Investigating the Mechanism of UBR-1, an Arg/N-end Rule Pathway E3 that Modulates Caenorhabditis elegans Motor Behaviour

by

Min Wu

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

© Copyright by Min Wu 2019

Investigating the Mechanism of UBR-1, an Arg/N-end Rule Pathway E3 Ligase that Modulates Caenorhabditis elegans Motor Behaviour

Min Wu

Master of Science

Department of Molecular Genetics University of Toronto

2019 Abstract

The Arg/N-end rule pathway regulates half-lives in . UBR-1 E3 ligase serves as a recognition component of this pathway in C. elegans, which recognizes N-terminal degradation signals of substrates and mediates their ubiquitination. The functional loss of UBR-1 results in a unique motor defect: ’ posterior body fails to establish normal bending due to synchronized A-type motor activation during backward locomotion. This motor defect was found to be caused by aberrant glutamate . Specifically, removing a glutamate synthesizing GOT-1 could restore ubr-1 mutants’ bending during reversals.

Locomotory patterns are regulated by sensorimotor circuits. These findings connect aberrant glutamate levels with neural circuit dysfunction in the UBR-1-dependent modulation of motor outputs. Extending from these findings, my studies further reveal UBR-1’s main sites of action to be premotor interneurons that regulate backward locomotion. Results of my studies help decipher a physiological role of UBR-1 in the nervous system.

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Acknowledgments

I am very grateful to have been mentored by my supervisor Dr. Mei Zhen. She has been guiding me to become a better thinker and researcher with her rich experience, keen scientific sense, and extraordinary enthusiasm for science. Without her, this project would not be possible. She encouraged and led me through difficulties in this project. I thank her for all these very rewarding experiences.

I thank my committee members, Dr. Peter Roy and Dr. William Ryu, for their very helpful criticism, directions, and suggestions.

I thank Dr. Jyothsna Chitturi for her important work on C. elegans UBR-1, which provides the essential basis for this project.

I thank Dr. Wesley Hung for his contributions throughout this project. His technical support and expertise walked me through many difficulties in this project.

I thank Kevin Zhang for his work on the ubr-1 suppressor screens.

I thank all other members of the Zhen Lab, Dr. Ben Mulcahy, Dr. Shangbang Gao, Dr. Yan Li, Ying Wang, Jun Meng, Yangning Lu, Daniel Witvliet, many undergraduate students and past members, who provided me with technical support, helpful discussions and encouragement.

I thank Dr. Quan Wen, who led me into this field of science and gave me lots of inspirations.

I thank Dr. Janine Harper for reading and editing the first draft of this thesis.

Last but not least, I thank my parents for their unconditional love and support. To you I dedicate this thesis.

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List of Specific Contributions

I was solely responsible for experiments, data collection and figure preparation for all figures in Chapter 2-4, unless specified as otherwise.

Confocal images in Figures 8 & 12 were taken by Dr. Wesley Hung.

Behaviour recordings in Figure 10 were acquired by Dr. Jyothsna Chitturi.

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Table of Contents

Acknowledgments...... iii

List of Specific Contributions ...... iv

Table of Contents ...... v

List of Tables ...... viii

List of Figures ...... ix

Chapter 1 ...... 1

Introduction ...... 1

1.1 ...... 1

1.1.1 The - system ...... 2

1.1.2 The N-end rule pathway in eukaryotes ...... 6

1.1.3 The N-end rule pathway in prokaryotes...... 8

1.1.4 The UBR family E3 ubiquitin ...... 12

1.2 The mechanisms of Ubr1 in different model organisms ...... 14

1.2.1 The discovery and characterization of Ubr1 in S. cerevisiae ...... 14

1.2.2 The Johanson-Blizzard syndrome (JBS), a multisystem disorder caused by loss-of-function in human UBR1 ...... 16

1.2.3 The mouse models ...... 17

1.2.4 The culture model ...... 20

1.2.5 The C. elegans model ...... 21

1.3 The locomotion of C. elegans ...... 22

1.3.1 The models for the mechanism of locomotion ...... 23

1.3.2 The neural mechanism underlying backward locomotion ...... 26

1.3.3 The neural mechanism underlying forward locomotion ...... 28

1.4 Objectives ...... 30

Chapter 2 ...... 31

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Materials and Methods ...... 31

2.1 Molecular biology and C. elegans strains ...... 31

2.1.1 Strains, constructs and oligos ...... 31

2.1.2 UBR-1 RING domain replacement allele ...... 31

2.2 Locomotory behaviour assay ...... 37

2.2.1 Image acquisition ...... 37

2.2.2 Motor activity quantification ...... 38

2.3 Fluorescence microscopy ...... 39

2.4 Optogenetic cell ablation ...... 39

2.5 Calcium imaging ...... 39

2.6 Statistical analysis ...... 40

Chapter 3 ...... 41

Results ...... 41

3.1 ubr-1 mutants exhibit reduced body bending during reversals ...... 41

3.2 UBR-1 is expressed in multiple tissues ...... 43

3.3 UBR-1 is critically required in mechanosensory and premotor INs to affect the reversal motor pattern ...... 43

3.4 Reduced bending results from synchronization of A-MNs’ intrinsic activities ...... 47

3.5 Reduced bending results from premotor IN-mediated dysregulation of the coordination of A-MNs’ intrinsic activities ...... 50

3.6 Neuronal toxicity and aberrant glutamate signaling may account for the reduced bending phenotype in ubr-1 mutants ...... 53

Chapter 4 ...... 56

Conclusions and Future Perspectives ...... 56

4.1 Conclusions ...... 56

4.2 Future Perspectives ...... 57

4.2.1 To examine interactions between UBR-1 and ubr-1 suppressors ...... 57

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4.2.2 To identify UBR-1’s substrates by genetic suppressor screens ...... 59

4.2.3 To identify UBR-1’s substrates by BioID ...... 59

4.2.4 To dissect UBR-1-dependent regulation of the A-MN phase coupling ...... 60

References ...... 62

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List of Tables

Table 1. Strains and constructs used in this study ...... 32

Table 2. Primers for genotyping strains in this study ...... 35

Table 3. Sequences of primers used for genotyping in this study ...... 36

Table 4. A list of transgenes that restore UBR-1 expression in different tissues tested for rescue effects of the ubr-1 reduced bending phenotype ...... 46

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List of Figures

Figure 1. The ubiquitin-proteasome system...... 5

Figure 2. The N-end rule pathway in eukaryotes and prokaryotes ...... 11

Figure 3. Mammalian UBR family E3 ligases ...... 13

Figure 4. Three proposed hypothetic models for the generation of locomotor patterns ...... 25

Figure 5. A model for backward locomotion ...... 27

Figure 6. A model for forward locomotion ...... 29

Figure 7. ubr-1 mutants exhibit reduced bending angles at posterior body during reversals ...... 42

Figure 8. UBR-1 is expressed in muscles, hypodermis and sub groups of neurons ...... 44

Figure 9. Transgenes that restore UBR-1 expression in subgroups of neurons rescue the ubr-1 reduced bending phenotype ...... 45

Figure 10. UBR-1 promotes body bending by preventing synchronized A-MN activation ...... 49

Figure 11. Ablation of premotor INs in ubr-1 mutants restores the phase coupling of A-MN activation ...... 51

Figure 12. The functional loss of UBR-1 results in morphological changes of specific premotor INs at the end of larval stage development ...... 54

Figure 13. UBR-1 may function through sensory neurons and premotor INs to affect the phase coupling among A-MNs so as to modulate motor outputs ...... 55

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Chapter 1 Introduction 1.1 Proteolysis

Proteolysis refers to a set of processes that lead to hydrolysis at one or more peptide bonds of a protein. Proteolysis plays a pivotal role in the maintenance of cellular and the regulation of many cellular processes. The major cellular functions of proteolysis include the elimination of misfolded and damaged , the regulation of cellular metabolism, the generation of active proteins, and the recycling of amino acids (Hershko et al., 1982; Hand and Hardewig, 1996; Dreher and Callis, 2007; Hershko and Rose, 1987).

Cells apply multiple mechanisms to execute protein degradation. These mechanisms mainly depend on two proteolytic systems: and . is the general designation of three distinct types of -dependent protein degradation, namely microautophagy, macroautophagy, and -mediated autophagy (Mizushima et al., 2008). The ubiquitin-proteasome system selectively degrades proteins that are modified by covalently bound ubiquitin with the aid of the 26S proteasome, an ATP-dependent multi-subunit protease complex (Hershko et al., 1983; Baumeister et al., 1998).

The two major catabolic systems, autophagy and ubiquitin-proteasome system, participate in the degradation of proteins with varying selectivity. Generally, autophagy is responsible for the degradation of long-lived cytosolic proteins and integral membrane proteins, as well as other macromolecules including lipids and entire organelles (Komatsu et al., 2007). Most short-lived cytoplasmic proteins and endoplasmic reticulum membrane proteins are degraded by proteasomes (Ciechanover et al., 1984).

The ubiquitin-proteasome system-mediated rapid and selective destruction of proteins is especially important for regulatory proteins whose concentrations must change quickly in accordance with changes in cellular states (Varshavsky, 2011). Therefore, the ubiquitin-proteasome system is central to the control of almost all aspects of cellular homeostasis, such as , cell differentiation, DNA repair, and cell signaling (Ciechanover, 1998; Haglund and Dikic, 2012; Schwarz and Patrick, 2012).

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1.1.1 The ubiquitin-proteasome system

The ubiquitin-proteasome system facilitates the tight spatial and temporal regulation of protein turnover in both the and the nucleus in eukaryotes (Bachmair et al., 1986; Gardner et al., 2005). The process of protein degradation through this system involves two successive steps: (1) a chain consisting of multiple copies of ubiquitin, a small conserved tag protein of 76 amino acids, are covalently attached to an ε amino group of a residue of the substrate protein (Hershko et al., 1983); and (2) ubiquitin is removed and the target protein is degraded into its constitutive peptides and amino acids by the 26S proteasome in an adenosine triphosphate (ATP)- dependent manner (Baumeister et al., 1998).

The first step, referred to as ubiquitination, or the conjugation of ubiquitin, is tightly regulated and catalyzed by three classes of : the ubiquitin-activating enzyme (E1), the ubiquitin- conjugating enzyme (E2), and the (E3) (Hershko et al., 1983). The most common form of the UPS involves polyubiquition of substrates. In the first of a series of enzymatic processes, an E1 activates ubiquitin in an ATP-dependent reaction, forming a ubiquitin-adenosine monophosphate (AMP) intermediate. The E1 binds to ubiquitin-AMP through a thioester bond between the C-terminus of ubiquitin and its (Haas et al., 1982). The E1 then passes activated ubiquitin onto an E2’s cysteine residue, where another transient thioester bond between ubiquitin and E2 is formed. An E3 recruits a substrate protein and transfers ubiquitin to a lysine residue of this protein from E2 directly or indirectly. The C terminus of ubiquitin is covalently ligated to the substrate’s lysine residue through an isopeptide bond (Hershko et al., 1983). Ubiquitin itself contains multiple lysine residues (Komander and Rape, 2012). A second- round process attaches another ubiquitin to a lysine residue of the first ubiquitin. By repeating the process, a polyubiquitin chain is tagged to the substrate protein at one of its lysine residues (Fig. 1).

E1 ubiquitin-activating enzymes are the least selective of the three classes of enzymes in the ubiquitination processes. There are only nine E1 ubiquitin activating enzymes in humans and one single E1 in C. elegans. These enzymes activate ubiquitin that is later used to tag target proteins for degradation with no substrate specificity.

E2 ubiquitin-conjugating enzymes are diverse both structurally and functionally. There are over 30 different E2s in humans and around 20 in C. elegans. Most of them harbour a core catalytic

3 domain for ubiquitin transfer. The core domains of different E2s are of similar sizes (around 150 amino acids) and high conservation (Haldeman et al., 1997; Huang et al., 1999). Other regions vary considerably. Some E2s have short N-terminal or C-terminal extensions that specify their subcellular localizations or their interactions with E1 ubiquitin activating enzymes and specific groups of E3 ubiquitin ligases (Huang et al., 2004; Haldeman et al., 1997; Merkley and Shaw, 2004). The diversity of E2s contributes to the selectivity of their ubiquitin-conjugating reactions.

E3 ubiquitin ligases selectively recruit protein substrates by recognizing their degradation signals, and facilitate the transfer of from E2s to the substrates. E3s have the highest protein diversity and substrate specificity among the three classes of enzymes catalyzing the ubiquitination processes. Hundreds of putative E3 ubiquitin ligases are encoded by the human and C. elegans genomes. They are classified into three major types based on their structures and mechanisms of ubiquitin transfer: HECT (Homologous to E6-AP Carboxyl-Terminus)-type, U-box-type, and RING (Really Interesting New )-type.

The HECT-type E3 ligases harbour a C-terminal catalytic HECT domain that is homologous to that of human papilloma virus (HPV) E6-associated protein (E6AP) (Huibregtse et al., 1995). A key feature of the HECT domain is a conserved cysteine residue that can form a thioester bond with the C terminus of ubiquitin. A HECT-type E3 ligase acts as an intermediary by accepting ubiquitin from an E2 through the thioester bond and then transferring it to the substrate (Rotin and Kumar, 2009).

The U-box-type E3 ligases share a cross-brace structure consisted of an alpha-helix with two surface-exposed loop flanks (U-box) (Aravind et al., 2000). The center of each loop consists of a network of hydrogen bonds and salt bridges; this network contributes to the stability of the overall structure. A U-box-type E3 ligase acts as a scaffold that brings the E2 and substrate into close proximity, allowing for the transfer of ubiquitin to a lysine residue of the substrate (Deshaies and Joazeiro, 2009).

