Identification of Novel Substrates of the Ubiquitin E3 Ligase RNF126 and Characterization of Its Role in Lipid Droplet Homeostasis

Identification of novel substrates of the ubiquitin E3 ligase RNF126 and characterization of its role in lipid droplet homeostasis

Zhongda Pan

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

Identification of novel substrates of the ubiquitin E3 ligase RNF126 and characterization of its role in lipid droplet homeostasis

Zhongda Pan

Master of Science

Department of Medical Biophysics University of Toronto

2016 Abstract

Ubiquitin E3 ligases confer specificity by recognizing target substrates and mediating the final step of the ubiquitination process. Despite this critical role, our understanding of their physiological partners and biological functions remains limited. Here we use a proximity-based protein screen called BioID to explore novel interacting proteins and substrates of the ubiquitin

E3 ligase RNF126. We demonstrate that RNF126 can associate with p97 as well as its co-factors

UBXD1 and UBXD8 and show that UBXD1 and UBXD8 are bonafide substrates of RNF126.

To determine a functional role, we explored the role of RNF126 in UBXD8 mediated functions.

We found that stable knockdown of RNF126 in HeLa cells results in smaller LDs following oleate stimulation and that this is due to defective expansion. Together, this work demonstrates the successful use of BioID to identify substrates of ubiquitin E3 ligases and furthers our understanding of the role of ubiquitin in LD biology.

Acknowledgments

I would like to thank my supervisor, Dr. Jane McGlade, for her outstanding support and mentorship throughout the years. Her patience and guidance came second only to her willingness to place my success as her number one priority. I would also like to thank my committee members, Dr. Brian Raught and Dr. Peter Kim, for their time, guidance, and support.

I would like to thank everyone in the McGlade lab for making this time an enjoyable journey through all the ups and downs of graduate school. I have not only learned a lot from everyone, but also truly appreciate the support when things turned sour. I wish you all the best.

Finally, I would like to thank my parents for sticking through this with me and listening to my over-detailed explanations about my project. I would also like to thank my best friend Annabelle Sumenap for keeping me afloat during difficult times.

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Table of Contents

Acknowledgments...... iii

Table of Contents ...... iv

List of Tables ...... vii

List of Figures ...... viii

List of Appendices ...... ix

Chapter 1 Introduction ...... 1

1.1 The ubiquitin regulatory system ...... 1

1.1.1 RING type ubiquitin E3 ligases ...... 3

1.1.2 The ubiquitin code ...... 4

1.1.3 RNF126 and RNF115 are two related ubiquitin E3 ligases ...... 5

1.2 The AAA ATPase p97 ...... 7

1.2.1 Biochemical properties of p97 ...... 7

1.2.2 Biological functions of p97 ...... 8

1.2.3 Control of p97 function by co-factors ...... 8

1.2.4 The UBX family is the largest family of p97 co-factors ...... 10

1.2.5 UBXD8 is an integral membrane protein ...... 10

1.3 Lipid droplets are dynamic cellular organelles ...... 13

1.3.1 Lipid droplet life cycle ...... 15

1.3.2 Regulation of lipid droplets by ubiquitin ...... 16

1.3.3 The role of UBXD8 and p97 in the regulation of lipid droplets ...... 17

1.4 Thesis rationale and objectives ...... 20

Chapter 2 Identifying interacting proteins and substrates of RNF126 and RNF115 ...... 21

2.1 Introduction ...... 21

2.2 Materials and Methods ...... 21

2.2.1 Cell culture, cDNA constructs, and reagents ...... 21

2.2.2 Forward and reverse transfection ...... 24

2.2.3 Generation of doxycycline inducible stable cell lines and BioID screen ...... 24

2.2.4 Production of GST fusion proteins ...... 25

2.2.5 Co-immunoprecipitations, GST pull-downs, in vivo ubiquitination assays, and western blotting ...... 25

2.3 Results ...... 26

2.3.1 Identification of proteins in near proximity to RNF126 and RNF115 by BioID...26

2.3.2 Validation of potential interacting proteins identified by BioID ...... 34

2.3.3 RNF126 associates with p97 and the UBX family of co-factors ...... 36

2.3.4 Over-expression of RNF126 promotes UBXD8 association with p97 ...... 40

2.3.5 UBXD8 and UBXD1 are ubiquitinated by RNF126 ...... 42

2.4 Discussion ...... 42

Chapter 3 RNF126 regulates lipid droplet size following fatty acid stimulation ...... 45

3.1 Introduction ...... 45

3.2 Materials and Methods ...... 45

3.2.1 Cell culture, cDNA constructs, and reagents ...... 45

3.2.2 Preparation of oleate-BSA conjugate...... 46

3.2.3 Generation of doxycycline inducible stable cell lines ...... 46

3.2.4 Immunocytochemistry ...... 46

3.2.5 Quantification of lipid droplets ...... 47

3.2.6 Lipid droplet purification and immunoprecipitation ...... 47

3.2.7 Statistical analysis ...... 48

3.3 Results ...... 48

3.3.1 RNF126 regulates lipid droplet size in oleate stimulated cells ...... 48

3.3.2 Re-expression of RNF126 in an inducible cell system can rescue decreased lipid droplet size ...... 51

3.3.3 RNF126 does not regulate lipid droplet turnover ...... 55

3.3.4 RNF126 is required for lipid droplet expansion ...... 58

3.4 Discussion ...... 60

Chapter 4 Conclusions and Future Directions ...... 62

4.1 Summary ...... 62

4.2 Regulation of p97 co-factors by RNF126 ...... 62

4.3 The role of RNF126 mediated ubiquitination of KEAP1 and CUL3 ...... 67

4.4 UBXD1, p97, and RNF126 in EGFR trafficking ...... 71

4.5 The role of RNF126 mediated ubiquitination of UBXD8 in lipid droplet regulation ...... 72

4.6 RNF126 and other lipid droplet regulatory proteins ...... 73

4.7 RNF126 regulation of lipid droplets beyond HeLa cells ...... 77

4.8 Concluding remarks ...... 78

Chapter 5 References ...... 79

List of Tables

Table 2-1 List of primers utilized for cloning 22

Table 2-2 List of cDNA constructs used 23

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List of Figures

Figure 1-1 The ubiquitin system 2

Figure 1-2 RNF126 and RNF115 are two related RING 6

type ubiquitin E3 ligases

Figure 1-3 The AAA ATPase p97 and its diverse co-factor family 12

Figure 1-4 The lipid droplet life cycle 14

Figure 1-5 The role of UBXD8 and p97 in lipid metabolism 19

Figure 2-1 The BioID method 27

Figure 2-2 Generation of stable cell lines for BioID 30

Figure 2-3 BioID proteome of RNF126 and RNF115 32

Figure 2-4 Validation of BioID candidates by co-immunoprecipitation 35

Figure 2-5 RNF126 associates with p97 and members of the UBX family 38

Figure 2-6 RNF126 ubiquitinates UBXD8 and UBXD1 41

Figure 3-1 Knockdown of RNF126 reduces lipid droplet size 49

Figure 3-2 siRNA knockdown of RNF126 decreases lipid droplet

size following oleate stimulation 53

Figure 3-3 Re-expression of mRNF126 in an inducible cell

model rescues lipid droplet size 54

Figure 3-4 Knockdown of RNF126 does not accelerate

lipid droplet turnover 56

Figure 3-5 RNF126 is required for lipid droplet expansion 59

Figure 4-1 Characterizing the interaction between RNF126

and the UBX family 65

Figure 4-2 RNF126 may associate and ubiquitinate KEAP1 and CUL3 69

Figure 4-3 RNF126 may ubiquitinate DGAT2 to enhance stability 75

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List of Appendices

Appendix I SAINT analyzed RNF126 HeLa BioID results 90

Appendix II SAINT analyzed RNF126 HEK293 BioID results 92

Appendix III SAINT analyzed RNF115 HEK293 BioID results 94

Chapter 1 Introduction 1.1 The ubiquitin regulatory system

Ubiquitin is a small, highly conserved protein involved in a wide variety of cellular processes. Like other post-translation modifications, the attachment of ubiquitin can alter the structure, function, and sub-cellular localization of a target protein. Originally discovered to mediate the proteolytic degradation of proteins(Ciehanover et al., 1978), ubiquitin has since been linked to cellular functions ranging from endocytosis to lipid droplet homeostasis. The ubiquitination of proteins is a sequential action requiring an ubiquitin-activating enzyme (E1), ubiquitin- conjugating enzyme (E2), and ubiquitin ligase (E3) (Figure 1-1). Through an ATP-dependent reaction, ubiquitin forms a thioester bond with an E1 enzyme through its C-terminal carboxy group(Hershko and Ciechanover, 1998). Following transfer of ubiquitin from E1 to E2, an E3 ubiquitin ligase determines the substrate and collaborates with the ubiquitin bound E2 to conjugate ubiquitin to the substrate(Hershko and Ciechanover, 1998). This final step creates an isopeptide bond between the ubiquitin carboxy terminal glycine and a lysine on the substrate(Deshaies and Joazeiro, 2009). Outside lysine, ubiquitin can occasionally be conjugated to cysteine, serine, threonine, and N-terminal methionine residues(Coyne and Wing, 2016). Conjugation of ubiquitin onto the target substrate by the E3 ligase can occur through two mechanisms. The RING (really interesting new gene) family catalyzes direct transfer of ubiquitin from the E2 to the substrate while HECT (homology to E6AP C terminus) family E3s undergo a two-step reaction where ubiquitin is transferred from the E2 to an active site cysteine on the E3 and then directly transferred from the E3 onto the substrate(Berndsen and Wolberger, 2014). Although some proteins are modified with a single ubiquitin moiety, poly-ubiquitin chains can be built by conjugating additional ubiquitin molecules onto a lysine of the preceding ubiquitin. The information encoded within these ubiquitin chains can then be interpreted by ubiquitin receptors to determine the protein’s fate.

1 2

Figure 1-1. The ubiquitin system. Ubiquitin (Ub) is activated for transfer by an E1 enzyme through the formation of a thioester bond and then transferred to the active-site cysteine of an E2 enzyme. The ubiquitin bound E2 then interacts with an E3 ubiquitin ligase. The left side depicts a RING type E3 ligase orienting a substrate molecule and E2 to facilitate direct transfer of ubiquitin onto the substrate. RBR and HECT type E3 ligases function by first forming a thioester bond with ubiquitin supplied by the E2. Ubiquitin is then directly transferred onto the substrate. This process can be repeated to generate ubiquitin chains. Adapted from (Woelk et al., 2007).

1.1.1 RING type ubiquitin E3 ligases

Ubiquitin E3 ligases play an essential role by determining the specificity of the ubiquitin pathway. There are three major families of E3 ligases including the RING family, HECT family, and RING-between-RING (RBR) family(Berndsen and Wolberger, 2014). In humans, there are two E1 enzymes, about 37 E2 enzymes, and over 600 E3 ligases with RING type E3s being the largest family(Deshaies and Joazeiro, 2009; Michelle et al., 2009). The large number of E3s allow for specific targeting of a broad array of substrates, encompassing nearly every biological pathway. Despite this, understanding the mechanism of action and biological function of many RING type E3s remains challenging.

Unlike HECT and RBR type E3s, RING type E3 ligases do not directly bind and transfer ubiquitin onto a target substrate(Berndsen and Wolberger, 2014). Instead, RING type E3s serve as a scaffold, binding both substrate and E2 enzyme in an orientation that facilitates the direct transfer of ubiquitin from E2 to substrate. Recruitment of E2s involves recognition of the conserved E2 UBC (ubiquitin-conjugating) domain by the RING domain of an E3 ligase(Metzger et al., 2014). Although variations exist, the canonical RING domain (C3HC4) contains eight Cys and His residues that maintain the RING structure by coordinating two zinc ions(Metzger et al., 2014). Since E2 enzymes need to shuttle between E1 and E3, the E2-E3 interaction is usually weak with dissociation constants in the micromolar range(Yin et al., 2009). This interaction can be supported by surrounding residues flanking the RING domain, possibly dictating E2-E3 specificity(Zheng et al., 2000). It is thought that by holding the ubiquitin bound E2 in a specific conformation, RING type E3s are able to accelerate the transfer of ubiquitin onto a substrate lysine(Berndsen and Wolberger, 2014). As E2 enzymes are primarily responsible for dictating the ubiquitin topology assembled on a target substrate, it is not surprising to observe a single RING being able to recruit multiple different E2s to facilitate assembly of different linkages(Deshaies and Joazeiro, 2009).

In addition to the RING domain, RING type E3s contain other substrate recruiting domains and may form homodimers or heterodimers by interacting with other RING domains. While RING homodimers have the ability to interact with E2s from both RING domains, some heterodimers lack this ability. Instead, these RING heterodimers involve other functions such as substrate selection, regulation of activity, or stabilization of the E3 complex(Metzger et al., 2014). For

4 example, the BRCA1-BARD1 heterodimer interacts with the E2 through the BRCA1 RING while recruitment of BARD1 stabilizes the complex(Brzovic et al., 2001; Joukov et al., 2001).

1.1.2 The ubiquitin code

The multi-functional characteristic of ubiquitin can be attributed to its ability to communicate through distinct ubiquitin chain topologies. Proteins can be modified by a single ubiquitin (monoubiquitination) or by a single ubiquitin on multiple lysines (multi-monoubiquitination). Ubiquitin itself contains seven lysine residues (K6, K11, K27, K29, K33, K48, and K63) and a free amino group and these residues can serve as acceptors for additional ubiquitin molecules, creating poly-ubiquitin chains. Although there can be significant cell line differences, the majority of conjugated ubiquitin in HEK293 cells are monoubiquitin chains while ~8.5% exist as poly-ubiquitin chains(Kaiser et al., 2011). Of the conjugated poly-ubiquitin chains, K48 chains are the most abundant followed by K63 chains(Kaiser et al., 2011). K48 linkages are classically known for their role in mediating proteasomal degradation while mono-ubiquitin and K63 poly-ubiquitin chains play non-proteolytic roles such as receptor endocytosis(Haglund and Dikic, 2012). The functions of poly-ubiquitin linkages outside of K48 and K63 chains are still poorly understood. With the exception of K63 chains, proteasomal inhibition increases the abundance of all other linkages suggesting a proteolytic role(Xu et al., 2009). Indeed, atypical K11 chains are recognized by proteasomal receptors and mediate proteasomal degradation(Jin et al., 2008; Komander and Rape, 2012). In addition to regulating protein turnover and sub- cellular localization, ubiquitination can also regulate protein-protein interactions and protein activity. For example, K63 chains can act as a scaffold to recruit a series of E3 ligases during DNA damage(Al-Hakim et al., 2010). Recently, linear ubiquitin chains were demonstrated to be formed by the E3 ligase complex LUBAC through the first methionine ubiquitin residue(Ikeda et al., 2011). These chains are required for the activation of the NF-kB pathway.(Ikeda et al., 2011).

Ubiquitination is a dynamic and reversible process. Cleavage of the isopeptide bond between ubiquitin and substrate is catalyzed by de-ubiquitinating enzymes (DUB). There are approximately 90 DUBs in the human genome and they can be classified into five families based on the identity of their catalytic domains(Hutchins et al., 2013). These include the ubiquitin C-terminal hydrolase (UCH), ubiquitin-specific protease (USP), ovarian tumor

5 domain (OTU), Machado-Joseph disease (MJD), and Jab1/Mpn/Mov34 (JAMM) enzymes(Coyne and Wing, 2016). The cellular functions of DUBs can be broadly classified into two categories(Komander et al., 2009). As de novo synthesis of ubiquitin occurs as a fusion protein, the first function is to maintain a pool of free ubiquitin through cleavage by DUBs. Removal of ubiquitin from proteins directed to destruction by the proteasome or lysosome also contributes to the maintenance of a free ubiquitin pool. The second function involves control of ubiquitin-mediated signaling events. As ubiquitin chains target proteins for specific destinations, removal of these chains by DUBs can redirect their fate. For example, ubiquitination of KEAP1 promotes dis-association with its scaffold CUL3 while deubiquitination by USP15 promotes complex formation between KEAP1 and CUL3(Villeneuve et al., 2013).

