Linear ion trap

MAP Microtubule associated protein

MS Mass spectrometry

MS/MS Tandem mass spectrometry

MTOC Microtubule-organizing center

NA numerical aperture

Nedd4 Neural precursor cell expressed, developmentally down-regulated 4

Nedd4L Nedd4-like protein

NHEJ Non-homologous end joining

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NIPSNAP3A 4-nitrophenylphosphatase domain and non-neuronal SNAP25-like protein (NIPSNAP) homolog 3A nLC-ESI- Nanoflow liquid chromatography electrospray ionization-tandem mass MS/MS spectrometry

NMR Nuclear magnetic resonance

NPC Nuclear pore complex

NUP Nucleoporin

OGT O-linked N-acetylglucosamine (GlcNAc) transferase pAb Polyclonal antibody

PABC Poly(A) binding protein C-terminal

PABP Poly(A) binding protein

PBS Phosphate buffered saline

PCNA Proliferative Cell Nuclear Antigen

PCR Polymerase chain reaction

PD Parkinson’s disease

PFA Paraformaldehyde

PHD Plant homeodomain

PMSF Phenylmethanesulfonylfluoride

Pol II RNA Polymerase II

PRR Post replication repair

PTM Post translational modification

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qRT-PCR Real time reverse transcription polymerase chain reaction

R6BD Rad6 binding domain

Rad18 Radiation sensitive 18 protein

Rad6 Radiation sensitive 6 protein

RING Really Interesting New Gene

RLD Regulator of chromosome condensation 1 (RCC1)-like domains)

RNF20/40 Ringer finger protein 20/40

S. Cerevisiae Saccharomyces cerevisiae

SAINT Statistical analysis of interactome

SARS Seryl-tRNA synthetase 2

SCF Skp1‐cdc53/cul1‐F‐box protein

SDS Sodiumdodecyl sulfate

SDS-PAGE sodiumdodecyl sulfate‐ polyacrylamide gel electrophoresis shRNA Small hairpin RNA

SMURF1 Smad ubiquitylation regulatory factor 1

SRM Selected reaction monitoring

SSBP1 Single stranded binding protein 1 ssDNA Single stranded DNA

TAP Tandem‐affinity purification

TEV Tobacco‐etch virus

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TLS Translesion Synthesis

TPP Trans‐proteomic pipeline

U-box Ubiquitin fusion degradation protein 2-homology box

Ub Ubiquitin

UBA Ubiquitin-associated domain

UBC Ubiquitin conjugating (domain)

UBE2 Ubiquitin E2 enzyme

UBE3A Ubiquitin protein ligase E3A

UBE3C Ubiquitin protein ligase E3C

UBL Ubiquitin-like modifiers

UBR1 Ubiquitin protein ligase E3 component n-recognin 1

UBR2 Ubiquitin protein ligase E3 component n-recognin 2

UBR4 Ubiquitin protein ligase E3 component n-recognin 4

UBR5 Ubiquitin protein ligase E3 component n-recognin 5

UBZ Ubiquitin binding zinc finger

UEV Ubiquitin E2 variant

UFD Ubiquitin fusion degradation pathway

UIM Ubiquitin-interacting motif

UPS Ubiquitin-proteasome pathway

WAC WW domain-containing adapter proteinwith coiled-coil

WWP2 WW domain containing E3 ubiquitin protein ligase 2

XLMR X-linked mental retardation

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Introduction

1 Introduction

Post-translational modifications (PTM) are chemical alterations of a protein after it has been synthesized, and are employed by the cell to regulate protein quantity, localization and activity. Ubiquitin (Ub) is one type of PTM. This thesis is focused on understanding the relationship between two important components of the Ub system, the Ub E2 conjugating enzyme and the Ub E3 ligase.

1.1 The ubiquitin system

Ubiquitin is a highly conserved 76 amino acid polypeptide that is found only in eukaryotes. Covalent attachment of ubiquitin(s) to protein substrates occurs through a coordinated chain of events that can alter the fate and function of these molecules. Ub conjugation (ubiquitylation) is a highly regulated process, consisting of a sequential series of E1-E2-E3 activation, conjugation and ligation reactions (Figure 1.1). Impairment of the Ub system can lead to many pathological effects, including cancer, neurodegenerative disorders and diabetes (reviewed in Petroski, 2008).

Ub is conjugated to target proteins via an isopeptide bond between the ε-amino group of a lysine residue in a target protein and the C-terminal carboxyl group of Ub (Behrends and Harper, 2011; Pickart, 2004; Schulman and Harper, 2009). Ubiquitylation regulates a range of critical cellular functions, and the function is dependent on the type of ubiquitin products attached to the substrate. Monoubiquitylation, the conjugation of a single Ub molecule to a target protein, has been implicated in transcriptional control, endocytosis, and DNA damage signaling (Mosesson and Yarden, 2006; Saksena et al., 2007). Mono or multiubiquitylation (monoubiquitylation at multiple lysine residues on a protein substrate) are also important mechanisms by which membrane proteins are internalized and sorted by the endocytic compartments (Bonifacino and Traub, 2003). Once internalized, membrane proteins may be either recycled by returning to the plasma membrane or degraded by sending it to the multivesicular body then to the lysosome. Newly synthesized proteins may also be monoubiquitylated, which targets them to the trans- Golgi network for sorting, either destined for the plasma membrane or the lysosome (Bonifacino and Traub, 2003)

Figure 1.1 The ubiquitylation cascade. (a) Ub is first activated by the E1 protein before it is transferred to the active-site Cys of an E2 Ub conjugating enzyme (b). (c) The “charged” E2 then interacts with an E3 Ub ligase, which transfers the Ub from E2~Ub to a Lys residue of a substrate protein. HECT E3s contain a conserved catalytic Cys, which forms an intermediate with Ub before transfer to a Lys residue in the substrate (S=substrate). RING E3s lack a catalytic Cys: they act as scaffolds by facilitating interactions between E2s and substrates. A monoubiquitylated substrate can either dissociate from the E3 (d) or can acquire additional Ub modifications in the form of Ub chains (e). Each Ub moiety in the chain is linked to the next through Lys residues. The biological outcome of ubiquitylation is dictated by Ub receptors, proteins that can bind to Ub(s) and interpret the Ub code. MonoUb and some types of chains (e.g. those assembled via Lys63 of ubiquitin) serve mainly to alter the function of the modified protein by changing its structure, binding partners, cellular localization, and so on (f). PolyUb chains assembled via the Lys48 residue of Ub typically direct the protein to the proteasome for degradation (g).

Ub chains of varying lengths and linkage types can confer very different biological outcomes to a targeted protein substrate. Ub itself contains seven lysine residues, all of which can be ubiquitylated (Deshaies and Joazeiro, 2009; Finley, 2009; Pickart and Eddins, 2004; Pickart and Fushman, 2004; Ye and Rape, 2009). The best characterized function of Ub chains is the

4 targeting of ubiquitylated proteins (in this case, consisting of at least four Ub polypeptides linked via K48 linked chains) to the 26S proteasome for degradation (Thrower et al., 2000). K11-linked Ub chains have also been reported to function as proteasomal targeting sequences (Dammer et al., 2011; Xu et al., 2009). Proteasome-dependent proteolysis regulates many other processes in the cell, including progression of the cell cycle (Koepp et al., 1999), induction of the inflammatory response (Ghosh et al., 1998), and antigen presentation (Rock and Goldberg, 1999). By contrast, K63-linked Ub chains play roles in the DNA damage response, epsin- mediated endocytosis and aggresome formation (Acconcia et al., 2009; Chin et al., 2010) (Figure 1.2). In addition to polyUb chains formed by the conjugation of Ub carboxy-terminal glycine to an internal lysine residue of another Ub, there is also linear-type Ub chains. Linear Ub chains are formed in which the C-terminal glycine is conjugated to the α-amino group of the amino-terminal methionine of another Ub (Iwai and Tokunaga, 2009). Linear Ub chains are generated by a specific Ub ligase complex, named linear ubiquitin chain assembly complex (LUBAC) (Kirisako et al., 2006). LUBAC has been shown to be important in nuclear factor- kappa B (NF-κB) activation (Iwai and Tokunaga, 2009).

Figure 1.2 The ubiquitin code. Ubiquitin is usually attached to the ε-amino group of Lys residues in substrates. The transfer of a single ubiquitin to one (monoubiquitylation) or multiple (multi‑monoubiquitylation) sites can affect protein interactions, alter protein localization or modulate protein activities (Kerscher et al., 2006; Mukhopadhyay and Riezman, 2007). Ubiquitin itself contains seven Lys residues, all of which can accept another Ub moiety during chain assembly. Depending on the connection between linked ubiquitin molecules, ubiquitin chains can differ in structure and function. Lys48- and Lys11- linked ubiquitin chains have a compact structure (Komander, 2009) and target proteins for degradation by the 26S proteasome. On the other hand, Lys63- linked chains are elongated in structure and play roles in DNA repair, targeting modified protein to lysosome and epsin-mediated endocytosis. The functions of other Ub chains are poorly defined.

1.1.1 E1 Ubiquitin Activating Enzyme

The E1 activating enzyme executes the first step in the ubiquitylation cascade. Upon first binding to MgATP, the E1 forms a ubiquitin adenylate intermediate (Pickart, 2001) which serves as a donor of ubiquitin to the active site cysteine in the E1 (Haas and Rose, 1982; Hershko et al., 1983). Each “loaded” E1 enzyme carries two activated ubiquitins, one as an adenylate and one as a thiol-ester. The thiol-linked ubiquitin is then transferred to the conjugating enzyme in the cascade, E2. In most organisms, a single E1 enzyme activates ubiquitin for all downstream conjugating reactions (McGrath et al., 1991). However, in mammals, there are two isoforms (e.g. UBE1 and UBA6) (Handley-Gearhart et al., 1994; Jin et al., 2007). UBE1 has been identified as

6 the predominant E1 for Ub activation in human cells (Pickart and Fushman, 2004) and shows broad E2 specificity. In contrast, UBA6 interacts mainly with the UBA6-specifc E2, UBE2Z (Jin et al., 2007).

1.1.2 E2 Conjugating Enzyme

The organization of the ubiquitin system is hierarchical. While there are only two human Ub E1 activating proteins, there are about 40 E2s (including both active and inactive E2 variants) encoded in the human genome, and over 600 E3 ligase proteins, each recognizing its own set of substrates (Hershko and Ciechanover, 1998; Hochstrasser, 1996). As determined by the relatively few that have been screened, most E3 ligases seem to be able to cooperate with multiple E2s (Sheng et al., 2012; van Wijk et al., 2009).

All E2 proteins share a conserved “core” ubiquitin conjugating (UBC) domain of ~150 amino acid residues which contains the active cysteine residue responsible for accepting the activated ubiquitin molecule from the E1. E2s are grouped into families based on the presence or absence of N- and/or C-terminal extensions (Pickart, 2001). These extensions can govern intracellular localization, confer regulatory properties, or provide specificity for interactions with particular E3 ligases (Pickart, 2001; Wenzel et al., 2010). The main function of E2 enzymes is to accept the activated ubiquitin from E1s, followed by binding to E3 ligases to transfer the Ub to substrates (Pickart, 2001). Some E2s function specifically in ubiquitin-like protein (UBL) pathways. For instance, SUMO, a ubiquitin-like protein, is exclusively conjugated via UBE2I, while NEDD8, another UBL, is conjugated via UBE2M and UBE2F(Huang et al., 2009a; Kawakami et al., 2001; Schwarz et al., 1998). Some E2-like proteins, such as UBE2V1/2 possess a UBC domain but lack the active-site Cys residue required for Ub coupling (Hofmann and Pickart, 1999). These proteins are referred to as the UEVs (Ubiquitin E2 Variants). Interestingly, the UEVs often associate with an active E2, such as Ubc13 with UBE2V1/2, in a heterodimeric complex.

The E2 protein family is structurally well characterized: The structures for 31 out of the 40 human E2s have been solved (Arai et al., 2006; Giraud et al., 1998; Huang et al., 2005; Lin et al., 2002; Mace et al., 2008; Miura et al., 2002; Ozkan et al., 2005; Sheng et al., 2012; Zhang et al., 2005; Zheng et al., 2000). Most E2s display a canonical E2 fold, which consists of four alpha helices along with an anti-parallel β-sheet and a short 310 helix. Residues involved in catalysis and ubiquitin binding are well conserved (Sheng et al., 2012). Residues in the two loop regions,

L1 and L2, are less conserved, conferring a high degree of variability in sequence, conformation and length. Residues on L1, L2, and α1 helix mediate binding with both the E1 and E3(Coccetti et al., 2008; Michelle et al., 2009; van Wijk and Timmers, 2010). The flexibility in the loop regions confers selection of specific E3s (Burroughs et al., 2008). The specificity of E2-E3 interactions allows E2s to execute distinct biological functions. However, the governing principles in determining the specificity between E2 and E3 interactions are not well understood, and the subset of E3 partners for each E2 has yet to be defined.

1.1.3 E3 Ubiquitin Ligases

Ub E3 ligases facilitate the transfer of Ub from an E2 to a substrate protein or another Ub molecule. E3s are also responsible for recognizing a multitude of substrates targeted for ubiquitylation. In most cases, the substrates are not constitutively recognized by the E3s. In general, either the E3 must undergo post-translational modification to become “activated” or the substrate must be modified in order to be recognized (Glickman and Ciechanover, 2002). Moreover, E3s can interact with the substrate directly or through ancillary proteins.

Most E3 ligases (e.g. RING finger domain E3s, see below) act as scaffolds, recruiting activated E2~Ub complexes to substrates to effect the efficient transfer of the Ub moiety (Deshaies and Joazeiro, 2009). In other cases (HECT domain E3s, see below), the E3 forms a thiol-ester linkage with Ub prior to its transfer to substrates (Bernassola et al., 2008; Rotin and Kumar, 2009). An additional subset of E3s (including U-box domain containing E3s) referred to as E4s, has a role in the elongation of polyUb chains, aiding in the transfer of Ub from the E2 to a previously conjugated Ub moiety (Glickman and Ciechanover, 2002).

1.1.3.1 RING E3 ligases

The RING (Really Interesting New Gene) type E3s comprise ~95% of all E3 ligases (Rotin and Kumar, 2009). The RING domain is characterized by a stretch of histidine and cysteine residues (the canonical sequence being Cys-X2-Cys-X(9-39)-Cys-X(1-3)-His-X(2-3)-Cys-X2-Cys-X(4- 48)-Cys-X2-Cys (Deshaies and Joazeiro, 2009) that coordinate zinc ions (Borden and Freemont, 1996). RING variants have also been noted, including the U-box (UFD2-homology) domain and the PHD (plant homeodomain) domain. Although they are structurally different from the RING domain, they can mediate ubiquitylation in a manner similar to the classic RING proteins

(Aravind and Koonin, 2000; Capili et al., 2001; Hatakeyama et al., 2001). It is believed that the binding of an activated E2 to a RING E3 triggers rearrangement of the E2’s active site, exposing a critical and conserved asparagine residue that is essential for catalysis by stabilizing the transition state of the oxyanion intermediate (Ozkan et al., 2005; Wu et al., 2003).

The underlying ubiquitin ligase activity of RING domains was first shown through the c-Cbl- UbcH7 (UBE2L3) interaction (Zheng et al., 2000). The RING domain commonly makes important contacts with the E2’s first α-helix and loops 1 and 2 to activate the E2 and aid in ubiquitin transfer (Deshaies and Joazeiro, 2009). Some RING domains do not exhibit E3 activity, such as Bard1 (Hashizume et al., 2001), Bmi1 (Wang et al., 2004a), and MdmX (Linares et al., 2003). Instead, they interact with a second RING domain protein, forming a heterodimer with Brca1, Ring1b, and Mdm2, respectively, which stimulates the E3 ligase activity of the latter. Some RING E3 ligases are components of multi-subunit complexes, such as the SCF type E3s (Skp1-Cullin-F-box) (reviewed in Deshaies and Joazeiro, 2009).

The affinity of isolated RING domains for E2s is usually quite low, with KD in the high micromolar range (Deshaies and Joazeiro, 2009). Some RING E3s associate with their respective E2s via another region, outside of the RING domain, to mediate higher affinity binding. For instance the tight binding between Rad6 and Ubr1 is mediated by the BRR (basic rich region) sequence N-terminal to the RING domain (Xie and Varshavsky, 1999). However, the RING domain was shown to confer ligase activity with Rad6. On the other hand, gp78, another RING E3, also has a motif distinct from the RING that recruits the E2, but this high-affinity E2-binding domain is essential for ubiquitylation (Chen et al., 2006).

1.1.3.2 HECT E3 Ligases

The first identified family of Ub E3 ligases was the HECT (Homology to E6-Associated Protein Carboxy Terminus) domain-containing proteins (Huibregtse et al., 1995). This~350kDa domain contains an active-site cysteine, utilized to form an intermediate thioester bond with Ub (Scheffner et al., 1995). The HECT domain can be divided into two lobes, the N-terminal lobe, containing the E2 binding site, and the C-terminal lobe, containing the active site Cys (reviewed in Kim and Huibregtse, 2009). E2s known to function with HECT E3s belong to the group most similar to human UBE2L3 (UbcH7). They have a conserved Phe residue in the loop between β3 and β4, directly at the E2-HECT interface. The flexible linker sequence between N and C lobes

9 is important to position the E2 and E3 active sites into an optimal orientation, and can restrict accessibility to certain Lys residues on the target protein or on a polyUb chain.

The human genome encodes ~28 predicted HECT E3 ligases, whose prototypic member, E6AP, associates with the E6 protein encoded by human papilloma virus (HPV), and effects the proteasomal degradation of the tumour suppressor protein p53, contributing to the development of HPV-associated cervical cancer (Scheffner et al., 1994). Near the N-terminus of HECT E3 ligases are one ore more protein–protein or protein–lipid interaction domains (Rotin and Kumar, 2009). Based on the architecture of N-terminal domain(s), the human HECT E3s can be grouped into 3 families: NEDD4, HERC and other HECTs (Rotin and Kumar, 2009).

