Role of Guanine Nucleotide Exchange Factors for Rab10 in Insulin-Regulated GLUT4 Trafficking

Role of guanine nucleotide exchange factors for Rab10 in insulin-regulated GLUT4 trafficking

Beverley Anne Murrow

A thesis in fulfilment of the requirements for the degree of Doctor of Philosophy

St. Vincent’s Clinical School, Faculty of Medicine UNSW Australia

January 2017

THE UNIVERSITY OF NEW SOUTH WALES Thesis/Dissertation Sheet Surname or Family name: Murrow First name: Beverley Other name/s: Anne Abbreviation for degree as given in the University calendar: PhD School: St Vincent’s Clinical School Faculty: Medicine Title: Role of guanine nucleotide exchange factors for Rab10 in insulin-regulated GLUT4 trafficking

Abstract 350 words maximum:

The insulin regulation of glucose uptake via translocation of the glucose transporter, GLUT4, is essential for maintenance of whole-body glucose homeostasis. GLUT4 traffic is a strikingly sophisticated biochemical process. The transporter must navigate its way throughout diverse endomembranous systems and multiple nodes along this path are exquisitely regulated by the concerted actions of Rab GTPases and their effectors. Understanding precisely how insulin orchestrates regulation of Rab activity appropriately at the various transport steps in the GLUT4 itinerary, however, has been a longstanding puzzle and the identities of key regulatory players have remained elusive. In pursuit of novel regulatory proteins functioning in GLUT4 traffic, I examined datasets from global quantitative mass spectrometric analyses of insulin- regulated protein phosphorylation in 3T3-L1 adipocytes, L6 skeletal muscle myotubes and murine liver tissue. This search uncovered DENND4A and DENND4C, two closely related members of the novel differentially expressed in normal versus neoplastic (DENN) domain- containing protein family of Rab guanine nucleotide exchange factors (GEFs), as highly insulin- sensitive phosphoproteins. Both DENND4A and DENND4C target Rab10, a Rab thought to regulate a single distal step in GLUT4 traffic. Using affinity pull-down and protein overlay methods, I demonstrated that DENND4A is an insulin-regulated 14-3-3β binding protein downstream of mTORC1 kinase. Point mutation of 35 serine/threonine phosphorylation sites in the DENND4A C-terminus eliminated its 14-3-3 binding capacity and enhanced the GEF activity of DENND4A towards Rab10 in vivo, as assessed by Rab effector pull-down. Live-cell microscopic analysis revealed that DENND4A inhibits insulin-stimulated GLUT4 translocation when overexpressed in adipocytes, whereas the phospho-dead '35P' mutant, in which 14-3-3 binding is lost, has no effect. DENND4C, previously implicated as a positive regulator of GLUT4 exocytosis, was found to localise to a distinct subpopulation of GLUT4 storage vesicles (GSVs) immunoisolated from 3T3-L1 adipocytes that exclude the well-characterised Rab10 GTPase activating protein (GAP), AS160. My findings indicate that there is at least one additional insulin-regulated step than previously thought in GLUT4 traffic involving DENND4A phosphorylation. I propose a model where multiple Rab10 GEFs have opposing actions in GLUT4 traffic and the simultaneous inhibition of DENND4A and AS160 by insulin-mediated phosphorylation confers net GLUT4 transfer to the adipocyte plasma membrane. Collectively, my work illuminates the potential of DENND4 proteins as novel therapeutic targets for insulin resistance and Type II diabetes mellitus.

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Beverley A. Murrow

January-2017

‘I hereby grant the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all proprietary rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation. I also authorise University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstract International (this is applicable to doctoral theses only). I have either used no substantial portions of copyright material in my thesis or I have obtained permission to use copyright material; where permission has not been granted I have applied/will apply for a partial restriction of the digital copy of my thesis or dissertation.'