1.1.3 RNF126 and RNF115 are two related ubiquitin E3 ligases

RNF126 and RNF115 are two closely related RING type ubiquitin E3 ligases containing a N- terminal zinc finger and a C-terminal RING domain (Figure 1-2)(Smith et al., 2013). The RING domain of both proteins is necessary for ubiquitin ligase activity while the zinc finger from both proteins can bind K48 and K63 ubiquitin chains(Amemiya et al., 2008; Smith et al., 2013; Zhi et al., 2013). Consistent with the role of E3 ligases mediating protein degradation, RNF126 and RNF115 have been found to ubiquitinate the cell cycle checkpoint protein p21 and target it for degradation as well as ubiquitinate the EGFR to target it for lysosomal degradation(Smith et al., 2013; Wang et al., 2013; Zhi et al., 2013). Although both RNF126 and RNF115 promote proliferation of a breast cancer cell line in culture, only RNF115 over-expression has been shown to be associated with increased invasiveness and progression of breast cancer as well as renal oncocytoma(Burger et al., 2005; Ehsani et al., 2013; Zhi et al., 2013). On the other hand, low RNF126 expression has been observed in a variety of human tumours(Wang et al., 2015). On a cellular scale, the two E3s have also been found to perform non-overlapping roles. RNF126, but not RNF115, was found to regulate the retrograde sorting of CI-MPR as well as participate with BAG6 to mediate the degradation of mis-localized hydrophobic proteins(Rodrigo-Brenni et al., 2014; Smith and McGlade, 2014). Recently, RNF126 has also been shown promote homologous recombination independent of its E3 ligase activity(Wang et al., 2015). The complexity of the two E3 ligases indicates that further understanding of the functions and regulatory mechanisms is required.

Figure 1-2. RNF126 and RNF115 are two related RING type ubiquitin E3 ligases. The domain architecture of human RNF126 and RNF115 are shown. Both proteins have a N- terminal ubiquitin binding zinc finger (Znf) and a C-terminal E2-recruiting RING finger domain. The amino acid sequence similarity between regions of the two proteins is indicated.

1.2 The AAA ATPase p97

Valosin-containing protein (VCP) or p97 is a hexameric AAA ATPase (ATPase associated with diverse cellular activities) that utilizes the energy of ATP hydrolysis to structurally remodel substrate proteins(Baek et al., 2013). This mechanism results in a physical change to the protein’s structure, disrupting protein-protein, protein-membrane, and even protein-DNA interactions (Figure 1-3A). p97 was initially discovered in yeast mutants defective in cell cycle progression(Moir et al., 1982). Today, p97 is known to be involved in a myriad of independent cellular processes and accounts for 1% of cytosolic protein in eukaryotic cells(Baek et al., 2013). The vast majority of p97 functions are intimately linked with the ubiquitin system. While ubiquitin can regulate the fate of a protein, p97 is able to regulate the ubiquitination of proteins as well as respond to the signals encoded within ubiquitin chains. This introduces an additional level of regulation and plasticity to ubiquitin mediated processes(Meyer et al., 2012). The sheer abundance of p97 hints at its greater role in cellular physiology, although it exhibits very little substrate specificity on its own. To maintain functional specificity, p97 relies on a plethora of co-factors that help to recruit substrates, regulate sub-cellular localization, and control the oligomeric state of p97(Buchberger et al., 2015a). Although research on p97 has greatly accelerated, the full extent of p97 mediated cell biology as well as how functional specificity is achieved is only beginning to be understood.

1.2.1 Biochemical properties of p97 p97 contains two ATPase domains, D1 and D2, as well as an N-domain and a C-terminal tail (Figure 1-3B). Structural work has shown that p97 exists as a homohexamer with a central core created by two separate hexameric rings formed through the D1 and D2 domains(Huyton et al., 2003). ATP hydrolysis appears to occur mainly in the D2 domain while binding of ATP to the D1 domain promotes hexamerization of p97 and is required for its biological function(Song et al., 2003; Wang et al., 2003). It is speculated that the ability for p97 to remodel proteins involves major structural changes to generate a pulling force. Recent advances in cryo-electron microscopy have revealed dramatic conformational changes that occur between the N, D1, and D2 domains due to binding of ATP(Banerjee et al., 2016). However, the precise mechanism of how these changes in conformation contribute to protein degradation or protein segregation is unknown.

1.2.2 Biological functions of p97

The best characterized p97 function is its role in endoplasmic reticulum associated degradation (ERAD). This pathway acts as a surveillance of ER proteins, ensuring only correctly folded proteins reach their final destinations. As misfolded proteins are transported to the ER membrane, they are recognized by resident ER ubiquitin E3 ligases and polyubiquitinated(Wolf and Stolz, 2012). Although the E3 ligase responsible for ubiquitination can vary, the pathways merge when p97 is recruited to the ER membrane by its co-factor Ubx2 (Human functional equivalents: UBXD2 and UBXD8)(Neuber et al., 2005; Wolf and Stolz, 2012). Here, p97 recognizes ubiquitinated misfolded proteins through its co-factor heterodimer UFD1-NPL4 and extracts them from the ER membrane through hydrolysis of ATP(Wolf and Stolz, 2012). Extracted ubiquitinated substrates are then handed off to proteasome shuttling factors for delivery to the proteasome(Richly et al., 2005). Interestingly, this mechanism has now emerged as a unifying theme for p97 and similar mechanisms can be found in the degradation of mitochondria membrane proteins and chromatin associated proteins (Figure 1- 3A)(Meyer et al., 2012). Outside its role in the ubiquitin-proteasome system, p97 has also been implicated in membrane trafficking, hypoxia response, and autophagy(Bug and Meyer, 2012). Given its broad cellular role, p97 is intimately linked with several human diseases ranging from cancer to degenerative diseases(Baek et al., 2013).

1.2.3 Control of p97 function by co-factors

The diversity and number of p97 co-factors is unique among the AAA ATPases(Buchberger et al., 2015b). While there are more than 40 known p97 co-factors, many of these co-factors are not essential for cell survival and growth. Functionally, p97 co-factors can be divided into two groups, substrate-recruiting factors and substrate processing factors (Figure 1-3B). For example, ubiquitinated HIF1α is recruited to p97 through the ubiquitin binding domain of UBXD7 while recruitment of the de-ubiquitinating enzyme VCPIP1 to p97 is required for reassembly of the golgi body following mitosis(Alexandru et al., 2008; Uchiyama et al., 2002). It has been suggested that p97 employs a distinct set of co-factors for each function(Meyer et al., 2012). How specificity is achieved during the assembly of functional p97 complexes is an area of intensive research. Currently, ordered cofactor binding has been shown to be regulated by a variety of mechanisms including competitive binding, hierarchical recruitment of co-factors, and post-translation modification of p97. A brief overview is given below.

To date, all known p97 co-factors bind to the N-domain or the C-tail (Figure 1-3B). Almost all N-domain interaction motifs including the VIM and VBM motifs as well as the UBX and UBXL domains bind to overlapping sites at the hydrophobic cleft between adjacent N- domains(Buchberger et al., 2015b). Thus, each unit of a p97 hexamer can only interact with one of these motifs at any given time, limiting the complexity of co-factor binding at the N-domain. Although competitive binding between co-factors has been shown experimentally in vitro, the outcome in living cells is likely to be much more complex and remains to be understood. Additionally, competitive binding between C-tail proteins has yet to be analyzed.

Assembly of p97 complexes through co-factor recruitment is a highly ordered process. To date, p97 is known to form mutually exclusive core complexes with p47, UFD1-NPL4, and UBXD1. This has given rise to the terms major and minor p97 co-factors. To understand how exclusivity is achieved, in vitro experiments demonstrated that UFD1-NPL4 can bind p97 in multiple different conformations, obstructing certain co-factors such as p47 through steric hindrance and allowing others to dock onto free p97 N-domains(Bebeacua et al., 2012; Bruderer et al., 2004). Core complexes formed by major co-factors can increase in complexity through the recruitment of additional minor co-factors, providing greater pathway specificity or additional enzymatic activities. Indeed, UBXD7 can only bind to p97UFD1-NPL4 complexes and not to free p97, thus forming a hierarchy of p97 binding partners where binding of certain co-factors precedes others(Hanzelmann et al., 2011). However, the hierarchal binding of FAF1 to p97UFD1-NPL4 was recently challenged when an alternate group found binding of FAF1 and naked p97 to occur in vitro(Ewens et al., 2014). Interestingly, siRNA knockdown of UFD1-NPL4 in HeLa cells abolished co-immunoprecipitation (co-IP) of p97 with FAF1, suggesting that UFD1-NPL4 is required for stable binding of FAF1 to p97, supporting a hierarchical binding model(Lee et al., 2013). This contrast may reflect an in vivo property that does not occur in vitro. The ability of p97p47 and p97UBXD1 to form higher order complexes remains to be explored.

Mammalian p97 was first found to be phosphorylated at tyrosine residues 796 and 805 following T-cell activation(Egerton et al., 1992). Phosphorylation of tyrosine 805 in yeast triggers nuclear import of p97 in late G1 phase while blocking binding of Ufd3 and PNGase (peptide:N-glycanase) to the C-tail of p97(Madeo et al., 1998; Zhao et al., 2007). Thus, tyrosine phosphorylation may be a conserved mechanism serving to regulate C-tail co-factor binding. In addition, acetylation of p97 has been reported to contribute to nuclear translocation of

10 p97(Koike et al., 2010). While other post-translational modifications of p97 have been described, the biological significance of these modifications is unknown and awaits investigation.

1.2.4 The UBX family is the largest family of p97 co-factors

The ubiquitin-regulator X (UBX) protein domain family is the largest p97 co-factor family identified (Figure 1-4C)(Meyer et al., 2012). The UBX domain is an ubiquitin-like domain that binds to the N-domain of p97 (Figure 1-4B)(Kloppsteck et al., 2012). Although this family of proteins has yet to be fully characterized biochemically and functionally, it has been shown that almost all 13 mammalian UBX proteins can bind p97, confirming the UBX domain as a general p97 interaction module(Kloppsteck et al., 2012). The sole exception is UBXD1, which lacks a conserved amino acid motif in its UBX domain. Instead, UBXD1 interacts with both N-domain and C-tail of p97 through two independent binding sites separate from its UBX domain(Kern et al., 2009). A sub-set of UBX family proteins also contain a N-terminal UBA domain that acts as a ubiquitin binding domain (Figure 1-3C). These proteins include UBXD7, SAKS1, FAF1, p47, and UBXD8 and are believed to direct p97 to ubiquitin-dependent cellular functions by recruiting ubiquitinated substrates. While non-UBA containing UBX proteins might be involved in ubiquitin-independent functions, UBXD2 is thought to direct p97 to ubiquitin- dependent functions by recruiting the UBA-containing protein ubiquilin to p97 complexes(Lim et al., 2009). Thus, it is likely that non-UBA containing UBX proteins also play important roles in ubiquitin associated pathways. The complexity behind p97 complex assembly is made more apparent when considering the abundance of substrate processing co-factors, such as E3 ligases, that UBX family proteins interact with(Alexandru et al., 2008).

1.2.5 UBXD8 is an integral membrane protein

UBXD8 was first described to be up-regulated in T cells of patients with atopic dermatitis(Imai et al., 2002). Further work has found that UBXD8 is inserted into the ER membrane via a short stretch of hydrophobic amino acids in the form of a hairpin loop(Lee et al., 2010). UBXD8 contains a N-terminal ubiquitin binding UBA domain, a central UAS domain that can interact with unsaturated fatty acids, and a C-terminal UBX domain that binds p97 (Figure 1-3C)(Kim et al., 2013). The best characterized function of UBXD8 is its role in targeting ER membrane proteins for degradation by ERAD(Mueller et al., 2008). Studies of the yeast orthologue UBX2

11 suggest that UBX2 acts to recruit p97UFD1-NPL4 to the ER where the p97 complex then recognizes ubiquitinated substrates and extracts them for degradation(Neuber et al., 2005). Recruitment of p97 appears to be essential as deletion of the UBX domain prevents degradation of at least one ERAD substrate(Phan et al., 2010). Local concentration of UBXD8 may also play an important regulatory role as both knockdown and over-expression of UBXD8 disrupted ERAD(Xia et al., 2014). In a non-proteolytic role, UBXD8 has been shown to cooperate with p97 to destabilize mRNA by removing ubiquitinated RNA binding protein, HuR, from its mRNA targets(Zhou et al., 2013). The final known function of UBXD8 is its role in maintaining lipid homeostasis, which will be described in greater detail below.

Figure 1-3. The AAA ATPase p97 and its diverse co-factor family. A) A general description of the mechanism of action. Following ubiquitination of a substrate-partner bound complex, p97 is recruited to the ubiquitinated substrate by a set of co-factors. The partner can be a protein, lipid, or DNA. Hydrolysis of ATP by p97 provides the energy necessary to physically remodel the substrate, separating it from its partner. The substrate can then be further modified through recruitment of substrate processing factors and recycled, or shuttled to the proteasome for degradation. B) Schematic depicting the domain architecture of p97 and binding of its co- factors. C) Domain architecture of all members of the UBX family. Adapted from (Meyer and Weihl, 2014).

1.3 Lipid droplets are dynamic cellular organelles

Lipids serve as sources of energy, signaling molecules, and membrane building blocks. However, high concentrations of lipids can be extremely damaging to membrane structure and can cause cellular toxicity. To safely maintain a lipid reservoir, cells compartmentalize lipids into cellular organelles called lipid droplets (LD). LDs consist of a phospholipid monolayer enclosing a neutral lipid core consisting mainly of triglycerides (TG) and cholesterol- esters(Thiam et al., 2013). Initially thought to be passive and inert structures, LDs became objects of interest when the first LD associated protein was discovered(Egan et al., 1990). Due to the rapid rise of global obesity, research in understanding the molecular mechanisms regulating the LD lifecycle and the functions of associated proteins has exploded. Now recognized as dynamic cellular organelles with complex biogenesis and functions outside of classic lipid storage, further understanding of LD functions and regulatory mechanisms will be essential for treatment of LD associated diseases such as obesity and its co-morbidities.

Figure 1-4. The lipid droplet life cycle. LD biogenesis occurs at the ER where ACSL enzymes activate FAs through esterification to coenzyme A. Through a series of enzymatic reactions, FA-CoA is incorporated into a glycerol backbone to form TG. TG is then packaged into forming pre-LDs in the ER membrane. As lipid accumulation begins to cause curvature of the ER membrane, the developing LD buds off the ER until it forms an independent structure. LDs can then continue to expand through localized synthesis of TG or mobilize its cargo. LD turnover occurs through two known pathways: enzymatic lipolysis catalyzed by ATGL and lysosomal hydrolysis of LDs in a process called lipophagy.

1.3.1 Lipid droplet life cycle

Biogenesis of cytosolic LDs (hereby referred to as LDs) occurs de novo from accumulation of neutral lipids in the ER. While LDs can accumulate cholesterol-esters, the LD life cycle has been best studied through the TG pathway. At the ER, inert fatty acids (FA) are first activated by esterification with coenzyme A by an ACSL family member (Figure 1-4). Although the precise roles of the various ACSL enzymes in LD biogenesis are unclear, ACSL3 was found to be rapidly recruited to LD assembly sites on the ER periphery, suggesting that neutral lipid synthesis may occur in specific regions(Kassan et al., 2013). These assembly sites, termed pre- LDs, are restricted ER microdomains where neutral lipids are packaged together (Figure 1-4). Activated FAs are then converted into diacylglycerol (DG) through an enzymatic cascade (Figure 1-4). The final step to synthesize triacylglycerol (TG) is catalyzed in mammals by DGAT1 and DGAT2(Harris et al., 2011). As pre-LDs accumulate TG, the structures grow until finally becoming distinct from the ER in a process resembling budding (Figure 1-4)(Thiam et al., 2013). Although this process can occur spontaneously, it is likely that proteins play a role in the development of forming LDs(Hashemi and Goodman, 2015). Mature LDs then move into the center of the cell while likely remaining attached to the ER(Pol et al., 2014). This connection with the ER is thought to facilitate the distribution of lipids and proteins between the two organelles.