N-terminal domains confer additional functions to HECT E3 proteins. In addition to ubiquitylating proteins for degradation by the 26S proteasome, HECT E3s regulate the trafficking of many receptors, channels, transporters and viral proteins (Rotin and Kumar, 2009). For instance, the Nedd4 family members contain a C2 domain, involved in targeting proteins to the plasma membrane and intracellular vesicles (Dunn et al., 2004; Plant et al., 2000), followed by two to four WW domains (the WW domain binds to proline (PY) motifs (Staub et al., 1996). NEDD4-like protein (NEDD4L), a member of NEDD4 family, binds through its WW domains to the PY motifs of the epithelial Na+ channel (ENaC), leading to ENaC ubiquitylation, endocytosis to endosomes and multivesicular bodies, and subsequent degradation(Gormley et al., 2003; Staub et al., 1996). Several other Nedd4 family members, such as NEDD4, SMURF1/2, and ITCH, are involved in signal transduction cascades regulating cellular growth, proliferation and immune response (reviewed in Rotin and Kumar, 2009). The HERC family members all contain the RLD (Regulator of Chromosome Condensation 1 (RCC1)-Like Domains) domain. This family can be further subdivided into the large HERCs (>500 kDa), which have multiple RLDs, and the small HERCs (~100 kDa), which contain a single RLD (Rotin and Kumar, 2009). It has been reported that many HERC E3 ligases play a role in membrane trafficking (Garcia- Gonzalo and Rosa, 2005; Hochrainer et al., 2008). Moreover, through its RLD domain, the large HERC (HERC1) can stimulate guanine-nucleotide dissociation from small ADP-ribosylation factor 1 (ARF1) and the small G protein Rab (Garcia-Gonzalo et al., 2005). The “other” HECT family members contain a spectrum of domains, from ubiquitin-associated domains to ankyrin repeats. EDD (or UBR5), one of the “other” HECT family members, contains a PABC (Poly(A) binding protein C-terminal) domain. Through this PABC domain, EDD can bind to PAIP2

(PABP-Interacting Protein 2), an inhibitor of Poly(A) binding protein(PABP), and target it for degradation, to regulate the translation initiation(Yoshida et al., 2006)

1.2 E2-E3 interactions and mechanisms of ubiquitylation

Proteins can be modified with various Ub architectures, including monoUb, multi-monoUb, and many different types of Ub chains. There are seven lysine residues in Ub (K6, K11, K27, K29, K33, K48 and K63), allowing for seven possible homotypic Ub polymers, and a large number of heterotypic polymers (Hershko and Ciechanover, 1998). To form Ub chains, glycine residue 76 of one Ub moiety is conjugated to a lysine residue in a second Ub molecule(Pickart and Fushman, 2004). The different Ub chains can be thought of as different “codes”, which are interpreted by a cohort of proteins containing Ub-binding motifs, including ubiquitin-interacting motif (UIM) and ubiquitin-associated (UBA) domains (Dikic et al., 2009; Hurley et al., 2006). These proteins also bind to downstream effectors of signaling pathways, thereby translating the Ub code into the desired biological outcome. Many of these interpreters have a preference for a distinct type of Ub conjugate, such as monoUb or specific Ub chain linkage types.

Much of the work in the ubiquitin field has been focused on the E3s, because of their specificity in recruiting substrates, but recent studies have revealed that the E2s can also determine the length and topology of Ub chains. Moreover, the outcome of substrate ubiquitylation can be influenced by specific E2-E3 partnerships. The rules governing E2-E3 pairing is an important part of current research, as well as determining physiological E2 partners for many E3s.

1.2.1 E2-E3 interaction

The interaction between E2 and E3 proteins is generally quite weak, making it difficult to identify the physiological partners of E3s through standard pull-down assays (Deshaies and Joazeiro, 2009). On the other hand, the weak E2-E3 interaction is advantageous for Ub chain formation, because the binding surface for the E1s and E3s overlap on the E2 proteins; an E2 thus needs to go through several rounds of E1 and E3 binding to effect Ub chain assembly (Huang et al., 2005; Ziad et al., 2005). As mentioned previously, some E3s possess additional E2 binding sites. This could allow for an increase in the capacity to stimulate chain formation, where the second binding site allows the E2 to disengage from the RING or HECT domain for recharging while remaining attached to the E3 (Ye and Rape, 2009). The assembly of most Ub

11 chains is promoted by the E2 alone, however when in the presence of a HECT E3 ligase, the HECT domain can determine the linkage specificity (Wang et al., 2006; Kim and Huibregtse, 2009; Maspero et al., 2011).

1.2.2 Selecting for the correct Ub linkage

1.2.2.1 E2s

The assembly of Ub chains usually begins with the initial conjugation of a Ub to a Lys on a substrate (initiation stage). Subsequently, the E2–E3 pair switches to chain elongation (elongation stage), during which additional Ub molecules are attached to the substrate-linked Ub (Ye and Rape, 2009). Recently some E2s have been shown to have dedicated roles in the chain initiation or elongation stages (Ye and Rape, 2009) (Figure 1.3 A). The segregation of the two roles is highlighted in the following example. The heterodimeric E3 ligase, BRCA1-BARD1 uses both UBE2W and UBE2E2 to initiate the Ub chain (monoubiquitylation), but uses the heterodimer UBE2N-UBE2V1 (UBE2V1 is an E2 variant that lacks the catalytic cysteine) and UBE2K for Ub chain elongation (Christensen et al., 2007). Some E2s involved in chain initiation (e.g. UBE2T) not only lack chain extension activity, but also do not cooperate with other chain- elongating E2s, thereby promoting only monoubiquitylation (Alpi et al., 2008). E2s involved in chain elongation often require a conjugated Ub on the substrate (Ye and Rape, 2009). Interestingly, some chain initiation E2s can extend short chains before the elongation process begins. For instance, UBE2C adds the first Ub moiety on human E3 anaphase-promoting complex (APC/C) and is responsible for extending short K11-linked Ub chains on APC/C. UBE2S then completes the full Ub chain extension process (Rape et al., 2006; Williamson et al., 2009). Furthermore, some E2s, like the yeast Cdc34 protein (UBE2R1 in human), can both extend and elongate chains. The acidic loop near the active site in Cdc34 has been suggested to be required for efficient elongation (Gazdoiu et al., 2007; Petroski and Deshaies, 2005)

Figure 1.3 Mechanisms of linkage-specific ubiquitin chain assembly. (A) E2s can dictate Ub chain linkage specificity by separating the role between chain initiation and elongation. Some E2s can only monoubiquitylate a substrate (initiator, E2a), while other E2s can polyubiquitylate only when there is a prior Ub moiety on the substrate (elongator, E2b). S=substrate. (B) The orientation between donor and acceptor Ub is important in selecting for the correct linkage type. UBE2S lacks a residue required for suppressing the pKa of the substrate Ub Lys. This function is provided by Glu34 of the Ub, which is in direct proximity of only Lys11. Other Lys residues do not have a suitably positioned acidic residue when docked into the active site, therefore a competent catalytic site is formed only when K11 of acceptor Ub is exposed to the active site of UBE2S. The orientation between the acceptor and donor Ub is important in selecting for linkage type (Wickliffe et al., 2011). Figure adapted from (Behrends and Harper, 2011). (C) Model for how the HECT domain directs chain specificity. The N lobe of the HECT domain contains the substrate binding site, while the C lobe containsthe catalytic Cys. Different HECT E3s could orient the thioester-linked Ub molecule in different orientations, so that only K63 (left) or K48 (right) of the distal Ub of a chain can approach the active site. Like the E2, the positioning between donor and acceptor Ub is key to linkage type selectivity. The relative positions of the N and C lobe, as well as the relative position of the substrate are also important. Figure adapted from (Kim and Huibregtse, 2009).

Connecting Ub molecules in a specific fashion is an intrinsic property of E2s (Ye and Rape, 2009). In the absence of an E3, some E2s, particularly the UBE2D family members, can synthesize Ub chains of varying linkage types in vitro, while other E2s, such as UBE2S, promote only a single linkage type (K11). Structural and mechanistic studies have dissected K63, K48 and recently K11-linked Ub chain formation. E2s determine linkage specificity through the noncovalent binding of donor Ub (a Ub molecule that accepts the Lys residue of another Ub) and substrate assisted catalysis, where residues from the E2, donor Ub and acceptor Ub (Ub molecule that donates the Lys residue to another Ub) create a competent active site for catalysis (Wickliffe et al., 2011). This linkage-determining mechanism is also shared by many E2s.

In the case of the K11-specific UBE2S, in addition to the thioester bond with the donor Ub, UBE2S requires an additional noncovalent interaction restricting accessibility of Lys residues on the donor Ub from the acceptor Ub (Wickliffe et al., 2011) (Figure 1.3 B). This noncovalent interaction optimally places the C-terminus of the donor Ub for nucleophilic attack by the acceptor Lys residues. In addition, a competent catalytic center in UBE2S is formed only when K11 of the acceptor Ub is exposed to the active site. This is because UBE2S cannot suppress the pKa of the substrate Lys, but the Glu34 residue of Ub, in proximity of K11 can compensate for this function (Wickliffe et al., 2011). Other E2s use a similar mechanism in specifying linkage type. UBE2R1 and UBE2G2 also tether donor Ub, while the HECTE3 NEDD4L binds to E2- linked donor Ub (Kamadurai et al., 2009; Wickliffe et al., 2011). Overall, how the donor Ub is tethered by the E2, and how the acceptor Ub is positioned determine how a Ub chain is assembled on a substrate. The need for substrate-assisted catalysis also highlights the importance of the interaction between the E2 the E3-associated substrate, and could account for substrate specificity.

1.2.2.2 HECT E3s

The HECT E3 ligases act like an E2, whereby the HECT domain can affect the topology of the chain and mode of chain assembly. In the presence of a HECT E3 ligase, the E2 plays a less important role in selecting for the correct linkage. When UBE2D1, a promiscuous E2 that promotes K11, K48 and K63 linked chains, binds to the HECT E3 KIAA10 (UBE3C), K29 and K48 linked chains are preferentially assembled (Wang et al., 2006). This is because HECT E3 ligases are charged with Ub while binding to its substrate. Hence, the HECT E3 ligase can

14 position the substrate, and orient specific Lys residues on the acceptor Ub to promote linkage- specific chains (Ye and Rape, 2009). Like the E2s, different HECT E3s also have specificities for different types of polyUb chains. E6AP preferentially assembles K48 linked chains (Kim et al., 2007; Wang and Pickart, 2005), consistent with the fact that its substrate, p53 is targeted for proteosomal degradation. In contrast, Nedd4 has a preference for K63 chains (Maspero et al., 2011).

Mechanistically, the molecular determinants of linkage selectivity for HECT E3s are similar to the E2s. The relative position of the donor and acceptor Ub are key to this process (Figure 1.3 C). The N lobe of a HECT domain encompasses the acceptor Ub binding site (Wang et al, 2006, Maspero et al., 2011). The N lobe can affect linkage-type preference by positioning the acceptor Ub in ways that allow for easier accessibility for certain Lys residues (Kim and Huibregtse, 2009; Wang et al., 2006). In addition, Kim and Huibregtse (2009) have shown that chain specificity is also a function of the C lobe. Through the use of chimeric enzymes, swapping the N lobe, C lobe or N-C lobe linker sequence between different HECT E3s, the replacement of ITCH’s C lobe with that of E6AP’s caused a switch to the K48 linkage type as opposed to K63. They propose that the C lobe determines the orientation of the thioester bound Ub, making only certain Lys of the acceptor Ub accessible to the active site (Kim and Huibregtse, 2009).

Residues around the targeted Lys in the acceptor Ub also contribute to linkage type determination. Using alanine scanning mutagenesis and an in vitro ubiquitylation assay, Wang and coworkers (2006) mapped the surface residues on the acceptor Ub that promotes linkage specificity in chain synthesis catalyzed by the HECT domain of K1AA10. Most of the residues affecting a given linkage are close to one another. Interestingly, mutations of these residues affect K48 usage of K1AA10 but not of other K48-specific HECT E3s, such as E6AP or E2-25K (Wang et al., 2006). This was proposed to be a result of the different mechanisms HECT E3s use in catalyzing Ub chains (Wang and Pickart, 2005). KIAA10 follows the sequential addition model of polyubiquitylation where the single Ub molecule is added sequentially to the distal end of a substrate-linked chain (Wang and Pickart, 2005). In contrast, E6AP lacks an acceptor Ub binding site, thus it is thought to polyubiquitylate via an “indexation” model, where the chain is first built up on the E3 active site through successive transfers of E2-bound Ub to E3-bound Ub chain. The entire chain is then transferred in bulk to the substrate (Verdecia et al., 2003; Wang and Pickart, 2005). Further structural and biochemical analyses are required to develop a better

15 understanding of the mechanism of polyubiquitylation. Moreover, the linkage preference for many HECT E3s and E2s need to be determined, shedding light on their respective physiological functions.

1.2.3 Mass spectrometry-based analysis of linkage types or ubiquitin products

The fate of an E3 substrate is determined by the structure of the conjugated Ub moiety. Thus, determining the mono-or polyubiquitylation status of the substrate, and the type of Ub linkages, are as important as identifying the substrate itself. Immunoblotting can be used to distinguish between mono-and poly-ubiquitylated forms of a protein because of the increase in total mass. Linkage type within a poly-Ub chain may also be determined through the usage of linkage- specific antibodies. However, the lack of specific antibodies directed against all linkage types makes it difficult to determine linkage frequency and topology of Ub chains(Mirzaei et al., 2010). Lysine to arginine Ub mutants have also been used to indirectly determine Ub chain architectures (Finley et al., 1994; Spence et al., 1995). Determined through immunoblotting, a mutant Ub can shorten Ub chains that are linked through the corresponding Lys residue. Recently, however, advancements in mass spectrometry have provided a more direct approach to characterizing Ub chain topology.

Tandem mass spectrometry (MS/MS) has proved to be a powerful tool in analyzing Ub-modified proteins. It has been used to identify modification sites, to characterize the topology of Ub chains, and to measure the level of linkages in a cell (Kaiser and Wohlschlegel, 2005; Kirkpatrick et al., 2006; Xu et al., 2009). In liquid chromatography-tandem mass spectrometry (LC-MS/MS), protein samples are digested with trypsin or other proteases into peptides before being fractionated by liquid chromatography. The peptides are then ionized and analyzed by a mass spectrometer based on its mass-to-charge (m/z) ratio. Tandem mass spectrometry further fragments these peptides (usually selecting a number of the most abundant peptides in an MS scan) to generate a set of peptide fragments. The data collected from an MS/MS scan can then be compared to theoretical spectra in a database to reconstruct the sequence of the peptide, and thus to identify the protein from which the peptides were derived (Chen et al., 2012; Volkel et al., 2010). For Ub-conjugates, including proteins conjugated to Ub or Ub chains, tryptic digestion yields a convenient di-glycine tag on the modified Lys residue (Goldknopf and Busch, 1977). This adds a characteristic monoisotopic mass addition of 114.0429 Da to the peptide, and is used

16 as a search parameter to identify ubiquitylation sites (Marotti et al., 2002; Peng, 2008; Peng and Gygi, 2001). Because a Ub chain linkage is just a specific example of a Ub attachment site in which Ub is attached to a second Ub moiety, the above method can also be used to identify linkage type in a Ub chain (Kaiser and Wohlschlegel, 2005). Nonetheless, MS/MS followed by database searching cannot determine from which ubiquitylated protein the Ub chain was conjugated to. One way to overcome this limitation is to identify Ub chain linkages from highly purified ubiquitylated proteins excised from SDS-PAGE gels (Kaiser and Wohlschlegel, 2005).

In addition to database searching, spectral matching can be a more sensitive and discriminating method in identifying ubiquitylation sites. Unlike database searching, spectral matching compares experimental spectra for a particular peptide against a library of actual spectra (an averaged representation of observed spectra for a particular peptide). Such method increases the sensitivity of detection by allowing for the identification of more “noisy” spectra (Craig et al., 2006; Frewen and MacCoss, 2007; Lam et al., 2007; Lam et al., 2008). To identify Ub chain linkages, a spectral library has been created by Srikumar et al. (2009). Purified human di-Ub proteins, linked through specific Lys residues, were digested with trypsin. Consensus spectra were generated for each linkage type, and used to identify Ub chain linkages in a number of different types of biological samples, using a spectral matching tool (SpectraST) (Srikumar et al., 2010). This method was shown to increase sequence coverage of Ub and Ub-like proteins. However, to successfully identify these types of peptides, each modified variant (the unmodified peptide, both types of singly modified peptide, and the doubly modified peptide) must be present in the spectral library.

1.3 Rad6 and its E3 ligases

The Rad6 gene encodes a ~17kDa Ub E2 protein that plays a number of critical roles in eukaryotes, including post-replication DNA damage repair (PRR), the N-end rule degradation pathway, and histone H2B ubiquitylation (transcriptional regulation and chromatin remodeling). In Saccharomyces cerevisiae, a Rad6 mutant shows defects in growth and mutagenesis, extreme sensitivity to UV, X-ray and chemical mutagens, and hypersensitivity to antifolate drug metabolites. All of these functions are a result of ubiquitylation, because an inactive rad6 produces a null phenotype (reviewed in Shekhar et al., 2002).

In humans, there are two homologues of Rad6 (RAD6A and RAD6B). Due to structural similarities with the yeast Rad6, the human homologues can complement the DNA repair and UV mutagenesis defects of yeast Rad6 mutant (Koken et al., 1991). However, the human homologues lack an acidic tail at its C-terminus, and cannot therefore complement functions in sporulation in yeast mutant Rad6 cells (Reynolds et al., 1985). RAD6A and RAD6B are localized on chromosomes Xq24-q25 and 5q23-q31 respectively, sharing 95% identity at the amino acid level(Koken et al., 1992). Their expression is cell cycle regulated, with maximal expression levels in late S/G2 phases of the cell cycle, consistent with their roles in PRR (Lyakhovich and Shekhar, 2004). RAD6A and RAD6B are also constitutively expressed in all tissues (although the ratio between these proteins varies in different cells and tissues), and have overlapping functions (Koken et al., 1996). In whole-animal studies, inactivation of the mouse RAD6B gene leads to male sterility(Roest et al., 1996), while male mice lacking the RAD6A gene are fertile (Roest et al., 2004). This could be a result of the high expression level of RAD6B in spermatids (Koken et al., 1996). When mice lack both Rad6 homologues, they are nonviable (Roest et al., 2004).

The roles that Rad6 play in PRR, transcriptional control and N-end rule protein degradation pathway have been well studied. Recently, a reconstruction of the ubiquitin network in yeast has actually identified Rad6 as a major hub, with many Golgi-, vesicle-, or endosome-associated roles for Rad6 that are not explained by currently known targets of Rad6 (Venancio et al., 2009). It will be therefore be important to use interactome studies to identify novel interactors of Rad6.