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Table of Contents

Table of Contents ...... i Abstract ...... v Acknowledgements ...... vi List of Figures ...... viii List of Tables ...... ix List of Abbreviations ...... x

Chapter 1: General Introduction 1 1.1 Insulin action ...... 1 1.2 GLUT4 ...... 1 1.3 The insulin signalling pathway ...... 2 1.4 The Akt/mTORC1 signalling hub ...... 4 1.5 Vesicle transport ...... 5 1.6 The GLUT4 lifecycle ...... 9 1.7 Rab GTPases in membrane traffic ...... 11 1.8 The Rab cycle ...... 12 1.9 Rab cascades ...... 15 1.10 Rab GTPases in GLUT4 traffic ...... 16 1.11 TBC domain proteins: GAPs for Rab GTPases ...... 18 1.12 The Rab-GAP, AS160 ...... 19 1.13 Rab-GEFs ...... 21 1.14 DENN domain proteins: a novel Rab-GEF family ...... 23 1.14.1 DENND1A-C (connecdenn1-3) ...... 24 1.14.2 Folliculin ...... 26 1.14.3 DENND2A-D...... 27 1.14.4 DENND3...... 27 1.14.5 DENND5A/B ...... 28 1.14.6 DENND6A/B ...... 28 1.14.7 MTMR5/13 ...... 28 1.14.8 DENND4A-C ...... 29 1.15 The present study ...... 31

Chapter 2: General Materials and Methods 33 2.1 Materials and antibodies ...... 33 2.2 Plasmids ...... 34 2.3 Methods ...... 35 2.3.1 Molecular cloning...... 35 2.3.2 Cell culture ...... 36 2.3.3 SDS-polyacrylamide gel electrophoresis (PAGE) and Western blot analysis ...... 36 2.3.4 Purification of recombinant GST fusion proteins...... 38 2.3.5 Colloidal Coomassie brilliant blue staining ...... 39 2.3.6 Immunoprecipitation of FLAG-tagged proteins ...... 40

Chapter 3: Insulin-regulated phosphorylation of DENN domain proteins

41 3.1 Introduction ...... 41 3.2 Materials and methods ...... 44 3.2.1 Phosphoproteomic analysis of insulin-stimulated 3T3-L1 adipocytes (Humphrey et al., 2013) ...... 44 3.2.2 Phosphoproteomic analysis of insulin signalling in murine liver (Humphrey et al., 2015a) ...... 45 3.2.3 Phosphoproteomic analysis of insulin-stimulated L6 myotubes (Hoffman et al., unpublished) ...... 47 3.2.4 In silico analysis of phosphoproteomic datasets ...... 48 3.2.5 DENN domain protein expression in murine tissues ...... 48 3.2.6 Phosphosite mapping of DENN-domain proteins ...... 48 3.3 Results ...... 52 3.3.1 Rab, Rab-GEF and Rab-GAP protein abundance in insulin target cell types and tissue...... 52 3.3.2 Insulin-regulated DENN protein phosphorylation in 3T3-L1 adipocytes ...... 60 3.3.3 Insulin-regulated DENN protein phosphorylation in murine liver tissue ...... 68 3.3.4 Insulin-regulated DENN protein phosphorylation in L6 myotubes ...... 71 3.3.5 DENND4A and DENND4C expression in murine tissues ...... 73 3.3.6 Mapping insulin-sensitive DENND4A and DENND4C phosphosites in HEK-293E cells ...... 74 3.4 Discussion ...... 80 ii

Chapter 4: Probing the insulin-regulated binding of 14-3-3 to DENND4A

84 4.1 Introduction ...... 84 4.2 Methods ...... 88 4.2.1 Molecular cloning...... 88 4.2.2 14-3-3 interactions studies ...... 89 4.3 Results ...... 91 4.3.1 14-3-3 binding to DENND4A is enhanced by insulin ...... 91 4.3.2 14-3-3 binding to DENND4A occurs slowly following insulin stimulation ...... 94 4.3.3 The DENND4A−14-3-3 interaction is sensitive to inhibitors of mTORC1 ...... 97 4.3.4 14-3-3 binding to a 6-site DENND4A phosphomutant is comparable to the wild- type protein ...... 99 4.3.5 Insulin-enhanced 14-3-3 binding to DENND4A is abolished in a 35-site phosphomutant ...... 103 4.4 Discussion ...... 108