To accommodate increased lipid burden, LDs accumulate greater amounts of cargo and increase in size and number. This process is rapid with some LDs being observed to increase in volume 30-fold within hours(Krahmer et al., 2011). LD expansion is thought to occur primarily through two mechanisms: LD fusion to form a larger LD or growth of individual LDs. Fusion of two smaller LDs to generate a larger LD is a rare event under normal conditions but can be induced(Murphy et al., 2010). Although slow, fusion is an essential LD expansion mechanism in white adipocytes and is responsible for the large, unilocular LD observed in the cell type(Sun et al., 2013). However, it is likely that the majority of LD expansion occurs from localized synthesis of TGs at the surface of LDs (Figure 1-4). TG synthesizing enzymes (ACSL3, GPAT4, AGPAT3, DGAT2) migrate together to growing LDs where they continue to synthesize cargo for deposition into LDs(Wilfling et al., 2013). Depletion of DGAT2 or GPAT4 prevented formation of large LDs while over-expression of DGAT2 increased LD size(Wilfling et al., 2013). Interestingly, depletion of the ER localized DGAT1 increased LD size while over-

16 expression of DGAT1 increased the total number of LDs without increasing size(Wilfling et al., 2013). Thus, the two terminal TG enzymes, DGAT1 and DGAT2, are responsible for generation of different LD sub-populations.

Mobilization of LD cargo can occur through the autophagic degradation of LDs in a process termed ‘lipophagy’ or through the enzymatic hydrolysis (lipolysis) of neutral lipids by LD membrane proteins such as ATGL (Figure 1-4)(Thiam et al., 2013). Originally discovered in hepatocytes, lipophagy has been demonstrated to occur in several different cell types although activation is likely context specific(Ward et al., 2016). For example, HeLa cells have been found to have low basal levels of lipophagy while mouse embryonic fibroblasts require serum starvation to activate lipophagy(Rambold et al., 2015; Velikkakath et al., 2012). The alternate pathway, lipolysis, occurs in all cell types and is primarily mediated by the rate limiting enzyme ATGL. Knockout of ATGL in mice results in systemic deposition of TGs and mice die from excessive accumulation of lipid in the heart(Haemmerle et al., 2006). Activation of ATGL is dependent on interaction with its co-activator CGI-58 as lipolysis is inhibited when ATGL and CGI-58 are separated(Wang et al., 2011). The role of the two pathways in LD turnover is unclear. Recently, it was shown in MEFs that enzymatic lipolysis is the major LD turnover mechanism used under carbon starvation conditions (low glucose, no amino acid, no serum) while lipophagy is activated under milder serum starvation conditions where glucose and amino acids aren’t limited(Rambold et al., 2015). How the availability of nutrients dictates activation of LD turnover remains to be addressed.

1.3.2 Regulation of lipid droplets by ubiquitin

Growing evidence has highlighted the importance of ubiquitin in LD homeostasis although mechanistic detail is lacking. PLIN2, a LD associated protein that inhibits lipolysis, has been shown to be ubiquitinated and degraded by the proteasome under lipid depleted conditions and stabilized by addition of FA(Xu et al., 2005). PLIN2 is also ubiquitinated at LDs by the HECT E3 ligase AIP4 and removed from LDs through proteasomal degradation or chaperone-mediated autophagy(Hooper et al., 2010; Kaushik and Cuervo, 2015; Tokunaga et al., 2013). Both over- expression and knockdown of the adapter protein that recruits AIP4 to PLIN2, SPG20, resulted in accumulation of LDs in cells for reasons unknown(Eastman et al., 2009). Spatial organization of LDs can also be regulated as mono-ubiquitination of LD associated AUP1 induced tight

17 clustering of LDs(Lohmann et al., 2013). Ubiquitin binding UBXD8 has also been shown to bind ubiquitinated LD proteins and will be described in further detail below.

1.3.3 The role of UBXD8 and p97 in the regulation of lipid droplets

UBXD8 was first linked to lipid metabolism when a group found that UBXD8 recruits p97 to mediate degradation of INSIG1 by ERAD(Lee et al., 2008). This results in activation of the transcription factor SREBP1 and upregulation of FA synthesis genes(Lee et al., 2008). Interestingly, unsaturated FAs block the interaction between UBXD8 and INSIG1, inhibiting synthesis of additional FAs during lipid rich conditions(Lee et al., 2008). In FA depleted cells, knockdown of UBXD8 inhibited FA synthesis while increasing conversion of DG into TG(Lee et al., 2010). This supports a role where UBXD8, under low cellular FA levels, promotes FA synthesis while simultaneously re-routing FAs to be potentially incorporated into phospholipids by inhibiting their storage in TGs. A potential mechanism to reduce TG synthesis may involve regulation of a DGAT enzyme by UBXD8 under low FA conditions. Interestingly, when Ubx2 deleted yeast were induced to form LDs, TG synthesis was inhibited through mis-localization of the enzyme Lro1, an orthologue of mammalian cholesterol synthesis enzymes(Wang and Lee, 2012). Although UBXD8 can rescue this defect, this function may be unique to yeast as the final TG synthesis step in mammalian cells its catalyzed by DGAT enzymes.

When FAs are abundant, UBXD8 migrates from the ER to LDs and returns to the ER as intracellular lipid levels drop(Zehmer et al., 2009). At the LD membrane, UBXD8 recognizes substrate proteins and recruits p97. In the liver cell line HUH7, UBXD8 recognizes LD associated ubiquitinated ApoB through its UBA domain and mediates the proteasomal degradation of ApoB through p97(Suzuki et al., 2012). In a non-proteolytic role, UBXD8 recruits p97 to the lipase ATGL and inactivates the protein by separating it from its co-activator CGI-58(Olzmann et al., 2013). How UBXD8 recognizes ATGL and the role of ubiquitin is unclear. Since ATGL is the rate-limiting enzyme of TG hydrolysis, over-expression of UBXD8 is thought to increase LD size by inhibiting LD turnover(Olzmann et al., 2013). Thus, UBXD8 appears to promote FA storage under lipid rich conditions while preventing FA storage under lipid depleted conditions. The complex role of UBXD8 in lipid metabolism is made more apparent when UBXD8 conditional liver knockout mice were found to have a slightly more severe phenotype of fatty liver when compared to controls after a high-fat diet(Imai et al.,

2015). Since UBXD8 has been proposed to inhibit ATGL, these findings contradict the prediction that UBXD8 liver knockout mice may have reduced lipid accumulation. As LDs only accumulated in a specific region of the liver in UBXD8 knockout mice, these findings may reflect differences in the in vivo regulation of UBXD8 function(Imai et al., 2015). Given the potential dual role of UBXD8 in regulating both TG hydrolysis and synthesis, further work needs to be done to elucidate the mechanisms that direct UBXD8 towards a specific function. A schematic depicting the known cellular roles of UBXD8 is below (Figure 1-5).

Figure 1-5. The role of UBXD8 and p97 in lipid metabolism. A proposed model depicting the known functions of UBXD8 and p97 in lipid metabolism. Under low FA conditions, UBXD8 remains anchored at the ER and activates SREBP1 and FA synthesis by recruiting p97 to mediate degradation of INSIG1. UBXD8 also inhibits conversion of DG to TG potentially through a DGAT enzyme. Stimulation with unsaturated FA (uFA) induces migration of UBXD8 to LDs. Here, UBXD8 recruits p97 and inhibits lipolysis by separating ATGL from its co-activator CGI-58.

1.4 Thesis rationale and objectives

Ubiquitin E3 ligases dictate functional specificity of the ubiquitin system by selecting the appropriate substrates. To further understand the functions and underlying mechanisms, it is essential to identify E3 substrates and study their fate. This thesis utilizes a proximity-based protein screening approach to explore the interactions and functions of RNF126 and RNF115. Chapter 2 characterizes RNF126 associating proteins and substrates identified by BioID while chapter 3 describes a potential role for RNF126 in lipid droplet homeostasis. This work demonstrates the effectiveness of a proximity-based screening approach and furthers our understanding of the functional roles of ubiquitin E3 ligases.

Chapter 2 Identifying interacting proteins and substrates of RNF126 and RNF115 2.1 Introduction

Although the ubiquitin system has been implicated in almost every cellular pathway, the mechanistic details are still poorly understood. One limiting factor is our lack of understanding of the specific functions of various ubiquitin E3 ligases. Recent techniques such as affinity purification mass spectrometry have greatly aided the identification of E3 functions, although these techniques are limited due to the often transient and weak nature of E3-substrate interactions as well as difficulty identifying insoluble proteins. In this chapter, I describe a novel protein screening approach that we performed in collaboration with Dr. Brian Raught’s lab called BioID. This technique identifies proteins that have been in close proximity to RNF126 or RNF115 in vivo. I then validate if the identified proteins can form a complex with RNF126 as well as which ones are bonafide substrates. These experiments begin to characterize the complex protein interaction network that RNF126 is involved in and suggest that RNF126 may play several roles in cell biology.

2.2 Materials and Methods

2.2.1 Cell culture, cDNA constructs, and reagents

HeLa cells stably expressing a shRNA targeting RNF126, RNF115, or a non-silencing sequence were generated previously(Smith et al., 2013). HeLa and HEK293 cells with a stable integration of a FRT site were a generous gift from Dr. Brian Raught. All cells were maintained in DMEM supplemented with 10% FBS and passaged prior to confluence.

Full length murine RNF126 cDNA cloned into pGEX4T3, pcDNA3.1, and 2x nHA pcDNA3.1 were previously generated(Smith et al., 2013; Smith and McGlade, 2014). The C13,16A point mutations in pcDNA3.1 was generated using a QuikChange site directed mutagenesis kit (Stratagene) as previously described(Smith et al., 2013; Smith and McGlade, 2014). Full length murine RNF115 cDNA in pGEX4T3 and pcDNA3.1, and the C229A and C22,25A mutations were generated previously(Smith et al., 2013). Human CUL3 cDNA was a generous gift from Dr. Gil Privé (University of Toronto, Toronto) and cloned into p3XFLAG-CMV-10 using the

21 22 restriction sites NotI and KpnI. The pcDNA5 FRT/TO Flag-BirA* expression vector and pOG44 were a generous gift from Dr. Brian Raught. Human RNF115 were PCR amplified and cloned into pcDNA5 FRT/TO Flag-BirA* using the restriction sites AscI and BsrGI. Murine RNF126 in pcDNA5 FRT/TO Flag-BirA* was generated previously by Aaron Prodeus and Hyeyeon Kim. The UBA domain (1-66), UAS domain (122-277), and UBX domain (336-445) of UBXD8 were PCR amplified and cloned into pGEX4T3 using the restriction sites BamHI and NotI. Primers and other constructs are listed below.

RNF126 and RNF115 antibodies have been previously described(Smith et al., 2013). The follow commercial antibodies were used: RNF126 (Atlas Antibodies, HPA043050), Flag-M2 (Sigma, F1804), HA (Covance, MMS-101P), HA (Roche, ROAHA), p62 (BD Biosciences, 610833), p97 (Novus, NB100-1558), α-tubulin (Sigma, T9026), V5 (ThermoFisher, R960-25), and myc (UBPBio, Y1091).

Table 2-1: List of primers utilized for cloning

Primer Name Forward Primer Reverse Primer

UBXD8- 5’ attggatccatggcggcgcctg 5’ aactgcggccgctgatggaggtgggttgaaa UBA-pGEX

UBXD8- 5’ attggatcccctgaccctcgcagcc 5’ aactgcggccgcgcgttctgacaccaggtaag UAS-pGEX

UBXD8- 5’ attggatcccggcagaatttacaggagga 5’ aactgcggccgcttcgtcagttaggtcctgaaca UBX-pGEX

RNF115- 5’ tataggcgcgccAatggcggaggcttcg 5’ Ttgtacatcagaaagtccatcggtcat BirA

Table 2-2: List of cDNA constructs used

cDNA Construct Tag Obtained from

Human UBXD8 pcDNA3.1-DEST nMyc SPARC (Hospital for Sick Children, Toronto)

Human UBXD1 pcDNA3.2-DEST cV5 SPARC (Hospital for Sick Children, Toronto)

Rat p97 pcDNA5-FRT-TO cMyc Addgene (#31837)

Human p62 pcDNA4-TO cHA Addgene (#28027)

Human VCPIP1 pDEST-LTR-IRES- nFlag and nHA Addgene (#22592) Puro

Human UFD1L pcDNA3.2-DEST cV5 SPARC (Hospital for Sick Children, Toronto)

Human NPLOC4 pcDNA3.1-DEST nFlag SPARC (Hospital for Sick Children, Toronto)

Human KEAP1 pcDNA3.1-DEST nMyc SPARC (Hospital for Sick Children, Toronto)

2.2.2 Forward and reverse transfection

For forward transfection of cDNA, cells were seeded on 10 cm2 dishes and transfected using a DNA-lipofectamine 2000 (Invitrogen) mixture at a ratio of 1 µg cDNA to 3 µL lipofectamine 2000. Cells were lysed 24 hours after transfection. For immunocytochemistry, cells were seeded on cover slips in a 24-well plate and forward transfected for 6 hours with lipofectamine 2000. After 6 hours, the media was replaced and cells were manipulated as required. For reverse transfection, 20 pmol of siRNA targeting RNF126: 5’-CCGGATTATATCTGTCCAAGA-3’ (Qiagen) or an All-Stars negative control (Qiagen) was incubated with 4 ul of lipofectamine 2000 in a 6-well dish for 20 minutes at room temperature. 1.5 mL of DMEM containing 10% FBS was then added to each well and cells seeded into the mixture. Experiments were conducted 48 hours post transfection.

2.2.3 Generation of doxycycline inducible stable cell lines and BioID screen

Doxycycline inducible HEK293 and HeLa stable cell lines were generated using the Flp-In system (ThermoFisher). HEK293 and HeLa cells with a stable FRT integration site were co- transfected with Flag-BirA*-hRNF115 and pOG44. Media was replaced the following day and the cells left to recover for 24 hours. Cells were then trypsinized and seeded into media containing DMEM, 10% FBS, and 400 µg/mL Hygromycin B (for HeLa) or 200 µg/mL Hygromycin B (for HEK293). Cells were left to grow in selection media until colonies were visible and then all cells were pooled together for future experiments. Doxycycline inducible HEK293 expressing Flag-BirA*-mRNF126 was generated by Dr. Donna Berry and the HeLa Flag-BirA*-mRNF126 cell line generated by Aaron Prodeus and Hyeyeon Kim.

To perform the BioID screen, inducible cell lines were seeded into ten 15 cm2 plates and grown to 70% confluency. Cells were then stimulated with 1 µg/mL doxycycline (Sigma-Aldrich) and 50 µM biotin (Bioshop) for 24 hours in the presence or absence of 1 µM MG132 (Peptide Institute). Cells were then collected by scraping, pelleted and washed 2 times with PBS, and frozen in a dry ice ethanol bath. HEK293 cells expressing Flag-BirA*-mRNF126 were prepared for BioID by Dr. Donna Berry. Cell pellets were given to Dr. Brian Raught’s lab to perform the mass spectrometry analysis. Statistical analysis of mass spectrometry data by SAINT was also performed by Dr. Brian Raught’s lab.