1.3.1 Functional relationships between Rad6 and its E3 ligases

1.3.1.1 Rad18 and PRR

The multitude of Rad6 functions are carried out via interactions with a number of different E3 ligases (Figure 1.4). In partnership with the RING E3 Rad18, Rad6 functions in the PRR pathway (reviewed in Shaheen et al., 2010). DNA damage, as a result of endogenous metabolism and exogenous insults, can lead to blockade at the replication fork. Cells use a process called translesion synthesis (TLS) or PRR to overcome this hurdle. The main regulator of PRR is the replication sliding clamp PCNA (Proliferative Cell Nuclear Antigen), which recruits DNA replication and repair factors to the stalled fork. The role of PCNA in PRR is governed by ubiquitylation. The Rad6-Rad18 complex mediates PCNA monoubiquitylation at a conserved

Lys164 residue, increasing its affinity for the PRR DNA polymerases ι and κ . PCNA then undergoes polyubiquitylation (K63 linked-Ub chains) on the same Lys residue, mediated by the Ubc13-Mms2/Rad5 E2-E3 complex in yeast to initiate error-free repair (template switching). The formation of the poly Ub chain on PCNA is highly dependent on the initial monoubiquitylation by Rad6-Rad18.

Recently, the Rad6-Rad18 complex has also been implicated in monoubiquitylation of the 9-1-1 checkpoint clamp in yeast (Huang et al., 2009b). The 9-1-1 complex has structural similarities to PCNA, and it recruits checkpoint proteins to exposed ssDNA at stalled forks to activate the replication checkpoint (Branzei and Foiani, 2010; Jackson and Bartek, 2009). Ubiquitylation of 9-1-1 by Rad6-Rad18 is involved in the control of global gene regulation (Huang et al., 2009b). Rad18 is also recruited to sites of DNA double stranded breaks (DSBs) through interaction of its Ub binding domain with Ub chains deposited at the DSB site (Huang et al., 2009b). It suppresses the NHEJ (Non Homologues End Joining) pathway, while promoting repair by HR (Homologous Recombination) (Saberi et al., 2007). Through the ubiquitylation of the DNA clamps PCNA and 9-1-1, the Rad6-Rad18 complex thus coordinates different DNA damage response pathways (Shaheen et al., 2010).

Figure 1.4 Biological functions of Rad6. Via interactions with a number of different E3 ligases, Rad6 carries out a multitude of functions in the cell. The different E2-E3 pairs also produce different types of Ub products. Rad6-Rad18 is responsible for the monoubiquitylation of PCNA, and plays a role in post-replication repair (PRR). Rad6-Bre1 monoubiquitylates H2B, and is involved in checkpoint activation and DNA damage repair. Rad6-Ubr1/2 polyubiquitylates N-degrons and targets them to the proteasome for degradation. Figure adapted from (Game and Chernikova, 2009).

1.3.1.2 Ubr1/2 and the N-end Rule Pathway

Another function of Rad6 is to mediate protein degradation via the N-end rule pathway, through an interaction with Ubr1 and Ubr2. The amino-terminal residue of some proteins can serve as a degradation signal, determining the half-lives of these proteins (Bachmair et al., 1986). The N- end rule is organized hierarchically, into three levels. Amino-terminal Asp and Glu are secondary destabilizing residues through their ability to be converted into Arg, the primary destabilizing residue. Asn and Gln are tertiary destabilizing residues because they can be converted into the secondary destabilizing residues (reviewed in (Dohmen et al., 1991). In Saccharomyces cerevisiae, Ubr1 is an important Ub ligase that recognizes N-degrons, and facilitates proteasome-mediated degradation by assembling K48-linked Ub chains (reviewed in Hwang et al., 2010). The initial affinity between Ubr1 and its substrates is moderate, but it becomes stronger when in complex with an E2. Substrates of Ubr1 include Cup9, a transcriptional repressor for amino acid import, Mgt1, a DNA repair protein, and misfolded proteins (Hwang et al., 2010; Varshavsky, 1996). Recently, it has been shown that Rad6-Ubr1 cooperates with the Ubc4-Ufd4 E2-E3 complex to increase the processivity of polyubiquitylation of their respective substrates (Hwang et al., 2010). Ufd4 is a HECT E3 involved in the ubiquitin fusion degradation system (UFD), which targets proteins with N-terminal Ub for degradation. In this case, Ufd4 acts as an E4, an enhancer of ubiquitylation efficacy, to Ubr1.

Ubr2 is a paralog of Ubr1,displaying 22% sequence identity and 45% sequence similarity at the amino acid level. Ubr2 physically interacts with Rad6 to mediate Ub-dependent degradation of proteins such as Rpn4, a transcription factor involved in regulating genes encoding proteasome subunits (Wang et al., 2004b). However, Ubr2 lacks the residues essential for degradation of N- end rule model substrates, suggesting its role outside of the N-end rule pathway (Wang et al., 2004). In fact, Ubr1 and Ubr2 have recently been implicated in the degradation of misfolded proteins under conditions where the N-end rule pathway is blocked, and serve as chaperones to suppress the aggregation of nonnative proteins (Nillegoda et al., 2010).

1.3.1.3 Bre1 (RNF20-RNF40) and histone modification

Histone modifications, including phosphorylation, acetylation, ubiquitylation and methylation are implicated in transcriptional regulation. In S. cerevisiae, H2B is frequently monoubiquitylated at K123 by the Rad6-Bre1 complex (reviewed in Game and Chernikova,

2009). H2B ubiquitylation in turn facilitates the di- and tri-methylation of H3 and H4 by COMPASS proteins, affecting downstream cellular processes including transcriptional regulation, silencing, DNA damage checkpoint activation and repair (reviewed in Game and Chernikova, 2009). Aside from histone methylation functions, H2B ubiquitylation has also been suggested to support transcriptional elongation (Xiao et al., 2005).

In human cells, RAD6 also serves as the cognate E2 for hBRE1 (the RNF20-RNF40 heterodimer), and is recruited to transcribed genes through the interaction with the hPAF1 complex, resulting in monoubiquitylation of H2B at K120, leading to subsequent methylation of H3-K4 and H3-K79 (Kim et al., 2009). Using affinity purification, the WW domain-containing adaptor with coiled-coil (WAC) has also been implicated as a functional partner of RNF20- RNF40 (Choi et al., 2011). Its N-terminal WW domain recognizes the Pol II CTD (C-terminal domain) while its C-terminal coiled-coil motif interacts with RNF20-RNF40 and RAD6. Thus, WAC helps to target RAD6-hBre1 to active transcription sites to modulate H2B ubiquitylation. (Choi et al., 2011).

1.3.2 Structural relationships between Rad6 and its E3 ligases

All of the known E3 ligase interactors of Rad6 contain a RING domain. In addition to the interaction between the RING domain and Rad6, Rad6 also makes contacts with other parts of the E3s, enhancing their affinity.

1.3.2.1 Rad6-Rad18

Rad18 recognizes Rad6 via its RING domain and a helix-loop-helix motif at its C-terminus (Bailly et al., 1997; Hibbert et al., 2011). The Rad18 protein has a C3HC4 zinc finger RING domain at its N-terminus, followed by a C2HC domain (ubiquitin binding zinc finger, UBZ (Masuda et al., 2012; Notenboom et al., 2007). It also has a SAP (SAF-A/B, Acinus and PIAS) domain, required for DNA binding (Notenboom et al., 2007). The RING-E2 interaction (mentioned earlier) is highly conserved among RING E3s and is required for ligase activity (Deshaies and Joazeiro, 2009). The RING domain of Rad18 binds to Rad6 through a canonical E2-E3 interface at the N-terminus of the E2 (Huang et al., 2011). It was shown that mutations in the RING domain of mouse Rad18 resulted in increased sensitivity to DNA-damaging agents (Tateishi et al., 2003). Recently, Matsuda et. al (2011) and Huang et. al (2011) reported that

Rad6 and Rad18 form a ternary complex, Rad6-(Rad18)2 ,where Rad18 forms a dimer through its RING domain. Interestingly, only one of the RING domains is necessary for ligase function, and binds to Rad6 (Matsuda et al., 2011). The function of this dimer is unclear, but could provide more space for substrate or have a regulatory role. The second RING domain could also function in other pathways (Huang et al., 2011).

The C-terminus of Rad18 contains an additional Rad6 binding domain (R6BD). Using a series of GST-deletion mutants of Rad18 and far-Western analysis, the region of Rad18 required for stable association with Rad6 was delimited to residues 371 to 410, and predicted to have a helix- loop-helix structure (Bailly et al., 1997). The loss of this site impairs Rad6 interaction in vitro but localization to sites of DNA damage is not affected (Watanabe et al., 2004). The R6BD binds to the backside of Rad6, involving residues of β1, β2, β3 and C-terminal residues on Rad6. Notably, this overlaps with the noncovalent Ub binding site required for chain formation (Hibbert et al., 2011). By competing with the interaction between Rad6 and the acceptor Ub, Rad18 thus effectively causes Rad6 to monoubiquitylate its substrate PCNA (Hibbert et al., 2011).

1.3.2.2 Rad6-Ubr1/2

In comparison to Rad18, Ubr1 makes similar contacts on Rad6. Ubr1 is a RING E3 that, in addition to the UBR box at its N-terminus (which confers substrate recognition), contains an auto-inhibitory domain at its C-terminus, and a Basic Rich Region (BRR) that serves as a secondary Rad6 binding site (reviewed in Choi et al., 2010). Sequence comparisons between Ubr1 and Ubr2 revealed that they have a conserved RING and an adjacent BRR domain (Kwon et al., 1998; Saurin et al., 1996). Mutagenesis studies have shown that mutations in the BRR domain can negatively affect the interaction between Rad6 and Ubr1. The same cannot be said for the RING domain (Xie and Varshavsky, 1999). However this high affinity interaction is not essential for the degradation of substrates, as only the RING-H2 finger of Ubr1 is required for ubiquitylation (Xie and Varshavsky, 1999). Like Rad18, the BRR binding site on Rad6 overlaps with the noncovalent Ub binding site (Hibbert et al., 2001). However, the interaction is of a lower affinity, which could explain why Rad6-Ubr1/2 complexes polyubiquitylate rather than monoubiquitylate substrates (Hibbert et al., 2011).

1.3.2.3 Rad6-Bre1 (RNF20-RNF40)

Like its human homolog (RNF20-RNF40), Bre1 has a RING-finger domain at its C-terminus, with many predicted coiled-coil regions throughout the rest of the protein (Kim et al., 2009). In yeast, the poly-acidic carboxy-terminal tail of Rad6 is essential for binding to Bre1. Mutant cells lacking this tail are defective in H2B ubiquitylation (Morrison et al., 1988). In humans, the RNF20-RNF40 heterodimer forms a stable complex through their N-terminal regions. The N- terminal coiled-coil regions of the complex are also important for Rad6 binding (Kim et al., 2009). Using an in vitro binding assay, it was shown that RAD6 interacts with RNF20 and RNF40 only when they are in a complex with each other, and not with the individual polypeptides (Kim et al., 2009). Interestingly, only the RING finger of RNF20 is critical for ubiquitylation (Kim et al., 2009). The Bre1 (or RNF20-RNF40) binding residues on Rad6 have yet to be characterized.

1.3.3 Rad6 and X-linked Mental Retardation (XLMR)

Impairment of the ubiquitin system has long been associated with neurodegenerative diseases such as Parkinson’s, Alzheimer’s, Huntington’s and others. Mutations in the RAD6A gene have been shown to cause a novel XLMR syndrome in several families (Nascimento et al., 2006; Budny et al., 2010). Monogenic XLMR has been estimated to affect ~10% of mentally retarded males (Mandel and Chelly, 2004; Ropers, 2006), and is a major socioeconomic burden. Severe forms stem from a number of different genetic causes, including chromosomal abnormalities, copy number variants and defects of single genes. So far, 82 X-linked genes have been implicated in XLMR, accounting for half of the cases (reviewed in Budny et al., 2010).

In 2006, it was reported that affected males from two generations of the same family developed a neurodevelopmental disorder due to a nucleotide substitution in the RAD6 coding sequence, corresponding to Q128X, resulting in a premature stop codon (Nascimento et al., 2006). The patients all had similar phenotypes, including seizures, obesity, hirsutism and a characteristic facial appearance. In 2010, another group reported two missense mutations in the highly conserved N-terminus of RAD6 (G23R and R11Q), where seven patients from two families developed a similar syndromic form of XLMR as the patients with the Q128X mutation (Budny et al., 2010). In the same year, a novel 0.4 Mb deletion at Xq24, that included the RAD6A gene, was detected in a 4-year-old and 10-month old boy of the same family (Honda et al., 2010).

These boys displayed phenotypic similarities with the other XLMR patients mentioned above. Mutations in RAD6A therefore mark the first description of an association between an E2 protein and neurological disease.

1.4 Affinity purification coupled to mass spectrometry and protein-protein interactions

Mass spectrometry coupled to affinity purification (AP-MS) represents a powerful tool for characterizing protein-protein interactions (e.g identification of the interactome of core components of the Ub system (Chen et al., 2012). A growing number of medium-scale AP-MS studies have been done to characterize human protein interaction networks, including the human deubiquitylating enzymes (Behrends et al., 2010; Ewing et al., 2007; Goudreault et al., 2008; Hutchins et al., 2010; Jeronimo et al., 2007; Malovannaya et al., 2011; Sowa et al., 2009).

Affinity purification can be done through two major approaches. The first approach is the use of antibodies directed against endogenous proteins. The main advantage to this approach is that proteins are purified in their native condition and that specific isoforms may be interrogated individually (Dunham et al., 2012). However, antibodies are costly, and not all proteins have a corresponding antibody. Moreover, antibodies can disrupt protein-protein interactions, or fail to recognize post-translationally modified forms of the protein of interest. Most importantly, the specificity and affinity of all antibodies are not equivalent, therefore the conditions of purification must be optimized individually (Dunham et al., 2012). Epitope-tagging, on the other hand, does not require the optimization of the purification protocol for each protein complex (Gingras et al., 2007).This approach involves fusing DNA sequences to the coding region of a protein of interest, where the tagged protein can be purified using a solid matrix that recognizes the epitope (Brizzard, 2008; Jarvik and Telmer, 1998). By using the same tag for multiple proteins, background contaminants are also consistent across all purifications (Dunham et al., 2012). However, the location and the nature of the tag could lead to mislocalization of the protein or disruption of protein-protein interactions, resulting in false-positives and/or false- negative interactors (Gingras et al., 2007). High throughput interactome studies have primarily used a single FLAG or tandem affinity purification (TAP) epitope tagging systems. The FLAG tag system consists of a single-step purification, which can be used to preserve weaker or more transient protein–protein interactions. By contrast, the TAP tag system, consisting of two

24 purification steps, can decrease the level of proteins binding non-specifically to solid phase support or epitope tag, but may result in the loss of weaker and transient protein interactors (reviewed in Gingras et al., 2007, Chen et al., 2012).

Protein tags can be inserted endogenously into the genome of cells using homologous recombination(Howson et al., 2005). Inducible expression systems, such as the Flp-In T-REx system (Invitrogen), is one such example, where isogenic lines are established and the expression of the protein of interest can be controlled by the addition of tetracycline. The Flp-In™ System takes advantage of a Saccharomyces cerevisiae-derived DNA recombination system, which uses a recombinase (Flp) and a Flp Recombination Target (FRT) site (Craig, 1988; Sauer, 1994)to mediate site-specific integration of the gene(s) of interest into the genome of mammalian cells. The T-REx™ System regulates the expression of gene(s) through the use of the E.coli Tn10- encoded tetracycline (Tet) resistance operon (Hillen and Berens, 1994; Hillen et al., 1983). The expression of the gene(s) of interest is thus repressed in the absence of tetracycline, and induced in its presence. Components of the Flp-In and T-Rex systems can be introduced into various mammalian host cells by standard transfection methods (Yao et al., 1998).

1.4.1 Analysis of protein interaction data

With any affinity purification approach, background proteins will be co-purified. Even with very stringent methods, such as the TAP system, non-specific binding proteins interacting with the epitope tag or resin can be identified. Development of software tools for the analysis of protein interaction data is thus crucial to visualize and determine non-specific vs. bona fide interactors. Recently, Breitkreutz and collegues (Breitkreutz et al., 2010)developed a novel software suite, consisting of ProHits (Liu et al., 2010) and SAINT (Choi et al., 2011), to visualize and identify non-specific interactors of the kinase and phosphatase protein network in yeast.

ProHits is a software tool that allows for easy comparison between different samples (Liu et al., 2010), and therefore allows the visualization between background interactors (those that are identified in control groups) and more specific interactors found only with the “bait” protein (i.e. protein of interest). ProHits has a data management module to store all MS data in an instrument-specific manner and searches data using Mascot or X!Tandem. It also has an Analyst module which allows the user to visually compare the identified proteins between different groups (displayed in separate columns; Figure 1.5). In the Analyst module, proteins that are

25 identified are displayed by row. Spectral count, unique peptides, peptide scores from search engines and protein coverage information can be displayed by clicking on the “Peptide” link. All search results can be filtered, allowing for removal of nonspecific background proteins defined by control groups, search engine score threshold or available contaminant lists. Here, the data are filtered based on the confidence score associated with each identified peptide and protein.

Figure 1.5 ProHits Analyst module-Comparison page. On the left are shown comparison results for three human baits, RAD6, KCMF1, and UBR2 (b) and merged negative controls (a). Negative controls include the FLAG tag protein alone, and other proteins that are functionally unrelated to the baits. Display, sort, filter and literature overlap options are listed on the top (c). Columns represent individual baits. Rows list the proteins identified with each protein sample (hits). The number in each cell represents the total peptide number for each protein identified. External links, including the NCBI Gene and Protein pages are available for a given hit (d-e). Peptide Comparison for each hit is also available (f), which allows the user to view all peptides identified for a given hit across all protein samples/experiments. Figure adapted from Liu et al., 2010.

The Significance Analysis of Interactome (SAINT) is a statistical algorithm used to identify high confidence protein-protein interactions, thereby also identifying background contaminants. SAINT uses data generated from AP-MS and constructs separate distributions for true and false interactions to derive the probability of bona fide interactors (Choi et al., 2011). The algorithm models the spectral counts for each prey-bait pair with mixture distributions of both true and false interactions. The parameters for these two distributions are based on the spectral counts for all interactions across the purifications/groups that involve the same prey-bait pair. From this, a probability of a true interaction between two proteins is calculated. Negative controls can be included in the modeling. In this case, spectral count distribution for false interactions is directly estimated from the negative controls (Choi et al., 2011).