Chapter 5: Exploring the function of DENND4A Rab10-GEF activity in

insulin-regulated cellular processes 111 5.1 Introduction ...... 111 5.1.1 Autophagy ...... 112 5.1.2 mTORC1 regulation of autophagy ...... 112 5.1.3 Autophagic roles of Rab GTPases and DENN domain proteins...... 113 5.1.4 Rab7 regulation of lipophagy ...... 117 5.1.5 Chapter aims ...... 118 5.2 Methods ...... 118 5.2.1 In vivo GEF assay ...... 119 5.2.2 Live-cell GLUT4 translocation assay ...... 120 5.2.3 Lipid droplet studies ...... 122 5.3 Results ...... 124 5.3.1 DENND4A 35-site phosphomutant exhibits enhanced Rab10-directed GEF activity ...... 124 5.3.2 Wild-type DENND4A inhibits insulin-stimulated GLUT4 translocation when overexpressed in 3T3-L1 adipocytes ...... 127

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5.3.3 Rab10 −/− mouse embryonic fibroblasts exhibit hallmarks of compromised autophagy under nutrient stress ...... 132 5.4 Discussion ...... 135

Chapter 6: Investigating the nature of DENND4C association with GLUT4

vesicles 139 6.1 Introduction ...... 139 6.2 Methods ...... 140 6.2.1 Subcellular distribution and protein interactions of DENND4C ...... 140 6.2.2 Molecular cloning...... 144 6.2.3 DENND4C truncation mutant localisation ...... 144 6.3 Results ...... 145 6.3.1 DENND4C is enriched in the low density microsome fraction of 3T3-L1 adipocytes ...... 145 6.3.2 DENND4C associates with GLUT4 vesicles in the non-stimulated state and dissociates following insulin stimulation in 3T3-L1 adipocytes ...... 147 6.3.3 AS160 and DENND4C occupy distinct GLUT4 vesicle populations ...... 147 6.3.4 DENND4C does not interact with a panel of established GLUT4 vesicle integral membrane proteins ...... 150 6.3.5 Localisation of DENND4C truncation mutants in HeLa cells ...... 151 6.4 Discussion ...... 153

Chapter 7: General Discussion 157

Chapter 8: References 166

Appendix: Supplementary Material ...... 210 Supplementary Figures ...... 210 Supplementary Tables ...... 211

Abstract

The insulin regulation of glucose uptake via translocation of the glucose transporter, GLUT4, is essential for maintenance of whole-body glucose homeostasis. GLUT4 traffic is a strikingly sophisticated biochemical process. The transporter must navigate its way throughout diverse endomembranous systems and multiple nodes along this path are exquisitely regulated by the concerted actions of Rab GTPases and their effectors. Understanding precisely how insulin orchestrates regulation of Rab activity appropriately at the various transport steps in the GLUT4 itinerary, however, has been a longstanding puzzle and the identities of key regulatory players have remained elusive. In pursuit of novel regulatory proteins functioning in GLUT4 traffic, I examined datasets from global quantitative mass spectrometric analyses of insulin-regulated protein phosphorylation in 3T3-L1 adipocytes, L6 skeletal muscle myotubes and murine liver tissue. This search uncovered DENND4A and DENND4C, two closely related members of the novel differentially expressed in normal versus neoplastic (DENN) domain-containing protein family of Rab guanine nucleotide exchange factors (GEFs), as highly insulin-sensitive phosphoproteins. Both DENND4A and DENND4C target Rab10, a Rab thought to regulate a single distal step in GLUT4 traffic. Using affinity pull- down and protein overlay methods, I demonstrated that DENND4A is an insulin-regulated 14-3-3β binding protein downstream of mTORC1 kinase. Point mutation of 35 serine/threonine phosphorylation sites in the DENND4A C-terminus eliminated its 14-3-3 binding capacity and enhanced the GEF activity of DENND4A towards Rab10 in vivo, as assessed by Rab effector pull-down. Live-cell microscopic analysis revealed that DENND4A inhibits insulin-stimulated GLUT4 translocation when overexpressed in adipocytes, whereas the phospho-dead '35P' mutant, in which 14-3-3 binding is lost, has no effect. DENND4C, previously implicated as a positive regulator of GLUT4 exocytosis, was found to localise to a distinct subpopulation of GLUT4 storage vesicles (GSVs) immunoisolated from 3T3-L1 adipocytes that exclude the well-characterised Rab10 GTPase activating protein (GAP), AS160. My findings indicate that there is at least one additional insulin-regulated step than previously thought in GLUT4 traffic involving DENND4A phosphorylation. I propose a model where multiple Rab10 GEFs have opposing actions in GLUT4 traffic and the simultaneous inhibition of DENND4A and AS160 by insulin-mediated phosphorylation confers net GLUT4 transfer to the adipocyte plasma membrane. Collectively, my work illuminates the potential of DENND4 proteins as novel therapeutic targets for insulin resistance and Type II diabetes mellitus.