2.2.4 Production of GST fusion proteins

An overnight culture of transformed BL-21 cells were grown to an O.D. of 0.6-0.8 and induced with 0.3 mM IPTG at 37oC for 3 hours. Bacterial pellets were sonicated in NP40 lysis buffer supplemented with complete protease inhibitors (Roche Applied Science). Bacterial lysates were cleared by centrifugation (15000 RPM for 15 minutes at 4oC) and incubated for 30 minutes at 4oC with glutathione-Sepharose beads (GE Healthcare). Beads were washed 4 times with NP40 wash buffer and re-suspended in NP40 lysis buffer and 1 mM DTT.

2.2.5 Co-immunoprecipitations, GST pull-downs, in vivo ubiquitination assays, and western blotting

For co-immunoprecipitations, forward co-transfected HEK293T cells were lysed in Nonidet P-40

(NP40) lysis buffer (50 mM HEPES (pH 7.5), 150 mM NaCl, 1.5 mM MgCl2, 1 mM EGTA (pH 8), 10% (v/v) glycerol, 1% (v/v) Nonidet P-40) supplemented with complete protease inhibitors (Roche Applied Science). Lysates were cleared by centrifugation (14000 RPM for 10 minutes at 4oC) and 1 mg of lysate was mixed with 1-2 µg of antibody and either protein-A or protein-G at 4oC overnight with gentle agitation. The next day, beads were washed three times with 1 mL of cold NP40 wash buffer (20 mM HEPES (pH 7.5), 150 mM NaCl, 10% (v/v) glycerol, 0.1% (v/v) Nonidet P-40) and proteins were eluted by boiling in SDS-Laemmli sample buffer. For GST pull-downs, 1 mg of cleared cell lysate was mixed with 5 µg GST fusion protein for 2 hours at 4oC, washed 3 times with NP40 wash buffer, and analyzed by western blotting.

In vivo ubiquitination assays were performed by lysing transfected HEK293T cells in boiling hot RIPA buffer (20 mM Tris (pH 7.5), 2 mM EDTA, 150 mM NaCl, 1% (v/v) Nonidet P-40, 1% sodium deoxycholate, 0.1% SDS) containing 1% SDS. Cell lysates were briefly sonicated and then boiled another 5 minutes. 1 mg of cell lysate was diluted 10-fold in NP40 lysis buffer and immunoprecipitated as above.

For western blotting, proteins were separated by SDS-PAGE and transferred to a PVDF membrane (Pall Corp) and blocked at room temperature for one hour with 5% skim milk powder in Tris-buffered saline with 0.05% Tween 20 (TBST). Membranes were probed overnight with primary antibody, washed 3 times with TBST, incubated at room temperature for 1 hour with corresponding secondary antibodies, and washed another 3 times with TBST. Antibody binding was detected using enhanced chemiluminescence reagent (Perkin Elmer). Antibody

26 concentrations are as follows: RNF126 (1:500, 5 µL for IP), RNF115 (1:500, 5 µL for IP), p62 (1:4000), p97 (1:5000), HA (Covance, 1:2000), HA (Roche, 2 µg for IP), myc (1:1000, 1-2 µg for IP), α-tubulin (1:5000), V5 (1:5000, 1 µg for IP), and Flag (1:1000, 1 µg for IP).

2.3 Results

2.3.1 Identification of proteins in near proximity to RNF126 and RNF115 by BioID

Although classical screening approaches such as affinity purification mass spectrometry (AP- MS) have proven to be extremely valuable, limitations exist. The two major limitations of AP- MS involve difficulty identifying insoluble proteins and loss of protein-protein interactions during the affinity purification step(Lambert et al., 2015). Since the success of AP-MS depends on the preservation of protein complexes, cells often have to be lysed under mild conditions. This can result in the poor extraction of membrane proteins and chromatin associated proteins(Gingras and Raught, 2012). Additionally, weak and transient interactions are often lost during the affinity purification step. One way to address these issues is to characterize proteins in the vicinity of a protein of interest. BioID is a novel approach that relies on biotinylation of proteins in a proximity-dependent manner (Figure 2-1)(Roux et al., 2012). In BioID, the Escherichia coli biotin ligase protein (BirA*) has been mutated to promiscuously biotinylate nearby primary amines. This is accomplished by the BirA* mutant prematurely releasing activated biotin, forming a cloud of activated biotin in vivo that can label nearby proteins(Gingras and Raught, 2012). The experimentally determined labelling radius of BioID is ~10 nm, providing enough resolution to resolve the topology of a protein complex in vivo(Kim et al., 2014). The in vivo nature of BioID provides an additional advantage as protein interactions post-lysis are not a source of false positives. Additionally, all detected candidate proteins are biotinylated under cell physiological conditions, avoiding a potential pitfall of AP-MS. As biotinylation of neighbouring proteins is a covalent modification, harsh lysis conditions can be used to solubilize most cellular proteins. Biotinylated proteins can then be enriched in a purification step and identified by mass spectrometry.

Figure 2-1. The BioID method. A promiscuous biotin ligase (BirA*) is fused to a desired bait protein and expressed in cells. In the presence of biotin, biotin is activated and released to create a cloud of activated biotin. Activated biotin then covalently labels proteins in close proximity to the bait of interest. Following a harsh lysis to solubilize proteins, biotinylated proteins are captured using streptavidin-conjugated beads and analyzed by mass spectrometry. Adapted from (Roux et al., 2012).

Since RNF126 and RNF115 have been shown to play important roles in regulating membrane proteins, it is likely that an affinity based mass spectrometry approach would not be optimal. Indeed, our previous affinity based mass spectrometry screens identified only a small cohort of potential interacting proteins when RNF126 was used as bait (Figure 2-2 A and B). To further understand the functions of these E3 ligases, we utilized the BioID method by first fusing a Flag- BirA* tag to the N-terminal of mouse RNF126 or human RNF115 and then stably expressing these proteins in HeLa or HEK293 T-REX Flp-In cells. Treatment with doxycycline induced the specific expression of Flag-BirA*-RNF126 or Flag-BirA*-RNF115 and no protein expression is observed in untreated cells (Figure 2-2 C and D). Importantly, Flag-BirA*-RNF126 was expressed at relatively low protein levels when compared to endogeneous RNF126 (Figure 2-2 C). Since fusion proteins are expressed at controlled levels in cells, it is likely that cellular ER stress is minimal and thus not a major contributor to noise in BioID data. Additionally, since our proteins of interest are both ubiquitin E3 ligases and have been implicated in protein degradation, we reasoned that potential substrates may be stabilized by inhibition of the proteasome with MG132. In collaboration with Dr. Brian Raught, biotinylated proteins were identified by mass spectrometry and the data analyzed by Significance Analysis of INTeractomes (SAINT)(Choi et al., 2011; Teo et al., 2014) to identify high confidence interacting partners. This was done by comparing BioID data with a set of controls including untransfected parental cells, cells expressing Flag-BirA* alone, and cells expressing an unrelated bait protein to determine endogenously biotinylated proteins and non-specific proteins. Proteins identified with a SAINT score ≥ 0.75 were considered high confidence interacting partners (See appendix for full list of high confidence candidates).

Two BioID experiments performed in HEK293 cells using RNF115 as bait yielded a total of 61 unique high confidence interacting proteins (Figure 2-3 A). The vast majority of these proteins were enriched by MG132 (58 proteins) while only 3 proteins were considered high confidence without MG132 treatment. These results reflect the low abundance of peptides detected in the absence of MG132 by mass spectrometry. As we have previously found evidence suggesting that RNF115 is rapidly degraded in cells (Data not shown), proteasome inhibition may be required to enrich for biotinylation. BioID of RNF126 was performed in HeLa cells twice and once in HEK293 cells and the combined data visualized (Figure 2-3 B). In HeLa cells, we identified a total of 68 high confidence interacting partners with 59 proteins that were identified without

MG132 and 9 proteins scoring positive after MG132 treatment. In HEK293 cells, we identified a total of 41 high confidence interacting partners with 8 proteins identified without MG132 and 33 scoring positive after MG132. Comparison of datasets showed that 10 proteins were identified in both cell lines when RNF126 was used as bait (Figure 2-3 C). This heterogeneity is likely due to variabilities between the two cell lines. To gain insight into the degree of overlap shared between the RNF115 and RNF126, we compared BioID data of all identified high confidence proteins and found that 26 proteins are shared between the datasets (Figure 2-3 C). These data indicate that in addition to sharing similar functions, these E3 ligases likely have independent roles as well.

In keeping with our previous work, BioID of RNF115 and RNF126 identified several endosomal sorting components such as EPS15 and HGS (Figure 2-3 A and B). We were also able to identify a previously known RNF126 interacting protein (BAG6)(Rodrigo-Brenni et al., 2014) and a member of its complex (UBL4A) (Figure 2-3 A and B). This provides confidence that our BioID screen is identifying biologically relevant components. Our highest spectra count was obtained from the autophagy adapter protein SQSTM1 for both E3 ligases. This novel connection to autophagy was further established by the identification of NBR1, another autophagy adapter protein, as a high confidence protein (Figure 2-3 A and B). Notably, our BioID screens identified several proteins that function as p97 co-factors (NPLOC4, UFD1L, VCPIP1, UBXN2A) for both E3 ligases as well as additional components (PLAA, UBXN1) for RNF126 (Figure 2-2 A and B). These data strongly suggest that RNF115 and RNF126 may form protein complexes with p97 and its associated co-factors.

Figure 2-2. Generation of stable cell lines for BioID. A and B) Data from affinity-purification mass spectrometry screens using RNF126 as bait. Stable HEK293T cells over-expressing Flag- tagged RNF126 were subjected to IP using anti-Flag M2-Agarose resins. Liquid chromatography-mass spectrometry was performed after A) competitive elution with Flag peptide followed by gel electrophoresis and in-gel tryptic digestion, or B) urea elution followed by in-solution tryptic digestion and reverse-chromatography on C18 resin. Affinity-purification mass spectrometry screens were performed by Dr. Donna Berry. C) HeLa cells stably expressing a doxycycline (dox) inducible form of Flag-BirA*-mRNF126 were stimulated with dox for 24 hours. Following cell lysis, a Flag IP was performed and then analyzed by immunoblotting with anti-RNF126 antibodies. D) HEK293 cells stably expressing a doxycycline inducible form of Flag-BirA*-hRNF115 were stimulated with dox for 24 hours. Following cell lysis, a RNF115 IP was performed and then analyzed by immunoblotting with anti-Flag antibodies.

Figure 2-3. BioID of RNF126 and RNF115. A and B) Mass spectrometry analysis of endogeneous proteins that are in close proximity to RNF115 (A) or RNF126 (B). Green lines represent proteins identified by BioID and the thickness of the line represents the number of peptides identified. STRING analysis connecting known protein-protein interactions is shown by the red dotted line. All candidates displayed have a SAINT score > 0.75 and STRING analysis was performed at the high confidence interval. C) Venn diagram showing the distribution of proteins between the 3 BioID data sets.

2.3.2 Validation of potential interacting proteins identified by BioID

Since BioID data encompasses both interacting proteins and non-interacting proteins in the vicinity of the bait, it is vital to validate protein interaction through an alternative method. Proteins that were determined by SAINT to be high confidence interacting partners were analyzed to create a list of top candidates. As we have previously shown RNF126 to be involved in endosomal trafficking(Smith et al., 2013), we were particularly interested in trafficking proteins from our candidate list. Additionally, we noticed that several proteins from our candidate list have been reported to form protein complexes together. The close proximity of proteins in a complex may result in the identification of several components by BioID, strongly indicating that these proteins may interact with RNF126. To determine if BioID candidate proteins could associate with RNF126, HEK293T cells were co-transfected with RNF126 and a cDNA of interest and cell lysates subjected to co-immunoprecipitation (co-IP) assays. Through these assays, NPLOC4, EPS15, and KEAP1 were found to co-IP with RNF126 while co- transfection with an empty vector control failed to pull down target proteins (Figure 2-4). These data suggest that RNF126 specifically associates with these proteins and provides evidence that BioID can successfully identify interacting partners. Furthermore, our data shows that RNF126 can associate with several proteins that function as p97 co-factors, suggesting that RNF126 may play an active role in p97 related functions.

Figure 2-4. Validation of BioID candidates by co-immunoprecipitation. A) RNF126 HEK293T cells were co-transfected with untagged RNF126 and Flag-NPLOC4. IPs were performed using anti-RNF126 antibodies and immunoblotted for Flag-NPLOC4 and RNF126, N = 1. B) HEK293T cells were co-transfected with untagged RNF126 and Myc-KEAP1. IPs were performed using anti-RNF126 antibodies and immunoblotted for Myc-KEAP1 and RNF126, N = 3. C) HEK293T cells were co-transfected with untagged RNF126 and untagged EPS15. IPs were performed using anti-RNF126 antibodies and immunoblotted for EPS15 and RNF126, N = 2.

2.3.3 RNF126 associates with p97 and the UBX family of co-factors p97 is an essential protein involved in a considerable number of cellular pathways. To provide functional specificity, p97 forms large complexes with a diverse group of co-factors that select substrates or recruit p97 to specific compartments. As our data indicated that RNF126 may be involved in p97 related functions, we first investigated if RNF126 could associate with p97. RNF126 and p97 were co-expressed in HEK293T cells and RNF126 IPs analyzed by immunoblot. To further characterize a potential interaction between RNF126 and p97, we utilized RNF126 constructs with inactivating mutations in the zinc finger or RING domain. These experiments found that p97 co-IPs with RNF126 but not with RNF126 constructs with inactivating mutations in the RING domain, suggesting association depends on the RING domain of RNF126 (Figure 2-5 A).

The largest group of p97 co-factors are defined by the presence of a C-terminal UBX domain. In addition to the UBX proteins identified by BioID, AP-MS experiments from our lab and by another group suggest that RNF126 can associate with UBXD8(Alexandru et al., 2008). Furthermore, it has been reported that p97 and UBXD1 play a role in endosomal trafficking and may also be involved in trafficking of the EGFR(Ritz et al., 2011), similar to RNF126. To test if RNF126 has a functional role with either of these proteins, we first determined that RNF126 associates with UBXD1 and UBXD8 through co-IP assays (Figure 2-5 B and C). Intriguingly, wild-type RNF126, but not the ligase dead RING domain mutant, was able to co-IP with large molecular weight forms of both UBXD1 and UBXD8 (Figure 2-5 B and C). As the RING domain is essential for ubiquitination, it is likely that these large molecular weight forms are post-translationally modified forms of UBXD1 and UBXD8. These data suggest that both proteins are substrates of RNF126.

As we have previously shown that the zinc finger of RNF126 binds ubiquitin(Smith et al., 2013), we rationalized that RNF126 may bind the ubiquitin-like UBX domain through its zinc finger. However, while UBXD1 binding is dependent on the zinc finger of RNF126 (Figure 2-5 B), inactivation of the zinc finger did not disrupt association between RNF126 and UBXD8 (Figure 2-5 C). As the UBX domain of UBXD1 is unique from all other UBX proteins(Kern et al., 2009; Madsen et al., 2008), it is possible that the zinc finger of RNF126 specifically recognizes the

UBX domain of UBXD1. The role of the zinc finger in mediating interaction with other UBX proteins remains to be determined.

To characterize the association between RNF126 and UBXD8, UBXD8 was first co-expressed with wild-type RNF126 or the zinc finger or RING domain mutant constructs. Following IP with anti-RNF126 antibodies, UBXD8 was found to co-IP with both the zinc finger and RING domain mutant constructs, suggesting that both domains are dispensable for association (Figure 2-5 C). Surprisingly, when co-IP experiments were performed in reverse by first IPing Myc- UBXD8, UBXD8 failed to co-IP with the RING domain mutant of RNF126 (Figure 2-5 F). As the role of the RING domain in mediating association to UBXD8 is unclear, we took an alternate approach and isolated the 3 main domains of UBXD8 and fused them to a GST tag. Through GST pull down experiments, we found that UBXD8 associates preferentially with RNF126 through its ubiquitin binding UBA domain and weakly through its UBX domain (Figure 2-5 D and Figure 4-1 B). As RNF126 has been shown to auto-ubiquitinate in vitro(Smith et al., 2013), UBXD8 may be associating with auto-ubiquitinated RNF126 through its UBA domain. Indeed, mutation of the RING domain abolished association with the UBA domain of UBXD8, suggesting a role for the auto-ubiquitination of RNF126 (Figure 2-5 E). Together, the data suggests that a functional RING domain is required for interaction with the UBA domain of UBXD8.