The RING-type E3 ligases are characterized by the presence of RING-type zinc-finger domains (Freemont et al., 1991). A RING domain differs from a U-box domain by having two zinc ions instead of salt bridges in the stabilizing centers. They are also involved in interactions with E2s and target proteins, and are critically required for the transfer of a ubiquitin to either substrates or RING-type E3s themselves. RING-type E3s can be further sorted into two subtypes: the multi-

4 subunit and monomeric RING-type E3s. Multi-subunit RING-type E3s are small proteins of 110- 135 amino acids. They act as integral components of multi-subunit complexes that facilitate the conjugation of ubiquitins to substrate proteins (Seol et al., 1999). Monomeric RING-type E3s are much larger; they function as monomeric scaffolds that bind both the E2 and substrate protein, expediting the transfer of ubiquitin moiety to the substrate (Lorick et al., 1999).

E3s provide high substrate specificity in the ubiquitin elongation process to ensure that the ubiquitin-proteasome system-mediated protein degradation is selective. A monomeric ubiquitin tag at one or multiple lysine residues of a substrate is typically not sufficient for the proteasome to recognize protein substrates. Rather, a polyubiquitin chain is required. After the first ubiquitin moiety is attached to the target protein, additional ubiquitins are successively linked to the lysine residue of the latest ubiquitin to elongate the tag. Ubiquitin has seven lysine residues and an N- terminus (K6, K11, K27, K29, K33, K48, K63 and M1) available for the linkage of another ubiquitin (Komander and Rape, 2012). The positioning of ubiquitin moiety linkages within a polyubiquitin chain determines how substrate proteins are processed. E3s target proteins tagged by K11, K29, and K48-linked polyubiquitin chains to the 26S proteasome for degradation (Jin et al., 2008; Johnson et al., 1995; Chau et al., 1989). K63-linked polyubiquitin modifications reversibly affect protein interaction, activity and localization (Xia et al., 2009; Deng et al., 2000; Plosky et al., 2006). E3s also regulate the affinity between polyubiquitinated substrates and 26S proteasomes through the lengths of polyubiquitin chains. For an example, a K48-linked polyubiquitin chain must be elongated to at least four ubiquitin molecules in order to be efficiently recognized by a 26S proteasome (Maurer and Wertz, 2016; Thrower et al., 2000). In short, E3 ubiquitin ligases facilitate the ubiquitin-proteasome system-mediated selective protein degradation by recruiting target proteins, determining the composition and length of the polyubiquitin chain tags.

The 26S proteasome recognizes and degrades proteins labelled with a polyubiquitin chain. This large multi-subunit complex consists of a 20S proteolytic core and a 19S regulatory cap at either end (Zwickl et al., 1999). The 19S regulatory complex recognizes the polyubiquitin chain and recruits the substrate protein. Deubiquitinating enzymes (DUBs) in the 19S complex remove the polyubiquitin chain from the substrate and disassemble it so as to recycle ubiquitin molecules. Deubiquitinated substrate is unfolded by the 19S regulatory complex in an ATP-dependent manner and translocated into the central cavity of the 20S proteolytic core, where it is broken down into a

5 set of peptides about 8 amino acids long. These short peptides are released for further use afterwards (Voges et al., 1999; Pickart et al., 2004) (Fig. 1).

Figure 1. The ubiquitin-proteasome system

A schematic diagram representing the process of the ubiquitin-proteasome system-mediated protein degradation. The ubiquitin activating enzyme (E1) activates a ubiquitin in an ATP- dependent reaction, forming an E1-ubiquitin-AMP intermediate. The activated ubiquitin is transferred to the ubiquitin conjugating enzyme (E2). The ubiquitin ligase (E3) binds to E2 and recruits a substrate protein, bringing them into close proximity. The ubiquitin is thereafter transferred from E2 to a lysine residue of the substrate protein. When these processes are repeated, a polyubiquitin chain is formed tagging this protein. A 19S cap of a 26S proteasome recognizes the polyubiquitin chain and recruits this protein. Deubiquitinating enzymes (DUBs) detach and disassemble the polyubiquitin chain, allowing ubiquitin molecules to be reused. The 19S cap then unfolds this protein. The unfolded protein is translocated into the central cavity of the 20S proteolytic core of the 26S proteasome, where it is degraded into oligopeptides in an ATP- dependent manner.

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1.1.2 The N-end rule pathway in eukaryotes

The N-end rule pathway refers to the process that proteins are targeted for selective degradation by an N-terminal degradation signal, named the N-degron. Proteins that recognize the N-terminal destabilizing amino residues are called the N-recognins.

In eukaryotes, the N-end rule pathway is a branch of the ubiquitin-proteasome system. N-recognins are E3s that bind to specific N-degrons. They facilitate the transfer of ubiquitin from E2s to substrates, targeting them to 26S proteasomes for degradation (Varshavsky, 2011). The eukaryotic N-end rule pathway consists of three sub-branches: the Arg/N-end rule pathway, the Ac/N-end rule pathway, and the Pro/N-end rule pathway (Bartel et al., 1990; Hwang et al., 2010a; Chen et al., 2017).

The Arg/N-end rule pathway is the classical form of the N-end rule pathway, which was initially identified in (Bartel et al., 1990), and later found in most species including mammals and plants (Tasaki and Kwon, 2007; Graciet and Wellmer, 2010). The UBR family E3 ubiquitin ligases are the N-recognins in this pathway. They harbour two domains for recognition of destabilizing N-terminal amino acids, either unmodified or covalently modified (Tasaki et al., 2009). The first type is called the UBR-box domain, or Type-1 substrate . It is a ~70 amino acid zinc finger motif that binds to three unmodified basic N-terminal residues: Arg, Lys, and His (Type-1 primary destabilizing residues) (Tasaki et al., 2005; Tasaki et al., 2009). The second type recognition domain, called the N-domain, or Type-2 substrate binding site, targets destabilizing bulky hydrophobic amino acids Leu, Phe, Tyr, Trp, and Ile (Type-2 primary destabilizing residues) (Tasaki et al., 2009). It is a ~80 amino acid motif that bears sequence similarity to ClpS, the bacterial N-recognin (Erbse et al., 2006). Apart from these N- degron recognition domains, the UBR family E3s also recognize internal degrons of target proteins via less well defined protein motifs (Du et al., 2002).

N-terminal primary destabilizing residues, or Ndp, can be directly recognized by Type-1 or Type- 2 binding sites (Tasaki et al., 2009). Other N-terminal residues of substrate proteins are covalently modified before binding to specific N-recognins. These other residues are termed secondary (Nds) or tertiary (Ndt) destabilizing residues, depending on their modifications. In mammalian cells, Nds residues include Asp and Glu. They are arginylated by the Arg-tRNA-protein (R- transferase) so as to be recognized by N-recognins (Hu et al., 2006). In contrast, R-

7 cannot conjugate Arg to Ndt residues, Asn, Gln, or Cys. However, specific modifications to these Ndt residues can facilitate arginylation. N-terminal Asn and Gln residues are converted to Asp and Glu, respectively, by N-terminal amidases, allowing for arginylation as Nds residues (Baker and Varshavsky, 1995; Kwon et al., 2000). N-terminal Cys can be non-enzymatically oxidized by nitric oxide (NO) and oxygen (O2). The oxidation product, N-terminal Cys-sulfinate or Cys-sulfonate, can be arginylated by R-transferase and thereby bound to N-recognins (Hu et al., 2005; Lee et al., 2005) (Fig. 2A).

The Arg/N-end rule pathway is complemented by a second proteolytic system: the ubiquitin-fusion degradation (UFD) pathway (Hwang et al., 2010b). The UFD pathway does not recognize the N- degrons in the Arg/N-end rule pathway, but targets substrates through their N-terminal ubiquitin moieties (Johnson et al., 1995). A HECT-type E3 ubiquitin ligase of the UFD pathway, Ufd4, can interact with S. cerevisiae Ubr1 physically and functionally. While it is not necessary for Ubr1 to form the Ubr1-Ufd4 complex to function as an N-recognin, this complex facilities Ubr1-mediated degradation by increasing the processivity of polyubiquitination of the Arg/N-end rule substrates (Hwang et al., 2010b).

The Ac/N-end rule pathway was discovered two decades after that of Arg/N-end rule pathway (Hwang et al., 2010a). It requires the unique and irreversible co-translational N-terminal acetylation of nascent proteins. These proteins have Met at the N-terminal, or have small uncharged amino acids, such as Ala, Val, Ser, Thr, Cys, Gly, and Pro, exposed after the N-terminal Met is co-translationally processed by Met-aminopeptidases. These N-terminal residues can be acetylated by Nt-acetylases and thereby recognized by specific E3 N-recognins (Hwang et al., 2010a). Because of the necessity of the Nt-acetylation, these destabilizing residues are termed secondary AcN-degrons (Fig. 2B). The first known Ac/N-end rule pathway E3 is a yeast protein Doa10 (Swanson et al., 2001; Hwang et al., 2010a), a RING-type ligase that is located in the endoplasmic reticulum (ER) membrane as well as the inner nuclear membrane (Deng et al., 2006), and targets both soluble and transmembrane proteins (Ravid et al., 2006). One of its human homologues Teb4 also serves as an Ac/N-end rule N-recognin to regulate the G protein signaling (Hassink et al., 2005; Hwang et al., 2010a).

Most cellular proteins, including more than 80% of mammalian proteins, are co-translationally Nt- acetylated (Arnesen et al., 2009). Thus, a bulk of nascent proteins may be targeted by the Ac/N-

8 end rule pathway, either during or after their synthesis (Shemorry et al., 2013). Other proteins are alternatively destructed by the Arg/N-end rule pathway, in part because Met-aminopeptidases cannot cleave off the N-terminal Met when the second-position residue is a large amino acid. Such amino acids include all the destabilizing residues in the Arg/N-end rule pathway except Cys. A second-position Cys can become the N-terminal amino acid by Met-aminopeptidases, after which it can be acetylated and then serves as an AcN-degron. However, in some cases, the acetylation of N-terminal Cys can be inhibited. In such cases, the protein is degraded through the Arg/N-end rule pathway after two modification steps. For example, a basic residue at the second position may block acetylation of a set of N-terminal residues including Cys (Arnesen et al., 2009). The mammalian G-protein regulator proteins RGS4, RGS5, and RGS16 have Lys, Lys, and Arg, respectively, at the third position following their second-position Cys (Cys-2). Their Cys-2 can be made N-terminal by methionine aminopeptidases (MetAPs), but cannot be acetylated by Nt- acetylases, preventing them from being recognized by the Ac/N-end rule pathway. Alternatively, these residues can be arginylated by R-transferases after NO/O2-mediated oxidation (Hu et al., 2005; Lee et al., 2005). These intricate processes allow RGS proteins to be targeted by the Arg/N- end rule pathway.

The Pro/N-end rule pathway is a newly found mechanism of selective protein degradation in S. cerevisiae (Chen et al., 2017). The GID E3 ubiquitin ligase serves as an N-recognin of this pathway. Through its Gid4 subunit, it targets proteins with Pro at the N-terminal and the second position with five specific adjoining residues: TLVNG and LVMNA, respectively (Fig. 2C). Substrates for GID include the gluconeogenic enzymes Fbp1, Icl1, Mdh2, and Pck1 (Menssen et al., 2012; Chen et al., 2017). The Arg/N-end rule and Ac/N-end rule pathways appear to be conserved in all eukaryotic organisms examined, while evidence for the presence of the Pro/N-end rule pathway in multicellular eukaryotes remains to be found.

In sum, the Arg/N-end rule, Ac/N-end rule, and Pro/N-end rule pathways can be responsible for regulated degradation of a majority of cellular proteins in eukaryotes.

1.1.3 The N-end rule pathway in prokaryotes

Prokaryotes do not have these branches of the N-end rule pathway. Instead, they employ the Leu/N-end rule pathway (Tobias et al., 1991; Mogk et al., 2007), and this branch does not involve ubiquitination. The N-recognin of the Leu/N-end rule pathway is ClpS, which recognizes

9 destabilizing N-terminal bulky hydrophobic residues, such as Leu, Phe, Trp, and Tyr (Ndp residues) (Tobias et al., 1991). These residues are also N-degrons targeted by the Type-2 substrate binding sites of the UBR family E3 ubiquitin ligases, suggesting a homology between bacterial and eukaryotic N-recognins of the N-end rule pathways. Other residues, including Arg, Lys, Asp, and Glu, may be leucylated or phenylalanylated so as to be bound to the ClpS N-recognin. They are therefore termed Nds residues. Escherichia coli has an L/F-transferase: Aat. It conjugates a Leu or Phe to the N-terminal Arg or Lys of a substrate protein. In addition to Aat, Vibrio vulnificus contains a second L-transferase called Bpt, which is a sequelogue to the eukaryotic Ate1 R- transferase and uses Leu-tRNA as a cosubstrate to facilitate the leucylation of N-terminal Asp, Glu, and possibly oxidized Cys (Graciet et al., 2006; Mogk et al., 2007) (Fig. 2D).

Proteins with Ndp or leucylated/phenylalanylated Nds are then processed by a proteolytic system that is similar to the ubiquitin-proteasome system. This system comprises two major components: (1) ClpAP, a proteasome-like protease complex; and (2) ClpS, the N-recognin of the Leu/N-end rule pathway (Schmidt et al., 2009; Román-Hernández et al., 2011). The ClpAP protease complex consists of a ClpA6 unfoldase and a ClpP14 protease (Guo et al., 2002; Hinnerwisch et al., 2005).

In this complex, ClpA6 has an axial pore that connects with the proteolytic chamber of ClpP14, permitting the efficient translocation of substrate proteins (Hinnerwisch et al., 2005; Effantin et al., 2010). The ClpS N-recognin is composed of a flexible N-terminal extension and a folded core binding domain (Schuenemann et al., 2009). The binding domain recruits a specific Leu/N-end rule pathway substrate by recognizing its N-degron. This binding enables the conformational change of the ClpS complex, which allows a part of the Nt-extension to be bound to the axial pore of the ClpA6 unfoldase and thereby increases the affinity between ClpS and ClpAP by ~10 folds (Zeth et al., 2002; Schuenemann et al., 2009; Román-Hernández et al., 2011). The substrate is then translocated from the ClpS binding domain to the ClpA6 axial pore, through which it is delivered to the ClpP14 proteolytic core for full breakdown in an ATP-dependent manner (Román- Hernández et al., 2011).