Acknowledgements

My sincerest thanks go to my PhD supervisor David James for allowing a Welsh country girl to pursue her science dreams in the big city Down Under. Thank you for your wisdom and wit, and for creating an invigorating research environment.

I would like to extend my thanks to my mentor Jacqueline Stöckli for her practical advice and extensive knowledge on Rab GTPases.

I thank those who contributed technical expertise: Himani Pant, Anagha Killedar, Kristen Thomas, Annabel Minard and James 'Burchy' Burchfield. Thank you also to Dr. Georg Ramm for conducting the TEM work.

Thank you to all members of the James laboratory past and present for your camaraderie.

Special mention has to be given to Sophie and Annabel (a.k.a. The Force Field). We shared our reagents and our secrets. I will cherish our memories forever.

Sean, my bench buddy (and now my UC buddy). Thank you for always believing in me.

Dougall, bru'h. Thank you for the music and conversation.

Thank you Mum and close friends for enduring my highs and lows.

Lastly, I am indebted to my medicine man, Dr. Phil van Zanden, for his care throughout my illness.

The James Laboratory (2012-2016)

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List of Figures

Figure 1.1. The P13K/Akt pathway ...... 3 Figure 1.2. Vesicular transport ...... 6 Figure 1.3. The GLUT4 lifecycle ...... 10 Figure 1.4. The Rab cycle ...... 14 Figure 1.5. Rab cascade model ...... 16 Figure 1.6. Human DENN domain-containing proteins ...... 25 Figure 3.1. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the 3T3-L1 adipocyte proteome ...... 54 Figure 3.2. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the L6 myotube proteome ...... 55 Figure 3.3. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the murine liver proteome ...... 56 Figure 3.4. Insulin-regulated phosphorylation of DENN domain-containing and DENN-related proteins in 3T3-L1 adipocytes ...... 62 Figure 3.5. Effect of Akt inhibition on insulin-sensitive DENND4A and DENND4C phosphorylation sites ...... 64 Figure 3.6. Insulin-regulated phosphorylation of DENN domain-containing and DENN-related proteins in murine liver tissue ...... 71 Figure 3.7. Insulin-stimulated phosphorylation of DENN domain-containing and DENN-related proteins in L6 myotubes ...... 72 Figure 3.8. DENND4A and DENND4C expression in murine tissues ...... 74 Figure 3.9. Insulin-sensitive phosphorylation sites in DENND4A ...... 76 Figure 3.10. Insulin-sensitive phosphorylation sites in DENND4C ...... 77 Figure 3.11. DENND4A and DENND4C phosphorylation sites ...... 78 Figure 3.12. Insulin-regulated DENND4A and DENND4C phosphosite dynamics ...... 79 Figure 4.1. 14-3-3 binding to DENND4A and DENND4C ...... 93 Figure 4.2. Time course of AS160−14-3-3 and DENND4A−14-3-3 interactions ...... 96 Figure 4.3. Kinase inhibitor sensitivity of AS160−14-3-3 and DENND4A−14-3-3 interactions ... 97 Figure 4.4. Direct binding of 14-3-3 to AS160 and DENND4A phosphomutants ...... 100 Figure 4.5. 14-3-3 binding to multi-site DENND4A phosphomutants ...... 102 Figure 4.6. Schematic diagram of 35-site DENND4A phosphomutant (4A-35P) ...... 103 Figure 4.7. 14-3-3 binding to DENND4A 35-site phosphomutant ...... 106