Figure 2-5. RNF126 associates with p97 and members of the UBX family. A) HEK293T cells were co-transfected with untagged RNF126 or the indicated mutant construct and Myc-p97. IPs were performed using anti-RNF126 antibodies and probed by immunoblot, N = 3. B) HEK293T cells were co-transfected with untagged RNF126 or the indicated mutant construct and UBXD1- V5. IPs were performed using anti-RNF126 antibodies and probed by immunoblot, N = 3. C) HEK293T cells were co-transfected with untagged RNF126 or the indicated mutant construct and Myc-UBXD8. IPs were performed using anti-RNF126 antibodies and probed by immunoblot, N = 3. D) The three known domains of UBXD8 were isolated and fused to a GST tag to generate fusion proteins. HEK293T cells transfected with untagged RNF126 were lysed and incubated with fusion protein for 2 hours and immunoblotted for RNF126. Membranes were then stained with Fast Green, N = 2. E) HEK293T cells were transfected with HA tagged wild- type RNF126, zinc finger mutant, or RING domain mutant. Cell lysates were prepared and incubated with the UBA domain from UBXD8 fused to a GST tag and immunoblotted for HA- RNF126. Membranes were then stained with Fast Green, N = 2. F) HEK293T cells were co- transfected with untagged RNF126 or the indicated mutant construct and Myc-UBXD8. IPs were performed using anti-Myc antibodies and probed by immunoblot for endogenous p97, N = 3.

2.3.4 Over-expression of RNF126 promotes UBXD8 association with p97

To ensure functional specificity, p97 forms large protein complexes with co-factors that recruit or modify substrate proteins. Although there is emerging literature detailing the functional roles of some of these p97 complexes, how formation of these complexes is regulated is still poorly understood. To test if RNF126 has a role in regulating p97-UBXD8 complex formation, we co- expressed UBXD8 and RNF126 and determined if co-IP of endogenous p97 with UBXD8 was altered. Interestingly, more p97 co-immunoprecipitated with UBXD8 when wild-type RNF126 was over-expressed than with empty vector control (Figure 2-5 F). This increased amount of p97-UBXD8 pull-down was abolished when the ligase dead RING domain mutant was expressed suggesting that the ubiquitination activity of RNF126 promotes complex formation (Figure 2-5 F). As mutation of the zinc finger did not alter p97-UBXD8 binding, it is likely that this domain does not play an essential role in mediating p97-UBXD8 complex formation. Indeed, this is not surprisingly as the zinc finger mutant is still capable of associating with both UBXD8 and p97 (Figure 2-5 A, C, F).

Figure 2-6. RNF126 ubiquitinates UBXD8 and UBXD1. HEK293T cells were co-transfected with untagged RNF126 or a mutant construct, HA-ubiquitin, and either Myc-UBXD8 (A) or UBXD1-V5 (B). Cell lysates were prepared by lysing cells in boiling hot RIPA buffer containing 1% SDS, sonicating, and boiling for another 5 minutes. Myc-UBXD8 and UBXD1-V5 IPs were performed by diluting lysates 10 fold in NP40 lysis buffer and analyzed by immunoblot. A) RNF126 ubiquitinates UBXD8, N = 3. B) RNF126 ubiquitinates UBXD1, N = 2.

2.3.5 UBXD8 and UBXD1 are ubiquitinated by RNF126

To determine if UBXD8 and UBXD1 are bonafide substrates of RNF126, we co-expressed each protein with HA-tagged ubiquitin and RNF126 and conducted the IP under denaturing conditions to abolish intermolecular interactions. This step ensures that any detected ubiquitin signal can be attributed solely to the immunoprecipitated protein as all protein interactions have been disrupted. Immunoblots with anti-HA antibodies of UBXD1 and UBXD8 IPs show a smear of large molecular weight bands when wild-type RNF126 is over-expressed (Figure 2-6 A and B). These signals correspond to ubiquitinated protein and likely represent poly-ubiquitinated UBXD1 or UBXD8. The high molecular weight smearing is reduced to levels similar to empty vector control when either protein is co-expressed with the ligase dead RING domain mutant, suggesting that the RING domain is required for ubiquitination (Figure 2-6 A and B). Since mutation of the zinc finger did not abolish co-IP of RNF126 and UBXD8 (Figure 2-6 C), it is not surprising that UBXD8 was ubiquitinated in the presence of the mutated zinc finger RNF126 construct (Figure 2-6 A). On the other hand, mutation of the zinc finger abolished co-IP of RNF126 with UBXD1 (Figure 2-5 B) and as expected, abolished ubiquitination of UBXD1 as well (Figure 2-6 B). This suggests that recognition of UBXD1 by RNF126 is necessary for ubiquitination.

2.4 Discussion

Identification of ubiquitin E3 ligase substrates has remained challenging. Due to the often transient and weak nature of the E3-substrate interaction, conventional screening approaches are often unsuccessful. We have previously characterized the E3 ligases RNF126 and RNF115 and demonstrated a role in membrane trafficking(Smith et al., 2013). As both E3 ligases can ubiquitinate the EGFR, it is likely that additional substrates may be membrane proteins. To identify these substrates, we utilized the proximity-dependent BioID approach to characterize neighbouring proteins surrounding the two E3s. Although RNF126 and RNF115 are closely related proteins with similar functions, both proteins have been shown to have separate functions(Rodrigo-Brenni et al., 2014; Smith et al., 2013; Smith and McGlade, 2014). In agreement with published data, our BioID data showed that RNF126 and RNF115 share a subset of potential interacting proteins as well as unique candidate proteins. Interestingly, BioID using RNF115 as bait required addition of MG132 to enrich for peptides. This may reflect rapid

43 turnover of RNF115 as proteasomal inhibition has been shown to stabilize the protein(Burger et al., 2005). Proteasomal inhibition also enhanced the abundance of specific proteins in RNF126 BioID screens, indicating that these proteins may potentially be targeted for proteasomal degradation by RNF126.

As expected, both E3s were found to be associated with endosomal trafficking components. We also found that both E3s may play roles in autophagy through p62 or NBR1. Given the importance of the lysosome in both these pathways, it would be interesting to explore the BioID interactome of both E3s following lysosome inhibition. This may further enrich for proteins targeted for lysosomal degradation by autophagy or endocytosis. Due to the nature of BioID detecting both physical and non-physical interactions, an alternative validation method is required. Through co-immunoprecipitation experiments, we demonstrated that BioID is an extremely effective method for identifying interacting proteins.

Our BioID data indicated that both E3s were strongly associated with p97 and its diverse group of co-factors. These proteins function in a broad array of cellular pathways in an ubiquitin dependent manner. To determine a potential role for RNF126 in this pathway, we explored if RNF126 could associate with p97 or its co-factors. Interestingly, RNF126 not only associated with p97 and several co-factors, but also ubiquitinates UBXD1 and UBXD8. Experiments using RNF126 mutants demonstrated that association with p97 requires the RING domain of RNF126 while association with UBXD1 requires the zinc finger. Interestingly, the zinc finger is not required for association with UBXD8 suggesting that RNF126 may have different modes of interaction with UBX proteins. Although co-IP experiments were unable to clearly establish the role of the RING domain in mediating association between RNF126 and UBXD8, further work using the isolated UBA domain of UBXD8 found that interaction depends on a functional RING domain. This data supports a model where the RING domain is required for binding to UBXD8. How the RING domain mutant is able to co-IP with UBXD8 is unclear. One possibility may be due to background resulting from the use of crude serum RNF126 antibodies. Future experiments should use epitope tagged RNF126 to increase specificity during the IP step. The UBX domain of UBXD8 was also found to weakly bind RNF126. Although the role of the UBX domain is unknown, this interaction may assume a bipartite binding model where UBXD8 interacts with ubiquitinated RNF126 through its UBA domain while the UBX domain binds an alternative location on RNF126. This mechanism may enhance specificity by ensuring UBXD8 binds

44 ubiquitinated RNF126 instead of other ubiquitinated cargo. One functional possibility may involve recruitment of ubiquitinated RNF126 by UBXD8 to p97 for proteasomal degradation similar to UBXD7 mediated recruitment of HIF1α(Alexandru et al., 2008). As p97 often forms large complexes, it remains to be seen if these interactions with RNF126 are direct.

The role of ubiquitinated UBXD8 is unclear. Co-expression of RNF126 with UBXD8 does not lower UBXD8 abundance, suggesting that RNF126 does not promote degradation of UBXD8. Interesting, co-expression of RNF126 promotes association between p97 and UBXD8, suggesting that RNF126 may regulate the binding of p97 co-factors. The regulatory mechanisms that determine how p97 selects its co-factors are poorly characterized. Further work examining the mechanisms of how RNF126 promotes UBXD8-p97 association may reveal a novel regulatory mechanism. Since RNF126 also ubiquitinates UBXD1 and may potentially interact with other UBX proteins, it will be interesting to explore if RNF126 can regulate p97 selection of other co-factors.

Chapter 3 RNF126 regulates lipid droplet size following fatty acid stimulation 3.1 Introduction

Cellular lipid levels reflect a constant balance between lipid synthesis and turnover. Excess lipids are converted to neutral esters such as triglyceride or ester-cholesterols and stored in lipid droplets. As lipid droplets function to protect cells from lipid toxicity as well as provide sources of energy, defects in lipid droplet growth or regression can lead to disease. Thus, the size of cellular lipid droplets is a strong indicator of lipid reservoirs. Lipid droplet size is regulated by a complex group of membrane proteins such as UBXD8. Previous work has shown that UBXD8 can bind fatty acids through a central UAS domain as well as regulate triglyceride hydrolysis through recruitment of p97(Kim et al., 2013; Olzmann et al., 2013). Since we have previously found that RNF126 associates and ubiquitinates UBXD8, we hypothesized that RNF126 may regulate lipid droplet size through UBXD8. Here, we provide evidence that RNF126 plays a role in regulating lipid droplet size in HeLa cells following oleate stimulation. We show that HeLa cells stably depleted of RNF126 form significantly smaller lipid droplets than control cells, when stimulated with oleate. We determined that the difference in lipid droplet size is not due to enhanced lipid droplet turnover in the absence of RNF126, but instead due to defective lipid droplet expansion. These data demonstrate the role of RNF126 in lipid droplet homeostasis through regulation of proteins involved in lipid droplet expansion.

3.2 Materials and Methods

3.2.1 Cell culture, cDNA constructs, and reagents

HeLa cells stably expressing a shRNA targeting RNF126, RNF115, or a non-silencing sequence were generated previously(Smith et al., 2013). The pcDNA5 FRT/TO Flag expression vector and pOG44 were a generous gift from Dr. Brian Raught. Murine RNF126 and human RNF115 were sub-cloned from pcDNA5 FRT/TO Flag-BirA* into pcDNA5 FRT/TO Flag using the restriction sites AscI and BsrGI.

The following commercial reagents were used: Triacsin C (Sigma-Aldrich), DEUP (Sigma- Aldrich), sodium oleate (Sigma-Aldrich), fatty acid free BSA (Sigma-Aldrich), and BODIPY 493/503 (ThermoFisher).

45 46

3.2.2 Preparation of oleate-BSA conjugate

Oleate was conjugated to fatty acid free BSA at a 6:1 oleate to BSA molar ratio for a final concentration of 12 mM. Briefly, 0.05 mmol of fatty acid free BSA was dissolved in 24 mL of 0.1 M Tris (pH 8.0) and 0.3 mmol of sodium oleate was dissolved in 1 mL of ultra-pure water at room temperature. Dissolved oleate was combined with the fatty acid free BSA solution and mixed gently at room temperature until the solution turned fully clear. The solution was then filter sterilized and stored at 4oC.

3.2.3 Generation of doxycycline inducible stable cell lines

Doxycycline inducible HeLa stable cell lines were generated using the Flp-In system (ThermoFisher). HeLa cells with a stable FRT integration site was co-transfected with either Flag-mRNF126 or Flag-hRNF115 and pOG44. Media was replaced the following day and the cells left to recover for 24 hours. Cells were then trypsinized and seeded into media containing DMEM, 10% FBS, and 400 µg/mL Hygromycin B. Cells were left to grow in selection media until colonies were visible and then individual colonies isolated and propagated to establish a clonal line. To induce expression, cells were stimulated with 1 µg/mL of doxycycline for 24 hours.

3.2.4 Immunocytochemistry

Cells seeded on glass coverslips were stimulated with 340 µM oleic acid conjugated to BSA for 24 hours. Cells were then fixed in a 4% PFA, 30 mM sucrose solution for 15 minutes at room temperature, permeabilized with 0.05% saponin for 10 minutes at room temperature, and blocked in 3% normal donkey serum (Jackson ImmunoResearch) supplemented with 0.025% saponin for 30 minutes at room temperature. Coverslips were then inverted onto primary antibody solution for 30 minutes in a humidified chamber at 37oC. Cells were washed 3x 10 minutes in PBS and then incubated with corresponding secondary antibodies for 30 minutes at 37oC in a humidified chamber. Cells were washed another 3x 10 minutes in PBS and lipid droplets labelled by incubating cells for 30 minutes at room temperature with 5 µg/mL BODIPY 493/503. Finally, nuclei were labelled with DAPI and cover slips mounted in Dako Fluorescence Mounting Medium. Images were acquired the next day using a Leica DMIRE2 inverted fluorescent microscope with a 63X/1.4NA objective, a step size of 0.23 µm, a Hamamatsu C9100-13 EM- CCD camera, and the imaging software Volocity (PerkinElmer). Representative images were

47 selected and adjusted in Volocity using the contrast enhancement function. Primary antibodies used are: RNF126 (Atlas Antibodies, 1:200), myc (1:500), and p97 (1:200).

3.2.5 Quantification of lipid droplets

Images were deconvoluted by Volocity software using the iterative restoration function set to 20 iterations and confidence interval of 100%. A region of interest was manually drawn around a single cell and average lipid droplet volume and number recorded. Lipid droplets were identified using the following parameters: Find objects: automatic, Separate objects: 0.08 µm3, exclude objects < 0.03 µm3 and > 15 um3, exclude objects touching edges of image.

3.2.6 Lipid droplet purification and immunoprecipitation

To isolate lipid droplets, ten 10 cm2 dishes of either RNF126 knockdown cells or non-silencing control were stimulated for 24 hours with 340 µM oleic acid. Plates were washed two times with 10 mL PBS and cells were scraped and pelleted by centrifuging 10 minutes at 1000 x g at 4oC. Cells were then washed one time with ice-cold hypotonic lysis medium (HLM) (20 mM Tris (pH 7.4), 1 mM EDTA) supplemented with protease inhibitors. To lyse, cells were re-suspended in 3 mL cold HLM and transferred to a Wheaton 7 mL dounce tissue grinder on ice. Cells were slowly homogenized on ice with 8 gentle strokes without twisting and centrifuged again for 10 minutes at 1000 x g at 4oC. Following centrifugation, a fat layer is visible near the surface of the lysate solution. Using a large tipped pipette, the fatty solution was collected starting from the surface into a 13.2 mL ultra-centrifuge tube (Beckman Coulter) suitable for a SW41 rotor. 1M sucrose was added to the fatty lysate to make a final concentration of 0.6 M sucrose and mixed by gentle pipetting to evenly disperse lipid droplets. To make the sucrose gradient, 4 mL of cold HLM containing 0.15M sucrose was gentle layered on-top of the fatty lysate followed by a final layer of 4 mL HLM. Tubes were then centrifuged in a SW41Ti swinging bucket rotor (Beckman Coulter) for 1 hour at 33000 RPM at 4oC and allowed to coast to a stop. To collect the floating fat layer at the surface of the solution, a 200 µL pipette tip was cut to make a large opening and used to collect floating lipid droplets in 20 µL aliquots to eliminate pipetting of excess volume. To concentrate lipid droplets, samples were centrifuged at 14000 RPM at 4oC for 5 minutes and the infranatant below the fatty layer removed carefully. Lipid droplets were then incubated with 1 volume of 10% SDS at 37oC for 1 hour and then centrifuged at 14000 RPM at 4oC for 5

48 minutes. The infranatant containing solubilized proteins was removed and analyzed by western blotting.