With novel software such as ProHits and SAINT, AP-MS can generate a list of high confidence protein interactors detected in a given sample. However, a single AP-MS analysis does not reveal the composition of individual protein complexes. To gather more information, iterative “high density” AP-MS (Gingras et al., 2007) can be employed, where the preys from one round of analysis become the baits in the next round (Gingras and Raught, 2012).

1.5 Thesis rationale and outline

Ubiquitylation plays a role in a wide variety of processes. This conjugation process requires the sequential actions of three key proteins, the E1, E2, and E3. The E2-E3 interaction is especially important in defining the type and topology of Ub products. The selectivity and specificity between E2-E3 interactions are not well understood, however, as the physiological E2 partners for many E3s are not defined. The mechanism for specifying different Ub products is also not well characterized. Here, in Chapter 2, I present the first comprehensive E2-HECT E3 structure- function screen, to begin to address these issues.

XLMR mutations in the RAD6A gene map to regions important for E2-E3 interactions. In Chapter 3, using AP-MS, I present the first Rad6 interactome in mammalian cells, and identify disruptions in protein-protein interactions for one of the XLMR RAD6A mutant proteins. In addition to all of the known E3 ligases, I also identified several novel Rad6 interactors, including the E3 ligase KCMF1 (potassium channel modulatory factor 1), which is specifically lost for one of the RAD6A mutant proteins. Consequently, I began to characterize the structural and

27 functional relationships between KCMF1 and RAD6A. My data implicates potential novel functions for RAD6.

A human ubiquitin conjugating enzyme (E2) - HECT E3 ligase function screen

Work presented in this chapter is from Sheng, Y., Hong, JH., Shloush, J., Doherty, R., Srikumar, T., Avvakumov, GV., Walker, JR., Sheng, X., Neculai, D., Wan, JW., Kim, SK., Arrowsmith, CH., Raught, B., and Dhe-Paganon, S. A human ubiquitin conjugating enzyme (E2) - HECT E3 ligase structure-function screen. Molecular & Cellular Proteomics.2012. Aug 11: 329-341

In this chapter, I will only be presenting the functional aspect of the screen. For structural analysis, please refer to the manuscript.

2 A human ubiquitin conjugation enzyme (E2)-HECT E3 ligase function screen 2.1 Chapter overview

Ub E3 ligases facilitate the transfer of Ub from an activated E2 to a substrate protein or to another Ub molecule. The human genome encodes hundreds of RING type E3s, and 28 HECT domain-containing E3 ligases (Rotin and Kumar, 2009). E3s demonstrate specificity for subsets of E2 proteins, and different E2-E3 combinations can generate very different types of Ub chains (e.g.Christensen et al., 2007; Jin et al., 2008; Ordureau et al., 2008; Wickliffe et al., 2011). However, the molecular determinants involved in E2-HECT E3 interaction specificity are not well defined, and the molecular mechanisms involved in the specification of different types of Ub linkages remain cryptic.

Due to the modest affinity of most E2-E3 interactions, techniques such as co- immunoprecipitation have been largely unsuccessful in characterizing functional E2-E3 pairs. While yeast two hybrid screening has identified a large number of putative E2-E3 functional interactions (e.g. Markson et al., 2009; van Wijk et al., 2009), this methodology does not provide information concerning processivity or Ub chain linkage types generated by each pair. Therefore, in vitro E2-E3 ubiquitylation reactions with purified proteins are needed to obtain this type of information (as shown in this chapter). The functional interactions between a large number of RING domain E3s and a smaller number of E2 proteins have been investigated previously (e.g.Christensen et al., 2007; Jin et al., 2008; Ordureau et al., 2008; Wickliffe et al., 2011) but no large-scale study has focused on the HECT E3 ligases. Here, I present the first comprehensive human E2-HECT E3 function analysis.

2.2 Contributions

Colleagues from the Structural Genomic Consortium and the Arrowsmith Lab (Toronto, ON) constructed the E2 library. I helped to purify some of the E2 proteins (E2D2, E2E2, E2J2, E2L3, E2K, E2R1, E2S, E2H, and E2T) and the E3 ligases (WWP2, NEDD4L, ITCH, HERC4, UBE3A, UBR5 and HERC2) needed for subsequent experiments. SGC members and Janet Wan (a previous graduate student from the Raught lab) conducted the autoubiquitylation reactions and the Ub loading assay. I helped to repeat some of these autoubiquitylation reactions (mainly

30 against the panel of nine E2s listed above). Tharan Srikumar (a Ph.D candidate in the Raught lab) did most of the mass spectrometric data analysis.

2.3 Materials and Methods

2.3.1 Plasmids

Human E2 open reading frames were amplified by PCR from various templates (see NP numbers in Table 2.1), and inserted using the Infusion system (BD Biosciences, San Jose, CA) into a pET28 vector with 6xHis-tag and a thrombin or TEV-cleavage site located upstream of the cDNA insert. Human HECT domain proteins (Table 2.1) and Ub were similarly cloned. The library of E.coli E2 expression clones will be made available through Addgene and/or other nonprofit sources.

2.3.2 Protein purification

Proteins were expressed in E. coli BL21 (DE3) Gold (Stratagene, LaJolla, CA) grown in TB medium in the presence of 50 µg/ml kanamycin at 37ºC to an OD600 of 4-5, induced with 2 mM isopropyl-1-thio-D-galactopyranoside (IPTG) and further grown for 16-18 h at 15ºC. Recombinant E2 and HECT domain proteins were purified using standard metal-affinity chromatography with TALON resin (Stratagene), according to manufacturer's instructions. All proteins for the Ub assays were dialyzed against 20 mM Tris-HCl pH 8.0, 150 mM NaCl, 10% glycerol, 2 mM dithiothreitol (DTT) and stored at -80°C.

2.3.3 Autoubiquitylation and E2 loading assays

E2 loading assays were carried out in a volume of 10µl containing 1µg E1, 1µg of E2, and 5µg of 6xHis tagged Ub in a buffer consisting of 10 mM HEPES pH7.5, 100 mM NaCl, 40 µM ATP and 2 mM MgCl2. Reaction mixtures were incubated for 10 min at 30ºC, stopped by the addition of non-reducing SDS-PAGE sample buffer, separated by 4-20% gradient SDS-PAGE gels (Invitrogen, Grand Island, NY), and visualized by Western blotting, using a mouse monoclonal antibody directed against the 6xHis epitope tag (Qiagen, Valencia, CA), an HRP-conjugated goat-anti-mouse secondary, and ECL (Bio-Rad, Hercules, CA).

Autoubiquitylation reactions (in the presence of E3s) were performed in a volume of 20 µL in a buffer of 50 mM Tris pH 7.6, 5 mM MgCl2, 2 mM ATP and 2 mM DTT, containing human-

31 recombinant E1 (50ng; Boston Biochem, Cambridge, MA), E2 (100ng), ubiquitin (5µg), and E3 (6xHis tagged HECT domain proteins, 0.5µg). After incubation at 30°C for 90 min, reactions were stopped by the addition of SDS-PAGE sample buffer and resolved on 7% SDS-PAGE gels. In the absence of E3, reaction conditions were 50 mM Tris pH 7.6, 50 mM NaCl, 50 mM KCl,

10 mM MgCl2, 5 mM ATP and 0.1mM DTT, with 100 ng E1, E2 (200ng of UBE2R1 or 1ug of UBE2K) and 5ug Ub. Reactions were incubated at 30°C for 3 hrs. Ubiquitylated proteins were evaluated by Western blotting using monoclonal antibodies directed against 6xHis (Qiagen), as above.

2.3.4 Mass Spectrometry

Autoubiquitylation reactions were scaled-up 3-fold for mass spectrometric analysis, and subjected to 4-12% gradient SDS-PAGE. For these reactions, a non-tagged human recombinant Ub was used in the reaction mixture (Boston Biochem). Gels were stained with Coomassie brilliant blue for visualization, and the region containing proteins migrating at >125kDa was processed as in(Lallemand-Breitenbach et al., 2008) for mass spectrometry. The digested peptide mixture was subjected to nLC-ESI-MS/MS, performed using an Orbitrap Velos instrument (Thermo Fisher Scientific, Waltham, MA) coupled to a Proxeon nanoHPLC system (Odense, Denmark). Reaction products from each E2-E3 pair were analyzed twice. Analytical columns were prepared in-house from 10 cm fused silica capillaries (75um inner diameter; InnovaQuartz,

Phoenix, AZ) and packed with C18 coated silica particles (300 Å pore size, 5um particle size; Michrom Bioresources, Auburn, CA). Peptides were first injected onto a 2cm (100um inner diameter) C18 pre-column, and chromatographic separation was achieved using a 120 min gradient, from 100% buffer A (5% acetonitrile with 0.1% formic acid) to 40% buffer B (95% acetonitrile with 0.1% formic acid) running at a constant flow rate of 250nl/min. The mass spectrometer was operated in data-dependent acquisition mode: one survey (400-1800 m/z) MS scan (at 60,000 resolution) was performed, and the forty most intense ions were chosen for fragmentation using collision-induced dissociation (CID) in the ion trap. Target ions for which two previous CID scans had been collected (within 30s) were dynamically excluded for 60s. Thermo .raw files were converted to the .mzXML format with ReadW software v.3.5.1(Pedrioli et al., 2004), and data were searched using both; (a) automated database search software X!Tandem(Beavis, 2006; Craig and Beavis, 2004) against the Homo sapiens ENSEMBL Genome Reference Consortium assembly GRCh37 database (75,126 entries), and (b) spectral

32 matching against our previously published Ub/Ubl spectral library (Srikumar et al., 2010), supplemented with additional consensus spectra derived from commercial (Boston Biochem) di- ubiquitin K6, K27, K29, and K33 linkages. Search parameters for X!Tandem specified a parent MS tolerance of 10ppm and an MS/MS fragment ion tolerance of 0.4 Da, with up to 2 missed cleavages allowed for trypsin. A +114.0429 Da modification of lysine was specified as a variable search parameter to identify the ubiquitin-derived diglycine motif. Oxidation of methionine (+15.995) and deamidation of Gln (+0.985) were also allowed as variable modifications. A GPM expect score of -2 was used as a cutoff, corresponding to a calculated false discovery rate of 0.80%. SpectraST (Lam et al., 2007; Lam et al., 2008) was used for spectral matching, with a dot product of ≥0.7 used as a cutoff, corresponding to a calculated false discovery rate of 0.66%. All .raw files will be submitted to Tranche.

To correct for inherent differences in Ub linkage signal intensity, Proteome Discoverer (Thermo, Ver. 1.3) was used to search and analyze three replicate MS analyses of trypsin digested equimolar mixes of all seven di-ubiquitin polypeptides (Boston Biochem). Files were searched using the Sequest search algorithm and the ipi.HUMAN.v3.83 database, with a10ppm precursor mass tolerance and 0.5 Da fragment mass tolerance. Oxidation of M, deamidation of N or Q, and ubiquitylation of lysines (+114.043) were also allowed. Sequest results were analyzed using the Percolator algorithm, with a target FDR of 0.01. Precursor ion area detection was enabled, using a precision of 4ppm, and the AUC for each Ub linkage was calculated (for all observed charge states). A “detection bias” ratio was then calculated as the ratio of the AUC for each linkage type with respect to the linkage type with the lowest AUC, K29. An average detection bias ratio was calculated from the 3 replicate MS runs, and this conversion ratio applied to all data.

2.4 Results

2.4.1 Construction of an E2 library

To define the E2 functional partners of HECT E3 ligases, we constructed a recombinant E2 protein library. Full-length and “core” (UBC domain) versions of the 40 H. sapiens E2 proteins were cloned into a 6xHis expression vector (Table 2.1). Using standard methods, both core and full-length polypeptides for 29 different E2s were successfully expressed in Escherichia coli, and purified to homogeneity. The remaining full-length proteins were not expressed or were insoluble, but the core domains of eight of the remaining E2s were successfully expressed

(Table 2.1). These 66 purified E2 proteins (covering 37 of the 40 UBC domains in the human genome) form the basis of our library, representing the most complete collection of recombinant human E2 enzymes available to date. Similar to the E2s, the HECT domain of a number of E3 ligases (representatives from each the HECT E3 families) was purified accordingly.

Table 2.1 Construction of a human E2 library. Protein information for the 36 E2s (including both active and inactive E2 variants) used in this chapter are listed in the above table. Information for HECT domain proteins is also included. Indicated are gene name, aliases, protein length, PDBID code (if structure has been solved) and NCBI protein and gene accession numbers. “Protein Note” indicates whether we were able to produce soluble full length or core recombinant protein. “Ub loading” indicates the ability of each recombinant E2 protein to form a thioester bond with Ub, in our assay. Bold PDB number represent new E2 structures reported in this study. ND: No data.

2.4.2 An E2-HECT E3 functional screen

To determine if the purified E2 proteins can load Ub, the entire E2 panel was subjected to an in vitro “Ub-loading” assay, assessing the ability of each conjugating enzyme to form a thio-ester- Ub intermediate. In the presence of Ub, ATP and the E1 enzyme Uba1, we found that 26 out of the recombinant human E2s were capable of loading Ub (Figure 2.1). This result is in accordance with previous studies (Jin et al., 2007), except for UBE2U. It was able to load Ub

(although weakly) in our assay. Our Ub loading screen also revealed that three E2s, UBE2K, UBE2R1 (Cdc34) and UBE2R2, do not require the presence of an E3 molecule to carry out Ub conjugation (i.e. the formation of an isopeptide bond with the epsilon amine group of a lysine residue in one of the proteins in the reaction) under our in vitro reaction conditions. As expected, we did not observe Ub loading for the known ubiquitin-like protein E2s UBE2I, UBE2F, UBE2M and UBE2L6, nor the inactive Ub E2 variants (UEVs, which lack a catalytic Cys) TSG101, TMEM189, UBE2V1, UBE2V2 or AKTIP. UBE2Z also had no activity in our assay, consistent with a previous report demonstrating that this E2 is loaded specifically by UBA6 (Jin et al., 2007).

Figure 2.1 Ub loading assays. Reactions consisting of His-tagged full-length or core E2 proteins incubated with E1 (Ube1) and ATP were incubated with and without His-tagged Ub, then subjected to non-reducing SDS- PAGE and anti-His Western analysis. Arrowhead denotes (thiol-ester) loaded E2.

To determine the functional interactions between the entire set of Ub loading E2s and a number of HECT E3 ligases, we used an in vitro autoubiquitylation assay. Ten HECT domains (Table 2.1), including representatives of the NEDD4 subfamily (NEDD4L, ITCH, SMURF1 and WWP2), the HERC subfamily (HERC2 and HERC4), and other more distantly related HECT domains (UBR5, UBE3A and UBE3C) were tested in an autoubiquitylation assay with each of the 26 Ub-loading E2 proteins. After 90 min at 30°C, reaction mixtures were resolved via SDS- PAGE, and ubiquitylated products were detected by Western blotting (Figure 2.2 A).

Based on Ub adduct migration in SDS-PAGE, we classified each of these 234 E2-E3 combinations into one of three groups: (i) those catalyzing long Ub chains (MW >125 kDa), (ii) those catalyzing shorter Ub oligomers or (multi-) monoubiquitylation (MW <125 kDa), and (iii) those with no apparent functional interaction (Figure 2.2 B). UBE2L3, UBE2D1-4, UBE2E1-3 and UBE2J1-2 displayed functional interactions with all of the NEDD4 subfamily HECT domains, as well as HERC4 and UBE3A (E6-AP).A majority of these interactions resulted in the synthesis of long Ub chains. UBE2S was able to catalyze the synthesis of long chains with WWP2 and ITCH, and short chains or multi-monoubiquitylation with several other E2s. UBE2C, UBE2G1, UBE2T and UBE2W also functionally interacted with several of the HECT domains to produce short chains or multi-monoubiquitylation. UBE2A, UBE2B, UBE2H, UBE2Q1 and UBE2U displayed low levels of activity in our screen, and no interactions were observed with UBE2G2. These data define the first human E2-HECT functional interaction landscape.

In a few cases, we observed significant differences in activity between the core and full-length versions of an E2. For example, the core domains of UBE2J1 and UBE2W displayed increased activity compared to their full-length counterparts, in some autoubiquitylation reactions. Conversely, the full-length UBE2S protein was more active in some reactions than the core domain. These data suggest that additional sequences (the N-terminal or C-terminal extensions) outside the core can act as positive or negative regulatory domains and/or provide additional E3 specificity.

Figure 2.2 Autoubiquitylation reactions. (A) Autoubiquitylation reaction was performed in the presence of recombinant E1, ATP, ubiquitin and E2. Reactions were subjected to SDS-PAGE and anti-Ub Western analysis. Shown is a representative screen, with the HECT E3 domains of WWP2 and ITCH. This is typical of results obtained in 3 independent experiments. Each lane represents an individual autoubiquitylation reaction with E2 indicated at top. (-) indicates negative control, lacking an E2 protein. The locations of unconjugated and oligomeric His-Ub are indicated to the right of each Western blot. (B) A heat map depicting Ub chain-building activity of each of the 234 E2-E3 pairs in in vitro autoubiquitylation reactions. Dark blue indicates long Ub chains (>125 kDa), light blue indicates short chains or (multi-) monoubiquitylation, and white indicates no functional interaction. E2s and E3s are hierarchically clustered according to activity.

2.4.3 Characterization of Ub chain linkages in E2 – HECT reactions

As shown through the E2-HECT functional screen, different E2-E3 pairs generated different Ub products. Using mass spectrometry, I characterized the Ub chain linkages generated by a number of E2-E3 pairs capable of generating long Ub oligomers. This included UBE2D2, UBE2E2, UBE2J2 and UBE2L3, with WWP2, ITCH, NEDD4L and HERC4. Autoubiquitylation reaction products were separated via SDS-PAGE, and proteins migrating at >125 kDa were subjected to trypsin proteolysis. The resulting peptides were analyzed using nanoflow liquid chromatography- electrospray ionization-tandem mass spectrometry (nLC-ESI-MS/MS). Ub-Ub linkages were identified using two methods: (i) standard database searching, with the inclusion of a +114.0429 Da mass shift (corresponding to the Ub tryptic GG remnant on lysine) as a variable modification (Kirkpatrick et al., 2005), and (ii) spectral matching, using a Ub/Ubl spectral library that we recently developed, and which now contains consensus spectra derived from all seven Ub chain linkage types (Jeram et al., 2009; Srikumar et al., 2010). To determine whether the HECT domain can dictate linkage specificity, I also characterized the Ub chain linkages generated by the same E2s with RING E3 proteins (MDM2 and RO52).