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Figure 4.8. Direct binding of 14-3-3 to DENND4A 35-site phosphomutant ...... 107 Figure 5.1. In vivo GEF assay ...... 126 Figure 5.2. Live-cell GLUT4 translocation assay in 3T3-L1 adipocytes...... 129 Figure 5.3. DENND4A localisation in 3T3-L1 adipocytes...... 131 Figure 5.4. Transmission electron microscopy (TEM) of wild-type and Rab10 knockout MEFs under fed and starved conditions ...... 133 Figure 6.1. Subcellular distribution of DENND4C and GLUT4 in 3T3-L1 adipocytes ...... 146 Figure 6.2. Immunoisolation of GLUT4-, DENND4C- and AS160-containing vesicles from 3T3-L1 adipocytes ...... 150 Figure 6.3. Binding of AS160 and DENND4C to GSV resident proteins ...... 151 Figure 6.4. DENND4C truncation mutant species ...... 152 Figure 6.5. Localisation of DENND4C truncation mutants in HeLa cells ...... 153 Figure 7.1. Model for the role of DENND4A and DENND4C in GLUT4 trafficking ...... 160

List of Tables

Table 3.1. Rab substrates of mammalian Rab-GEF and DENN domain proteins ...... 58 Table 3.2. Rab substrates of mammalian Rab-GAP protein ...... 59 Table 3.3. Orthologous DENND4A phosphosites and protein kinase prediction ...... 66 Table 3.4. Orthologous DENND4C phosphosites and protein kinase prediction ...... 67 Table 4.1. Candidate 14-3-3-binding sites spanning the DENND4A C-terminus ...... 104

List of Abbreviations

~ approximately 2-DG [3H] 2-deoxy-D-glucose ADAM a disintegrin and metalloproteinase ADP adenosine diphosphate AICAR 5-aminoimidazole-4-carboxamide ribonucleotide AMP adenosine monophosphate AMPK AMP-activated protein kinase Arf ADP-ribosylation factor AS160 Akt substrate of 160 kDa AU arbitrary units BAD Bcl-2-associated antagonist of cell death BCA bicinchoninic acid BLOC biogenesis of lysosome-related organelles complex bp base pair(s) BSA bovine serum albumin C carbon

C- carboxy CaM calmodulin CAMKII Ca2+/calmodulin-dependent protein kinase II Cdk cyclin-dependent kinase CHAPS 3-((3-cholamidopropyl)dimethylammonio)-1-propanesulfonate CK casein kinase CNBr cyanogen bromide DENN differentially expressed in normal versus neoplastic DMEM Dulbecco’s Modified Eagle Medium DMSO dimethyl sulfoxide DNA deoxyribonucleic acid DTT dithiothreitol ECL enhanced chemilumescent EDC3 enhancer of mRNA-decapping protein 3 EDTA ethylenediaminetetraacetic acid EE early endosome(s)

eGFP enhanced green fluorescent protein EH Eps15 homology EHBP1 EH domain-binding protein 1 EHD2 EH domain protein 2 ES embryonic stem ESCRT endosomal sorting complexes required for transport EtBr ethidium bromide EVI5 ecotropic viral integration site 5 FBS fetal bovine serum FC fold change FDR false discovery rate Fig. figure FLCN folliculin FNIP folliculin-interacting protein FOXO forkhead box O GAP GTPase-activating protein GDI guanine nucleotide dissociation inhibitor GDP guanosine diphosphate GEF guanine nucleotide exchange factor GFP green fluorescent protein GLUT4 glucose transporter 4 gp glycoprotein GSK glycogen synthase kinase GST glutathione S-transferase GTP guanosine triphosphate GTPase guanosine triphosphatase GSV GLUT4 storage vesicle GWAS genome-wide association study Gyp GAP for Ypt H hydrogen HDM high density microsomes HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HOPS homotypic fusion and vacuole protein sorting HPLC high-performance liquid chromatography