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Figure 2. The N-end rule pathway in eukaryotes and prokaryotes

Schematic diagrams of four major branches of the N-end rule pathway. (A) The Arg/N-end rule pathway. Proteins with primary/secondary/tertiary N-terminal destabilizing residues (Ndp/Nds/Ndt) are unstable. These residues can be exposed at the N-terminal by , calpains, and other non-processive proteases. Two Ndt residues, Asn and Gln, can be amidated into Nds, Asp and Glu, respectively, by Nt-amidases. The other Ndt, Cys, can be oxidized into Cys-sulfinate or Cys-sulfonate (C*) by NO/O¬2. The three Nds can be arginylated by R-transferase, forming a Ndp residue. Type-1 and Type-2 Ndp residues can be directly recognized by respective Type-1 and Type-2 substrate binding sites of UBR family E3 ligases. Ubr1 can also recognize some internal degrons. Substrate proteins are then tagged with a polyubiquitin chain and degraded into oligopeptides by 26S proteasomes. Red: processes or components identified in the mammalian system and absent in S. cerevisiae; ₠: a processe also identified in C. elegans; Φ: Leu, Phe, Tyr, Trp, and Ile. (B) The Ac/N-end rule pathway. Small uncharged amino acids, Ala, Ser, Thr, Val, Cys, Gly, and Pro, at the second position can be made N-terminal by methionine aminopeptidases (MetAPs). These N-terminal residues, as well as Met, can be acetylated by Nt-acetylases. The Doa10 E3 ligase recognizes these acetylated residues and mediates the degradation of substrate proteins by 26S proteasomes. (C) The Pro/N-end rule pathway. Pro at the second position following Met or third position following Met-Cys can be made N-terminal or second position after MetAPs cleave off the original N-terminal Met. The N-terminal Pro with five specific adjoining residues, TLVNG, and the second position Pro with LVMNA can be recognized by the GID E3 ligase. (D) The Leu/N-end rule pathway. Proteins with primary Ndp residues, Leu, Phe, Trp, and Tyr, can be directly targeted by the ClpS N-recognin for degradation by the ClpAP protease. Two Nds residues, Asp and Glu, can be leucylated by the Bpt L-transferase. The other two, Lys and Arg, can be leucylated or phenylalanylated by the Aat L/F-transferase. Panel (A) modified from Kim et al., 2015; panels (B) and (C) modified from Chen et al., 2017; panel (D) modified from Varshavsky, 2011.

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1.1.4 The UBR family E3 ubiquitin ligases

Ubiquitin-dependent proteolysis was first discovered in a reticulocyte extract (Hershko et al., 1980). Later, the first specific E3 ubiquitin ligase, Ubr1, was cloned in S. cerevisiae (Bartel et al., 1990). Ubr1 is the sole UBR family E3 ubiquitin ligase in yeast. It is a RING-type E3 ligase with two N-recognin domains: the UBR-box and the N-domain, which recognize Type-1 (Arg, Lys, and His) and Type-2 (Leu, Phe, Tyr, Trp, and Ile) N-terminal destabilizing residues, respectively.

The mammalian genome encodes seven UBR-box proteins, termed UBR1 through UBR7 (Tasaki et al., 2005). UBR1, UBR2, and UBR3 are the RING-type E3 ubiquitin ligases. They are referred to as canonical UBR family members, by and conserved UBR-box and RING domains. UBR4 through UBR7 are classified as non-canonical. They are divergent, and share limited homology among each other. With the exception of UBR4, each contains a unique signature domain. UBR5 is a HECT-type E3 ubiquitin ligase; its HECT domain facilitates substrate ubiquitination. UBR6 has a F-box domain that mediates protein-protein interaction. UBR-7 harbours a PHD (plant homeodomain finger) domain, a conserved zinc finger domain that resembles the RING domain in UBR1. UBR4 has an exceptionally large protein size of 570kDa. The complete open reading frame (ORF) of this gene is not clear (Fig. 3). It has no known ubiquitination domain, but does contain a UBR-box domain. The knockdown of Ubr4 using RNA interference (RNAi) in Ubr1-/-Ubr2-/- fibroblasts was reported to abolish the degradation of proteins with Type-2 N-terminal destabilizing residues, suggesting that UBR4 may play a role in the Type-2 pathway (Tasaki et al., 2009).

The presence of a UBR-box domain signifies the role of a UBR family E3 ligase as an N-recognin. However, the UBR-box domain alone is insufficient for an E3 ligase to function as an N-recognin. Mouse UBR3, UBR6 and UBR7 do not act as N-recognins, in spite of the presence of a UBR-box domain, possibly due to the lack of specific residues required for appropriate conformation (Wang et al., 2004; Tasaki et al., 2009). The N-degron-binding properties of the other UBR-box proteins have been validated (Tasaki et al., 2009). Among them, UBR1, UBR2, and possibly UBR4 contain an N domain. The physiological functions of UBR4 and UBR5 as N-recognins remain to be investigated. Similarly, yeast Ubr2 does not exhibit N-recognin activities despite its significant homology to Ubr1. It lacks some residues that are conserved in Ubr1 homologues from yeast to mammals; they may be essential for N-recognins (Wang et al., 2004).

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Figure 3. Mammalian UBR family E3 ligases

Schematic diagrams of mammalian UBR family E3 ligases. All seven members, named UBR1 through UBR7, contain a UBR-box domain. UBR1, UBR2, and UBR3 are RING-type E3 ligases of similar sizes, featured by a RING domain. Besides, UBR1 and UBR2 contain an N domain. UBR4 is an extraordinarily large protein with a largely unknown ORF. UBR5 is a large HECT- type E3 ligase harbouring a C-terminal HECT domain. UBR6 is a relatively small E3 ligase with a F-box domain near the N-terminal. UBR7 is the smallest one among the seven E3 ligases and contains a PHD domain. Figure modified from Tasaki et al., 2008.

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1.2 The mechanisms of Ubr1 in different model organisms

Ubr1 is the canonical Arg/N-end rule pathway E3 ubiquitin ligase. It was first cloned and analyzed by Varshavsky and colleagues with genetic and biochemical approaches in S. cerevisiae. Since then, a great number of Ubr1-like E3 ligases have been investigated in different model organisms. Studies on Ubr1 reveal substantial diversity of Ubr1’s physiological functions. Reported substrates of Ubr1 display extreme diversity in structures. The pathways in which they participate cover a wide range of cellular processes.

1.2.1 The discovery and characterization of Ubr1 in S. cerevisiae

In 1978, Hershko and colleagues first observed ubiquitination and successive degradation of some proteins added to a reticulocyte extract. This finding suggested the existence of a selective degradation system. They went on to identify and characterize components of this system that conjugate ubiquitins to target proteins, termed E1, E2, and E3.

Contemporaneously, Varshavsky and colleagues discovered and defined the ubiquitin system through genetic and biochemical studies with mammalian cells and S. cerevisiae. They cloned and analyzed the first E3 ligase, yeast Ubr1, in 1990. Through in vitro binding and in vivo degradation tests with artificial protein substrates engineered to bear destabilizing or stabilizing N-terminal residues, Bartel et al. (1990) concluded that Ubr1 binds to specific substrates and determines the rate of their degradation.

Yeast Ubr1 is, however, non-essential, at least under the laboratory culturing condition. Yeast null mutants are viable but unable to degrade engineered substrates of the Arg/N-end rule pathway, which are normally short-lived in wildtype cells. Physiologically, the ubr1 null leads to a slight (~2%) increase of doubling time (Bartel et al., 1990).

In 2001, the first physiological substrate of yeast Ubr1, Scc1, was identified by Varshavsky’s laboratory (Rao et al., 2001). Scc1 is a subunit of the sister chromatid cohesion named cohesin. The cohesin complex holds the sister chromatids of a replicated together until the onset of (Uhlmann et al., 1999). At the to anaphase transition, the Scc1 subunit is cleaved by the separin protease, exposing an Arg residue at the amino-terminal of its carboxy-terminal fragment (Nasmyth et al., 2000). The resulting Scc1 fragment is then rapidly degraded by the Arg/N-end rule pathway, with a half-life of about 2min,

15 allowing the microtubules to pull the sister chromatids apart. In ubr1Δ cells, the absence of the Arg/N-end rule pathway leads to a significantly increased frequency of chromosome loss (Rao et al., 2001). In sum, Ubr1 promotes proper separation of the sister chromatids through efficient degradation of the Scc1 fragment.

A recent study revealed that the Type-2 binding site of yeast Ubr1 can recognize, in addition to the N-terminal hydrophobic residues (Leu, Phe, Tyr, Trp, and Ile, denoted as Φ), the unacetylated N-terminal Met of Met-Φ proteins (Kim et al., 2014b). Nascent polypeptides bearing N-terminal Met are usually Nt-acetylated and targeted by the Ac/N-end rule pathway for degradation. However, many proteins are only partially Nt-acetylated, which means a fraction of one such protein would remain inaccessible to the Ac/N-end rule pathway. Therefore, Met-Φ proteins may be open to both the Ac/N-end rule and Arg/N-end rule pathway. Nt-acetylated Met-Φ proteins are degraded through the Ac/N-end rule pathway, while their unacetylated counterparts may be targeted by Ubr1 for the Arg/N-end rule pathway-mediated turnover.

In both yeast and mammalian cells, a significant ratio (~15%) of encode Met-Φ proteins, many of which may be substrates of Ubr1. Varshavsky’s laboratory picked, mostly by random, four Met-Φ proteins that have been reported to be Nt-acetylated in wildtype yeast, and tested whether they are degraded through the Ubr1-dependent Arg/N-end rule pathway. The four proteins are Msn4, a stress-responsive transcriptional activator; Sry1, a 3-hydroxyaspartate dehydratase; Arl3, a Golgi-associated cytosolic GTPase; and Pre5, a subunit of the 20S proteasome. They turned stable in naa30Δ ubr1Δ cells, which lacked both the NatC Nt-acetylase and the Ubr1 E3 ligase.

They could also be stabilized after the second position Φ residue being mutated to Lys in naa30Δ cells (Kim et al., 2014b). These results suggest that all four proteins are potential physiological 22-519 substrates of Ubr1. Moreover, some misfolded proteins, including MI-ΔssC Leu2myc and MI-

22-58 Φ ΔssC Ura3ha, were targeted by Ubr1 through their Met /N-degrons (Eisele and Wolf, 2008; Kim et al., 2014b).

Apart from the Type-1 and Type-2 substrate binding sites, yeast Ubr1 contains recognition domains for internal degrons. Two Ubr1 substrates, Cup9, a transcriptional repressor, and Mgt1, a DNA alkyltransferase, bear no N-degrons (Du et al., 2002; Hwang et al., 2009).

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Cup9 is a transcriptional regulator that represses over 30 genes, including the main dipeptide/tripeptide importer gene PTR2 (Turner et al., 2000; Alagramam et al., 1995; Byrd et al., 1998). Imported dipeptides and tripeptides bearing Type-1/2 destabilizing N-terminal residues can act as competitive inhibitors of the Type-1/2 binding sites of Ubr1. Concurrently, they may also allosterically activate the otherwise auto-inhibited third substrate binding site. Ubr1 molecules that bear an active Cup9-binding site target Cup9 for fast degradation, and thereby mitigate the transcriptional repression of PTR2. Elevated Ptr2 level then leads to the import of more di/tripeptides, forming a loop (Du et al., 2002). Ubr1 thus helps to increase the uptake of extracellular peptides upon detection through the degradation of Cup9.

Mgt1 is an O6-alkylguanine DNA alkyltransferase. It repairs O6-alkylguanine lesions in double- stranded DNA by irreversibly transferring methyl and other alkyl adducts from an alkylated O6 in guanine to its active site (Cys residue) (Xiao and Samson, 1992). The resulting alkylated Cys in Mgt1 cannot be restored to Cys, making the S-alkylated form of Mgt1 the dead-end product of the DNA repair pathway. To reset it, the rapid selective degradation of the S-alkylated Mgt1 is required (Srivenugopal and Ali-Osman, 2002). Interestingly, Mgt1 is targeted by both Ubr1 and Ufd4 through its internal degron. The two E3 ubiquitin ligases cooperate to facilitate the degradation of Mgt1 with high efficacy (Hwang et al., 2009). In short, Ubr1, together with Ufd4, ensures the continuous functioning of the DNA repair circuit through the timely degradation of S- alkylated Mgt1.

1.2.2 The Johanson-Blizzard syndrome (JBS), a multisystem disorder caused by loss-of-function mutations in human UBR1

The JBS is an autosomal recessive multisystem disorder first reported by Ann Johanson and Robert Blizzard in 1971. The clinical hallmarks are hypo-/aplasia of the alae nasi and exocrine pancreatic insufficiency. Other common features of this disease include absent permanent teeth, hypothyroidism, aplasia scalp cutis, short stature, urogenital and anorectal malformations, sensorineural deafness, and mental retardation (Vieira et al., 2002). In 2005, through a genome- wide linkage scan using microsatellite makers, as well as sequencing of genomic DNA from 14 patients, Zenker et al. determined that JBS is caused by loss-of-function mutations in the human homologue of yeast Ubr1, termed UBR1.

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With the gene identified, Sukalo et al. (2014) investigated the correlation between the genotypes and phenotypes of JBS patients. With the assumption that truncating (nonsense or frameshift) mutations most likely result in complete functional loss of UBR1, while non-truncating (missense or small in-frame deletion) mutations may retain residual protein function, they divided patients into two groups. The first group comprised patients with biallelic truncating mutations; the second comprised patients with a non-truncating mutation in at least one allele. In both groups, all individuals manifested pancreatic insufficiency, dental defects, and nasal wing aplasia. However, the severity of patients’ facial phenotypes may be associated with their UBR1 genotypes. Patients in group 1, with biallelic truncating alleles, showed complete aplasia of the alae nasi, while much milder patterns of hypoplasia were observed in group 2. Other phenotypes that exhibited different distributions between the two groups include cognitive impairment (100% vs. 39%, group 1 vs. group2), hearing impairment (100% vs. 50%), and short stature (88% vs. 48%). The most evident difference lay in patients’ cognitive impairment. All patients in group 1 showed cognitive impairment of various degrees; half were further reported to have moderate to severe intellectual disability. In contrast, more than half of the patients in group 2 had normal intellectual abilities. These findings suggest that different cells/tissues have different levels of tolerance for the functional loss of UBR1. Some residual UBR1 function may be sufficient for normal cognitive development or, more generally speaking, normal nervous system functions within the brain. In contrast, residual UBR1 function seems insufficient to prevent pancreatic dysfunction, dental defects, and nasal wing hypo-/aplasia.