For immunoprecipitation of lipid droplet associated proteins, concentrated lipid droplets were incubated in 10 volumes of HLM containing 1% triton-X for 30 minutes on ice. Samples were then centrifuged at 14000 RPM at 4oC for 5 minutes and the infranatant removed. The infranatant was quantified and subjected to immunoprecipitation as outlined previously.

3.2.7 Statistical analysis

Data were analyzed using Prism statistical analysis software (GraphPad Software) and a p < 0.05 was considered statistically significant.

3.3 Results

3.3.1 RNF126 regulates lipid droplet size in oleate stimulated cells

To test if RNF126 has a role in regulating LD size or numbers, we stimulated a clonal HeLa cell line stably knocked down of RNF126 (Figure 3-1 D) with the monounsaturated fatty acid oleate (OA) and labelled LDs with the fluorescent dye BODIPY 493/503. As expected, oleate stimulated cells increased LD size and numbers when compared to untreated cells (Figure 3-1 A). Following 24 hours of OA stimulation, RNF126 knockdown cells had significantly smaller LDs than control (NS) cells (Figure 3-1 B) while LD numbers per cell did not change (Figure 3-1 C). Further analysis of LD size distribution shows that knockdown of RNF126 decreases the number of large LDs (LD volume > 0.6 um3) while increasing the abundance of small LDs (LD volume < 0.6 um3) (Figure 3-1 E). As larger LDs appear to be dramatically reduced in the absence of RNF126, it is possible that RNF126 is essential for the formation of larger LDs in HeLa cells.

Figure 3-1. Knockdown of RNF126 reduces lipid droplet size. A) HeLa cells stably expressing a shRNA targetting RNF126 or non-silencing (NS) control were incubated with 340 uM oleic acid (OA) for 24 hours. Cells were then stained for neutral lipids using BODIPY 493/503 (green). A single Z-section is shown. B) Quantification of average LD volume from OA treated cells in (A). C) Quantification of average LD numbers from OA treated cells in (A). D) Immunoblot of RNF126 knockdown in stable cell lines. E) Histogram representation of LD size distribution from three independent experiments. All values are mean ± s.e.m with N = 3 and at least 20 cells measured per experiment. ns is not-significant, * p < 0.05, ** p < 0.01, *** p < 0.001, Scale bars: 5 µm.

To ensure that our observed phenotype was specific to knockdown of RNF126, we transfected a siRNA targeting an alternate region of RNF126 into parental HeLa cell lines and measured LD size and number after OA stimulation. Following knockdown of RNF126 by siRNA (Figure 3-2 A), LD volume was again significantly decreased (Figure 3-2 B), providing further evidence that our phenotype is specific to knockdown of RNF126. While there was a trend towards increased LD numbers in RNF126 knockdown cells, this difference was not significant when compared to non-silencing controls (Figure 3-2 C).

3.3.2 Re-expression of RNF126 in an inducible cell system can rescue decreased lipid droplet size

To confirm that decreased LD size is specific to RNF126 and not due to off-target effects, we transfected shRNA resistant mRNF126 in stable knockdown HeLa cells to test if we could rescue the effect on LD size. Transient over-expression of RNF126 resulted in unexpected effects on cell adhesion (data not shown) and did not show any signs of rescue. To avoid the consequences of excessive over-expression of RNF126, we constructed a clonal doxycycline inducible stable cell line expressing the siRNA resistant Flag-mRNF126. Treatment of cells with doxycycline induced expression of a single band corresponding to Flag-mRNF126 that was not detected in untreated cells (Figure 3-3 A). To perform the rescue experiment, cells were reverse-transfected with siRNA targeting hRNF126 and stimulated with OA and doxycycline 48 hours later. Immunostaining of RNF126 shows successful knockdown in non-induced (no dox) cells transfected with RNF126 siRNA while NS siRNA transfected cells and doxycycline induced cells all show the presence of RNF126 (Figure 3-3 B). As expected, knockdown of RNF126 by siRNA resulted in significantly smaller LDs than control cells (Figure 3-3 C). Re-expression of RNF126 by doxycycline stimulation was sufficient to fully rescue LD size in RNF126 siRNA knockdown cells back to control levels, confirming that the phenotype observed is specific to the knockdown of RNF126 (Figure 3-3 C). Interestingly, doxycycline induced over-expression of RNF126 was unable to further increase LD size (Figure 3-3 C). This may suggest that the rate limiting step regulating LD size is downstream of RNF126 and that excess RNF126 may have no additional effect on.

While knockdown of RNF126 by siRNA reduced LD size, we also observed a significant increase in LD numbers per cell (Figure 3-3 D). However, re-expression of RNF126 was unable to reverse the increase in LD numbers suggesting that this phenotype may be due to non-specific

52 effects of the siRNA. Indeed, we have previously observed a similar trend towards increased LD numbers when using this siRNA (Figure 3-2 C). Additional siRNA targeting different regions of RNF126 should be tested to determine if this is a non-specific effect.

Figure 3-2. siRNA knockdown of RNF126 decreases lipid droplet size following oleate stimulation. HeLa cells were transfected with siRNA targetting RNF126 or a non-silencing (NS) control for 48 hours and then seeded onto cover slips and stimulated with OA for 24 hours. Cells were imaged at 100X on a oil lens with a 1.4 NA A) Knockdown of RNF126 by siRNA B) Quantification of LD volume C) Quantification of LD numbers. N = 3 with at least 20 cells measured per experiment. All values are mean ± s.e.m. ns is not-significant, * p < 0.05, ** p < 0.01, *** p < 0.001.

Figure 3-3. Re-expression of mRNF126 in an inducible cell model rescues lipid droplet size. A) A clonal HeLa cell line stably expressing a doxycycline (dox) inducible form of mRNF126 was generated and expression induced with 1 ug/mL dox for 24 hours. B) Inducible cells were transfected with siRNA targetting hRNF126 or a non-silencing control (NS) for 48 hours and then stimulated with 1 ug/mL dox and 340 uM OA for 24 hours. A single Z-section is shown. Cells were stained for RNF126 (red) and BODIPY 493/503 (green). C) and D) Quantification of average LD volume (C) and average LD numbers (D) from (B). N = 3 with at least 20 cells measured per experiment. All values are mean ± s.e.m. ns is not-significant, * p < 0.05, ** p < 0.01, *** p < 0.001, Scale bars: 5 µm.

3.3.3 RNF126 does not regulate lipid droplet turnover

The decrease in LD size could be due to defective LD growth or enhanced LD turnover. Since UBXD8 and p97 have previously been shown to inhibit the lipolytic enzyme ATGL by segregating its co-activator CGI-58(Olzmann et al., 2013), we initially hypothesized that knockdown of RNF126 may result in over-active ATGL and greater LD turnover. To test this, we took three experimental approaches. First, we treated OA stimulated cells with the lipase inhibitor DEUP to block LD turnover. If lipase activity is increased, we predicted that inhibition of lipase activity would rescue LD size in RNF126 knockdown cells. DEUP treatment increased LD volume in both control and knockdown cells relative to non-DEUP treated cells (Figure 3-4 A). However, DEUP treatment was unable to rescue LD volume as knockdown cells continued to have significantly smaller LDs even after DEUP treatment (Figure 3-4A). Analysis of the fold change in volume between DEUP treated and untreated cells shows that both knockdown and control LDs increase in size with similar magnitude after lipase inhibition (Figure 3-4 B). These results suggest that increased lipase activity is not the cause of small LDs in RNF126 knockdown cells. This conclusion is further supported by biochemical analysis of purified LDs from knockdown and control cells. Protein abundance of LD associated ATGL and its co-activator CGI-58 are not different in RNF126 knockdown cells compared to controls (Figure 3-4 C and D). We also performed a co-IP experiment to indirectly examine the activity of ATGL by assessing the amount of associated CGI-58. In support of our previous conclusions, a preliminary experiment showed that similar amounts of ATGL co-IP’d with CGI-58 in knockdown cells and controls (Figure 3-4 D). In addition to enzymatic lipolysis, it is known that LD turnover can also occur through alternate pathways such as autophagic degradation of LDs. To test if an alternate turnover pathway is accelerated, we performed a pulse-chase assay to monitor LD turnover. Cells were first pulsed with OA to induce the growth of LDs and then chased in Triacsin C, a drug that prevents TG synthesis(Tomoda et al., 1987), to promote LD turnover. After monitoring regression of LDs over time, we observed that LD volume and number from both knockdown and control cells decreased at an equal rate (Figure 3-4 E and F). Altogether, these data demonstrate that decreased LD size is not due to enhanced LD turnover.

Figure 3-4. Knockdown of RNF126 does not accelerate LD turnover. A) RNF126 knockdown and control cells were stimulated with OA in the presence of absence of the lipase inhibitor DEUP for 24 hours. Average LD volume was quantified. B) Quantification of the fold change of LD volume when cells are treated with DEUP. LD volume from DEUP treated cells were normalized to LD volume from untreated cells. C) RNF126 knockdown and control cells were stimulated with OA for 24 hours and LDs isolated. Two LD purifications (LD1 and LD2) were loaded and the lysate corresponds to LD2. LD1 (unquantified), LD2 (8 ug of protein). Immunoblot was performed with the indicated antibodies. D) LDs from RNF126 knockdown and control cells were immunoprecipitated with antibodies against CGI-58 and immunoblotted. 2.8 ug of LD protein was loaded and 10 ug of protein was used for IP, N = 1. E) and F) Pulse-chase experiment to assay LD turnover in RNF126 knockdown and control cells. LD size and number were normalized against the 0 hour time point. All values are mean ± s.e.m with N = 3 and at least 20 cells measured per experiment unless indicated. ns is not-significant, * p < 0.05, ** p < 0.01, *** p < 0.001.

3.3.4 RNF126 is required for lipid droplet expansion

In addition to enhanced turnover, smaller LDs can also result from defective LD biogenesis or expansion. UBXD8 has previously been reported to inhibit synthesis of TG during low lipid conditions by inhibiting conversion of DG to TG(Lee et al., 2010). De-regulation of this process could result in smaller LDs due to decreased synthesis of TG. To test if LD biogenesis or expansion is disrupted, RNF126 knockdown and control cells were first pulsed with Triacsin C to deplete all existing LDs. Cells were then chased with OA and LD numbers and volume measured at different time points. To assay biogenesis and LD growth without the added complexity of LD turnover, some cells were treated with DEUP to inhibit lipolysis of LDs. As expected, RNF126 knockdown cells continued to have significantly smaller LDs than control cells regardless of lipase inhibition after 24 hours of OA (Figure 3-5 A and B). LD numbers increased with time and were unchanged between knockdown and control cells, except for a slight difference when cells were treated for 24 hours with OA and DEUP (Figure 3-5 C and D). As cellular LDs have previously been depleted, LD biogenesis can be observed at early time points. After 4.5 hours, RNF126 knockdown cells had significantly smaller LDs than controls (Figure 3-5 B). Surprisingly, this phenotype was rescued after treatment with DEUP (Figure 3-5 B). Although statistically significant, this difference may not be biologically significant as DEUP treatment is unable to rescue LD size at any other time point (Figure 3-5 A and B). More importantly, the similarities in LD volume and numbers after DEUP treatment at 4.5 hours (Figure 3-5 B and D) suggests that biogenesis of LDs is unaffected by the depletion of RNF126.

While control LDs continued to increase in volume with time, growth of knockdown LDs was minimal (Figure 3-5 A). Though DEUP treatment restored LD expansion, the volume of LDs in DEUP treated knockdown cells was significantly smaller than control cells starting at 9 hours and continuing to 24 hours (Figure 3-5 A and B). Since LD turnover is not a factor, this data further suggests that LDs in knockdown cells increase in size slower than control cells. As biogenesis is unaffected, the data indicates that expansion of pre-formed LDs is disrupted by knockdown of RNF126.

Figure 3-5. RNF126 is required for lipid droplet expansion. RNF126 knockdown and control cells were depleted of LDs with an overnight treatment of Triacsin C. Cells were then stimulated with OA in the presence or absence of DEUP and LD size and number quantified. A) Progression of LD growth over time. B) Statistical analysis of (A). B) Progression of LD numbers per cell over time. D) Statistical analysis of (C). All values are mean ± s.e.m with N = 3 and at least 20 cells measured per experiment, 12 hour timepoint N = 4. ns is not-significant, * p < 0.05, ** p < 0.01, *** p < 0.001.

3.4 Discussion

Recent work has focused heavily on understanding the mechanisms regulating LD turnover(Kaushik and Cuervo, 2015; Olzmann et al., 2013; Rambold et al., 2015). However, our understanding of how LDs form and expand requires additional research. Here, we provide evidence supporting a role for RNF126 in the regulation of LD size. Depletion of RNF126 decreases LD size following stimulation with the unsaturated FA oleate. This defect is specific to RNF126 as re-expression of mouse RNF126 is sufficient to restore LD size. Although transient knockdown of RNF126 by siRNA increased LD numbers, this is likely a non-specific effect of the siRNA as re-expression of mouse RNF126 could not rescue LD numbers. To confirm, additional siRNA targeting alternate regions of RNF126 should be tested. We are also interested in understanding the role of ubiquitin ligase activity. Due to technical difficulties involving over- expression of RNF126 in our stable knockdown cell lines, we have been unable to test the role of ubiquitin ligase activity. Generation of additional doxycycline inducible stable cell lines expressing the ligase dead mutant form of RNF126 may provide information on the role of ubiquitin. Additionally, as HEK293T and HEPG2 cells appear to not be noticeably affected by over-expression of RNF126 (Data not shown), knockdown and rescue experiments performed in alternate cell lines may be an attractive option.

While we have shown that OA stimulation in RNF126 knockdown cells leads to smaller LDs, LD morphology under resting conditions have not been quantified. This is due to technical challenges separating weak LD signal from background signal. Furthermore, LDs in HeLa cells under resting conditions are very small and may be difficult to resolve accurately. A potential approach could be to perform electron microscopy to increase resolution or to measure intracellular TGs with a more sensitive method such as thin-layer chromatography.

Analysis of the size distribution of LDs found that knockdown of RNF126 decreases the abundance of larger LDs while increasing the abundance of small LDs. This may be due to enhanced turnover of large LDs or a failure of smaller LDs to grow larger. Although UBXD8 and p97 have been shown to regulate lipolysis, our data suggests that RNF126 does not play a role in LD turnover(Olzmann et al., 2013). Instead, depletion of RNF126 inhibits LD growth without affecting biogenesis of LDs. Since continual LD expansion requires a re-distribution of TG synthesizing machinery from the ER to LDs, a potential regulatory mechanism may involve

RNF126(Wilfling et al., 2013). The rate limiting enzyme of TG synthesis, DGAT2, is one of these proteins that localizes to LDs during LD expansion. Since depletion of DGAT2 prevents formation of large LDs, RNF126 may be involved in regulating the function of DGAT2 at LDs.

In addition to its role in lipolysis, UBXD8 has been shown to regulate TG synthesis under lipid depleted conditions(Lee et al., 2010). Thus, it appears that UBXD8 plays a dynamic role in lipid homeostasis by promoting lipid storage in LDs under lipid rich conditions and re-directing free lipids to be used in alternate pathways under lipid depleted conditions. Although we have shown that RNF126 ubiquitinates UBXD8 and regulates LD size, the role of UBXD8 in this phenotype is unknown. One possibility is that RNF126 regulates recruitment of p97 to UBXD8 at specific organelles. Future work will need to determine if RNF126 regulates LD size through UBXD8 and p97 and the mechanism involved.