HPS Hermansky–Pudlak syndrome h hour(s) HRP horseradish peroxidase HTP high-throughput iBAQ intensity based absolute quantification IBMX isobutyl methylxanthine IgG immunoglobulin G IP immunoprecipitation IPTG isopropyl thiogalactopyranoside IQR interquartile range IR insulin receptor IRAP insulin-regulated aminopeptidase IRSE interferon-stimulated response element kb kilobase(s) KIF5B kinesin family member 5B KRP Krebs-Ringer phosphate LAMP1 lysosomal-associated membrane protein 1 LB Luria broth LC liquid chromatography LC3 microtubule-associated protein 1 light chain 3 alpha LD lipid droplet LD longin domain LDM low density microsomes LE late endosome(s) LKB1 liver kinase B1 LRP1 low density lipoprotein receptor-related protein 1 LSB Laemmli sample buffer MABP MVB12-associated β-prism MADD MAP kinase activating death domain MAPK mitogen-activated protein kinase MBR match between runs MEF mouse embryonic fibroblast MEM Minimum Essential Medium MICAL-L2 molecule interacting with CasL-like 2

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min minute(s) mito mitochondria MS mass spectrometry MS/MS tandem mass spectrometry mTOR mammalian target of rapamycin MTMR myotubularin-related MVB multivesicular body MycPBP c-myc promoter-binding protein N nitrogen N- amino NA numerical aperture NESCA new molecule containing SH3 at the C-terminal NDR nuclear Dbf2-related nm nanometre NSF N-ethylmaleimide-sensitive fusion factor nuc nuclei PAK1 p21 protein (Cdc42/Rac)-activated kinase 1 PAS phospho Akt substrate PAS phagophore assembly site PBS phosphate-buffered saline PCR polymerase chain reaction PFA paraformaldehyde PI3K phosphatidylinositol 3-kinase PIP2 phosphatidylinositol-4,5-bisphosphate PIP3 phosphatidylinositol-3,4,5-triphosphate PKA protein kinase A PKD protein kinase D PM plasma membrane PPR pentatricopeptide repeat PRAS40 proline-rich Akt substrate of 40 kDa PTB phosphotyrosine-binding PTM post-translational modification PVDF polyvinylidene fluoride px pixel

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RE recycling endosome(s) RIN1 Ras and Rab interactor 1 RME receptor-mediated endocytosis RNA ribonucleic acid RPIP8 Rap2-interacting protein 8 RT room temperature RTK receptor tyrosine kinase RUN RPIP8, UNC-14 and NESCA RUTBC3 RUN and TBC1 domain-containing 3 SAX strong anion exchange SCX strong cation exchange SDM site-directed mutagenesis SDS sodium dodecyl sulfate SDS-PAGE SDS-polyacrylamide gel electrophoresis SE sorting endosome(s) SEM standard error of the mean sec second(s) SILAC stable isotope labelling by amino acids in cell culture siRNA small interfering RNA SNAP soluble NSF attachment protein SNARE SNAP receptor SNX sortin nexin STAGE stop and go extraction sup. Supernatant T2D Type II diabetes mellitus TBC Tre-2/Cdc16/ Bub2 TBS Tris-buffered saline TBS-T Tris-buffered saline, 0.1% Tween-20 TCEP Tris (2-carboxyethyl) phosphine TEM transmission electron microscopy TFA trifluoroacetic acid TFE trifluoroethanol TGN trans-Golgi network TIRF total internal reflection fluorescence

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TRAPP transport protein particle Trs TRAPP subunit TUSC5 tumour suppressor candidate 5 U units UNC-14 uncoordinated family member 14 USP6 ubiquitin specific peptidase 6 USP6NL USP6 N-terminal like UV ultraviolet V volts VARP VPS9 ankyrin repeat protein vp165 vesicle protein of 165 kDa Vps9 vacuolar protein sorting 9 WB Western blot WT wild-type Ypt yeast protein transport

Chapter 1: General Introduction

1.1 Insulin action

Blood glucose homeostasis is essential for normal mammalian function. Following ingestion of a carbohydrate-rich meal, digestive enzymes in the small intestine liberate glucose and other monosaccharides, which are promptly absorbed across the intestinal epithelium into the bloodstream. The consequential elevation in plasma glucose levels rapidly stimulates pancreatic β-cells to secrete insulin (Flatt, 1995), an anabolic peptide hormone which acts to maintain euglycaemia. Insulin promotes glucose uptake and assimilation in adipose (fat) and skeletal muscle tissues whilst simultaneously suppressing hepatic glucose efflux (Lomberk and Urrutia, 2009), thus restoring blood glucose concentration to within its normal, relatively narrow physiological range of 5-6 mM (Lanner et al., 2008). The combined effects of ageing, obesity, genetic factors and a sedentary lifestyle may diminish the insulin sensitivity of liver, muscle and fat tissue, impairing glucose clearance from the bloodstream after feeding. This state of insulin resistance predisposes to Type II diabetes mellitus (T2D), a metabolic disorder characterised by β-cell dysfunction, hyperinsulinemia and chronic glucose intolerance (Taylor, 1999).