Despite many case reports and genetic studies, the molecular pathophysiology of JBS remains unclear and there is no causal treatment available so far (Sukalo et al., 2014). Considering the relevance of UBR1 genotypes to JBS phenotypes, the identification of possible modifiers of particular phenotypes is of great interest. It may be possible, for example, to identify the modifiers for nervous system disorders that are associated with complete functional loss of UBR1. Discoveries of modifying factors for each phenotype may lead to a deeper understanding of these human Arg/N-end rule pathway disorders, from which treatment prospects may be found.

1.2.3 The mouse models

In order to decipher the mammalian Arg/N-end rule pathway, Varshavsky’s laboratory carried out a series of studies on the mouse homologues of the UBR family proteins. In 2001, they constructed

18 and analyzed the Ubr1-/- mouse strains. Similar to yeast Ubr1, mouse UBR1 is non-essential. The Ubr1-/- mice were viable, fertile, and healthy. No morphological abnormalities were observed in any of the examined tissues in the UBR1-/- strains from embryonic stages to adulthood. They showed normal size, overall behaviour, limb movements, and motor coordination. The only detectable defect in Ubr1-/- mice was a mild reduction of mass, ranged from ~12% at 1 year for both genders to ~32% for males and ~26% for females at the time of weaning. Further measurement of the weights of different tissues and organs revealed that a decrease in the mass of skeletal muscles and adipose tissues made up the majority of the total mass reduction. Using artificial substrates, the UBR1-/- extracts were shown to lack Arg/N-end rule proteolytic activity; but such an activity was found to remain active both in embryonic fibroblast (EF) cell lines and in EF extracts (Kwon et al., 2001).

The active Arg/N-end rule pathway in Ubr1-/- EF cells indicated the presence of additional N- recognin(s) aside from Ubr1. Mouse UBR2 exhibits significant sequence similarity to UBR1 (47% identity, 67% similarity); it also serves as an N-recognin of the Arg/N-end rule pathway (An et al., 2006; Tasaki et al., 2009). Ubr2-/-females exhibited high prenatal lethality, whereas the males were viable but infertile (Kwon et al., 2003). In contrast to the viability of Ubr1-/- and the conditional viability of Ubr2-/-, Ubr1-/-Ubr2-/- embryos exhibited neurogenetic and cardiovascular developmental defects, and died at midgestation (An et al., 2006). At embryonic day 9.5 (E9.5), Ubr1-/-Ubr2-/- embryos were indistinguishable from their wildtype littermates. By E10.5, the growth of double-mutant embryos has ceased and developmental defects - kinked neural tubes, thin neuroepithelium, absence of interatrial and interventricular septa, disorganized myocardial wall, and poorly developed blood vessels - were apparent. By E12.5, all Ubr1-/-Ubr2-/- embryos were dead. Decreased cell proliferation and increased were responsible for these defects in neurogenesis and cardiovascular development in Ubr1-/-Ubr2-/- embryos (An et al., 2006).

Interestingly, impaired neurogenesis and cardiovascular development was also observed in mouse embryos that lacked components in the Notch signaling pathway, suggesting a potential relationship between Ubr1-/-Ubr2-/- and reduced Notch signaling. The level of Notch1, a - activated receptor is significantly reduced in Ubr1-/-Ubr2-/- embryos (An et al., 2006). Notch1 plays a critical role in embryonic development through regulating multiple cellular processes, including proliferation, differentiation and apoptosis (Yoon et al., 2005). The decreased Notch1 level may correspond with the embryonic developmental defects and the resulting lethality

19 observed in Ubr1-/-Ubr2-/- mice. However, substrates through which UBR1 and UBR2 influence the level of Notch signaling remain unknown.

Several physiological substrates of mouse UBR1 have been identified, including the RGS (regulator of G protein signaling) family proteins RGS4, RGS5, and RGS16; the Parkinson’s disease-associated mitochondrial /threonine kinase PINK1; and, possibly, the susceptibility type 1 gene product BRCA1 (Hu et al., 2005; Lee et al., 2005; Yamano and Youle, 2013; Xu et al., 2012).

RGS4, RGS5, and RGS16 downregulate G protein signaling by binding to specific Gα subunits of G proteins and increasing their intrinsic GTPase activity (Wieland and Mittmann, 2003). All of them bear a Cys-2 residue, which can be made N-terminal by MetAPs. The exposed Cys residue can be thereafter oxidized into Cys-sulfinate or Cys-sulfonate by NO/O2 and further arginylated by R-transferase, allowing for binding of UBR1, and subsequent degradation by the UPS (Hu et al., 2005; Lee et al., 2005). UBR1 thereby participates in the regulation of G protein signaling in a NO/O2-dependent manner. UBR2 was later shown to also participate in the binding and polyubiquitination of RGS4 and RGS5 (Lee et al., 2011).

PINK1 and are involved in mitochondrial quality control, and which are mutated in Parkinson’s disease. In dysfunctional mitochondria, newly synthesized mitochondrial kinase PINK1 precursor accumulates on the outer membrane, which recruits cytosolic PARKIN and promotes PARKIN-mediated (Narendra et al., 2008). In contrast, healthy mitochondria import the N-terminal part of the PINK-1 precursor into the inner membrane via two translocator complexes, TOM and TIMM23, at the outer and inner membranes, respectively (Chacinska et al., 2009). The mitochondrial intramembrane protease PARL can then cleave off the N-terminal transmembrane segment of the PINK1 precursor and retrotranslocate the C-terminal fragment to the (Deas et al., 2010). PARL-cleaved PINK1 harbours an N-terminal Phe, which is a Type-2 N-terminal destabilizing residue of the Arg/N-end rule pathway. It can thus be targeted by UBR1, UBR2 and UBR4 for rapid degradation (Deas et al., 2010; Yamano et al., 2013). In brief, UBR family E3 ligases protect healthy mitochondria from PARKIN-mediated autophagic elimination through rapid PINK1 turnover.

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1.2.4 The cell culture model

Mammalian cell culture models were employed in the identification and/or validation of substrate proteins of the mammalian Arg/N-end rule pathway. In fact, the RGS family protein RGS4 was first identified as a candidate substrate of mammalian UBR1 in an expression-cloning screen in a reticulocyte lysate (Davydov and Varshavsky, 2000). The relative abundance of the protein encoded by the mouse Rgs4 cDNA was elevated after the Arg/N-end rule pathway was selectively inhibited by dipeptides bearing N-terminal destabilizing residues. The stability of RGS4 was further examined in mouse L cells. In L cells, RGS4 was short-lived, with a half-life of 40-50min, and it was significantly stabilized when L cells were treated with the proteasome inhibitor MG132 (Davydov and Varshavsky, 2000). RGS5 and RGS16 are members of the same family as RGS4; they share similar N-terminal sequences. They were validated as the substrates of the Arg/N-end rule pathway in mouse embryos (Hu et al., 2005).

The property of PINK1 as a substrate of the Arg/N-end rule pathway was examined in both HeLa cells and mouse embryonic fibroblast (MEF) cells. A destabilizing residue, Phe, was exposed after the PINK1 precursor was cleaved by the protease PARL between A103 and F104 (Deas et al., 2010). A truncated form of PINK1104Δ which lacked the N-terminal 104 residues and bore a N- terminal Met, and a full length PINK1F104M in which Phe at residue 104 was mutated into a stabilizing Met exhibited prolonged half-lives in HeLa cells. PINK1 could also be significantly stabilized in Ubr1-/-Ubr2-/-Ubr4RNAi MEF cells (Yamano and Youle, 2013). Together, these results suggest that PINK1 is a substrate of the mammalian UBR1/UBR2/UBR4-dependent Arg/N-end rule pathway.

BRCA1 was examined as candidate substrates of the mammalian Arg/N-end rule pathway in HEK 293T, MDA-MB-468, MDA-MB-231, and MEF cells. The C-terminal fragment of BRCA1 generated by a caspase cleavage, which bears an N-terminal Asp residue, was short-lived with a half-life of ~3h in HEK293T, MDA-MB-468, and MDA-MB-231 cells. The C-terminal fragment could be significantly stabilized by treating the cells with the proteasome inhibitor MG132, or by mutating the Asp residue to a stabilizing Val one. In ate-/- MEFs, which lack the R-transferase ATE1, the BRCA1 C-terminal fragment was stable regardless of the N-terminal stabilizing or destabilizing residues (Xu et al., 2012). In short, BRCA1 can be degraded through the mammalian Arg/N-end rule pathway independent of cell type.

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1.2.5 The C. elegans model

C. elegans has three UBR family proteins by sequence homology: UBR-1, UBR-4, and UBR-5. UBR-1 is the only homologue to yeast Ubr1 in C. elegans. It contains conserved key functional domains, the UBR-box domain, the N domain, and the RING domain. The physiological functions of C. elegans UBR-1 are not fully understood (Weaver et al., 2017; Chitturi et al., 2018), but its in vitro and in vivo N-recognin property in the Arg/N-end rule pathway has been reported (Weaver et al., 2017).

The first and only reported substrate of the C. elegans UBR-1 is LIN-28 (Weaver et al., 2017). LIN-28 is an RNA-binding protein expressed in hypodermal seam cells, which are a -like cell type in C. elegans. It contains two potential RNA-binding motifs: a cold shock domain and a C-terminal domain with two retroviral-type zinc finger motifs (Moss et al., 1997). It binds pre-let- 7 to prevent Dicer processing-mediated maturation. This repression can be enhanced by an alternative step in which LIN-28 recruits the poly(U) polymerase PUP-2 to uridylate blockaded pre-let-7, which leads to the degradation of the LIN-28-pre-let-7 complex (Lehrbach et al., 2009). let-7 is a 21-nucleotide microRNA, also known as a small temporal RNA (stRNA). It negatively regulates several transcriptional factors to repress cell proliferation and promote terminal differentiation in seam cells at the larval to adult transition (Reinhart et al., 2000). Reinhart et al. (2000) showed that let-7(lf) mutants reiterated larval patterns of cell proliferation, and failed to produce alae at the L4-to-adult molt. At a permissive temperature (25°), these mutants went through an additional larval stage, in which seam cells stopped cell division and fused with neighboring seam cells to generate alae. After the extra larval stage, let-7 mutants reached the adult stage and died with vulval bursting (Großhans et al., 2005). Similarly, lin-28(gf) mutants, in which let-7 was downregulated, reiterated the L2 stage-specific pattern of cell division (Moss et al., 1997). In wildtype animals, L2 is the only stage in which midbody seam cells undergo two rounds of cell division, resulting in an increase in the number of seam cells. lin-28(gf) mutants showed an accumulation of seam cells caused by a greater increase in number (Moss et al., 1997). In contrast to lin-28(gf) mutants, lin-28(lf) mutants skipped the double-division pattern of seam cells, as well as other L2-specific developmental events, exhibiting a decreased number of seam cells and precocious development (Rougvie and Moss, 2013).

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UBR-1, coupled with the CED-3 caspase and ATE-1 R-transferase, limits the LIN-28 protein level to regulate seam cell proliferation and developmental timing (Weaver et al., 2017). CED-3 cleaves LIN-28 and exposes an Asn residue that can be arginylated by ATE-1, which allows the C-terminal fragment targeted by UBR-1 for rapid degradation. ubr-1(lf) and ced-3(lf) mutations could increase, in a non-additive manner, both LIN-28 protein levels and the number of L4 seam cells (Weaver et al., 2017).

LIN-28 is unlikely the only substrate of UBR-1, because UBR-1 exhibits robust and persistent expression in neurons and muscles, and only transition expression in the seam cells (Chitturi et al., 2018). This work (from our lab) revealed that ubr-1(lf) mutants exhibited a unique motor defect: reduced bending during backward locomotion. Interestingly, this motor phenotype was unrelated to LIN-28 regulation, and was caused by aberrant glutamate metabolism. Specifically, removing GOT-1 (a glutamate synthesizing enzyme), EAT4 (a vesicular glutamate transporter), or AVR-15 (a glutamatergic chloride receptor) restored ubr-1 mutants’ bending during reversals. UBR-1 and GOT-1 are expressed in multiple somatic tissues, including the hypodermis, muscles, and subgroups of neurons. Restoring UBR-1 in subgroups of neurons, including the premotor interneurons (INs), rescues ubr-1 mutants’ bending defect. Expressing GOT-1 in the same set of neurons reverts the bending pattern of ubr-1; got-1 to that of ubr-1 mutants. These results suggest that UBR-1 affects glutamate metabolism in a subgroup of neurons, which in turn modulates motor outputs. However, the total protein level of GOT-1 was not changed in ubr-1 mutants. Instead, GOT-1’s enzymatic activity was elevated in ubr-1 mutants.

These studies suggest that GOT-1 may not be a substrate directly targeted by UBR-1 for degradation. Nevertheless, this study offered the first insight into the physiological role of C. elegans UBR-1 in the nervous system. Since glutamate is also an essential neurotransmitter and metabolite, the mechanism by which UBR-1 affects glutamate metabolism may have direct implications for the pathophysiology of JBS symptoms.

1.3 The locomotion of C. elegans

Locomotion is fundamental and essential to behaviour. C. elegans is the first animal with a nearly complete adult neural connectome. The investigation of the neural mechanisms underlying the locomotion in C. elegans may thus contribute to a fully integrated model explaining the functions of the nervous system from sensory inputs to behavioural outputs.