Chapter 4 Conclusions and Future Directions 4.1 Summary

The goal of this work was to further characterize the biological functions of RNF126 and RNF115 by identifying novel interacting partners and substrates. Using a proximity based screening approach, we identified novel connections with cellular components involved in a broad array of cell biology. Further validation experiments demonstrated the effectiveness of this screening approach and uncovered an intimate relationship between RNF126 and the p97 family of co-factors. In addition, several proteins were found to be substrates of RNF126 including the co-factor UBXD8. UBXD8 has been shown to regulate LD homeostasis and we found that knockdown of RNF126 was sufficient to decrease LD size but not number following fatty acid stimulation. Further evidence suggested that smaller LDs are due to defective LD expansion. Though these data have uncovered a novel function of RNF126, the mechanism and the role of UBXD8 remains unknown. In this chapter, questions that have arisen from this work will be discussed.

4.2 Regulation of p97 co-factors by RNF126

The diverse functions of p97 are tightly controlled by a host of regulatory co-factors. Importantly, mutations in human p97 can cause a fatal protein aggregation disease called inclusion body myopathy with Paget disease of bone and/or frontotemporal dementia (IBMPFD)(Nalbandian et al., 2011). Furthermore, disease mutant forms of p97 have altered co- factor binding and can disrupt autophagy and endolysosomal trafficking(Ju et al., 2009; Ritz et al., 2011). Thus, it is important to understand the conditions that regulate how p97 selects its co- factors. Currently, little is known about the cellular mechanisms that regulate p97 complex assembly.

We have found that RNF126 is associated with p97 and a diverse group of its co-factors. RNF126 associates with both UBXD1 and UBXD8 and may also associate with the heterodimer UFD1-NPL4 and UBXN1 (Figure 4-1 A). This relationship also appears to be specific as preliminary data suggests that RNF126 associates weakly with the UBX protein p47 (Figure 4-1 A). In addition, RNF126 was found to ubiquitinate UBXD1 and UBXD8 and potentially

UBXN1. Co-IP of UBXN1 and RNF126 revealed a ladder of larger molecular forms of UBXN1 (Figure 4-1 A). These results are similar to those observed with UBXD1 and UBXD8 that were subsequently confirmed to be bonafide substrates. It will be interesting to further explore the relationship between RNF126 and the UBX family. To this end, we assembled UBX family expression constructs in order to examine which proteins RNF126 associates with and which are substrates. Though we previously hypothesized that RNF126 may bind the UBX domain through its ubiquitin binding zinc finger, it appears that this may not be the case. Mutation of the zinc finger did not abolish interaction with isolated GST-UBX domains from UBXD8 and UBXN1 (Figure 4-1 B). This agrees with our co-IP data where the zinc finger was not required for association with UBXD8. Instead, UBXD8 may bind ubiquitinated RNF126 primarily through its UBA domain. UBXN1 may also adopt a similar mode of interaction as it contains a UBA domain as well. Although, we found that the zinc finger is required for association with UBXD1, we have been unable to generate GST fusion proteins of its UBX domain for testing. As the UBX domain of UBXD1 is unique(Kern et al., 2009), future experiments using the isolated UBX domain may determine if the zinc finger can specifically recognize the UBX domain of UBXD1. It will be important to continue characterizing how RNF126 interacts with other UBX family proteins to determine if there is a conserved mode of interaction. Of particular interest will be understanding how RNF126 interacts with proteins like UBXD1 which do not contain UBA domains. These experiments will characterize the relationship of RNF126 and UBX proteins and may provide insight into additional functions of RNF126. For example, UBXN1 has been shown to regulate the ubiquitin ligase activity of BRCA1(Wu-Baer et al., 2010).

An obvious question remaining is the role of RNF126 mediated ubiquitination of p97 co-factors. Since co-expression of RNF126 did not alter the abundance of any associating proteins tested, it is likely that RNF126 does not promote the ubiquitin dependent degradation of p97 co-factors. Instead, our data leads us to believe that RNF126 mediated ubiquitination may promote binding between UBXD8 and p97. As this regulatory mechanism has not been demonstrated with this family of proteins, it will be interesting to determine if UBXD8 and p97 binding is altered by ubiquitin and if this mechanism is conserved with other UBX proteins. UBX proteins are also known to select ubiquitinated cargo for degradation through p97(Alexandru et al., 2008). Thus, UBXD8 may bind ubiquitinated RNF126 through its UBA domain and target it for degradation. Co-expression of RNF126 with UBXD8 or an empty vector control may indicate if UBXD8 has

64 a role in regulating RNF126 stability. We could also determine the ubiquitin chain topology that is assembled on UBXD8 by RNF126. Ubiquitin chains of different topology have defined functional roles, for example, K48 chains are well known mediators of proteasomal degradation(Woelk et al., 2007). This can be examined by performing ubiquitination assays with exogeneously expressed HA tagged KtoR ubiquitin mutants in vivo. If mutation of a lysine residue on ubiquitin prevents ubiquitination of UBXD8, this would suggest that RNF126 mediates the formation of specific ubiquitin chains. It would also be interesting to test if UBXD8 is a direct substrate of RNF126 though in vitro ubiquitination assays. These experiments may provide clues on a potential role for RNF126 mediated ubiquitination of UBXD8.

After assembly of a core p97 complex, additional co-factors can be recruited to provide greater substrate specificity or additional enzymatic activities(Meyer and Weihl, 2014). To test if RNF126 has a role in regulating the assembly of higher order p97 complexes, a proteomics approach would be optimal. Through affinity purification mass spectrometry analysis of p97 interacting partners in RNF126 knockdown and over-expressing cells, we can characterize how p97 complexes change relative to RNF126 expression. These experiments may uncover a novel regulatory mechanism governing p97 complex assembly.

Figure 4-1. Characterizing the interaction between RNF126 and the UBX family. A) HEK293T cells were co-transfected with untagged RNF126 or empty vector and UBXD1-V5, UBXN1-V5, or p47-V5. IPs were performed with anti-V5 antibodies and immunoblotted, N = 1. B) HEK293T cells were transfected with HA-RNF126 or the HA tagged zinc finger mutant construct and cell lysates incubated with GST, or GST-UBX proteins from UBXD8 and UBXN1 and immunoblotted. Below is a Fast Green stain of the membrane showing fusion proteins, N = 1.

4.3 The role of RNF126 mediated ubiquitination of KEAP1 and CUL3

The most abundant hit from the BioID screen was the autophagy adapter protein p62. Although we found that p62 associates with RNF126, p62 is not a substrate and mapping the interaction has been technically challenging (Data not shown). BioID experiments also suggested that KEAP1 may be a candidate interacting protein of RNF126. Since p62 has been shown to mediate the lysosomal degradation of KEAP1 by autophagy in what is likely an ubiquitin dependent manner, we reasoned that KEAP1 might be a substrate of RNF126 (Lau et al., 2010; Taguchi et al., 2012). Although it received a weak SAINT score, it is possible that BioID with lysosomal inhibition will enrich KEAP1 peptides sufficiently to become statistically significant.

KEAP1 acts as the substrate adaptor for a cullin-RING ubiquitin E3 ligase complex, CUL3- RBX1, and its best characterized role is the poly-ubiquitination of the antioxidant response protein NRF2(Bryan et al., 2013; Hayes and Dinkova-Kostova, 2014). NRF2 serves as the master regulator of the antioxidant response in cells, regulating approximately 200 genes involved in cytoprotection, metabolism, and transcription(Hayes and Dinkova-Kostova, 2014). Due to this role, KEAP1 has become a very attractive drug target. In the canonical mechanism, cells mediate the constant proteasomal degradation of NRF2 through ubiquitination by KEAP1- CUL3. The presence of electrophiles or reactive oxygen species is sensed by KEAP1 and impairs the ubiquitination and degradation of NRF2, leading to translocation of NRF2 into the nucleus and activation of target genes(Harder et al., 2015). The non-canonical mechanism occurs via p62 disruption of the KEAP1-NRF2 interaction through competitive binding and mediating the lysosomal degradation of KEAP1. This pathway is thought to occur slower than the canonical pathway and believed to prolong the antioxidant response of NRF2(Jiang et al., 2015).

We found that RNF126 associates with KEAP1 through its RING domain and that KEAP1 is a bonafide substrate (Figure 4-2 A and B). Auto-ubiquitination of KEAP1 by the CUL3 complex has been shown to mediate the non-proteasomal degradation of KEAP1 and to activate NRF2(Zhang et al., 2005). Thus, it may be interesting to explore if RNF126 mediated ubiquitination of KEAP1 regulates the interaction of KEAP1 and NRF2. Since NRF2 activates the antioxidant response, we could first test if RNF126 depleted cells are more resistant to oxidative insults than controls. If so, we could test whether co-IP of KEAP1 and NRF2 is altered

68 by the depletion or over-expression of RNF126 and if this correlates with decreased ubiquitination of NRF2. These experiments may lead to the development of novel therapeutics targeting RNF126 as a way to regulate NRF2 signaling.

Outside of its role in targeting NRF2 to the proteasome, KEAP1-CUL3 can inhibit NF-κB signaling by ubiquitinating IKKβ and targeting it for proteasomal degradation(Lee et al., 2009). This suggests that KEAP1-CUL3 can ubiquitinate additional substrates and may participate in other cellular functions. In light of this, we wanted to determine if ubiquitination of KEAP1 by RNF126 affected assembly of the KEAP1-CUL3-RBX1 E3 ligase complex. Intriguingly, co- expression of RNF126 and CUL3 resulted in a distinctive banding pattern resembling ubiquitination (Figure 4-2 C). To test if RNF126 could associate with CUL3, we performed co- IP experiments and found that RNF126 immunoprecipitates were highly enriched with modified forms of CUL3 (Figure 4-2 C). Notably, mutation of either the zinc finger or RING domain of RNF126 did not abolish association with CUL3 although mutation of the zinc finger dramatically altered the banding pattern of CUL3 (Figure 4-2 C). As we have previously found the zinc finger to specify ubiquitin chain topology, the zinc finger may also control the ubiquitin topology built on CUL3(Smith et al., 2013). Further ubiquitination experiments suggested that CUL3 is indeed a substrate of RNF126 (Figure 4-2 D). While cullin-E3 ligases are known to be activated by neddylation, it has also been demonstrated in vitro that mono-ubiquitination is sufficient for activation(Duda et al., 2008). Accordingly, it has been reported that CUL3 is ubiquitinated on residue K414 although the E3 ligase is unknown(Wimuttisuk et al., 2014). Thus, it would be interesting to test if mutation of CUL3 at K414 can disrupt ubiquitination by RNF126. As it appears that RNF126 assembles poly-ubiquitin chains on CUL3, the modification is likely not serving as an activation signal. Interestingly, preliminary data suggests that KEAP1 may compete with RNF126 for binding to available CUL3 (Figure 4-2 E). When transfected amounts of Flag-CUL3 and RNF126 are kept constant, increasing amounts of Myc-KEAP1 results in loss of RNF126 from CUL3 IPs (Figure 4-2 E). This may result from KEAP1 abundance reaching a critical level and out-competing RNF126 for association with CUL3. This is of interest as RNF126 may function as a novel substrate adapter for CUL3. It will be interesting to explore the role of RNF126 mediated ubiquitination in CUL3 and KEAP1 functions and the mechanisms involved.

Figure 4-2. RNF126 may associate and ubiquitinate KEAP1 and CUL3. A) HEK293T cells were co-transfected with untagged RNF126 or mutant construct and Myc-KEAP1. IPs were done with anti-RNF126 antibodies and immunoblotted, N = 1. B) HEK293T cells were co-transfected with untagged RNF126 or mutant construct, HA-ubiquitin, and Myc-KEAP1. Cells were lysed in boiling hot RIPA buffer containing 1% SDS, diluted 10 fold, and IPs done with anti-myc antibodies. IPs were immunoblotted for HA-ubiquitin, N = 2. C) HEK293T cells were co- transfected with untagged RNF126 or mutant construct and 3x Flag-CUL3. IPs were done with anti-RNF126 antibodies and immunoblotted, N = 1. D) HEK293T cells were co-transfected with untagged RNF126 or mutant construct, HA-ubiquitin, and 3x Flag-CUL3. Cells were lysed in boiling hot RIPA buffer containing 1% SDS, diluted 10 fold, and IPs done with anti-Flag antibodies. IPs were immunoblotted for HA-ubiquitin, N = 1. E) HEK293T cells were co- transfected with equivalent amounts of untagged RNF126, Flag-CUL3, and either empty vector or increasing amounts of myc-KEAP1. Total DNA transfected was made up to equivalent amounts with empty vector. IPs were done using anti-Flag antibodies and immunblotted, N = 1.

4.4 UBXD1, p97, and RNF126 in EGFR trafficking

We have shown that RNF126 associates with UBXD1 through its ubiquitin binding zinc finger and that UBXD1 is a bonafide substrate. Although the functions of UBXD1 are poorly defined, UBXD1 has been shown to play a role in endolysosomal trafficking(Ritz et al., 2011). UBXD1 recruits p97 to mono-ubiquitinated CAV1 to promote trafficking of CAV1 to lysosomes. Additionally, inhibition of p97 activity resulted in delayed lysosomal degradation of the EGFR, similar to the phenotype described in RNF126 knockdown cells(Ritz et al., 2011; Smith et al., 2013). Thus, exploring a potential role for RNF126 in this pathway is very attractive.

The role of UBXD1 in EGFR trafficking is unclear. Thus, we would have to first examine if EGFR trafficking is regulated by UBXD1. This could be done by depleting cells of UBXD1, stimulating with EGF ligand, and then assaying by western blot if degradation of the EGFR is delayed. If UBXD1 indeed has a role, we could re-express UBXD1 mutants that do not bind RNF126 or are not ubiquitinated in UBXD1 depleted cells. If these mutants fail to rescue the trafficking of the EGFR, this may suggest that RNF126 functions with UBXD1 and p97 to regulate EGFR trafficking. We have previously proposed a model where RNF126 binds ubiquitinated EGFR and modifies the ubiquitin chains to promote sorting to the lysosome (Christopher Smith, Ph.D thesis). A potential mechanism may involve UBXD1 binding to RNF126 to bias p97 towards substrates of active E3 ligases. Following binding of UBXD1 to RNF126, ubiquitination of UBXD1 may function as a sensor for ligase activity and recruit p97 to the EGFR. Additionally, UBXD1 may regulate the ligase activity of RNF126 to specify the ubiquitin chain topology to specifically recruit p97. UBXD7 functions in a similar manner by restricting ubiquitin chain length on HIF1α to specifically recruit p97 and reduce ubiquitin receptor promiscuity(Bandau et al., 2012). To determine if this may be a general trafficking mechanism, we could test if trafficking of GFP-CAV1 to the lysosome is disrupted in RNF126 knockdown cells and if this process also requires UBXD1 and p97. These experiments may further describe the molecular mechanisms of RNF126 mediated trafficking of membrane proteins.

4.5 The role of RNF126 mediated ubiquitination of UBXD8 in lipid droplet regulation

The finding that UBXD8 is ubiquitinated by RNF126 prompted us to explore the role of RNF126 in lipid droplet biology. Our data suggests that depletion of RNF126 decreases LD volume but does not have an effect on LD numbers. This phenotype can be rescued by re-expression of RNF126 although we have been unable to test if ligase activity is required. To test the role of ligase activity, we could generate additional inducible cell lines that stably express the ligase dead mutant of RNF126. A previous report showed that UBXD8 inhibits lipolysis, however we found that depletion of RNF126 did not affect LD turnover(Olzmann et al., 2013). Instead, depletion of RNF126 inhibited LD expansion without noticeably disrupting LD biogenesis. Although these findings have established a novel function for RNF126 in LD homeostasis, the mechanistic details of how RNF126 regulates LD size is unclear. Specifically, whether RNF126 affects LD size through ubiquitination of UBXD8 is unclear. To determine if UBXD8 function is regulated by RNF126, we can take advantage of the specificity of UBXD8 to unsaturated FAs(Lee et al., 2010). As UBXD8 does not respond to saturated FAs, we predict that stimulation of RNF126 knockdown cells with saturated FAs may show no difference in size when compared to controls. Further experiments could over-express and knockdown UBXD8 in RNF126 depleted cells as well as our inducible over-expressing cell lines and examine LD size and number.