1.2 GLUT4

Glucose disposal into peripheral tissues is the rate-determining step for the utilisation of glucose as an energy source or for storage (Leto and Saltiel, 2012). Therefore, an efficient mechanism for the import of glucose into fat and muscle cells is vital for maintenance of whole-body glucose homeostasis. In the human genome, the solute carrier family 2 (SLCA2) gene family encodes 14 integral membrane proteins that belong to a superfamily of facilitative (energy-independent) sugar transporters termed GLUTs (Thorens and Mueckler, 2010). GLUT proteins are predicted to possess 12 transmembrane domains and are grouped into three classes based on their sequence similarities (Mueckler and Thorens,

2013). One or more GLUTs are expressed in virtually every human cell type. In adipocytes (white and brown), cardiomyocytes and skeletal muscle cells, insulin- stimulated glucose uptake is mediated by the Class I facilitative glucose transporter, glucose transporter 4 (GLUT4/SLCA24) (James et al., 1988), whose expression is predominantly restricted to these insulin-sensitive tissues (James et al., 1989).

Under basal (low insulin) conditions, GLUT4 is primarily sequestered in intracellular, perinuclear membrane compartments with the majority residing in small, 50-70 nm vesicles (referred to as GLUT4 storage vesicles, GSVs) (Martin et al. 2000a). As circulating insulin levels rise following ingestion of a carbohydrate-rich meal, GSVs translocate to and fuse with the plasma membrane (PM), allowing for glucose uptake along its concentration gradient and preventing chronic elevations in blood glucose levels (Slot et al., 1991a; Bryant et al., 2002). In skeletal muscle, contraction can stimulate GLUT4 redistribution to the PM independently of insulin (Lund et al., 1995; Ploug et al., 1998). GLUT4 heterozygous knockout mice develop severe peripheral, but not hepatic, insulin resistance and diabetes (Rossetti et al., 1997; Stenbit et al., 1997), whereas re- expression of GLUT4 in the skeletal muscle of these animals prevents the onset of hyperinsulinemia and restores glucose tolerance (Tsao et al., 1999). Adipose-specific GLUT4 knockout mice develop insulin resistance in adipose, muscle and liver (Abel et al., 2001). It is therefore evident that GLUT4-mediated glucose uptake into muscle and adipose tissue is essential for the maintenance of whole-body glucose homeostasis. Since one of the earliest defects contributing to insulin resistance in humans occurs at the level of GLUT4 translocation (Garvey et al., 1988; Maianu et al., 2001; Lizunov et al., 2013; Tan et al., 2015), identifying the major regulatory nodes in this process could yield novel translational therapies for metabolic disease.

1.3 The insulin signalling pathway

Insulin signals to GLUT4 via the canonical phosphatidylinositol 3-kinase (PI3K)/Akt pathway, a complex cascade of intracellular protein phosphorylation events (reviewed in Rowland et al., 2011) [Fig. 1.1]. An overview of insulin signalling is given here.