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C. elegans locomotes by propagating dorso-ventral body bends. Under standard laboratory conditions, animals exhibit sinusoidal body postures on the agar surface during both forward and backward locomotion. In water or other liquids with a similar viscosity, they swim at higher frequencies of undulation with longer wavelengths, adopting C-shape conformations (Pierce- Shimomura et al., 2008). Quantitative analysis of undulation, from wavelength, frequency, curvature to velocity, through automated image processing has developed (Sawin et al., 2000; Pierce-Shimomura et al., 2008; Kawano et al., 2011). Automated image processing has facilitated the development of algorithms that reduce dimensionality of parameters of locomotion (Stephens et al., 2008; Gomez-Marin et al., 2016; Broekmans et al., 2016).

1.3.1 The models for the mechanism of locomotion

Two key aspects of the C. elegans locomotion are the generation of dorso-ventral bends and their directional propagation, which involve central pattern generation and sensory feedback (Wen, Gao and Zhen, 2018).

The concept of Central Pattern Generator (CPG) refers to a neuron or a neural network that generates autonomous rhythmic electric activity patterns, independent of inputs from descending neural networks or sensory organs. Different CPG modules have been found in the vertebrate spinal cord and its invertebrate counterpart, the ventral nerve cord (Juvin et al., 2007; Grillner et al., 1984; Friesen and Cang, 2001). The CPG rhythm is adaptive to the environment by incorporating sensory feedback (Cang and Friesen, 2002; Grillner, 2006; Gjorgjieva et al., 2013). To explain the machinery of C. elegans locomotion, three competing hypothetic models have been put forward, proposing different types of CPGs and functions of sensory feedback (Gjorgjieva et al., 2014).

In the first model, a head CPG initiates undulatory body bends that are propagated along the body to drive forward locomotion (Niebur and Erdös, 1991). The environment provides sufficient lateral forces to support the sinusoidal body posture and generate forward thrust. The discovery that B- type motor neurons (B-MNs) transduce proprioceptive signals to drive and organize forward locomotion in a viscosity-dependent manner may support the function of proprioception-driven propagation in this model (Wen et al., 2012).

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In the second model, multiple oscillators are present along the body to generate rhythmic body bends and act as phase-coupled CPGs to propagate them. Sensory feedback modifies their activities to produce various motor patterns. This model is derived from other organisms that were found to contain a set of coupled CPGs, such as lampreys, and stick insects (Williams et al., 1990; Bässler and Büschges, 1998; Marder and Bucher, 2001). CPGs of this type are modulated by sensory feedback to generate flexible motor outputs (Cang and Friesen, 2002; Grillner et al., 2006; Gjorgjieva et al., 2013).

In the third model, sensory feedback is responsible for the generation and propagation of body bends completely independent of CPGs. It was first suggested by Niebur and Erdös when they put forward a theory on the control of somatic MNs by INs in 1993. Instead of an autonomously oscillating neuron or neural network, this model assumes a constant input from an IN. Sensory feedback, specifically proprioceptive feedback, integrates this input to generate oscillations in modules along the body and set phase lags between them for forward crawling or swimming behaviours (Bryden and Cohen, 2008).

Recent studies reveal evidence that strongly favor the first two models. The A-type motor neurons (A-MNs) and mid-body B-type motor neurons (B-MNs) exhibit coupled oscillatory activities independent of inputs from premotor INs, serving as at least part of the CPGs for backward and forward locomotion, respectively (Xu et al., 2018; Fouad et al., 2018; Gao et al., 2018; Wen, Gao and Zhen, 2018).

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Figure 4. Three proposed hypothetic models for the generation of locomotor patterns

(A) A model containing one single CPG: a single head CPG generates rhythmic dorsoventral body bends; sensory feedback propagates these bends along the body from anterior to posterior to drive forward locomotion. (B) A model containing a set of coupled CPGs: coupled CPGs along the body generate and propagate rhythmic body bends; sensory feedback modulates the oscillations to produce flexible motor outputs. (C) A model containing no CPG: premotor INs produce a constant input; sensory feedback along the body generates and propagates rhythmic body bends. Figure panels modified from Gjorgjieva et al., 2014.

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1.3.2 The neural mechanism underlying backward locomotion

Gao et al. (2018) showed that the second model, in which coupled CPGs generate and propagate body bends while sensory feedback modulates the motor outputs, captures backward locomotion. A-MNs serve as a set of coupled CPGs along the body, driving backward locomotion. They exhibit intrinsic oscillatory activities independent of inputs from premotor INs. The activities are phase- coupled and are sufficient to facilitate rhythmic alternating dorso-ventral body bends to drive slow backward locomotion. The removal of either all premotor INs or all chemical synaptic transmission in the nervous system failed to abolish the rhythmic activities of A-MNs (Gao et al., 2018).

The rhythmic activity pattern of oscillatory neurons requires coupled activity of a plethora of sodium, calcium and potassium channels (Izhikevich, 2007; Harris-Warrick, 2002). UNC-2, the C. elegans orthologue of the P/Q/N type high-voltage-activated calcium channel, is involved in the process of rhythm generation. UNC-2 conductance in A-MNs is positively correlated with the amplitude and frequency of calcium oscillation and the resulting muscle rhythmic postsynaptic current (rPSC) bursting (Gao et al., 2018). Other channels contributing to the oscillations of A-MNs remain to be explored.

The oscillatory activities of A-MNs are phase-coupled, exhibiting appropriate lags between them, which allows for the propagation of alternating dorso-ventral bends along the body. The mechanism underlying phase coupling remains to be investigated. B-MNs’ ability to transduce proprioceptive signals (Wen et al., 2012) strongly suggests that proprioceptive entrainment in A- MNs establishes this phase coupling. While direct experimental evidence for proprioception in A- MNs is lacking, several lines indirect evidence support the notion. Sparse removal of a small number of mid-body A-MNs blocked local body bending, leaving the oscillations of the anterior and posterior body regions uncoupled in phase (Gao et al., 2018). Constraining the mid-body in a straight microfluidic channel also disrupted the coordination between the unconstrained anterior and posterior body regions, resulting in different oscillatory frequencies (Wen, Gao and Zhen, 2018).

AVA premotor INs, which project along the ventral nerve cord to connect with all A-MNs through both gap junctions and chemical , modulate A-MN’s oscillatory activities to generate reversal locomotory patterns. Through gap junctions, AVA premotor INs shunt the spontaneous

27 oscillation of A-MNs, setting forward locomotion as the default motion state. Upon depolarization, AVA premotor INs overcome the shunting effect and use chemical synapses to potentiate A-MNs to sustain backward locomotion (Kawano et al., 2011; Gao et al., 2018). AVA premotor INs are downstream to a number of sensory neurons, and are depolarized upon various sensory inputs.

Figure 5. A model for backward locomotion

(A) A-MNs harbour intrinsic oscillatory activities. Proprioceptive feedback establishes the phase coupling among them. AVA premotor INs connect with A-MNs through both gap junctions and chemical synapses. Through gap junctions, they can shunt A-MN activities; through chemical synapses, they can potentiate A-MN activities. (B) A-MN exhibit oscillatory activities when premotor INs are ablated. Figure panels modified from Gao et al., 2018.

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1.3.3 The neural mechanism underlying forward locomotion

Mechanisms underlying forward locomotion also involve CPG and proprioceptive feedback (Xu et al., 2018; Fouad et al., 2018). A few mid-body B-MNs exhibit oscillatory activities, acting as a part of the underlying force for forward motor rhythms (Xu et al., 2018; Fouad et al., 2018), but their intrinsic activities are low when compared to A-MNs (Gao et al., 2018). It remains to be explored whether channels that underlie A-MNs’ intrinsic membrane potential oscillation, such as UNC-2, also contribute to the CPG properties of B-MNs.

Oscillating B-MNs also require proprioceptive phase-coupling to organize forward locomotion. Proprioception in B-MNs is local and directional, from anterior to posterior (Wen et al., 2012). Inhibiting head muscles or anterior B-MNs by optogenetic approaches resulted in the mid-body undulating at higher frequencies, suggesting that anterior-posterior proprioception entrains the activities of B-MNs to generate coherent propagating undulation (Xu et al., 2018; Fouad et al., 2018).

A pair of premotor INs AVB project along the ventral nerve cord to connect with all B-MNs through gap junctions. Optogenetic depolarization of AVB potentiates forward locomotion, while hyperpolarizing AVB halts forward locomotion, both in an AVB-B electrical coupling-dependent manner. Without the AVB-B gap junctions, bending amplitudes decayed rapidly from anterior to posterior, suggesting a reduction in B-MN oscillatory activities, and/or effectiveness of proprioceptive feedback (Xu et al., 2018). Sensory inputs may regulate the depolarization and hyperpolarization of AVB, thus modulating oscillations and possibly proprioceptive feedback of B-MNs to produce flexible motor outputs.

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Figure 6. A model for forward locomotion

(A) To drive forward locomotion, B-MNs’ intrinsic oscillations, transduction of proprioceptive signals, and electrical coupling with AVB premotor INs work synergistically to generate and propagate rhythmic dorsoventral bends along the body from anterior to posterior. (B) When the oscillations of anterior B-MNs are inhibited, the proprioceptive signals from anterior body are absent; mid-body exhibits high frequency undulations that are induced by AVB-B gap junction coupling. (C) When inputs from AVB premotor INs through gap junctions are abolished, proprioceptive feedback is weakened and becomes less efficient in propagating body bends, leading to a significant decay in bending amplitude towards the tail. Figure panels modified from Xu et al., 2018.

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1.4 Objectives

My study extends from the previous work of Dr. Chitturi that uncovered motor defects caused by aberrant glutamate metabolism in ubr-1 mutants. I applied genetic, behavioural and calcium imaging approaches to investigate the circuit mechanism by which UBR-1 modulates motor outputs. I reveal a critical requirement of UBR-1 in mechanosensory neurons, in addition to premotor INs, to establish body bending. My work further reveals that ubr-1 mutations cause reduced bending by disrupting phase coupling among A-MNs through premotor INs. The hypothesis derived from the results is that ubr-1 mutations affect sensory neurons through aberrant glutamate signaling and premotor INs through glutamate toxicity. This hypothesis may provide hints for the pathogenesis of the neurological symptoms of JBS, such as deafness and cognitive impairment.

Chapter 2 Materials and Methods 2.1 Molecular biology and C. elegans strains 2.1.1 Strains, constructs and oligos

See Table 1-2 for complete lists of strains, constructs and oligos used in this study. All strains were cultured under standard conditions, on Nematode Growth Medium (NGM) plates seeded with OP50 Escherichia coli bacteria and maintained at 22°C (Brenner, 1974). Transgenic strains used in this study include the ones with extra-chromosomal multi-copy arrays (hpEx), integrated multi- copy arrays (hpIs) and one ubr-1 RING domain replacement allele (hp). Transgenic strains carrying extra-chromosomal arrays (hpEx) were generated by co-injecting the target DNA plasmid and a co-injection marker at ~10ng/μL. Extra-chromosomal arrays were integrated into the genome through UV irradiation (hpIs). All integrated strains were outcrossed against the wildtype (N2) strain for several times before experiments.

2.1.2 UBR-1 RING domain replacement allele

The UBR-1 RING domain replacement allele, hp865, was generated by the CRISPR-Cas9 system- mediated homologous recombination system as described (Norris et al., 2015). The replacement template for UBR-1 RING domain replacement (pJH3806) contains a upstream homology arm (800bp sequence upstream to the UBR-1 RING domain), a SL2-NLS::GFP reporter with a loxP- Pmyo-2-GFP-Prps-27-NeoR-loxP (NeoR denotes a neomycin resistant gene) dual-marker cassette inserted in an intron region of GFP, and a downstream homology arm (800bp sequence downstream to the UBR-1 RING domain), allowing the cassette to be inserted and replace the entire sequence encoding the RING domain. N2 (wildtype) animals were injected with the replacement template, two plasmids expressing sgRNA located near the 5’- and 3’-ends of the RING domain-coding sequences, one Cas9 protein expressing plasmid and two co-injection markers. After injection, the animals were allowed to recover and lay eggs at 25°C for 24h before G418 neomycin treatment. Candidate progenies for insertion were selected based on the pharyngeal GFP signal, the G418 resistance and the absence of co-injection markers. Animals with co-injection markers were carrying extrachromosomal arrays. Animals with a correct insertion

31 32 were confirmed by phenotyping, genotyping and sequencing. They were then injected with a Pelt- 3::Cre plasmid to remove the dual-marker cassette between the two loxP sites. The resulting allele, hp865, was selected based on the loss of the GFP marker and again confirmed by phenotyping, genotyping and sequencing. It was outcrossed against N2 wildtype animals prior to any further studies.

Table 1. Strains and constructs used in this study

Strain Genotype Plasmid Host strain Description

ubr-1 alleles

VC3668 ubr-1(gk3640)I pJH3804 N2 RING domain replaced by multiple copies of a SL2- /ZM9492 pJH3805 NLS::GFP cassette (with a pJH3806 dual-marker cassette in an intron region). The selection markers cannot be excised possibly due to the multicopy insertion.