When UBE2D2, UBE2E3 or UBE2J2 were combined with the RING E3s, a mix of Ub linkages was generated, in which K63 ≈ K48 ≈ K11 >> all other linkage types (Figure 2.3). This is surprising, because so far, only UBE2N-UBE2V1/2 has been shown to generate K63-linked Ub chains. UBE2L3 was not functional with the RING proteins, because UBE2L3 is preferentially utilized by HECT E3s (Eletr and Kuhlman, 2007; Wenzel et al., 2011). In the presence of the same E2s (and UBE2L3), the HECT domains derived from ITCH, WWP2 and NEDD4L (NEDD4 subfamily) also catalyzed the formation of a mix of K63-, K48- and K11-linked Ub products, where rank order of abundance was K63 >K48 > K11 >> all other linkage types (Figure 2.2). In contrast, the HERC4 HECT domain (HERC subfamily) gave rise to a very different spectrum of Ub chains with the same set of E2s, synthesizing Ub oligomers that were markedly enriched in K48 linkages (K48 >> K63 > all other linkages). Thus, with the same E2s, the HECT domains from different HECT E3 subfamilies different HECT domains display very different linkage preferences.

To further delineate the role of HECT domains in Ub linkage specification, I examined the relationship between HECT domains and linkage-specific E2s. UBE2R1 (Cdc34) and UBE2K

40 have both been shown to generate Ub chains enriched in K48 linkages (Haldeman et al., 1997; Petroski and Deshaies, 2005). These E2s were analyzed in reactions containing; (i) no E3, (ii) the RING E3s MDM2 or RO52, and (iii) HECT domains derived from WWP2, ITCH, NEDD4L and HERC4. Both of these E2s catalyzed the synthesis of Ub oligomers highly enriched in K48 linkages in the absence of E3, or when combined with RING E3s (Figure 2.3). A similar linkage distribution was observed in reactions containing HERC4. However, when UBE2R1 or UBE2K were combined with NEDD4 subfamily HECT E3s (n.b. NEDD4L was inactive with UBE2K), the composition of Ub linkages in the reactions were similar to those observed with UBE2D2, UBE2E3, UBE2J2 and UBE2L3; i.e. mixed chain products, with significant levels of K63 and K48 linkages, along with a low level of K11 linked Ub oligomers (Figure 2.3). Consistent with earlier studies (Kim and Huibregtse, 2009; Wang and Pickart, 2005), this data confirms that HECT domains can govern linkage composition in autoubiquitylation reactions, largely independent of E2 linkage specificity.

Figure 2.3 Ub linkage analysis. Autoubiquitylation reactions were subjected to SDS-PAGE, and reaction products analyzed by mass spectrometry (see Methods for details). Ub linkage composition for each E2-E3 reaction are presented as a fraction of the total number of linkages detected. See Legend inset for color codes. 2.5 Discussion

Here, I present a comprehensive functional screen between the entire set of ubiquitin-loading E2s found in the human genome and members of different HECT subfamilies. As predicted, E3s

41 show selectivity toward specific E2s, and the resulting Ub products from different E2-E3 pairs vary. Members of the NEDD4 family interact with many of the E2s, and the E2-E3 interactions result in mostly long Ub chain products. It is important to note that functional E2-HECT pairs that can only extend Ub chains on monoubiquitylated targets, or those that require one or more cellular co-factors, would not be detected in our assay.

Previous publications have suggested that HECT E3s determine linkage specificity in autoubiquitylation reactions (Kim and Huibregtse, 2009; Wang and Pickart, 2005), and that different E2s can synthesize specific Ub chain types, alone or in combination with RING E3s (e.g. Christensen et al., 2007; Jin et al., 2008; Ordureau et al., 2008; Wickliffe et al., 2011). Here, I show that in the presence of linkage-specific E2s, HECT domains can actually alter the linkage specificity, further indicating that the HECT domain is the primary determinant of Ub-Ub linkages in this context.

I also show that HECT domains from different HECT E3 subfamilies display different linkage specificity. Sequence and structural information highlights differences between the E2 binding surfaces on HERC4 versus the NEDD4 subfamily E3s (NEDD4L, WWP2, ITCH, SMURF1). This could explain the variation in chain linkage types synthesized by the NEDD4 subfamily vs. HERC4 with the same E2 proteins.

Identification of a novel RAD6 associated E3 ligase, KCMF1, and specific loss of KCMF1 binding by an X-linked mental retardation-associated mutant R11Q RAD6A protein

3 Identification of a novel RAD6-associated E3 ligase, KCMF1, and specific loss of KCMF1 binding by an X- linked mental retardation-associated mutant R11Q RAD6A protein 3.1 Chapter overview

Rad6 is an evolutionarily conserved Ub E2 protein (coded for by two genes in humans, RAD6A and RAD6B) that plays a number of critical roles in eukaryotes, including the degradation of proteins via the N-end rule pathway, histone H2B ubiquitylation, and post-replication DNA damage repair (Game and Chernikova, 2009; Lee and Myung, 2008). Via its interaction with a number of different E3 ligases, Rad6 is targeted to a variety of substrates, modulating their activities and mediating downstream functions. To date, Rad6 has been reported to interact with a number of RING E3 ligases, including the ubiquitin recognin E3 ligases Ubr1, Ubr2 and Ubr3

(N-end rule pathway), Rad18 (PRR) and Bre1 (RNF20-RNF40 in mammals, implicated in histone monoubiquitylation) (Bailly et al., 1997; Kim et al., 2009; Wang et al., 2004; Watkins et al., 1993). Interestingly, several different RAD6A coding sequence mutations have been detected in X-linked mental retardation (XLMR) patients (Budny et al., 2010; Nascimento et al., 2006).

The effects of these mutations on RAD6 protein-protein interactions have yet to be assessed.

MS has emerged as a powerful tool to identify proteins in complex biological samples. In this chapter, I present data from an affinity-purification coupled with mass spectrometry (AP-MS) study to characterize the RAD6 interactome in human cells. This study identified a number of novel Rad6 interactors, including the ubiquitin E3 ligase KCMF1. I used the same technique to analyze the effect of XLMR-associated RAD6A mutations on protein-protein interactions, and found that the Kcmf1 interaction is specifically lost. Using an in vitro binding assay and AP-MS,

I mapped the region of KCMF1 that is critical for RAD6A binding, and using NMR

44 spectroscopy, I mapped the regions on RAD6A that are important for KCMF1 binding. Like many of the E3 ligases associated with RAD6, KCMF1 utilizes a C-terminal region (CT) outside of its RING domain to interact with the E2. In contrast to the other E3s, KCMF1 also binds to the N-terminal α1 helix (including the R11 residue) on RAD6A. These data can explain why the

RAD6A R11Q protein specifically loses the interaction with KCMF1, but not with any of the other E3 ligases (UBR1/2, RAD18 and RNF20-40). Interestingly, KCMF1 also interacts with the region around the catalytic Cys88 of RAD6A,and very effectively inhibits RAD6A ubiquitylation activity in vitro. Finally, to gain insight into the biological role of KCMF1, I have begun to characterize its intracellular localization, along with some of its interactors.

3.1.1 Contributions

Janet Wan (a previous graduate student) generated some of the AP-MS data for mutant RAD6A proteins. I also repeated these experiments myself. Dr. Etienne Coyaud (post-doctoral fellow) did the cellular fractionation experiment to examine the localization of KCMF1 proteins. Dr. Lilia Kaustov (Arrowsmith lab) helped me to perform Nuclear Magnetic Resonance (NMR) experiments. She also processed and analyzed all of the NMR data.

3.2 Materials and Methods

3.2.1 Expression Constructs

RAD6A/B plasmids (in pET15b vector) were a gift from the Arrowsmith lab (Structural and Genomic Consortium, Toronto, ON). Point mutants of RAD6A were introduced by QuikChange Site-Directed Mutagenesis kit (Stratagene, LaJolla, CA). Using standing cloning procedures, RAD6, and its mutants were also cloned into the FLAG-, GFP-, Cherry-pcDNA5/FRT/TO vectors (gifts from Dr. AC Gingras; Samuel Lunenfeld Research Institute, Toronto, ON) for mammalian expression. The tags in these clones are all fused to the N-terminal of the open reading frame. Full-length protein coding cDNA clones for KCMF1, UBR2, ABHD10, NIPSNAP3A, and SSBP1 were obtained from the Mammalian Gene Collection (National Institute of Health). Subsequently, full-length KCMF1, KCMF1 truncation mutants and Ubr2

(aa. 1010-1267, encompassing Basic Rich Region and RING domain) were subcloned into the pGEX-6p-1 vector (N-terminal GST tag; GE Healthcare, Piscataway, NJ) and/or the pET15b vector (N-terminal 6xHis tag; Novagen, San Diego, CA) for bacterial expression. Full-length KCMF1 and its truncation mutants, ABHD10, NIPSNAP3A, and SSBP1 were cloned into the FLAG-pcDNA5/FRT/TO vector. For immunofluorescence purposes, full-length KCMF1 was cloned into GFP- and Cherry-pcDNA5/FRT/TO vectors for mammalian expression. Full-length ABHD10 was cloned into the pAcGFP1-N3 vector (C-terminal GFP tag; Clonetech). Primers used for cloning are listed in Table 3.1.

3.2.2 Stable Cell lines

Tetracycline-inducible, FLAG-, GFP-, Cherry-tagged proteins were expressed in human Flp-In T-REx 293 cells (Invitrogen, Carlsbad, CA). Protein expression was induced by adding 1 µg/ml tetracycline to the culture medium (DMEM + 10% fetal calf serum) for 24 h. HEK293 cells were maintained at 37°C in DMEM supplemented with 10% fetal bovine serum, 10 mM HEPES (pH 8.0), and 1% penicillin–streptomycin.

3.2.3 Transfection and shRNA interference

293 T-REx cells were maintained at 37°C in DMEM supplemented with 10% fetal bovine serum, 10 mM HEPES (pH 8.0), and 1% penicillin–streptomycin. Cells were transfected with appropriate plasmids using Lipofectamine reagent and PLUS reagent according to the manufacturer’s protocol (Invitrogen). For shRNA interference, 3µg of shRNA (pGIPZ lentiviral shRNAmir; Open biosystems, Thermo Fisher Scientifc) construct against RAD6A (Catalogue No. V3LHS_333991) was used to generate stable knock-down cell lines. Cells were then selected using 1µg/mL puromyocin (Bioshop, Burlington, ON).

3.2.4 Affinity purification

2 For MS analysis of interacting proteins, 6 × 150 cm dishes of sub-confluent (80%), stable, FLAG-tagged protein expressing cells were scraped into PBS, pooled, washed twice in 25 ml PBS, and collected by centrifugation at 1000 × g for 5 min at 4°C. Cell pellets were stored at −80°C. The cell pellet was weighed, and 1:4 pellet weight : volume lysis buffer was added. Lysis buffer consisted of 50 mM HEPES-NaOH (pH 8.0), 100 mM KCl, 2 mM EDTA, 0.1% NP40, 10% glycerol, 1 mM PMSF, 1 mM DTT, and 1:500 protease inhibitor cocktail (Sigma-Aldrich,

St. Louis, MO). On resuspension, cells were incubated on ice for 10 min, subjected to one additional freeze–thaw cycle, and then centrifuged at 27,000 × g for 20 min at 4°C. Supernatant was transferred to a fresh 15-ml conical tube, and 1:1000 benzonase nuclease (Novagen) plus 30 µl packed, preequilibrated Flag-M2 agarose beads (Sigma-Aldrich) were added. The mixture was incubated for 2 h at 4°C with end-over-end rotation. Beads were pelleted by centrifugation at 1000 × g for 1 min and transferred with 1 ml of lysis buffer to a fresh centrifuge tube. Beads were washed once with 1 ml lysis buffer and twice with 1 ml ammonium bicarbonate rinsing buffer (50 mM ammonium bicarbonate, pH 8.0, 75 mM KCl). Elution was performed by incubating the beads with 150 µl of 125 mM ammonium hydroxide (pH 11.0). The elution step was repeated twice. Eluate was centrifuged at 1000 × g for 1 min, transferred to a fresh centrifuge tube, and lyophilized.

3.2.5 Mass Spectrometry

One microgram MS-grade TPCK trypsin (Promega, Madison, WI) dissolved in 70 µl of 50 mM ammonium bicarbonate (pH 8.3) was added to the FLAG eluate and incubated at 37°C overnight. The sample was lyophilized and brought up in buffer A (0.1% formic acid). LC analytical columns (75-mm inner diameter) and precolumns (100-mm inner diameter) were made in-house from fused silica capillary tubing from InnovaQuartz (Phoenix, AZ) and packed with

100 Å C18–coated silica particles (Magic, Michrom Bioresources, Auburn, CA). Peptides were subjected to nanoflow liquid chromatography - electrospray ionization - tandem mass spectrometry (nLC-ESI-MS/MS), using a 90 min reversed phase (10-40% acetonitrile, 0.1% formic acid) buffer gradient running at 250 nL/min on a Proxeon EASY-nLC pump in-line with a hybrid linear quadrupole iontrap (Velos LTQ) Orbitrap mass spectrometer (Thermo Fisher Scientific, Walthma, MA). A parent ion scan was performed in the Orbitrap, using a resolving power of 60,000. Simultaneously, up to the forty most intense peaks were selected for MS/MS (minimum ion count of 1000 for activation) using standard CID fragmentation. Fragment ions were detected in the LTQ. Dynamic exclusion was activated such that MS/MS of the same m/z (within a 10ppm window, exclusion list size 500) detected three times within 45sec were excluded from analysis for 30 sec.

For protein identification, Thermo .RAW files were converted to the .mzXML format using Prot- eowizard (Kessner et al., 2008), then searched using X!Tandem (Craig and Beavis, 2004) against

47 the human (Human RefSeq Version 37) database. Search parameters for X!Tandem specified a parent MS tolerance of 10ppm and an MS/MS fragment ion tolerance of 0.4 Da, with up to 2 missed cleavages allowed for trypsin. Oxidation of methionine was allowed as a variable modification, and a +114.0429 Da modification of lysine was specified as a variable search parameter to identify any ubiquitin-derived diglycine motif. Each immunopurification sample was analyzed using multiple technical replicates. Statistical validation of the results was performed using Peptide Prophet and Protein Prophet (Keller et al., 2002; Nesvizhskii et al., 2003) as part of the trans-proteomic pipeline. For each search, the Protein Prophet probability at 1% error rate was used as a cutoff value to generate SAINT-compatible input files. SAINT parameters were as follows: 5000 iterations, low mode On (1), minFold 1 and no normalization (0) (Choi et al., 2011). Interactors with a 95% confidence level are reported, and the average peptide counts per two technical runs are shown. All AP-MS data are analyzed as indicated above, unless otherwise noted.

3.2.6 Protein Expression and Purification

GST-tagged and His-tagged protein constructs were transformed into Escherichia coli BL21 0 (DE3) cells (Invitrogen, Carlsbad, CA), and were grown in TB medium at 37 C to an OD600nm of ~1.0, induced with 0.5 mM or 1mM IPTG at 40C overnight. Cells were harvested by centrifugation and frozen overnight at -800C. For His-tagged proteins, cell pellets were thawed briefly on ice and resuspended in 50mM Tris (pH7.5), 500mM NaCl, 10% glycerol, 5mM imidazole, 1mM Benzamide, and 1mM PMSF. The proteins were then purified using Talon beads (BD Biosciences, San Jose, CA) and eluted with buffer contain 500mM Imidazole. For GST-tagged proteins, thawed cell pellets were resuspended in phosphate buffered saline (PBS, pH 7.3), 0.1% Triton, 10% glycerol, 2mM DTT, 1mM PMSF, and 1mM Benzamide. Proteins were purified using MagneGSTTM glutathione particles (Promega, Madison, WI), and stored on beads at 40C, or purified using Glutathione-Sepharose 4B beads (GE Healthcare) and eluted with 10mM reduced glutathione, pH 8.0 in native buffer.

3.2.7 In vitro RAD6A binding assay

2 μg-4 μg of GST-KCMF1 proteins on MagneGSTTM glutathione particles were incubated with 2 μg of purified His-tagged RAD6A for 1 h 4 °C in 50 μl of binding buffer (50mM Tris pH 8.0,

150mM NaCl, 2mM DTT, 25µM ZnCl2, and 0.05% Triton). For competition assays, GST-

KCMF1 or GST-Ubr2 protein was first incubated with His-RAD6A for 30 min 4 °C. Increasing amounts of competition proteins (up to 10 molar excess) were then added to the complex and incubated for an additional 1h 4 °C. In both cases, the magnetic particles were washed six times with binding buffer, and bound proteins were eluted with SDS sample buffer. Western blot analysis using monoclonal anti-Penta His antibody (Qiagen, Valencia, CA) was used to detect for the presence of RAD6A. Primary antibody was visualized with the aid of secondary horseradish peroxidase (HRP) anti-mouse antibody (Bio-Rad, Hercules, CA) using enhanced chemiluminescent (ECL) (Immuno-Star HRP, Bio-Rad).

3.2.8 NMR sample preparation

His-tagged KCMF1 protein (CT, aa. 302-281) and His-tagged RAD6A protein (aa. 1-152) was grown at 370C until A600 nm of ~1.0, and then induced with 0.5mM (IPTG) at 40C overnight. For RAD6A expressing cells, the bacteria were grown in M9-defined medium supplemented 15 13 15 15 13 with N-ammonium chloride (0.8 g/L) and/or C6-D-glucose (0.4 g/L) for N or N / C- labeled RAD6A samples. These highly expressed proteins were purified by HisPur Cobalt Resin (Fisher) affinity chromatography under native conditions and eluted with buffer containing 500mM Imidazole. The proteins were further purified by size exclusion chromatography using a HiLoad 26/60 Superdex-75 column (GE Healthcare). The proteins were monomeric in solution as determined by size exclusion chromatography. The final NMR samples were prepared in buffer containing 50mm KH2PO4 pH 8.0, 200mM NaCl, 5mM β-mercaptoethanol,1mM benzamide, 0.5mM PMSF, and 5% D2O. The RAD6-KCMF1 complex was prepared for NMR by titrating aliquots of unlabeled KCMF1 into the 15N labeled RAD6A (500µM) in molar ratios from 1:1 to 1:20 until no further changes in chemical shifts were detected in the 1H-15N HSQC spectrum. The weighted chemical shift displacements were calculated using the following formula: Δppm=((ΔδHN)2 +(ΔδN/5)2)1/2.