One potential mechanism may involve regulation of UBXD8 sub-cellular localization by RNF126 mediated ubiquintation. For example, ubiquitinated UBXD8 could be prevented from localizing to LDs by being sequestered at the ER by the UBA domain containing protein UBAC2(Olzmann et al., 2013). As p97 is recruited to LDs through UBXD8(Suzuki et al., 2012), we can test if UBXD8 or p97 distribution is altered by immunostaining OA stimulated RNF126 knockdown cells with anti-UBXD8 and anti-p97 antibodies. UBXD2 has also been found on LDs and may recruit p97 as well(Suzuki et al., 2012). Thus, if UBXD8 and p97 localization to LDs is unaffected by depletion of RNF126, it is possible that UBXD8-p97 interaction at LDs is regulated by RNF126. We can test this by proximity ligation assay (PLA), a technique used to identify protein-protein interactions in situ by immunofluorescence(Soderberg et al., 2006). Through this assay, we can test if UBXD8-p97 association at LDs is altered by depletion of RNF126 by quantifying the difference in fluorescence between control and knockdown cells.

Additionally, PLA of UBXD8 and p97 can be examined in our doxycycline inducible cell lines. Another possible regulatory mechanism may occur through selection of p97 substrates by UBXD8. Ubiquitination of UBXD8 by RNF126 may regulate protein-protein interactions and promote selection of specific substrates for removal by p97. This may occur by inhibiting polymerization of UBXD8, which has previously been shown to inhibit the TG synthesis inhibitory function of UBXD8(Lee et al., 2010). In a similar manner, ubiquitination of AXIN disrupts its interaction with Wnt co-receptors LRP5/6 which attenuates Wnt signalling(Fei et al., 2013). Since UBXD8 is a membrane protein, we could explore if UBXD8 substrate specificity is regulated by RNF126 by performing BioID and analyzing UBXD8 interacting proteins while modulating RNF126 expression. These experiments may begin to elucidate the mechanisms of how RNF126 and UBXD8 regulate LD size.

4.6 RNF126 and other lipid droplet regulatory proteins

Conversion of diacylglyceride to TG, the final step of TG synthesis, is catalyzed by either DGAT1 or DGAT2. Although they have similar functions, they are structurally and evolutionarily unrelated. While both DGAT proteins reside in the ER, only DGAT2 can localize to LDs(Wilfling et al., 2013). DGAT2 appears to be responsible for the majority of TG synthesis as DGAT1 knockout mice are healthy with a mild decrease in overall TG levels while DGAT2 knockout mice die shortly after birth and have drastically reduced TG levels(Stone et al., 2004). On a cellular level, DGAT1 appears to regulate formation of small LDs while DGAT2 is required for formation of large LDs(Wilfling et al., 2013). DGAT2 is thought to mediate LD expansion by re-localizing to LDs in a large complex with other TG synthesizing enzymes to mediate local synthesis of TG at LDs(Wilfling et al., 2013). DGAT2 has also recently been shown to be ubiquitinated and degraded by the proteasome(Jin et al., 2014). Studies in animal models have found that knockdown of DGAT2 can reduce liver TG content and body weight, making DGAT2 an attractive therapeutic target to combat obesity(Choi et al., 2007). In addition to its role as an inhibitor of lipolysis, UBXD8 has also been implicated as a potential regulator of TG synthesis(Lee et al., 2010). Knockdown of UBXD8 promotes conversion of DG to TG, suggesting that UBXD8 may inhibit a DGAT enzyme under low fatty acid conditions(Lee et al., 2010). Since the depletion of RNF126 also inhibits TG expansion, it may be worth investigating whether RNF126 regulates other proteins involved in TG expansion. Of particular interest is DGAT2 as it may be regulated by UBXD8 and is essential for LD expansion.

To test if RNF126 may potentially regulate DGAT2, we first co-expressed RNF126 and Flag- tagged DGAT2 and examined DGAT2 abundance. Interestingly, co-expression of wild-type RNF126 increases Flag-DGAT2 abundance and this may require the ubiquitin ligase activity of RNF126 (Figure 4-3 A and B). This may suggest that RNF126 can stabilize DGAT2 by degrading an E3 ligase or through a direct ubiquitination dependent stabilization mechanism. Indeed, ubiquitination of the oncoprotein Myc has been shown to stabilize the protein(Popov et al., 2010). Preliminary experiments suggest that RNF126 can associate with DGAT2 (Figure 4-3 B). Furthermore, RNF126 IPs contain highly modified forms of DGAT2, suggesting that DGAT2 may also be ubiquitinated by RNF126 (Figure 4-3 B). Subsequent ubiquitination assays have proven to be technically challenging due to the low expression of transfected Flag-DGAT2 and the quick turnover rate(Choi et al., 2014). To attempt to enrich for DGAT2, transfected cells were treated with MG132 for 6 hours before performing ubiquitination assays. A promising preliminary experiment suggests that RNF126 can indeed ubiquitinate DGAT2, although signal was very weak (Figure 4-3 C). Thus, it is tempting to speculate that RNF126 regulates LD size by ubiquitinating DGAT2 to promote its proteasomal degradation. To date, attempts to detect endogenous DGAT2 or Flag-DGAT2 abundance in RNF126 knockdown cells by western blot have been unsuccessful. Additionally, expression of Flag-DGAT2 in HEK293T cells is very low and has prevented cycloheximide chase assays to test the role of RNF126 on DGAT2 stability. As other groups have been able to assay DGAT2 half-life, it is possible that we have not determined the optimal conditions for this assay(Choi et al., 2014). Indeed, we have found that a polyclonal antibody targeting DGAT2 is more efficient at detecting Flag-DGAT2 than a Flag monoclonal antibody (Data not shown). If RNF126 is indeed regulating LD size by promoting stabilization of DGAT2, then over-expression of DGAT2 should rescue LD size in RNF126 knockdown cells. We can test this by over-expressing DGAT2 in our RNF126 knockdown cells and assaying LD morphology. UBXD8 may also play a role in this mechanism. ER associated DGAT2 has been shown to associate with p97 and this association is enhanced when proteasomal degradation of DGAT2 was inhibited(Jin et al., 2014). Since p97 is actively involved in extracting proteins from membranes for proteasomal degradation, RNF126 may have a role in targeting UBXD8 and p97 to DGAT2. UBXD8-DGAT2 association could be assayed in OA stimulated RNF126 knockdown cells to determine if RNF126 has a role in regulating this interaction. Together, these experiments may provide further insight into the mechanism by which RNF126 regulates LD size.

Figure 4-3. RNF126 may ubiquitinate DGAT2 to enhance stability. A) HEK293T cells were co-transfected with either empty vector or untagged RNF126 or a mutant construct and Flag- DGAT2. Cells were lysed in NP40 lysis buffer and subjected to immunoblot, N = 2. B) HEK293T cells were co-transfected with HA-RNF126 or a HA tagged mutant construct and Flag-DGAT2. IP was performed using anti-HA and anti-Flag antibodies and immunoblotted, N = 1. C) HEK293T cells were co-transfected with untagged RNF126 or mutant construct, HA- ubiquitin, and Flag-DGAT2. Cells were treated with 10 µM MG132 for 6 hours and lysed in boiling hot RIPA buffer containing 1% SDS. Following sonication and another boiling step, lysates were diluted 10 fold in NP40 lysis buffer and Flag-DGAT2 immunoprecipitated and analyzed by immunoblot, N = 1.

4.7 RNF126 regulation of lipid droplets beyond HeLa cells

Lipid droplets are ubiquitous organelles with diverse roles across all cell types. Our current work has focused on the role of RNF126 in lipid droplet biology in HeLa cells. Although HeLa cells are attractive cell biology models due to being easily manipulated, it will be important to understand the role of RNF126 in a more physiologically relevant cell model. As the majority of an organism’s lipids are stored in adipocytes, it will be interesting to determine if RNF126 has a role in regulating LD size in adipocytes. This could be achieved by differentiating 3T3-L1 cells into adipocytes, depleting RNF126 by siRNA, and assaying LD size and number. Since 3T3-L1 adipocytes have large LDs under resting conditions, we may be able to quantify LDs under resting and OA stimulated conditions. We can also test the role of RNF126 in specialized lipid transporting cells such as liver and intestinal cells. In addition, UBXD8 has been shown to regulate lipolysis in the liver cell line HUH7 as well as regulate lipoprotein assembly(Olzmann et al., 2013; Suzuki et al., 2012). Although preliminary experiments in HUH7 suggested that depletion of RNF126 does not affect LD size in hepatocytes, the role of RNF126 in enterocytes remains to be investigated. One commonly used cell line that we could test is CACO2. Other cell types of interest are cardiac and skeletal muscle cells. As excessive accumulation of TG in the heart can cause death, conversion of lipotoxic intermediates such as DG to TG can prevent heart failure(Liu et al., 2009). Thus, build-up of TGs can prevent heart damage by limiting the amount of reactive DG available. Similarly in skeletal muscle, increased TG synthesis improved insulin sensitivity, suggesting that lipid conversion to TG may help prevent development of diabetes(Bosma et al., 2012). We can study the role of RNF126 in these cell types by using HL-1 mouse cardiomyocytes and differentiating C2C12 myoblasts into skeletal muscle cells. These experiments will begin to elucidate the physiological role of RNF126 in regulating lipid homeostasis.

4.8 Concluding remarks

The ubiquitin system plays an essential role in almost all aspects of cell biology. Although we now realize the sheer abundance of ubiquitin E3 ligases, characterization of their functions has been lacking. We have used a novel proximity based protein screen to identify several substrates of the E3 ligase RNF126. Furthermore, we have established a novel role for RNF126 in the regulation of lipid droplet size. With the growing global obesity epidemic, understanding the mechanisms of lipid storage and mobilization will be critical to treatment of obesity and co- morbidities. Thus, future work may identify RNF126 as an attractive therapeutic target to lower intracellular lipid content. Additionally, the identification of other substrates of RNF126 suggests that novel functions remain to be explored. This work has shown the effectiveness of BioID to identify substrates of E3 ligases and furthers our understanding of the role of ubiquitin in lipid biology.

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Appendix I – SAINT analyzed RNF126 HeLa BioID results

Gene Average spectra count without Average spectra count with MG132 MG132 BAG6 21.5 4.0 UBL4A 5.3 4.5 SQSTM1 60.5 163.5 EPPK1 6.8 6.8 ANXA2 62.4 78.8 PLEC 10.4 10.5 PLOD1 6.2 8.0 PDIA3 15.0 11.8 RAB2A 8.6 11.5 CPS1 128.6 150.5 ANXA1 20.6 20.5 ASS1 22.7 15.0 HIST1H1B 17.4 12.8 SLC16A3 4.6 9.3 CLU 6.0 9.3 RAP1A 5.5 6.5 PLA2G4A 9.3 6.5 NPLOC4 4.5 16.8 ZFAND5 5.0 UFD1L 3.3 30.0 SLC3A2 7.0 21.3 ACOT9 2.4 4.3 LOC100510073 4.0 3.0 SERPINH1 29.6 40.0 TRIML2 13.0 3.3 LMNA 16.4 20.5 NSA2 1.6 3.0 HMGCR 3.0 GSTP1 5.8 7.3 RAB7A 8.0 10.3 SLC7A5 4.5 6.5 PSMD4 8.3 12.0 PLAA 1.0 2.7 SLC38A2 1.0 2.3 SPCS3 2.7 2.5

MTHFD1L 2.9 2.0 CLPTM1L 2.1 2.0 SEC11A 1.3 2.0 RAB6A 4.3 5.3 LGALS1 2.5 2.0 LMO7 2.5 2.0 PLP2 2.1 2.0 CYP51A1 2.5 2.0 PTDSS1 2.0 G6PD 3.0 4.3 NNT 2.8 4.0 AFG3L2 6.8 2.0 FSCN1 6.6 3.8 TPM2 7.0 6.0 C19orf21 8.5 2.5 DAB2 4.6 1.7 LIMCH1 8.1 1.0 RGPD1 20.3 RAB32 7.5 NUP98 4.0 1.7 CBX5 3.0 1.0 CENPE 4.0 1.0 SEC23A 2.8 PSMB6 2.5 2.0 PSMB7 2.5 2.5 HMGCS1 2.5 1.5 PSMB1 2.0 1.0 MUC13 2.3 DDX24 1.8 1.5 YIF1A 1.5 1.7 CSRP1 1.8 1.0 CALU 2.0 1.0 SDHA 3.3 1.5

Appendix II – SAINT analyzed RNF126 HEK293 BioID results

Gene Average spectra count without Average spectra count with MG132 MG132 BAG6 3.5 53.0 UBL4A 3.5 13.0 SQSTM1 19.0 126.5 NPLOC4 15.5 UFD1L 1.5 37.0 ZFAND5 4.5 HMGCR 4.5 EPS15 6.5 48.0 UBB 39.5 437.5 VCPIP1 29.0 53.0 FAM21A 37.0 MAGED1 33.0 PJA2 6.5 GGA3 7.5 PSMA1 8.0 UBXN1 6.5 N4BP1 3.0 16.5 MAGED2 9.5 TBK1 4.0 TNIP1 4.0 LOC100510073 4.5 NBR1 3.5 POLR2B 4.0 PSMA6 3.5 CALR 1.0 4.5 CNOT10 4.5 SMG9 7.5 TRAFD1 3.5 PSMB1 2.5 NME2 32.0 19.5 HGS 2.0 NSA2 2.0 SPAG5 2.0 UBXN2A 2.0 TOM1L1 2.0

ATIC 4.5 5.5 LOC652147 6.5 3.0 PDZRN3 36.5 3.0 RYR2 9.0 2.0 KLC2 5.0 TNRC6B 9.0 6.5

Appendix III – SAINT analyzed RNF115 HEK293 BioID results

Gene Average spectra count without Average spectra count with MG132 MG132 BAG6 5.0 32.5 UBL4A 1.5 6.3 SQSTM1 6.3 93.8 NPLOC4 28.0 EPS15 2.5 63.3 VCPIP1 22.3 53.5 FAM21A 23.8 PJA2 10.3 NBR1 3.5 HGS 7.8 SPAG5 7.8 EPPK1 1.3 5.5 PSMB6 4.8 SMC1A 3.0 23.3 RABEP1 21.3 HUWE1 18.5 KIF11 16.8 ASCC3 15.3 PSMB4 11.0 PSMB3 8.3 GET4 2.0 7.0 MLF2 6.8 PSMB2 6.0 ALMS1 4.8 TMEM160 4.8 SMC2 4.7 MAGED1 20.3 NME7 4.7 HECTD1 4.5 AK2 1.0 3.8 STAM2 3.5 USP24 3.5 PSMB1 3.3 POLR2B 1.0 3.0 CNOT10 3.8

UBE2A 4.5 PSMB5 3.8 N4BP2 15.0 GGA3 3.3 ZFAND5 3.0 PSMB7 2.8 AP2A1 8.3 PEG10 6.3 PMPCB 1.0 3.0 GLUL 3.0 TBK1 2.7 PSMD2 1.5 21.0 PLD3 2.0 MAGED2 6.0 UBXN2A 1.8 UBAC1 2.0 CAD 7.3 11.0 STAM 6.5 UFD1L 20.0 AFG3L2 1.0 2.7 COPS6 3.0 VPS37A 2.7 PSMD1 2.7 15.5 PSMA1 2.8 ECH1 8.0 6.5 SLC25A10 2.7 2.0 BIRC6 3.0 4.0