ZM9621 ubr-1(hp861)I pJH3804 N2 RING domain replaced by a SL2-NLS::GFP cassette pJH3805 (with a dual-marker pJH3806 cassette in an intron region)

ZM9671 ubr-1(hp865)I pDD104 ZM9621 RING domain replaced by a SL2-NLS::GFP cassette

ZM8230 ubr-1(hp684)I Q1864X

Transgenes for rescuing ubr-1

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ZM9647 ubr-1(hp865)I; pJH2913 ZM9671 Pubr-1-ubr-1 hpEx3898

ZM9782 ubr-1(hp865)I; pJH3380 ZM9671 Prgef-1-ubr-1 hpEx3535

ZM9742 ubr-1(hp865)I; pJH4050 ZM9671 Pift-20-ubr-1 hpEx3929

ZM9767 ubr-1(hp865)I; pJH4071 ZM9671 Pmec-3-ubr-1 hpEx3949

ZM9770 ubr-1(hp865)I; pJH4078 ZM9671 Pser-2prom3-ubr-1 hpEx3952

ZM9781 ubr-1(hp865)I; pJH3436 ZM9671 Pnmr-1-ubr-1 hpEx3564

ZM9787 ubr-1(hp865)I; pJH3381 ZM9671 Pmyo-3-ubr-1 hpEx3536

ZM9761 ubr-1(hp865)I; pJH4051 ZM9671 Pmec-4-ubr-1 hpEx3943

ZM9762 ubr-1(hp865)I; pJH4066 ZM9671 Psra-6-ubr-1 hpEx3944

ZM9807 ubr-1(hp865)I; pJH3436 ZM9767 Pmec-3-ubr-1 hpEx3949; hpEx3960 + Pnmr-1-ubr-1

ZM9809 ubr-1(hp865)I; pJH4071 ZM9781 Pnmr-1-ubr-1 hpEx3564; hpEx3963 + Pmec-3-ubr-1

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ZM9836 ubr-1(hp865)I; pJH4050 ZM9781 Pnmr-1-ubr-1 hpEx3564; hpEx3976 + Pift-20-ubr-1

Strains for optogenetic cell ablation and calcium imaging

ZM8428 hpIs459 pJH3137 lin-15 Punc-4- GCaMP6::wcherry

ZM8429 hpIs460 pJH3137 lin-15 Punc-4- GCaMP6::wcherry

ZM9176 hpIs603 pJH3449 lin-15 Pacr-5- ::miniSOG-UrSL- pJH3453 ebfp;

Plgc-55- tomm20::miniSOG-UrSL- ebfp

ZM9228 hpIs603; hpIs459 ZM8428× ZM9176

ZM10047 ubr-1(hp865)I; ZM9671× ZM9176 hpIs603

ZM10124 ubr-1(hp865)I; ZM9671× ZM9228 hpIs603; hpIs459

ZM8749 ubr-1(hp684)I; ZM8230 × ZM8429 hpIs460

ZM8846 ubr-1(hp684)I; pJH3476 ZM8749 Popt-3-ubr-1 hpIs460;hpEx3676

ZM8750 ubr-1(hp684)I;got- ZM8226 × ZM8429 1.2(hp731)X;hpIs460

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Other strains used in this study

ZM9603 ubr-1(gk3640)I; From a genetic suppressor hp856X screen in ZM9492

ZM9971 ubr-1(hp865)lin- 28(n947)I

ZM10005 ubr-1(hp865)lin- 28(n719)I

ZM8226 ubr-1(hp684)I; got- From a genetic suppressor 1.2(hp731)X screen in ZM8230

Table 2. Primers for genotyping strains in this study

Gene and allele Primer Wildtype Band Mutant Band

ubr-1(hp861) OZM4814/1717 none 1467bp

ubr-1(hp861) OZM4814/3518 1773bp None

ubr-1(hp861) OZM2796/5243 none 1015bp

ubr-1(hp861) OZM3458/5243 823bp None

ubr-1(hp865) OZM5246/5243 none 1698bp

ubr-1(hp865) OZM4814/2875 none 2201bp

ubr-1(hp684) OZM3533/3534 262bp 217bp

+OZM3535/3536

got-1.2(hp731) OZM3259/3261 748bp none

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got-1.2(hp731) OZM3260/3261 none 749bp

lin-28(n719) OZM5472/5473 296bp 269bp

+OZM5474/5475

Table 3. Sequences of primers used for genotyping in this study

Primer Sequence Genotyping

OZM1717 GGAACAGGTAGTTTTCCAGTAGTGC ubr-1(hp861)

OZM2796 TTAGACACAACATTGAAGATGGAA ubr-1(hp861)

OZM2875 TTTGTATAGTTCATCCATGCCATGTG ubr-1(hp865)

OZM3458 GCCGTTGAGTAGTCAAATCCAGCATG ubr-1(hp861), ubr-1(hp865)

OZM3518 ACGTGTAGATATTGTCCCACTTG ubr-1(hp861), ubr-1(hp865)

OZM4814 ACCTTCTTCATCGATTCCGTCT ubr-1(hp861), ubr-1(hp865)

OZM5243 TAGTGGAGGTACGATGAAATGG ubr-1(hp861), ubr-1(hp865)

OZM5246 AGTAAAGGAGAAGAACTTTTCACTGG ubr-1(hp865)

OZM5472 AATGGAAAAGGTAGAGAAGCTTATGAGA lin-28(n719)

OZM5473 GCTGTTAGAAAAAATAAAAATAAAACGCAC lin-28(n719)

OZM5474 AAGAAGATGTCAATTAATAACTGCGTCA lin-28(n719)

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OZM5475 CATTCAAAATATAGGTTAAAGCAGCGTT lin-28(n719)

OZM3533 TCTGGGTTTACCGCATAAGATTGAAGCAT ubr-1(hp684)

OZM3534 CAACTTTTCGAGCATACTTCCACTGATGTG ubr-1(hp684)

OZM3535 TCAGACGATCAGGAAGCTTTACGCTCTT ubr-1(hp684)

OZM3536 TCAATCGGTATCGTTGGACATTTGAATG ubr-1(hp684)

OZM3259 AAAGAACGTATGGAGTTCAGTG got-1.2(hp731)

OZM3260 GAAAGAACGTATGGAGTTCAGTA got-1.2(hp731)

OZM3261 TGTAAAGTCCAAAGTTCTTGGC got-1.2(hp731)

2.2 Locomotory behaviour assay 2.2.1 Image acquisition

The video recordings of the locomotion of C. elegans were acquired from a Hamamatsu Orca-R2 camera on a Zeiss Axioskop 2 Plus microscope equipped with an ASI MS-40000 motorized stage. The image was taken for one single one-day old young adult worm each time on a seeded 35mm NMG plate. L4 stage worms were prepared one day prior to the experiments. In order to acquire images with high contrast between the outline of the worm body and the background bacterial lawn, the bacterial lawn was scrapped off by a sterilized bended glass rod, leaving only a thin layer. Under these conditions, the worms spend the vast majority of the time performing forward locomotion. To make the quantitative analysis of backward locomotion possible within a reasonable length of recording, copper chloride (CuCl2) was used to increase the fraction of backward locomotion without affecting other characteristics of the locomotion patterns of animals with different genetic backgrounds. A CuCl2 ring of 15mm in diameter was created by applying a fixed amount (100μL) of 100mM CuCl2 solution evenly dropped around the center of the NGM plate (Fig. 7B). The CuCl2 ring was let dry at room temperature for 10 min to fix before transferring

38 the animal to the center of the plate. The animal was let habituate for 1min before imaging. The behaviour of one single animal was recorded for 3min and analyzed.

2.2.2 Motor activity quantification

The recordings were processed and analyzed using ImageJ plugins and R scripts developed by previous members of Zhen Lab, Dr. Taizo Kawano and Dr. Michelle Po, respectively. The scripts have two major outputs: the bending angles and the speed. The outline of the worm body was recognized automatically, with the head and tail tips determined based on the sharpness of the shape, which two points were later manually curated. The images without a clear body outline, wherein the worm touched themselves or the edge of the imaging field, were not processed. The centerline was then calculated from the outline and evenly divided into 32 segments by 33 marker points including the head and tail tips. The angles between adjacent segments were calculated. In total, 31 bending angles were derived from 33 centerline points (Fig. 7C). The head and tail parts of a hermaphrodite worm of normal body shape are always thinner than the mid body. Consequently, the head and tail tips were frequently out of focus during the imaging process, resulting in large systematic errors in the position recognition step during the analyzing process. Therefore, the first and last bending angles, calculated with the positions of the head and tail tips respectively, were not included in the assessment of the body bending. The rest 29 bending angles presented the body bending of the imaged worm at the time point of the frame. The other major output, the moving velocity, was calculated by integrating the velocity of the motorized stage and the displacement of the center point of the body image. The directionality of the movement (forward or backward) was determined by the angle between the absolute velocity and the anterior- posterior axis of each frame.

In this study, the reversal events were of primary interest. A period with 6 or more continuous frames exhibiting “backward” speed was defined as a reversal event. I extracted the bending angle and velocity stats to calculate three major parameters: (1) backing initiation frequency, the frequency of the reversal events across of the 3min recording; (2) backing duration, the length of each reversal event; (3) body bending, the 29 bending angles averaged across each reversal event.

In order to assess rescue effects of transgenes that restored UBR-1 expression in different tissues, the term normalized posterior body bending difference was created. It was defined as the difference between the mean of the three most posterior bending angles of a rescue strain during each reversal

39 event and that of the ubr-1 mutant control over all reversal events, normalized to the difference of the mean of these three angles over all reversal events between the wildtype and ubr-1 mutant controls.

2.3 Fluorescence microscopy

Animals carrying a GFP::UBR-1 reporter or a RFP marker were fixed and stained by anti-GFP or anti-RFP antibody before imaging. Images of young adult/L4 larvae and L2/L3 larvae were acquired on a Nikon Eclipse 90i confocal microscope using a 60X and a 100X objective, respectively.

2.4 Optogenetic cell ablation

MiniSOG (mini singlet oxygen generator) is an optogenetic cell ablation tool, which can produce singlet oxygen in the mitochondria upon blue light illumination to induce programmed cell death at single-cell resolution (Qi et al., 2012). The L3 stage worms that express mitochondrially targeted miniSOG specifically in premotor INs (AVA, AVE, AVD, PVC, AVB) and B-MNs (hpIs603) were subjected to blue light illumination for 45min and allowed for recover overnight in darkness at 22°C. EBFP was co-expressed in the same set of neurons for examining the morphologies of these neurons before and after the blue light illumination. Ablated neurons showed an unstructured soma without neurites, while peripheral tissues, most prominently muscles exhibited scattered EBFP signals. Animals lacking this set of neurons can only conduct continuous slow backward locomotion or stay still. The fluorescence and locomotion patterns of animals after ablation were visually examined before the calcium imaging experiments.

2.5 Calcium imaging

Animals of different genotypes with the A-MN calcium imaging reporter hpIs460 (Punc-4- GCaMP6::wcherry) were used for calcium imaging experiments. Animals were placed in a drop (~2μL) of M9 buffer on a 2.5% agar pad and gently covered with a cover slip. Images were acquired under a 40X objective on a Zeiss Axioskop 2 Plus Microscope equipped with an ASI MS-40000 motorized stage, a Photometrics dual-view beam splitter and a Hamamatsu Orca-R2 CCD camera. The fluorescent imaged were split by dual-view with a GFP/RFP filter on the camera. The GFP and RFP channels were calibrated for good alignment before imaging. Each recording was acquired at 10 fps for 3min.

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The analysis of A-MN calcium imaging was processed with Matlab scripts developed by Yangning Lu, a Ph. D. candidate of Zhen Lab, based on the method initially from Dr. Andrew Leifer (Leifer et al., 2011). The first step of the analysis of A-MN calcium imaging was to select a minimum region of interest (ROI) that covers the soma of an A-MN in the RFP channel. The corresponding area in the GFP channel targeting the same neuron was registered with fine adjustments in the meantime. Then the neuron was traced throughout the recording. Manual correction was required in cases when the auto-tracking went wrong due to interference on the signal of the target neuron. The signals were presented as the average intensities within the ROIs (selected and auto-registered). The ratio of the GFP signal to its corresponding RFP signal was used to represent the activity of the neuron in question. The displacement of the ROI was also recorded and combined with the stage velocity to calculate to the absolute velocity of the animal.

This project studied the relation between A-MN activities during backward locomotion. Three A- MNs, VA10, DA7 and VA11, were selected in consideration of the high signal quality of and the appropriate physical distance between them. The activities of these A-MNs were analyzed as described above one by one. The directionality of the movement could then be determined by the angle between the absolute velocity and the anterior-posterior axis derived from the positions of two A-MNs. Traversing the directionality changes throughout the recording, reversal events that lasted for more than 2s were extracted. Subsequently, the time lags between the activities of the three A-MNs in each reversal event could be calculated based on the cross correlations between the time-dependent activity traces of them.

2.6 Statistical analysis

For bending angles in behavioural assays, comparisons were performed using Mann-Whitney tests. For normalized posterior body bending, Mann-Whitney tests were applied in the comparisons. For time lags between A-MN activities in calcium imaging, comparisons were conducted among three or four groups (N2, ubr-1, experimental group(s)) with One-way ANOVA Kruskal-Wallis tests. Differences with P-values smaller than 0.05 were considered statistically significant. All statistical tests were performed with the statistical tools in the GraphPad Prism5 software.

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Chapter 3 Results 3.1 ubr-1 mutants exhibit reduced body bending during reversals

UBR-1 has several functional domains including two putative degron recognition domains and a ubiquitination domain (Fig. 7A). All previously identified alleles of ubr-1 selectively lesion protein sequences outside these functional domains, raising a possibility that phenotypes observed in previous ubr-1(lf) alleles may not reflect the consequence of the lost UBR-1 E3 ligase activity. The RING domain is a critical for the ubiquitin transfer property of E3 ligases (Budhidarmo et al., 2012). I used the CRISPR-Cas9 system to replace the whole RING domain with a SL2-NLS::GFP cassette, and used the resulting allele (ubr-1(hp865)) for phenotypic analyses as well as mechanistic studies (Methods).

Under standard conditions, a wildtype C. elegans crawls on agar surface by propagating sinusoidal body bending in either forward or backward direction. ubr-1 mutant adults appeared normal in forward locomotion. They were able to perform backward locomotion; however, their body failed to exhibit full bending during reversals (Fig. 7D). The reduced bending phenotype was progressive as worms developed from larval stages to adults.

I performed behavioural assays on wildtype (N2), ubr-1(hp865) and ubr-1(hp865); hpEx3898[Pubr-1-UBR-1] one-day-old young adults (Methods). My quantification of their bending angles during reversals showed that ubr-1(hp865) adults exhibited statistically significant reduction of mean bending angles when compared to N2 animals. The reduction was most prominent at the posterior segments. Restoring the expression of wildtype UBR-1 from its endogenous rescued the reduced bending phenotype (Fig. 7E).