3.2.9 NMR Spectroscopy and Data Analysis

The assignment of RAD6A has been done using the ABACUS approach (Lemak et al., 2008) from NMR data collected at high resolution from nonlinearly sampled spectra and processed using multidimensional decomposition (Gutmanas et al., 2002; Orekhov et al., 2003). All of the NMR spectra were recorded at 310K on Bruker Avance 600 and 800-MHz spectrometers equipped with cryoprobes. The sequences specific assignment of 1H, 13C and 15N resonances

49 were assigned using the ABACUS protocol (Lemak et al., 2008) from peaks lists derived from manually peak picked spectra using SPARKY. Figures were prepared using PyMOL (DeLano Scientific).

3.2.10 Autoubiquitylation assay and mass spectrometry analysis

Autoubiquitylation reactions were performed in a volume of 30µL in a buffer of 50mM Tris pH 8.0, 50mM NaCl, 50mM KCl, 10mM MgCl2, 5mM ATP, 0.1mM DTT, with 5ug Ub (human recombinant; Boston Biochem), 3µM E2, 90nM E1 (Human Recombinant, UBE1; Boston Biochem), and E3 3µM. After incubation at 30°C for 3 h, reactions were stopped by the addition of SDS-PAGE sample buffer and resolved on Criterion TGX Precast Gel 4-20% (Bio-Rad) Ubiquitylated proteins were evaluated by Western blotting using mouse monoclonal antibody against ubiquitin (P4G7, equivalent to P4D1; Covance, Emeryville, CA), an HRP-conjugated goat-anti-mouse secondary (Bio-Rad), and Immun-Star HRP ECL (Bio-Rad).

Autoubiquitylation reactions were scaled-up 3-fold for mass spectrometric analysis, and subjected to 4-12% gradient SDS-PAGE. Gels were stained with Coomassie brilliant blue for visualization, and the region containing proteins migrating at >125kDa was processed as in(Lallemand-Breitenbach et al., 2008) for mass spectrometry. See 2.3.4 for MS details.

3.2.11 Immunofluorescence and Image Acquisition and Processing

Stable cell lines expressing tagged proteins or cultured HEK293 cells were grown on coverslips and fixed with 4% formaldehyde for 15 min and washed in PBS with 0.25% Triton X-100. For mitochondria staining, MitoTracker Red CMXRos dye was added to the cells before fixing at a concentration of 300nM for 15min at 370C. To treat cells with MG132 (Calpain inhibitor IV, Z- Leu-Leu-Leu-CHO; American Peptide Company, Sunnyvale, CA), 10µM of MG132 was added to cells for 12h, before fixing the cells with 100% methanol at -200C for 10min.. The cells were blocked in 5% bovine serum (BSA) in PBS for 30min before incubating in appropriate primary antibody mixtures for 1h at RT. Primary antibody dilutions used included anti-KCMF1 polyclonal antibody (Rabbit, 1:450; Sigma prestige Antibodies), anti-ubiquitin Rabbit pAb (1:200; Calbiochem, San Diego, CA), anti-GFP mouse monoclonal antibody (clones 7.1 and 13.1, 1:1000; Roche, Mississauga, ON), anti-FLAG M2 antibody (1:500; Sigma-Aldrich), anti- EEA1 (1:200), anti-CD63 (1:500), anti-GM130 (1:500), anti-Lamp1 (1:100), and anti-KDEL

(1:100). The latter five antibodies were all mouse antibodies, and a gift from J. McGlade lab (Hospital for Sick Children, Toronto, ON). Secondary antibodies conjugated to Alexa 488, and Texas Red were used at 1:500, and incubated at RT for 1h. After removing from the antibody solution, cells were incubated with 1µg/mL of 4’,6-dia-midino-2-phenylindole (DAPI) in PBS for 5 min. After washing with PBS three times for 5 min each, the coverslips were mounted with the ProLong Gold Antifade reagent (Thermo Fischer Scientific). Cells were imaged using PlanApo 60X oil lens, NA 1.40 on Olympus FV1000 confocal microscope (zoom factor between 3-5; Olympus America, Melville, NY). Images were then processed using FluoView10ASW 3.0 Viewer, version 3.1.1.9 (Olympus), and assembled using Adobe Illustrator CS5 (Adobe Systems Inc.)

3.2.12 Live-cell confocal microscopy

For live-cell studies, cultured cells were stained with 1µg/mL of Hoechst 33342 dye (Fisher scientific) for 15min at 370C. Cells were washed twice with warm PBS, before incubating in DMEM media with no pH indicator (GIBCO, Invitrogen). MG132 treatment, mitochondria staining and image processing were as described as above. Aggresome formation was detected as a single perinuclear inclusion and was confirmed by the indented shape of the nucleus. For each cell condition, more than 200 transfected cells from randomly selected fields were analyzed. Two-tailed student’s t-test was used for statistical analysis (p=0.05). For CFTR localization, 0.4µg of the GFP-CFTR-Δ508 plasmid was transfected into stable cell line expressing protein of interest. 24h post transfection, cells were treated with 10µM of MG132 for 12h before imaging. The GFP-CFTR-Δ508 plasmid was a gift from Dr. R. Grace Zhai (University of Miami Miller School of Medicine, Miami, Florida)

3.2.13 qRT-PCR

Cultured cells were grown to 80% confluency before proceeding to RNA purification using the total RNA extraction kit from Qiagen. RNA quantity was estimated on a Nanodrop 1000 (Thermo Fisher Scientific). RT-qPCR primers were designed using the cDNA for RAD6A with qPCR settings in Primer3Plus (Untergasser et al., 2007). 40 ng template RNA and 500nM of each primer were used with the Power SYBR® Green RNA-to-CT™ 1-Step Kit (Applied Biosystems, Carlsbad, CA) in 20µl reactions, as per manufacturer’s protocols, on a Stratagene

Mx3000P. GAPDH was used as a control for ΔΔCt-based relative quantification (Livak and Schmittgen, 2001).

Table 3.1 Primers used for cloning. Restriction sites are indicated by the primer’s name.

3.3 Results

3.3.1 A mammalian RAD6 interactome

The functions of Rad6 have been well characterized in yeast. To learn more about the functions of RAD6 in mammalian cells, I characterized the interactome of both RAD6A and RAD6B in Human Embryonic Kidney (HEK) 293 cells. Tetracycline-inducible HEK293 cell lines (Flp-In T-REx system), stably expressing FLAG-tagged RAD6A and RAD6B proteins were established. Following induction in two independently isolated cell pools, FLAG-tagged RAD6A/B was immunopurified under non-denaturing conditions. Interacting proteins were identified using nanoflow liquid chromatography-electrospray ionization-tandem mass spectrometry (nLC-ESI- MS/MS). Cells expressing the FLAG tag alone, and a number of unrelated FLAG-tagged proteins, were analyzed simultaneously, to control for background polypeptides that interact non- specifically with the antibody or solid phase support. MS data were analyzed using the trans- proteomic pipeline (TPP) suite, and subjected to SAINT analysis (Choi et al., 2011). Those proteins assigned a SAINT confidence value >0.95 (and therefore predicted to be bona fide RAD6 interacting partners) are reported in Table 3.2.

Using AP-MS, I was able to identify all of the known RAD6-interacting E3 ligases (UBR1, UBR2, RAD18, RNF20-RNF40) (Bailly et al., 1997; Kim et al., 2009; Wang et al., 2004b; Watkins et al., 1993; Zhang and Yu, 2011). I also identified the WW domain-containing adaptor with coiled-coil (WAC), which regulates Histone 2B ubiquitylation through its functional interaction with RNF20-RNF40 (Zhang and Yu, 2011). Interestingly, a number of previously unreported high-confidence interacting partners were also found for RAD6. This includes two E3 ligases, potassium channel modulatory factor 1 (KCMF1) and ubiquitin protein ligase E3 component n-recognin 4 (UBR4); an abhydrolase domain-containing protein, ABHD10; two related proteins implicated in vesicle trafficking, 4-nitrophenylphosphatase domain and non- neuronal SNAP25-like protein (NIPSNAP) homolog 3A; and two mitochondria-associated proteins, single stranded binding protein 1 (SSBP1), and a seryl-tRNA synthetase, SARS2. RAD6A and RAD6B shared the same binding partners. These data define for the first time a RAD6 interactome in human cells.

Table 3.2 A RAD6 interactome. FLAG-tagged RAD6A and RAD6B were expressed in Flp-In T-REx 293 cells. Cells were lysed under non-denaturing conditions and FLAG-tagged RAD6A/B and associated proteins were immunopurified using FLAG-M2 agarose beads. Immunopurified proteins were eluted with ammonium hydroxide, digested with trypsin, and analyzed by nLC-MS/MS. For protein identification, Thermo .RAW files were converted to the mzXML format using Proteowizard, then searched using X!Tandem against the human (Human RefSeq Version 37) database. Data were analyzed using the ProHits software tool (Liu et al., 2010) and SAINT (Choi et al., 2011). Each pool represents a different biological replicate. For each pool, two technical runs were run on the MS. The average number of spectral counts (the number of times a peptide from each protein was observed) resulting from the two technical runs are indicated. Proteins detected with a>95% confidence level (TPP) and determined to be bona fide RAD6 interactors (SAINT score >0.95) are shown. Blue colored interactors are known E3 ligases/interactors of RAD6. Orange colored interactors are novel.

3.3.2 A RAD6A XLMR mutant protein loses the interaction with the newly identified RAD6 interactors

Several studies have identified mutations in the RAD6A coding sequence that cause XLMR in patients from three families. (Budny et al., 2010; Nascimento et al., 2006). Two of the point

55 mutations (R11Q, G23R) are at the N-terminus, while the other mutation (Q128X) is at the C- terminus. Both regions of RAD6 are highly conserved. They have been implicated to interact with different domains of the E3 ligases. To determine if these RAD6A mutations disrupt interactions with the E3 ligases, the different RAD6A point mutants were expressed as FLAG- tagged proteins in 293 T-REx cells, and then conducted AP-MS as above. The RAD6A Q128X mutant, which lacks the 25 C-terminal amino acids (Nascimento et al., 2006) did not express well in the system. The G23R mutation had no apparent effect on protein-protein interactions (Table 3.3). On the contrary, the RAD6A R11Q mutant maintained interactions with UBR1, UBR2, RAD18, RNF20/40, and WAC, but specifically lost the interactions with KCMF1, UBR4, ABHD10, NIPSNAP3A, SSBP1, and SARS2 (Table 3.3). In contrast to known RAD6A interactors, the novel interactor(s) may be binding to a different region on the E2 protein.

Table 3.3 RAD6A R11Q mutation disrupts protein-protein interactions. FLAG-tagged RAD6A mutant proteins (R11Q, G23R and Q128X) were expressed in Flp-In T- REx 293 cells. Interacting proteins were identified viaAP-MS, as above (see Materials and methods for details). Each immunopurification sample was analyzed using two technical replicates. Table shows the average spectra counts resulting from the two technical replicate. Each pool corresponds to a biological replicate.Proteins detected with a>95% confidence level (TPP) and determined to be bona fide RAD6 interactors (SAINT score >0.95) are shown. The RAD6A R11Q mutation disrupts the interactions with KCMF1, UBR4, ABHD10, NIPSNAP3A, SSBP1 and SARS.

3.3.3 KCMF1 interact with all the newly identified RAD6 interactors

To further examine some of the novel RAD6 interactors, I generated stable cell lines (293 T- Rex) expressing FLAG-tagged proteins, and conducted AP-MS as above. The first cell line that I generated was KCMF1, a Ub E3 ligase (Jang et al., 2004). Interestingly, RAD6A was identified as a high-confidence interactor for KCMF1 (Table 3.4). Moreover, all of the newly identified RAD6A interacting partners (UBR4, ABHD10, NIPSNAP3A, SSBP1, and SARS2) also interacted with KCMF1. These data point to the possibility of one or more novel RAD6A- containing complexes.

KCMF1 is an evolutionarily conserved protein, with a homology of 95% between human and zebrafish. The protein is organized into seven exons spanning an 88.38 MB region on human chromosome 2 (Beilke et al., 2010). Northern blot analysis of KCMF1 revealed that it has higher mRNA expression levels in human spleen, small intestine, ovary, peripheral blood, lung, skeletal muscle, kidney, and pancreas, and lower mRNA expression levels in thymus, prostate, testis, colon, heart, brain, placenta, and liver (Jang, 2004). KCMF1 has been implicated in a number of cancers. Its mRNA expression level is downregulated in Ewing’s sarcoma cell lines after overexpression of CD99 (cluster of differentiation 99) (Kreppel et al., 2006), while its mRNA expression level is upregulated through fibroblast growth factor (FGF) receptor2 signaling pathways in gastric cancer cells (Jang, 2004). KCMF1 proteins are overexpressed in epithelial cancers, enhancing proliferation, migration and invasion (Beilke et al., 2010). Using AP-MS, several other bona fide interactors were identified for KCMF1, including components of the nuclear pore complex (NPC), NUP93 and NUP188 (components of the spoke ring), NUP214 and NUP88 (cytoplasmic filaments), as well as the O-linked N-acetylglucosamine (GlcNAc) transferase (OGT) protein.

Table 3.4 Interactome of KCMF1. FLAG-tagged KCMF1 was expressed in 293 T-REx cells and subjected to AP-MS. Each pool represents a different biological replicate. For each pool, two technical runs were run on the MS. Spectral counts (the number of times a peptide from each protein was observed) resulting from each technical run are indicated. High confidence interactors (SAINT score >0.95) for KCMF1 include RAD6A, and all the additional novel RAD6A interactors. KCMF1 can also interact with nucleoporins and a nucleoporin-associated protein (OGT). PTM = Post-translational modification.

3.3.4 KCMF1 directly binds to RAD6A in vitro

To test whether KCMF1 can bind directly to RAD6A, a GST-tagged KCMF1 protein was expressed in E.coli and immobilized on MagneGST™ glutathione particles. This was followed by incubation with recombinant His-tagged Rad6A. The GST-tag alone was used as a negative control, and a GST-Ubr2 fusion protein (consisting of the BRR and RING domains) was used as a positive control for RAD6A binding. Like Ubr2, the full-length KCMF1 can also bind directly to RAD6A in vitro (Figure 3.1 A lanes 3 and 5).

1 2 3 4 5 6 7 8 9 10 A C 2% RAD6A RAD6A 2% input Ubr2 -beads GST KCMF1 KCMF1A KCMF1B KCMF1C KCMF1CT KCMF1D 3 54 78 101 213 287 381 KCMF1 RING C2H2 Coiled Coil PRR RAD6A KCMF1A RING

KCMF1 B RING C2H2 B KCMF1 C C2H2 Coiled Coil PRR KCMF1A KCMF1A KCMF1B KCMF1C KCMF1CT 2% RAD6A RAD6A 2% input KCMF1 KCMF1D Ubr2 -beads GST KCMF1 D Coiled Coil PRR

RAD6A- KCMF1 CT R11Q PRR

Figure 3.1 The KCMF1 C-terminus can bind to RAD6A (A) In vitro RAD6A binding assay. Recombinant GST-KCMF1 proteins (including the indicated truncation mutants), GST-Ubr2 (BRR+RING) and His-tagged RAD6A were produced in E. coli and captured on glutathione magnetic particles. Each protein was incubated with purified His- RAD6A protein for 1h at 40C followed by extensive washing. Proteins were eluted with SDS- sample buffer, resolved by SDS-PAGE, and analyzed by immunoblotting using an anti-His antibody. GST-Ubr2 (lane 2) was used as a positive control, while GST alone (lane3) was used as a negative control. KCMF1 CT is necessary and sufficient to bind to RAD6A. (B) In vitro RAD6A R11Q binding assay. Rad6 binding assay conducted as above. The RAD6A R11Q does not bind to KCMF1. (C) Schematic representation of KCMF1 truncation mutants. The indicated KCMF1 truncation mutants were generated using PCR and subcloned into the pGEX-6p-1 vector. Features and domains of KCMF1, including the RING, C2H2 and Coiled-coil domains, and proline rich region (PRR) are indicated. Amino acid residues assigned to each domain are based on the Jpred3 secondary structure prediction program (Cole et al., 2008).

3.3.5 The C-terminus of KCMF1 is necessary and sufficient to bind to RAD6A

To further characterize the physical interaction between RAD6A and KCMF1, I generated a series of recombinant GST-KCMF1 truncation mutants in E. coli to identify the region(s) of KCMF1 necessary for RAD6A binding in vitro. In silico analyses indicate that KCMF1 contains two N-terminal zinc-finger domains (a RING domain followed by a C2H2-type zinc finger), a coiled-coil domain, and a proline-rich region (Beilke et al., 2010) (Figure 3.1 C). Removal of the two N-terminal zinc-finger domains (KCMF1 D) had no effect on RAD6A binding in vitro (Figure 3.1 A lane 9). In contrast, KCMF1 mutants lacking the C-terminus lost binding to RAD6A (KCMF1 A and B truncation mutants, lanes 6 and 7). The residues that are important for RAD6A binding were thus delimited to those between 302 and 381 (KCMF1 CT, lane 10).

Consistent with my AP-MS results, the RAD6A R11Q mutant did not bind to KCMF1 (and its truncation mutants) in vitro, but was able to interact with Ubr2 (Figure 3.1 B)

A similar set of truncated KCMF1 mutant proteins were stably expressed in 293 T-REx cells (FLAG-tagged). Consistent with my in vitro data, deletion of the N-terminus did not affect the interaction with RAD6A in vivo, whereas the loss of KCMF1’s C-terminus (KCMF1 B) resulted in a loss of Rad6A binding (Table 3.5). Moreover, only the KCMF1 CT was required for RAD6A interaction in vivo. Interestingly, when KCMF1 lost its affinity for RAD6A (see KCMF1 B), interactions with UBR4, ABHD10, NIPSNAP3A, SSBP1, and SARS2 appeared to be unaffected. These proteins were also shown to interact with RAD6A (Table 3.2). This suggests that KCMF1 may be bridging these proteins to RAD6A.