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Figure 7. ubr-1 mutants exhibit reduced bending angles at posterior body during reversals

(A) A ubr-1 loss-of-function mutation allele, ubr-1(hp865), was generated by the CRISPR-Cas9 system, where the key functional domain for substrate ubiquitination is completely replaced by a SL2-NLS::GFP cassette. (B) Behaviour assay was performed on a low-food plate circled by a CuCl2 ring, with one single worm recorded each time. (C) The image of the worm body is evenly divided into 32 segments. The angles between adjacent segments are calculated. (D) Representative body posture during reversals. Left panel: a wild type animal exhibits a sinusoidal body posture. Middle panel: a ubr-1(hp865) mutant shows reduced bending angles at posterior body. Right panel: a ubr-1(hp865);hpEx3898 animal, which has restored expression of the ubr-1 genomic fragment under ubr-1(hp865) background, exhibits sinusoidal bending along the body. (E) Mean bending angles of different body segments across reversal events. Left panel: mean bending angles along the body; right panel: five mean bending angles at the most posterior body. Data is presented as mean ± SEM. n=22, *P<0.05, **P<0.01, ***P<0.005 by Mann-Whitney Test.

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3.2 UBR-1 is expressed in multiple tissues

I generated the hp865 allele to examine the behavioral consequence of the functional loss of UBR- 1, as well as to determine the endogenous expression pattern of UBR-1 (Fig. 7A). However, the knock-in GFP did not display detectable signal, possibly due to low expression levels. To determine the expression of UBR-1, Dr. Wesley Hung constructed a functional GFP::UBR-1 reporter driven by its endogenous promoter, and generated a multi-copy transgenic array (ubr- 1(hp865);hpEx3922) to examine the expression pattern.

I examined this strain, and identified persistent cytoplasmic GFP expression in the head and body wall muscles and a subgroup of neurons (Fig. 8). UBR-1::GFP expressing neurons include sensory neurons (FLP, PVD, and others) and premotor INs (AVA, AVE, AVD, and others) (Fig. 8D, 8F). UBR-1::GFP was absent from most motor neurons, with a few rare exceptions (SIA, SIB and DVB) (Fig. 8D-8F). I also observed weak and transient GFP expression in the hypodermal seam cells (Fig. 8C).

3.3 UBR-1 is critically required in mechanosensory neurons and premotor INs to affect the reversal motor pattern

I further examined the tissue requirements of UBR-1 by restoring the expression of the UBR-1 genomic fragment specifically in tissues of interest, and quantified these transgenes’ rescuing capacities of the reduced bending at the posterior body in ubr-1 mutants (Fig. 9; Table 4). Transgenes that express UBR-1 from the endogenous promoter (Pubr-1), or exogenous promoters that drive expression in all neurons (Prgef-1), or in a subset of premotor INs (AVA, AVE, AVD, PVC, and RIM; Pnmr-1) restored the posterior bending of ubr-1 mutants to wildtype levels (Fig. 9; Table 4). Exogenous promoters that drive UBR-1 expression in ciliated sensory neurons (from Pift-20), mechanosensory neurons (PVD, FLP, AVM, ALM, PVM, and PLM; from Pmec-3), or two mechanosensory neurons (PVD and OLL; from Pser-2p3) partially rescued the reduced bending phenotype of ubr-1 mutants (Fig. 9; Table 4).

Based on UBR-1’s expression pattern and rescuing results, I conclude that UBR-1 is critically required in premotor INs (AVA, AVE and AVD) and mechanosensory neurons (PVD and FLP) to modulate the motor outputs (Fig. 13A).

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Figure 8. UBR-1 is expressed in muscles, hypodermis and sub groups of neurons

Confocal images of tissues that expressed a GFP::ZF::UBR-1 reporter from the endogenous promoter of ubr-1. The expression of the reporter is detected in head and pharyngeal muscles (A), body wall muscles (B), hypodermal seam cells (C), subgroups of neurons at the pharynx including sensory neurons (FLP) and premotor INs (AVA and AVE) (D), premotor interneuron along the ventral nerve cord (E), and two tail neurons (PVD and DVB) (F). Animals were fixed and stained by anti-GFP antibody. Neurons were identified based on relative positions. Panels (A) (B) (E) were acquired in adult worms under 60X. Panel (C) was acquired in a L3 worm under 100X. Panels (D) (F) were acquired in a L2 worm under 100X. Credit: Wesley Hung.

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Figure 9. Transgenes that restore UBR-1 expression in subgroups of neurons rescue the ubr- 1 reduced bending phenotype

Normalized posterior bending differences (Methods) of wildtype, ubr-1 mutants, and rescue strains are shown. Wildtype and ubr-1 mutants controls show normalized differences with a mean of 1 and 0, respectively. Rescues strains that express UBR-1 from Pubr-1, Prgef-1 (all neurons), and Pnmr-1 (premotor INs AVA, AVE, AVD, RIM, and PVC) show normalized differences that are significantly greater than 0 and not significantly difference from 1. Rescue strains that express UBR-1 from Pift-20 (all ciliated sensory neurons including PVD), Pmec-3 (mechanosensory neurons FLP, PVD, ALM, AVM, PLM, and PVM), and Pser-2p3 (mechanosensory neurons PVD and OLL) show normalized differences that are significantly greater than 0 and smaller than 1. Rescue strains that express UBR-1 from Pmyo-3 (body wall muscles), Pmec-4 (mechanosensory neurons ALM, AVM, PLM, and PVM), and Psra-6 (ASH, ASL, and PVQ) show normalized differences that have no significant difference from 0. Error bar: SEM. n=13-22, *P<0.05, **P<0.01, ***P<0.005 by Mann-Whitney test.

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Table 4. A list of transgenes that restore UBR-1 expression in different tissues tested for rescue effects of the ubr-1 reduced bending phenotype

Transgene Tissue Rescue

Pubr-1-UBR-1 Neurons and muscles ●

Prgef-1-UBR-1 All neurons ●

Pift-20-UBR-1 Ciliated neurons. ASH, OLL, FLP and others

Pmec-3-UBR-1 FLP, PVD, ALM, AVM, PLM, PVM

Pser-2 promp3-UBR-1 PVD, OLL

Pnmr-1-UBR-1 AVA, AVE, AVD, PVC, RIM ●

Pmyo-3-UBR-1 Body wall muscles ○

Pmec-4-UBR-1 ALM, AVM, PLM, PVM ○

Psra-6-UBR-1 ASH, ASI, PVQ ○

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3.4 Reduced bending results from synchronization of A-MNs’ intrinsic activities

In wildtype C. elegans, A-MNs harbour intrinsic oscillatory activities. These activities are phase- coupled, exhibiting lags between activation of adjacent A-MNs, probably through proprioceptive modulation among A-MNs (Gao et al., 2018).

By simultaneous calcium imaging of three adjacent A-MNs, VA11, DA7, and VA10, which innervate body wall muscles at the posterior body segments (Fig. 10E), in freely moving wildtype animals (Methods), I observed time lags between oscillatory activities of the A-MNs (Fig. 10A). Specifically, VA10 and VA11, two A-MNs that innervate adjacent ventral muscles (Fig. 10E) were phase-coupled in activation (Fig. 10A; left panel in Fig. 10A’), exhibiting various time lags (Fig. 10F). DA7 likely innervates dorsal muscles that are opposed to ventral muscles targeted by VA10 (Fig. 10E). This activity pattern captured the sequential contraction of adjacent muscles during backward locomotion at different velocities. Consistent with the alternating dorso-ventral muscle contraction during backward locomotion, VA10 and DA7 exhibited asynchronized activation (Fig. 10A; middle panel in Fig. 10A’), with various lags (Fig. 10G). DA7 and VA11 were activated with shorter lags (Fig. 10H), showing relative synchrony compared to the other two pairs (Fig. 10A; right panel in Fig. 10A’). This indicates a less direct dorsal-ventral opposition between DA7 and VA11 than that between DA7 and VA10.

In ubr-1 mutants, I observed a significantly altered activation pattern of A-MNs: all three A-MNs, VA11, DA7, and VA10, exhibited synchrony in activation (Fig. 10B, 10B’), with drastically reduced time lags between VA10 and VA11(Fig. 10F), and between VA10 and DA7 (Fig. 10G). The short time lags between DA7 and VA11 were not significantly changed (Fig. 10H).

Consistent with the results that the ubr-1 reduced bending phenotype could be rescued by both restored UBR-1 expression in premotor INs (AVE and RIM), and the got-1 mutation (Chitturi et al., 2018), the phase coupling among A-MNs was restored in the two respective strains (Fig. 10C- D’, 10F, 10G). Hence, the reduced bending phenotype in ubr-1 mutants results from increased synchronization in A-MN activation, rather than lack of A-MNs’ activities.

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Figure 10. UBR-1 promotes body bending by preventing synchronized A-MN activation

(A) An example trace of simultaneous calcium imaging of three A-MNs in a moving wildtype animal. Top panel: activities of VA11, DA7, and VA10 neurons, reflected by the GCaMP6/RFP ratio; bottom panel: the instantaneous velocity of the animal, reflected by the displacement of DA7 soma position (positive values indicate moving towards the head; negative values indicate moving towards the tail), during a period of 40s from a 3min recording. Boxed period denotes the reversal period applied for cross-correlation analyses. (A’) The cross-correlation of the activity profiles between VA10 and VA11 (left), VA10 and DA7 (middle), and DA7 and VA11 (right), respectively. Dotted vertical line denotes the lag time. (B, B’) An example trace of simultaneous calcium imaging (B) and cross-correlation analyses (B’) of VA11, DA7, and VA10 in a moving ubr-1 mutant. (C, C’) An example trace of simultaneous calcium imaging (C) and cross-correlation analyses (C’) of VA11, DA7, and VA10 in a moving transgenic ubr-1 mutant animal with restored UBR-1’s expression in neurons that include AVE/RIM premotor INs. (D, D’) An example trace of simultaneous calcium imaging (D) and cross-correlation analyses (D’) of VA11, DA7, and VA10 in a moving ubr-1; got-1 mutant animal. (E) Schematic diagram of three adjacent A-MNs (VA11, DA7, and VA10) and their predicted ventral and dorsal muscle targets, drawn based on Haspel and Donovan, 2011. (F-H) Phase lags between activities of VA10 and VA11 (F), VA10 and DA7 (G), and DA7 and VA11 (H) in animals of respective genotypes. The asynchrony between VA10 and VA11 (F), and between VA10 and DA7 (G) are significantly reduced in ubr- 1 mutants compared to wildtype animals, and restored by both UBR-1 expression in premotor INs, and the got-1 mutation. The activation of DA7 and VA11 (H), with higher synchrony than the other two pairs in wildtype animals, was not significantly altered in ubr-1 mutants. Horizontal lines represent mean values. n=10-15, *P<0.05, **P<0.01, ***P<0.005 by Kruskal-Wallis test.

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3.5 Reduced bending results from premotor IN-mediated dysregulation of the coordination of A-MNs’ intrinsic activities

UBR-1 is absent from the ventral cord motor neurons, including the A-MNs. A-MN activities are regulated by AVA premotor INs through a mixed configuration of gap junctions and chemical synapses (Gao et al., 2018). Hence, I speculated that in ubr-1 mutants, synchronized A-MN activation might be a consequence of dysregulation from premotor INs.

To test this hypothesis, I ablated all premotor INs using an optogenetic cell ablation tool, miniSOG (Methods), to remove the potential dysregulation to A-MNs (Fig. 11B), and imaged the activities of three adjacent A-MNs that innervate adjacent and opposing body wall muscles (Fig. 11A). Indeed, the phase coupling among these A-MNs was restored after the ablation (Fig. 11C-G). These results further support the notion that UBR-1 modulates the phase coupling of A-MN activities through premotor INs (AVA, AVE, and AVD) in the reversal motor circuit.

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Figure 11. Ablation of premotor INs in ubr-1 mutants restores the phase coupling of A-MN activation

(A) Schematic diagram of three adjacent A-MNs, VA11, DA7 and VA10, and the predicted ventral and dorsal body wall muscles that they innervate, drawn based on Haspel and Donovan, 2011. (B)

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Schematic diagram of premotor INs and MNs in the neural circuits controlling forward and backward locomotion. Grey box represents optogenetic cell ablation by miniSOG (Methods). (C) Diagram of the neural circuit through which UBR-1 regulates motor outputs, with premotor INs ablated as shown in (B) in wildtype animals. (C’) An example trace of simultaneous calcium imaging of three A-MNs in a moving wildtype animal after the ablation shown in (B). Boxed period denotes the reversal period extracted for the calculation of time lags. (D, D’) Diagram of the neural circuit through which UBR-1 regulates motor outputs (D) and an example trace of calcium imaging (D’) in a moving ubr-1 animal after the ablation shown in (B). (E, E’) Diagram of the neural circuit through which UBR-1 regulates motor outputs (E) and an example trace of simultaneous calcium imaging (E’) in a moving ubr-1 animal after mock ablation. (F, G) The time lags between the activity traces of VA10 and VA11 (F), and VA10 and DA7 (G). The asynchrony between VA10 and VA11 (F), and between VA10 and DA7 (G) were restored in ubr-1 mutants after ablating the premotor INs, compared to ubr-1 mutants with mock ablation. n=8-10, *P<0.05, **P<0.01, ***P<0.005 by Kruskal-Wallis test.

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3.6 Neuronal toxicity and aberrant glutamate signaling may account for the reduced bending phenotype in ubr-1 mutants ubr-1 mutations result in aberrant glutamate metabolism, significantly elevating the total glutamate level (Chitturi et al., 2018). High levels of glutamate may be harmful for cell metabolism, and in neurons, an increased glutamate level may directly affect glutamate signaling, leading to (Dong et al., 2009).

Together with Dr. Wesley Hung, a research associate in the lab, we used a cytosolic RFP marker (Pnmr-1-RFP) to examine the morphology of premotor INs from the L1 larva up to the 1-day old adults. In wildtype animals, neuron somata maintain a round and smooth surface throughout the developmental stages. In young larva (L1 to L3 stages) of ubr-1 mutants, the morphologies of all three examined premotor INs (AVA, AVE, and RIM) appeared similar to those in wildtype animals of the same stages (Fig. 12A). Starting from the L4 larva stage, in ubr-1 mutants, the somata of AVA and AVE premotor INs developed rough edges with short branches (Fig. 12A), whereas those of other premotor INs, such as RIM, appeared normal as in wildtype animals (Fig. 12C). The onset of the reduced bending phenotype in ubr-1 mutants - the end of the larval development - coincides with the timing of the morphological changes in AVA and AVE becoming prominent. Moreover, similar to the reduced bending phenotype, these morphological abnormalities in ubr-1 mutants could be rescued by removing GOT-1 (Fig. 12B), a metabolic enzyme that converts Aspartate to Glutamate, the got-1 mutations led to decreased glutamate level (Chitturi et al., 2018). These results suggest that there may be a causal relationship between the glutamate level-induced toxicity in premotor INs (AVA, AVE, and AVD) and the reduced bending phenotype in ubr-1 mutants. In sum, the functional loss of UBR-1 may cause the reduced bending phenotype through glutamate neural toxicity in premotor INs (AVA, AVE, and AVD), which affects premotor INs’ morphology and function in regulating A-MN activities; the dysregulation from premotor INs in turn disrupts the phase coupling among A-MNs, preventing the animals from establishing normal bending during backward locomotion (Fig. 13B).

A former graduate student in the lab, Dr. Jyothsna Chitturi identified loss-of-function mutations in eat-4 and avr-15 as suppressors of ubr-1 mutants’ bending defects (Chitturi, 2015). They encode EAT-4, a vesicular glutamate transporter, and AVR-15, a glutamate-activated chloride channel, respectively. Their genetic interactions with ubr-1 suggest that UBR-1 may also participate in the

54 regulation of glutamate signaling. Two other UBR-1-active sites, FLP and PVD, are glutamatergic sensory neurons. Therefore, aberrant glutamate signaling may also, in part, account for the ubr-1 reduced bending phenotype.

Figure 12. The functional loss of UBR-1 results in morphological changes of specific premotor INs at the end of larval stage development

(A) Confocal images of somata of AVA and AVE premotor INs in wildtype and ubr-1 mutants, which were labeled with cytosolic RFP (Pnmr-1-RFP), from L3 larva to adult stages. Top panels: AVA and AVE premotor IN somata showed round and smooth surface in both wildtype and ubr- 1 mutants at the L3 larva stage. Middle and bottom panels: AVA and AVE premotor IN somata showed round and smooth surface in wildtype animals, and rough and branched surface in ubr-1 mutants at the L4 larva (middle panels) and adult stages (bottom panels). (B) A confocal image of somata of AVA and AVE premotor INs in a ubr-1;got-1 adult animal. AVA and AVE premotor IN somata showed round and smooth surface. (C) Confocal images of the RIM soma in a wildtype and a ubr-1 adult animals. The RIM soma exhibited round morphology under both wildtype and ubr-1 background. Arrows denote roughness and short branches of the surface. Stars denote axons. Credit: Wesley Hung.

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Figure 13. UBR-1 may function through sensory neurons and premotor INs to affect the phase coupling among A-MNs so as to modulate motor outputs

(A) A model for the neural requirements of UBR-1 for modulating motor outputs. UBR-1 is critically required in mechanosensory neurons (FLP and PVD) and premotor INs (AVA, AVE, and AVD) to affect the activities of A-MNs, and thereby modulate motor outputs during backward locomotion. Neural connections are drawn based on White et al., 1986. (B) A model for the generation of backward locomotory patterns in wildtype and ubr-1 mutants. Top panel: in wildtype animals, A-MNs harbour intrinsic oscillatory activities. Sensory feedback establishes phase coupling between them, which enables the propagation of body undulations to drive backward locomotion. AVA premotor INs regulate the activities of A-MNs through gap junctions and chemical synapses to modulate backward locomotory patterns. This panel is modified from Gao et al., 2018. Bottom panel: in ubr-1 mutants, A-MNs harbour oscillatory activities as normal. Premotor INs (AVA, AVE, and AVD) are affected by glutamate neural toxicity, showing morphological changes. Through gap junctions and/or chemical synapses, premotor INs (AVA, AVE, and AVD) disrupt the phase coupling among A-MNs, leading to synchronized A-MN activation and the consequent reduced bending phenotype.

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Chapter 4 Conclusions and Future Perspectives 4.1 Conclusions

An animal’s nervous system consists of three major types of neurons, sensory neurons, interneurons and motor neurons, that form neural circuits to regulate behavioural outputs. Internal or external signals are perceived by sensory neurons, which are integrated by the downstream interneurons. Integrated signals are passed on to motor neurons, which innervate muscles and control their contraction to generate motor outputs. Crosstalk exists between and within different layers of neurons. Disruption of the function of any component in the circuit may affect normal motor outputs.

C. elegans locomotes by generating and propagating dorso-ventral bends along the body in either forward or backward direction. Both forward and backward locomotor programs are regulated by sensorimotor circuits. The reduced bending motor defect in ubr-1 mutants indicates a malfunctioning neural circuit. Previous studies revealed that ubr-1 loss-of-function mutations result in aberrant glutamate metabolism and signaling (Chitturi et al., 2018; Chitturi, 2015). However, which neural circuit and how this circuit is affected in ubr-1 mutants was only partially understood. This study aimed at addressing two main questions: (1) which neurons are involved in the neural circuit through which UBR-1 modulates the motor outputs; (2) how is the neural circuit affected in ubr-1 mutants to generate the reduced bending motor pattern.

Here I present results from these two aims. First, my results reveal that UBR-1 is critically required in sensory neurons (FLP and PVD) and premotor INs (AVA, AVE, and AVD) to establish bending. Second, extending from previous findings that ubr-1 mutations lead to synchronized A-MN activation, I show that the functional loss of UBR-1 disrupts the phase coupling of A-MN activation through toxicity in premotor INs (AVA, AVE, and AVD). With these results and the identification of glutamate signaling mutants (eat-4 and avr-15) as ubr-1 genetic suppressors, I propose that UBR-1 may regulate both glutamate signaling (in glutamatergic sensory neurons such as FLP and PVD) and glutamate metabolism (in non-glutamatergic neurons such as premotor INs AVA, AVE, and AVD). Aberrant glutamate signaling and metabolism both contribute to the reduced bending phenotype in ubr-1 mutants.

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Glutamate is a universal neurotransmitter and essential metabolite. Abnormal glutamate homeostasis is associated with several neurological diseases such as Parkinson’s disease and Schizophrenia (Kashani et al., 2007; Caudle and Zhang, 2009; Goff and Coyle, 2001). The spectrum of JBS, caused by loss of function mutations in human UBR1, includes neurological symptoms such as sensorineural deafness and cognitive impairment. Our discovery of the C. elegans UBR-1’s regulatory role in glutamate metabolism and signaling may hold implications for the pathophysiology of JBS.

A deeper understanding of the molecular and circuit mechanisms through which UBR-1 regulates the C. elegans motor outputs will facilitate the investigation of the relationship between the physiological role of UBR family proteins and JBS. Below I summarize and discuss the remaining questions and future perspectives.

4.2 Future Perspectives

4.2.1 To examine interactions between UBR-1 and ubr-1 suppressors

UBR-1 targets substrate proteins for the UPS-mediated degradation. The substrates of UBR-1 have some common features that can be used as criteria to tell whether a protein is a substrate of UBR- 1, including (1) the physical interaction with UBR-1; (2) the ubiquitination by UBR-1; (3) the protein level negatively related with that of UBR-1. The substrate of UBR-1 should meet at least these three criteria.

Previously, the total GOT-1 protein level and total GOT activity were measured and compared in wildtype and ubr-1 mutants. However, the possibility exists that UBR-1 may target GOT-1 only in some specific tissues, making the change in protein level in ubr-1 mutants undetectable when only the total level is measured. I would examine the interaction between UBR-1 and GOT-1 as well as other identified suppressors with more precise measurements.

I would examine their potential physical interactions by expressing tagged UBR-1 and GOT-1 constructs in HEK293T cells, and performing reciprocal co-immnoprecipitation. Proteasome inhibitors, MG132, would be applied to prevent substrates from fast degradation, allowing for the detection of the stabilized complex. If a physical interaction can be detected, I would examine the UBR-1-mediated ubiquitination. Similarly, constructs expressing tagged GOT-1, UBR-1, and ubiquitin K63R variant will be transfected into HEK293T cells, which would be treated with

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MG132 before harvesting. Tagged GOT-1 would then be isolated and purified. Its ubiquitination status would be checked by mass spectrometry. Following these, I would examine its protein level and expression pattern in ubr-1 mutants. I would use the CRISPR-Cas9 system to insert a GFP tag in frame at the endogenous C-terminal loci before the stop codon of got-1 in wildtype C. elegans. The engineered endogenous GOT-1::GFP fusion can visualize the expression pattern of GOT-1 and may show the relative protein level changes through the GFP intensity. If the GOT-1::GFP marker shows increased intensity in specific tissues in ubr-1 mutants, I would express RFP markers specifically in these tissues and isolate the cells with RFP signals by cell sorting. Then I would quantify the GOT-1 protein level change in these cells by western blot analysis. Similar processes can be employed for investigating relations with EAT-4, AVR-15 and other suppressors (see below).

4.2.2 To identify UBR-1’s substrates by genetic suppressor screens

It is crucial to identify substrates through which UBR-1 modulates the motor outputs. Although we have identified several genetic suppressors of ubr-1 mutants’ motor defects, they may not be direct substrates of UBR-1 (Chitturi et al., 2018; Chitturi, 2015).

To identify UBR-1’s substrate(s), I may increase the number of genomes screened. With an undergraduate student Kevin Zhang, we identified another ubr-1 suppressor, hp856. hp856 complements the suppression effects of, known suppressors, confirming that it represents a novel suppressor. Molecular cloning and functional characterization of hp856 may lead to new insights of the cellular function of UBR-1.

To clone hp856, I would first identify the single nucleotide polymorphisms (SNPs) in hp856 by whole genome sequencing (WGS). The causative SNP would be identified by restoring wildtype genomic fragments of the respective locus in ubr-1; hp856 mutants and examining whether they can revert the bending pattern to that of ubr-1.

To characterize this suppressor, I would examine its expression pattern and site of action as described for UBR-1. If this gene is expressed in and functions through a same set of neurons as UBR-1, I would proceed to determine whether it is a potential UBR-1 substrate using several criteria: it should exhibit elevated protein levels in ubr-1 mutants, interact physically with UBR- 1, and undergo UBR-1-dependent ubiquitination. I would further investigate its potential involvement in glutamate metabolism or signaling. For example, whether its functional loss changes glutamate levels and the expression of GOT-1, EAT-4, or AVR-15.

4.2.3 To identify UBR-1’s substrates by BioID

The isolation of hp856 implicates the presence of additional ubr-1 suppressors. By scaling up the genetic suppressor screens, more genetic suppressors of ubr-1 that functionally interact with UBR- 1 can be identified. However, genetic suppressor screens do not distinguish substrates or components of substrate-mediated functional pathway.

Substrates of UBR-1 should exhibit physical interaction with UBR-1. It is however challenging to identify substrates of an E3 ligase by conventional biochemical methods, due to binding-induced fast ubiquitination and degradation of the substrates. Proximity-based biotinylation (BioID) allows

59 60 efficient characterization of protein-protein interactions in living cells (Roux et al., 2012; Kim et al., 2014a). This method can be adapted in C. elegans (Waaijers et al., 2016; Branon et al., 2017). By expressing UBR-1 fused with a biotin-conjugating enzyme BirA in C. elegans, proteins that reside close to UBR-1 become biotinylated. This facilitates their purification by streptavidin, generating UBR-1’s “interactome”.

BioID may recover many proteins in close proximity to UBR-1 in addition to its substrates. To improve the chance of identifying substrates among them, I would perform Bio ID in the presence of proteasome inhibitors. I would further examine their overlap with genetic suppressors, or whether they have been implicated in biological pathways, such as glutamate metabolism and signaling known to interact with ubr-1, to identify the most promising candidates.

4.2.4 To dissect UBR-1-dependent regulation of the A-MN phase coupling

In wildtype animals, A-MNs harbour intrinsic oscillatory activities. They fire sequentially to control the alternating muscle contractions for propagating dorso-ventral body bends in backward locomotion. Sparse removal of A-MNs alters backward locomotion. Specifically, ablating a small number of mid-body A-MNs resulted in uncoupled anterior and posterior body bends, indicating that the phase coupling of A-MN activation is dependent on the connections between adjacent or neighboring A-MNs (Gao et al., 2018). The anterior-posterior proprioceptive coupling of B-MNs facilitates the propagation of rhythmic alternating body bends to drive forward locomotion (Wen et al., 2012; Xu et al., 2018). Although the proprioception in A-MNs has not been reported, several lines of indirect evidence support its existence and necessity for the phase coupling of A-MNs (Wen, Gao and Zhen, 2018). The removal of mid-body A-MNs may block the posterior-anterior transduction of proprioceptive signals in A-MNs and thereby lead to the uncoupling between the anterior and posterior body bends.

My study reveals that ubr-1 mutations disrupt the phase coupling of A-MN activation through neural toxicity in premotor INs. The AVA premotor INs modulate the autonomous oscillations of A-MNs to generate flexible motor outputs (Gao et al., 2018). However, how they eliminate the phase coupling among A-MNs when affected by aberrant premotor INs remains poorly understood. One hypothesis is that the premotor INs aberrantly activate A-MNs in ubr-1 mutants, the effect of which may overwhelm the proprioception in A-MNs that A-MNs fail to establish appropriate phase lags.

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To test this hypothesis, I will first examine the activity patterns of premotor INs, AVA, AVE, and AVD, to identify potential changes in ubr-1 mutants. I will further examine if I could recapitulate ubr-1’s bending pattern by optogenetically activating premotor INs, or disrupting A-MN proprioception by sparse ablation of A-MNs in wildtype animals. If so, I will directly assess the effect of activity coordination between A-MNs that are anterior and posterior to the region of ablation.

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