The KCMF1 truncation mutants provided further insights into to the domain(s) involved in binding to some of its interactors. Deletion of the N-terminal zinc fingers resulted in a loss of interaction for ABHD10, NIPSNAP3A, and SARS2 proteins (Table 3.5). The C2H2 domain alone was able to interact with these proteins to varying degrees. The coiled-coil domain mainly interacts with nucleoporin proteins (NUP188, NUP93, NUP214, NUP88) and with OGT (Figure 3.2). OGT is a glycosyltransferase that can modify nucleoporins and plays a role in nuclear transport (Butkinaree et al., 2010; Yu et al., 2012). It is unclear where UBR4 and SSBP1 bind on KCMF1. The UBR4 interaction appears to be mediated by more than one region of the KCMF1 protein, through both the zinc-finger domains and the C-terminal region (Table 3.5). Together these data define the interactome of KCMF1, as well as the regions on KCMF1 necessary to interact with each of its binding partners.

Table 3.5 KCMF1’s C-terminus interacts with RAD6A in vivo. FLAG-tagged KCMF1 and truncation proteins were expressed in 293 T-REx cells and subjected to AP-MS. Each pool represents a different biological replicate. For each pool, two technical runs were run on the MS. The average spectral counts resulting from two technical runs are indicated. Proteins detected with >95% confidence level (TPP) and determined to be bona fide KCMF1 interactors (SAINT score >0.95) are shown. KCMF1’s C-terminus (CT) retains binding to RAD6A.

Figure 3.2 KCMF1 interacting domains RAD6A binds on C-terminus of KCMF1. NIPSNAP3A and ABHD10 interact with KCMF1 through its zinc-fingers. Nucleoporin proteins and OGT interact with KCMF1 through its coiled- coil domain.

3.3.6 RAD6A binds to KCMF1 through its N-terminal helix and residues around its active site

NMR spectroscopy has been used previously to identify the binding residues on Rad6 for several different E3 ligases. In addition to the conserved RING-E2 interface, Ubr1 and Rad18 recognize the C-terminus of RAD6, and the noncovalent Ub binding site (antiparallel βstrands) via a secondary RAD6-interacting region (BRR and R6BD for Ubr1 and Rad18, respectively) (Hibbert

61 et al., 2011; Worthylake et al., 1998). To identify the RAD6A residues that mediate KCMF1 binding, a series of 1H-15N heteronuclear single quantum coherence (HSQC) spectra of 15N- labeled RAD6A was recorded, to which increasing amounts of unlabeled KCMF1 CT were added (up to a final molar ratio of 1:20) (Figure 3.3 A). Based on our own assignment of RAD6A, the chemical shift perturbations (CSPs) can be grouped into two regions: 1) the N- terminal helix α1 (R8, R11, K14, Q17), a part of the canonical RING-E2 interaction surface (Huang et al., 2011; Miura et al., 1999); 2) the region surrounding the catalytic Cys88, with shifts on helix α2, and 310 helix residues (Figure 3.3 C). Notably, a few of the residues (L89, V102, Q110, A122) in this second region displayed a significant decrease in peak intensity (>90%) upon KCMF1 titration (Figure 3.3 B). The affinity between KCMF1 CT and RAD6A was unable to be determined, since the titration did not approach saturation under the concentrations used. The pattern of CSPs shows a weaker interaction than Rad18 R6BD (Hibbert et al., 2011), therefore we estimate an affinity in the mM range. Interestingly, in comparison to the binding surface between R6BD and BRR to RAD6, KCMF1 CT binds to a different surface on RAD6 (Figure 3.4). This could help to explain why the RAD6A R11Q mutant specifically loses the interaction with KCMF1.

Figure 3.3 The interaction of KCMF1 CT with RAD6A. NMR spectra (1H-15N-HSQC) of RAD6A alone (black), and with increasing amounts of KCMF1 CT to a final RAD6A:CT molar ratio of 1∶20 (red). Significant (>0.04) CSPs of RAD6A are indicated in blue (residues R6, R11, A38, F41, G45, 51, L55, K66, T69, F77, V81, G85, I109, D114, Q131), and peaks with decreasing intensity are shown in green (L9, S29, W36, F59, V73, Y82, I87, L89, W96, V102, A122, N123, A127). R11 was one of the peaks that displayed a small chemical shift, highlighted in orange. (B) CSP data from the titration of KCMF1 CT into 15N-RAD6A. Histogram displaying weighted chemical shift changes with respect to RAD6A residue number. Each amide nitrogen along the polypeptide backbone of RAD6A has a spectroscopic label (15N) plotted on the x axis. The magnitude of the spectra change (reflecting a change in local environment upon binding) is plotted on y axis. Observed changes are indicative of subtle structural rearrangements in RAD6A upon binding to the KCMF1. (C) Mapping of 15N-1H chemical shift changes for RAD6A upon titration with KCMF1 CT on the Rad6b structure (PDB code 2Y4W). Significant (>0.04) CSPs of RAD6A are highlighted in red. Blue color indicates resonances that disappeared. Green color represents the active Cys88.

Figure 3.4 A comparison of the RAD6 binding sites for Rad18, Ubr1 and KCMF1. (A) Rad6 binding residues involved in theRad18 R6BD interaction. Residues that showed significant chemical shift when 15N-Rad6 was titrated with R6BD synthetic peptide are shown in blue (Hibbert et al., 2011). Active Cys88 is colored green. (B) RAD6 binding residues for KCMF1 CT interaction. Red indicates residues showing significant chemical shifts, blue indicates residues showing decreasing intensity, and green indicates the active Cys88. (C) Rad6 binding residues for Ubr1 BRR interaction. Residues that showed significant chemical shift when 15N-Rad6 was titrated with BRR synthetic peptide are shown in blue (Hibbert et al., 2011). Active Cys88 is colored green. Unlike Rad18 and Ubr1, KCMF1 binds to a different region on RAD6. The mapping of all 15N-1H chemical shift changes for RAD6A upon titration with Rad18 R6BD, Ubr1 BRR, and KCMF1 CT are all done on the Rad6b structure (PDB code 2Y4W).

3.3.7 KCMF1 inhibits RAD6A chain formation activity in vitro

KCMF1 was previously reported to act as an E3 ubiquitin ligase with the E2 protein UbcH5b (UBE2D2) (Jang, 2004). To determine if KCMF1 functionally interacts with RAD6A, I conducted RAD6 in vitro autoubiquitylation reactions, as described in (Hibbert et al., 2011). Briefly, affinity purified GST-tagged KCMF1 protein was incubated at 300C with ATP, E1, Ub, and purified His-tagged RAD6A protein for 3 hours. As previously shown, in the absence of an E3, RAD6A can synthesize high molecular weight Ub oligomers in vitro (Figure 3.5 A). Mass spectrometric analysis of these reaction products revealed a mix of K11-, K48- and K63-linked

Ub chains (Figure 3.5 C). The linkage composition of Ub chains was not altered in the presence of full-length KCMF1 or Ubr2 (BRR and RING domain) (Figure 3.5 C).

Figure 3.5 RAD6A autoubiquitylation reactions. (A) Full-length KCMF1 inhibits RAD6A-mediated ubiquitin chain formation. Autoubiquitylation reactions were performed with ATP, recombinant human E1 (90 nM), recombinant human ubiquitin (5 µg), and His-RAD6A (3 µM) alone, or reconstituted with stoichiometric GST-tagged KCMF1, GST-tagged KCMF1 truncation proteins or GST-Ubr2 (BRR+RING) for 3h at 300C. This was followed by anti-Ub Western blotting. A KCMF1 mutant lacking the N-terminus, KCMF1 D (lane 3) had no effect on RAD6A activity. Like the full- length KCMF1 protein, a KCMF1 mutant lacking the N-terminus (lane 4) had some inhibitory effect on RAD6A Ub chain building activity. (B) KCMF1 CT can inhibit RAD6A-mediated Ub

65 chain formation. Autoubiquitylation reactions were performed as indicated above. The concentration of GST-KCMF1 CT used was 100µM. The R11Q point mutation impairs RAD6A polyubiquitylation activity (lane 5). While the addition of KCMF1 protein had no effect in this assay, the addition of Ubr2 resulted in increased RAD6A R11Q autoubiquitylation activity. (C) Ub linkage analysis. WT RAD6A autoubiquitylation reaction products were subjected to SDS- PAGE, and Ub multimers (migrating at >100 kDa) analyzed by mass spectrometry. Ub linkage composition for each E2-E3 reaction is presented as total spectral counts seen from a MS run. See Legend inset for color codes. Ubr2 and KCMF1 do not alter the linkage composition of RAD6A mediated Ub chains. (D) Schematic diagram depicting KCMF1 truncation mutants used in the in vitro autoubiquitylation assays.

Although the addition of the E3 ligases had no effect on linkage composition, the addition of the GST-Ubr2 protein (BRR and RING domain) stimulated Ub chain-building activity of RAD6A (Figure 3.5 A lane 5), and the GST-KCMF1 protein had an inhibitory effect (lane 2). Ubr1 BRR was previously shown to stimulate Rad6 polyubiquitylation activity (Hibbert et al., 2011). It is not surprising that Ubr2 has the same effect, since both of these E3 ligases are involved in protein degradation. Hibbert and collegues (2011) have also shown that Rad18 can inhibit the intrinsic polyubiquitylation activity of RAD6, in order to promote monoubiquitylation of PCNA (Hibbert et al., 2011). This is similar to the effect of KCMF1 on RAD6A. Consistent with my in vivo and in vitro interaction data, the addition of the KCMF1 N-terminus had no apparent effect on RAD6 activity (KCMF1 D, Figure 3.5 A lane 3), while the addition of the KCMF1 C- terminus (KCMF1 B, lane 4; KCMF1 CT, Figure 3.5 B lane 3) inhibited RAD6A Ub chain formation activity in vitro (albeit not to the same extent as the full-length protein, suggesting that other regions of the protein may be required for full activity). Together these data suggest that Ubr2 acts to increase the chain-forming activity of RAD6, while KCMF1 inhibits it.

I also tested the chain-formation activity of the RAD6A R11Q mutant protein (found in XLMR patients). In the absence of any E3, RAD6A R11Q did not efficiently catalyze the assembly of Ub chains (Figure 3.5 B lane 5). The addition of full-length KCMF1 had no effect on this result, but the addition of Ubr2 significantly enhanced RAD6A R11Q Ub chain-building activity (Figure 3.5 B lanes 7 and 8). The R11Q mutation thus not only disrupts protein-protein interactions, but also appears to affect RAD6’s ability to build Ub chains.

3.3.8 Intracellular localization of KCMF1

To investigate the subcellular localization pattern of the KCMF1 and RAD6A proteins, I generated stable cell lines (293 T-REx) expressing N-terminal green fluorescent protein (GFP), Cherry- or FLAG-tagged protein. Immunofluorescence staining (for FLAG-tagged proteins) and live-cell confocal microscopy (for GFP-tagged proteins) for KCMF1 revealed its localization in the cytoplasm (Figure 3.6 A, B). An identical pattern of localization was observed for a KCMF1-specific antibody (Sigma-Aldrich), suggesting that the addition of N-terminal epitope tags to KCMF1 does not alter its subcellular localization (Figure 3.6 C). Indeed, immunohistochemical staining of pancreatic cancer tissue sections revealed mainly cytoplasmic localization for KCMF1 (Beilke et al., 2010). KCMF1’s staining pattern detected by the anti- KCMF1 antibody overlaps with the staining pattern detected by the anti-FLAG antibody (Supplementary Figure 1), demonstrating that the anti-KCMF1 antibody is specific to KCMF1. Subsequent studies were conducted with cells expressing either GFP-, Cherry- or FLAG-tagged KCMF1.

As reported earlier (Gerard et al., 2011; Shekhar et al., 2002), RAD6A displayed both nuclear and cytoplasmic localization (Figure 3.7 A). The RAD6A R11Q mutant displayed the same localization pattern (Figure 3.7 B). When Cherry-KCMF1 protein was transiently expressed in cells stably expressing RAD6A, KCMF1 co-localized with RAD6A in the cytoplasm (Figure 3.8).

Figure 3.6 Intracellular localization of KCMF1. 293 T-REx cells stably expressing GFP- or FLAG-tagged KCMF1 were viewed using live-cell imaging (A) or immunofluorescence staining with the anti-FLAG M2 antibody (B). KCMF1 is expressed in the cytosol. An N-terminal tag on KCMF1 did not affect its localization. (C) Endogenous KCMF1 staining using an anti-KCMF1 antibody was performed using untransfected 293 T-REx cells. DNA was stained with DAPI for IF studies, Hoecst for live-cell image studies (blue). Scale bars: 10 µm.

Figure 3.7 Intracellular localization of the RAD6A and RAD6A R11Q proteins. 293 T-REx cells stably expressing GFP- or FLAG-tagged RAD6A (A)orRAD6A R11Q (B) proteins were viewed using live-cell imaging (for GFP-tagged proteins) and immunofluorescence imaging (for FLAG-tagged proteins). DNA was stained with DAPI or Hoechst (blue). The mutant RAD6A protein shared the same intracellular localization as the WT protein. Scale bars: 10 µm.

Figure 3.8 KCMF1 colocalizes with RAD6A in the cytosol. 293 T-REx cells stably expressing GFP-RAD6A were transfected with a Cherry-KCMF1 construct. The cells were imaged live 48h post transfection. DNA was stained with DAPI (blue). Scale bar: 10 µm.

Many of KCMF1’s interactors have been reported to localize and/or function within membranous organelles, including the mitochondria. For instance, ABHD10 has recently been identified to be responsible for the deglucuronidation of an immunotoxic metabolite, acyl glucuronide (AcMPAG) in both human liver cytosol and microsomes (Iwamura et al., 2012). Analysis of the primary sequence using mitoprot (http://ihg.gsf.de/ihg/mitoprot.html) shows that ABHD10 has cleavable mitochondrial import sequences, suggesting that ABHD10 may be localized to the mitochondria. NIPSNAP3A belongs to a conserved family of proteins with putative vesicular transport functions (Buechler et al., 2004). It can also be found in the

69 mitochondria, and has been suggested to play a role in promoting apoptosis (Verhagen et al., 2007). SARS2 is a mitochondrial Seryl-tRNA synthetase that attaches amino acids to the 3’ends of its cognate tRNA (Belostotsky et al., 2011).To determine if KCMF1 protein can be found in the mitochondria, I conducted immunofluorescence for KCMF1 along with a mitochondria- directed dye (using the Mitotracker Red CMXRos probe (Invitrogen) in 293 T-REx cells. FLAG- tagged KCMF did not localize to mitochondria (Figure 3.9 A). I obtained an identical result using an anti-KCMF1 antibody (Sigma-Aldrich) (Figure 3.9 B). Using a biochemical fractionation kit (Qproteome Mitochondria Isolation Kit, Qiagen), followed by anti-FLAG Western analysis, FLAG-KCMF1 was also not detected in the mitochondrial fraction (Figure 3.9 C lane 4). Notably, FLAG-KCMF1 was detected in both the cytoplasmic and microsomal fractions (Figure 3.9 C lanes 1 and 3). Together these data suggest that the KCMF1-RAD6A complex is cytoplasmic, and possibly within vesicular compartments.

Figure 3.9 KCMF1 proteins are in both the cytoplasmic and microsomal cellular fractions. (A) 293 T-REx cells stably expressing FLAG-tagged KCMF1 were stained for both the mitochondria, using the mitoTracker Red CMXRos dye (Invitrogen) and KCMF1, using the anti- FLAG M2 antibody. KCMF1 proteins are not found in the mitochondria. (B) Colocalization between KCMF1 and mitochondria was also determined in untransfected 293 T-REx cells using an anti-KCMF1 antibody. DNA was stained with DAPI (blue). Scale bar: 10 µm. (C) Results from biochemical fractionation. 293 T-REx cells stably expressing FLAG-tagged KCMF1 were fractionated using the Qproteome mitochondria fractionation kit (Qiagen). Proteins were then ran on SDS-PAGE gel, and visualized using anti-FLAG M2 antibody (1:1000) followed by ECL. FLAG-tagged KCMF1 proteins can be found in the cytoplasmic and microsomal fractionations.

3.3.9 KCMF1 may localize to endoplasmic reticulum and lysosomes

To determine if KCMF1 localizes to membranous organelles in the cell, such as the endoplasmic reticulum (ER), Golgi, endosome and lysosome, I used a panel of antibodies marking these compartments withIF against endogenous KCMF1. The panel of antibodies includes a GM130 (Golgi-Matrix protein, 130 kDa)-specific antibody, cis-Golgi marker; a KDEL (ER targeting sequence)-specific antibody, ER marker; an EEA1 (Early Endosome Antigen 1)-specific antibody, early endosomal marker; a CD63 (Lysosomal associated membrane protein 3) and a LAMP-1 (Lysosomal associated membrane protein 1) specific antibodies, lysosomal markers. Interestingly, fluorescence microscopy revealed that KCMF1 co-localizes with the ER marker (KDEL) and lysosome markers (CD63, LAMP-1) (Figure 3.10). UBR4, which was shown to interact with both KCMF1 and RAD6, has been identified as a novel endoplasmic reticulum (ER)-associated microtubule associated protein (MAP) important for ER dynamics, and regulates neuronal differentiation and migration (Shim et al., 2008). Therefore KCMF1 may have ER- and lysosome-associated roles. The localization of RAD6 to the ER and lysosome remain to be determined.

Figure 3.10 KCMF1 localizes to ER and lysosomes. 293 T-REx cells were cultured and fixed in 4% paraformaldehyde. Using immunofluorescence staining and fluorescence microscopy, colocalization between endogenous KCMF1 (using an anti-KCMF1 antibody) and a number of membraneous organelle markers (antibodies) wasanalyzed. Immunofluorescence was conducted using antibodies directed against GM130 (a cis-Golgi marker, A), EEA1 (an early endosomal marker,B), KDEL (an ER marker,C), LAMP-1 (D) and CD63 (E) (lysosomal markers). DNA was stained with DAPI (blue). Scale bar: 10 µm.

3.3.10 KCMF1 localizes to aggresomes in response to proteasome inhibition

In addition to the diffuse localization pattern, I found that GFP-KCMF1 showed small spherical inclusions in <7% of the cells. This localization pattern was reminiscent of aggresomes

(Johnston et al., 1998). I therefore examined whether the localization pattern for KCMF1 is altered upon addition of the proteasome inhibitor MG132. After six or 12 hours of MG132 treatment, a majority of 293 T-REx cells showed perinuclear cytoplasmic inclusions of GFP- KCMF1 (52%) (Figure 3.11 A, B). Immunofluroscence staining using endogenous KCMF1 antibody yielded the same results (Figure 3.11 C). The mutant cystic fibrosis transmembrane conductance regulator (CFTR) ΔF508 found in cystic fibrosis patient (Johnston et al., 1998) can be used as a marker for aggresomes. Indeed, KCMF1 colocalizes with GFP-CFTR-ΔF508, demonstrating aggresome localization (Figure 3.11 D).

Cytosolic inclusions containing disease-specific proteins are a hallmark of neurodegenerative diseases, including Parkinson’s disease (PD), Alzheimer’s disease (AD), Huntington’s disease (HD) and Lafora disease (Mittal et al., 2007; Olzmann et al., 2008). When the Ub-proteasome pathway is disrupted or overloaded, misfolded proteins are compartmentalized into aggresomes and facilitate their clearance through autophagy (Garcia-Mata et al., 2002; Johnston et al., 1998; Kopito, 2000). Aggresomes are dynamic structures that recruit chaperones and proteasomes to aid in the disposition of misfolded proteins (Garcia-Mata et al., 2002). Since KCMF1 is a Ub E3 ligase, I also checked for colocalization of Ub and KCMF1 at aggresomes. In the absence of MG132 treatment, Ub showed a diffuse staining pattern in the nucleus and cytoplasm (Figure 3.11 E). After the addition of MG132, some of the Ub can be found at aggresomes positive for KCMF1 (Figure 3.11 E). Using the MitoTracker Red CMXRox dye (Invitrogen), fluorescence microscopy also revealed mitochondrial clustering around KCMF1-perinuclear inclusions, but KCMF1 was not localized to these structures (Figure 3.11 F). Together these data suggest that upon proteasome impairment, KCMF1 has an intrinsic tendency to localize to aggresomes.

Figure 3.11 Proteasomal inhibition induces KCMF1 to form perinuclear aggregates. (A) KCMF1 forms aggregates upon MG132 treatment. 293 T-REx cells stably expressing GFP- KCMF1 were treated with 10µM MG132 for 12h. The cells were then visualized using live-cell confocal microscopy. DNA was stained with Hoechst (blue). (B) Bar diagram show percent cells, expressing GFP-KCMF1, developing aggresome in the absence of MG132 and with MG132. The values represent the mean average of three independent experiments, with a minimum of 200 cells scored each time from five random fields. Values represent means ±S.E. A difference in the number of cells having KCMF1 aggregates between cells treated with MG132 and cells with no treatment was calculated for statistical significance by paired Student’s t-test. * denotes P-value less than 0.05. (C) Endogenous KCMF1 showed to have identical pattern as GFP-KCMF1 upon MG132 treatment. Immunofluroscence staining using an anti-KCMF1 antibody reveals identical localization pattern in 293 T-REx cells. DNA was stained with DAPI (blue). (D) KCMF1 was colocalized with GFP-CFTR-F508 aggregates. 293 T-REx cells stably expressing Cherry-KCMF1 were transfected with GFP-CFTR-F508. 24h post transfection, the cells were treated with were transfected with 10µM MG132 for 12h. The cells were then imaged using live-cell confocal microscopy. DNA was stained using Hoechst (blue). (E) KCMF1 colocalizes with ubiquitin upon MG132 treatment. 293 T-REx cells stably expressing GFP-

KCMF1 were treated with 10µM MG132 for 12h, before fixing and staining with anti-GFP antibody and anti-Ubiquitin antibody. DNA was stained with DAPI (blue). (F) Mitochondria show clustering around KCMF1-positive aggregates. 293 T-REx cells were cultured and stained with mitoTracker Red CMXRos dye and anti-KCMF1 antibody. Cells were either treated with DMSO (control) or 10µM MG132 for 12h. DNA was stained with DAPI (blue). (G) RAD6A proteins do not localize to KCMF1-positive aggregates. 293 T-REx cells stably expressing GFP- RAD6A were transfected with Cherry-KCMF1. 24h post transfection, the cells were treated with 10µM MG132 for 12h, and then imaged using live cell confocal microscopy. DNA was stained with Hoechst (blue). Scale bars: 10 µm.

As reported for malin, a Ub E3 ligase associated with Lafora disease, its E2 partner, UbcH5 (UBE2D1), is also found at malin-positive aggresomes (Mittal et al., 2007). Therefore, I also checked for the localization of RAD6A at KCMF1-positive aggresomes. However, following proteasomal inhibition, GFP-RAD6A was not detected at aggresomes (Figure 3.11 G). I have also begun to characterize the localization of KCMF1 interactors at aggresomes. Using live-cell imaging, C-terminal GFP-tagged ABHD10 shows localization to the mitochondria (Figure 3.12 A). Upon MG132 addition, GFP-ABHD10 proteins, like the mitochondria, are clustered around the nucleus (Figure 3.12 B). The colocalization between ABHD10 and KCMF1 remain to be determined.

Figure 3.12 ABHD10 localizes to mitochondria. (A) HEK293 cells were transfected with C-terminal GFP-ABHD10 construct. 48h post transfection, (B) HEK293 cells transfected with C-terminal GFP-ABHD10 construct were cultured for 36h before adding 10µM MG132 for 12h. Cells were then stained for mitochondria using mitoTracker red CMXRos dye, before imaging using live-cell confocal microscopy. DNA was stained with Hoechst (blue). Upon MG132 treatment, like the mitochondria, ABHD10 shows clustering centrosomally. Scale bar: 10 µm.

3.3.11 Loss of RAD6A in cells results in a lower number of KCMF1- positive aggresomes

The R11Q mutation in the RAD6A gene was found in XLMR patients, and disrupts the interaction between RAD6 and KCMF1 (Table 3.2). Since aggresome formation is linked to many neurological disorders, I wished to examine the role of RAD6A in aggresome formation. To this end, I generated stable cell lines expressing pGIPZ shRNA (Open biosystems) directed against RAD6A. The level of knockdown in mRNA expression was determined by qRT-PCR. For RAD6A, I detected a 85% knockdown level in mRNA expression(Figure 3.13 A). In the absence or presence of MG132 treatment, I next determined the number of KCMF1-positive aggregates in five separate microscopic fields (~250 cells). As a negative control, I used cells transfected with scrambled pGIPZ shRNA. When RAD6A was knocked down, I observed a significant decrease in the number of KCMF1-positive aggresomes (Figure 3.13 B). The loss of RAD6A thus appears to prevent KCMF1 from localizing to the aggresome.

Figure 3.13 Loss of RAD6A expression leads to decreasing amount of KCMF1-positive aggregates.

(A)shRNA-mediated silencing of RAD6A was assessed using a ΔΔCq method to determine relative gene expression from qPCR data with GAPDH as an endogenous reference gene. In comparison to cells transfected with non-Targeting shRNA, there was a 85% knockdown of RAD6A mRNA level in cells transfected with RAD6A shRNA. (B) Bar diagram showing percentage of cells with cherry-KCMF1-containing aggresomes in the presence or absence of MG132, and/or following treatment with a Rad6A shRNA.Values are the mean of three independent experiments (±S.E), with a minimum of 250 cells scored each time from five random fields. The difference in the number of cells displaying KCMF1 aggregateswas calculated between cells transfected with scrambled shRNA and RAD6A-directed shRNA, using a paired Student’s t-test. * denotesp-value less than 0.05.

3.4 Discussion

3.4.1 A novel RAD6 associated E3 ligase, KCMF1

In this chapter, I show the first RAD6 interactome in human cells. In addition to identifying all of the known E3 ligases associated with RAD6, I was able to identify several additional high- confidence interactors, including the E3 ligase KCMF1. Interestingly, my AP-MS results show that KCMF1 can also interact with all the newly identified interactors (e.g. UBR4, ABHD10, NIPSNAP3A, SSBP1), and seems to bridge them to RAD6. This suggests a novel RAD6 containing complex. Many of these proteins are mitochondrial and/or membrane-associated, and therefore propose new functions for RAD6. This is consistent with the recent reconstruction of the Ub network in yeast (by integrating genetic and protein-protein interaction data obtained from public databases and specific case-studies in the literature on the Ub-system) which identified Rad6 as a major hub, with many Golgi-, vesicle-, or endosome-associated roles that are not explained by currently known targets of Rad6 (Venancio et al., 2009). Moreover, a

XLMR-associated RAD6A mutation disrupts the interaction between RAD6A and the newly identified interactors. The functional importance of the novel RAD6 complex(s) may also be important in neuronal physiology.

Similar to the other RAD6 associated E3 ligases, KCMF1 is shown to interact with RAD6 through a region that is located outside of its RING domain. I show that KCMF1 binds to RAD6A through a region that is C-terminal to its RING domain (termed CT in this thesis). This is similar to RAD18, which also interacts with RAD6 through a C-terminal motif (R6BD) (Hibbert et al., 2011). Moreover, the structure for R6BD and CT appear to be similar. As shown through X-ray crystallography, R6BD forms a helix-loop-helix (Hibbert et al., 2011). Using a structural prediction program, the secondary structure for KCMF1 CT is predicted to also be a helix-loop-helix motif. However, unlike R6BD and the Ubr1/2-BRR, I show that the KCMF1 CT binds to the Rad6 N-terminal α1 helix (including the R11 residue) and residues around the active site Cys88. This reveals a new E2-E3 interaction interface for RAD6. Notably, the KCMF1CT binds to a region on RAD6A that is commonly used to interact with the RING domain of E3 ligases. The N-terminal α1 of RAD6 is one of the three regions that are part of the canonical E2- E3 RING binding interface (Deshaies and Joazeiro, 2009; Dye and Schulman, 2007). R11 is also one of the residues directly at the canonical E2-E3 RING interface (Huang et al., 2011). This poses an interesting question as to why the XLMR-associated point mutation R11Q specifically disrupts the interaction between RAD6 and KCMF1, but none of the other RING E3 ligases. It is likely that since the other RING E3 ligases also bind to a separate region of RAD6 with higher affinity, the R11Q mutation is not enough to disrupt their interactions with RAD6. Even though the RAD6A R11Q protein can still bind to most of its E3 ligases, it is unclear whether the mutation has any effects on ubiquitylation of substrates or downstream functions. The RING domain is often required for ubiquitylation by activating the E2 enzyme (Deshaies and Joazeiro, 2009). For the RAD18 RING domain, the R11 residue is one of the important residues for RAD6 binding (Huang et al., 2011). If the R11Q mutation can disrupt the RAD6-KCMF1 interaction, it could also affect RING domain binding, and therefore affect downstream biological functions. In summary, my results demonstrate a novel RAD6-containing complex involving KCMF1. KCMF1 binds to a different region on RAD6 than the other E3 ligases, and a disease-specific point mutation in RAD6 specifically disrupts the RAD6-KCMF1 interaction.

3.4.2 Functional relationships between RAD6 and KCMF1

RAD6 is an abundant E2 enzyme, present in micromolar concentrations in vivo(Siepmann et al., 2003). E3 enzymes could activate or inhibit this activity toward specific targets. A role of RAD6-mediated Ub chain formation in the Rad18 and RNF20/40 pathway is unlikely since both monoubiquitylate their targets (Kim et al., 2009; Parker and Ulrich, 2009). On the other hand, the Rad6-Ubr1 complex polyubiquitylates substrates to be degraded by the N-end rule pathway (Turner et al., 2000). I show that UBR2 can also activate RAD6’s activity toward polyubiquitylation. In contrast to N-recognins, KCMF1 acts like RAD18 by inhibiting the intrinsic chain building activity of RAD6A. This points to the possibility of substrate monoubiquitylation. As shown in this chapter, many of KCMF1’s interactors are membrane associated and have roles in protein trafficking (e.g. NIPSNAP3A). This is compatible with one of the downstream effects of monoubiquitylation (Hicke, 2001). Mono- or multi- monoubiquitylation plays a role in vesicular trafficking, endocytosis and membrane-receptor recycling (Mosesson and Yarden, 2006; Saksena et al., 2007).

Rad18 promotes monoubiquitylation by competing for access to the noncovalent Ub binding site on the backside of Rad6 with the acceptor Ub (Hibbert et al., 2011). KCMF1 could also promote monoubiquitylation through catalytic interference. I show that KCMF1 CT binds to residues around the catalytic cysteine, mainly involving residues on helices. These residues overlap with the donor Ub binding site on Rad6 (Figure 3.14; Hibbert et al., 2011). NMR spectroscopy has also indicated that both α2 and 310 helices are involved in E2-Ub thioester formation (Hamilton et al., 2001). Moreover, an allosteric inhibitor of hCdc34 (CC0651) produced wholesale shifts in the α2 and 310 helices, leading to secondary conformational changes in the active site region (Ceccarelli et al., 2011). CC0651 was shown to promote monoUb or di-Ub products. Therefore unlike Rad18, KCMF1 could be competing with the donor Ub rather than the acceptor Ub.

Figure 3.14 A comparison of the RAD6 binding sites for ubiquitin and KCMF1. (A) Rad6 binding residues for interacting with ubiquitin. Residues that showed significant chemical shift when 15N-Rad6 was titrated with Ub are shown in blue and red (Hibbert et al., 2011). Active Cys88 is colored green. (B) RAD6 binding residues for KCMF1 CT interaction. Red indicates residues showing significant chemical shifts, blue indicates residues showing decreasing intensity, and green indicates the active Cys88. KCMF1’s binding site overlaps with Ub-Rad6 binding site.

I also show that the point mutation R11Q leads to a loss of RAD6A polyubiquitylation activity. Another XLMR-associated mutation, G23R, has also been reported to result in reduced chain- formation activity, by disrupting the backside noncovalent Ub binding (Hibbert et al., 2011). It is thus possible that all of these XLMR patients develop the associated symptoms because of impairment in the activity of RAD6.

I demonstrate here that proteasomal inhibition induces peri-centrosomal accumulation of KCMF1. Cytoplasmic protein inclusions are found in many neurodegenerative diseases and are formed by retrograde transport on microtubules to the Microtubule Organizing Center (MTOC) (Garcia-Mata et al., 2002; Kopito, 2000). KCMF1 localizes to aggresomes, along with of ubiquitin. I also show preliminary data suggesting that KCMF1 localizes to lysosomes and ER. Lysosomes accumulate around the periphery of the aggresome, playing a role in the clearance of aggresomes (Fortun et al., 2003; Garcia-Mata et al., 2002) through the aggresome-autophagy pathway. Autophagy is an evolutionarily conserved catabolic process in which cytosolic constituents, including organelles (including mitochondria) and macromolecules, are sequestered

80 into a double-membrane structure and delivered to the endosome and lysosome (Tooze and Yoshimori, 2010). This degradative pathway is essential for neuron survival (Lee et al., 2010). Emerging evidence has suggested that autophagosomes are generated from pre-exiting membrane compartments, including the ER and the Golgi (reviewed in Tooze and Yoshimori, 2010). Since KCMF1 may localize to the ER, and interacts with a Microtubule/ER-associated protein, UBR4, KCMF1 may be important in clearing aggregated proteins or organelles. Furthermore, although KCMF1 does not localize to the mitochondria, it can interact with mitochondrial proteins. It is possible that KCMF1 interacts with these mitochondrial proteins through the aggresome-autophagy pathway, which is required to discard damaged mitochondria (Eskelinen and Saftig, 2009). As shown for Parkinson’s disease, autophagy of mitochondria is implicated in the disease’s pathogenesis (Geisler et al., 2010).

It is interesting to note that KCMF1 was recently shown to interact with brain cytosolic and synaptic 26S proteasome in rat cortex (Tai et al., 2010). UBR4 and SSBP1 (both interactors of KCMF1 and RAD6) were also identified as interacting partners of the proteasome. Although KCMF1’s function with the proteasome is still unclear, the other E3 ligases associated with the brain proteasome, UBE3A and HUWEI, have been linked to mental retardation in humans. UBE3A mutations cause Angelman syndrome (a neuro-genetic disorder), while HUWE1 mutations are found in XLMR (Froyen et al., 2008; Lalande and Calciano, 2007). It thus seems that proteasome-associated E3 ligases are highly linked to neurological diseases.

The biological significance of the KCMF1-RAD6 complex is still unclear. The identification of KCMF1 as a novel RAD6-associated E3 ligase opens up the possibility that RAD6 indeed has vesicle-associated roles. Additional experiments are needed to investigate this. KCMF1 also seems to have aggresome-associated role. A better understanding of this function, as well as the importance of RAD6 activity in aggresome clearance or formation could eventually help to provide an explanation for the link between RAD6A and XLMR.

3.5 Future Directions

To further characterize the RAD6-KCMF1 complex, it is important to assay for any direct interactions between ABHD10, NIPSNAP3A and UBR4 to KCMF1. The possibility of a direct interaction between RAD6 and UBR4 should also be characterized. This could lead to the

81 identification of a possible E2 for UBR4. NMR spectroscopy or in vitro binding assays can be used to examine whether the R11Q point mutation affects RING domain binding.

Since KCMF1 inhibits the chain-building activity of RAD6A, KCMF1 may be able to bind to Ub. To determine whether KCMF1 can bind to Ub, NMR spectroscopy can be used to examine the formation of KCMF1-Ub complex. This will give additional insights to the mechanism in which KCMF1 inhibits RAD6A polyubiquitylation activity.

The colocalization between KCMF1 and all of its interactors (ABHD10, NIPSNAP3A, etc.) also remains to be characterized. I have begun to characterize their localization by using N-terminal tagged fusion proteins. Notably, however, in some cases this proved to disrupt their mitochondrial localization. C-terminal tagged fusion proteins have now been generated. The biological significance of the subcellular localization of KCMF1 is still unclear. One of the areas that could be further examined is KCMF1’s relationship with the aggresome-autophagy pathway (and/or mitophagy). It is also important to look into the significance of RAD6-KCMF1 interaction in this pathway. The localization of RAD6 at ER and lysosomes should also be examined.

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Supplementary Information

Supplementary Figure 1. The anti-KCMF1 antibody is specific to KCMF1 proteins. 293 T-REx cells stably expressing FLAG-tagged KCMF1 werecultured and stained using anti- FLAG antibody (immunofluorescence). Cells were also stained with anti-KCMF1 antibody. DNA was stained with DAPI. As determined by the two antibodies, the localization pattern of KCMF1 proteins is identical. Scale bar 10µm.

Supplementary Table 1. Raw data showing aggresome-counting. 293 T-Rex cells expressing cherry-KCMF1 were transfected with either Non-Targeting shRNA control or RAD6A-directed shRNA. Aggresome formation was detected as a single perinuclear KCMF1-positive inclusion and was confirmed by the indented shape of the nucleus. For each cell condition, ~250 cells were counted from randomly selected fields were analyzed with a fluorescence microscope (Olympus Fluoview). Paired t-test was used to evaluate the significance level of the difference in % of cells with phenotype between control and RAD6A K.D. cells.