Figure 1.1. The P13K/Akt pathway

Schematic diagram of insulin signal transduction via the PI3K/Akt pathway. Activation of the transmembrane insulin receptor tyrosine kinase by insulin (blue diamond) leads to receptor auto-phosphorylation, creating docking sites for IRS proteins, which are themselves tyrosine phosphorylated. Phosphorylated IRS proteins recruit the regulatory PI3K subunit, p85, leading to activation of the PI3K catalytic subunit, p110. Active PI3K converts PIP2 to PIP3 at the plasma membrane, and PIP3 then acts as a platform for membrane recruitment of Akt and PDK1. Dual phosphorylation of Akt on residues Thr308 and Ser473 by PDK1 and the mTORC2 complex, respectively, leads to full Akt activity, allowing for the Ser/Thr phosphorylation of various downstream substrates with diverse cellular roles, for example involvement in GLUT4 exocytosis. A positive feedback loop exists whereby Akt-mediated phosphorylation of SIN1 enhances mTORC2 activity. Abbreviations: IR, insulin receptor; IRS, insulin receptor substrate; mTOR, mammalian target of rapamycin; PDK1, phosphoinositide dependent kinase-1; PI3K, phosphatidylinositol 3-kinase; PIP2, phosphatidylinositol-4,5-bisphosphate; PIP3, phosphatidylinositol-3,4,5-triphosphate; PM, plasma membrane; S, serine; SIN1, stress-activated map kinase-interacting protein 1; T, threonine.

At the cell surface, insulin binds to the extracellular α-subunit of the transmembrane heterotetrameric insulin receptor (IR), stimulating autophosphorylation of the intracellular β-subunits and activation of the IR intrinsic tyrosine kinase activity (Gammeltoft and Van Obbergehn, 1986). Insulin receptor substrate (IRS) proteins dock on to the IR via their phosphotyrosine-binding (PTB) domains and are themselves tyrosine phosphorylated (Myers and White, 1996). Phosphorylated IRS proteins then recruit Class IA PI3K via its regulatory subunit, p85, which leads to activation of the PI3K catalytic subunit, p110 (Backer et al., 1992; Myers et al., 1992). Active PI3K catalyses the conversion of phosphatidylinositol-4,5-bisphosphate (PIP2) to phosphatidylinositol-3,4,5-triphosphate (PIP3) on the cytosolic leaflet of the PM. PIP3 acts as a platform for the membrane recruitment of the pleckstrin homology (PH) domain- containing serine/threonine kinases, Akt and phosphoinositide dependent kinase-1 3

(PDK1), members of the AGC kinase family (Andjelković et al., 1997; Lietzke et al., 2000). Dual phosphorylation of Akt on residues Thr308 and Ser473 by PDK1 and the mammalian target of rapamycin (mTOR) C2 complex, respectively, leads to full Akt activity, allowing for the Ser/Thr phosphorylation of various downstream substrates with diverse cellular roles (Alessi et al., 1996; Alessi et al., 1997; Sarbassov et al., 2005). mTORC2 is generally considered to lie upstream of Akt. However, it was recently demonstrated that Akt and mTORC2 are involved in a positive feedback loop, whereby Akt-mediated phosphorylation of stress-activated map kinase-interacting protein 1 (SIN1), an indispensable component of the mTORC2 complex, enhances mTORC2 activity (Humphrey et al., 2012; Yang et al., 2015).

1.4 The Akt/mTORC1 signalling hub

The Akt kinase family is comprised of three homologous isoforms: Akt1, Akt2 and Akt3 (Dummler and Hemmings, 2007). Akt2, the predominant isoform expressed in insulin- responsive tissues, is responsible for insulin-regulated glucose uptake (Bae et al., 2003; Jiang et al., 2003; Bouzakri et al., 2006). Indeed, activation of Akt2 alone is sufficient to recapitulate insulin-stimulated GLUT4 translocation in 3T3-L1 adipocytes (Kohn et al., 1998; Ng et al., 2008). However, despite a thorough understanding of the upstream events in the insulin signal transduction cascade, the precise pathway linking Akt2 activation to the distal GLUT4 trafficking machinery is uncertain. Akt2 is a central node connecting insulin signalling to several other downstream cellular processes, including glucose metabolism, protein synthesis, gene transcription and cell survival (Manning and Cantley, 2007). One of the major outcomes of Akt signalling is the indirect activation of the mTORC1 kinase, a master regulator of cell growth, metabolism and autophagy (Laplante and Sabatini, 2009). Autophagy is a catabolic process that recycles damaged proteins and organelles to maintain cellular homeostasis, and provides a supply of substrates for energy production when nutrients are scarce (Yang and Klionsky, 2010). mTORC1 substrates include the p70 ribosomal protein S6 kinase (S6K) and UNC-51 like autophagy activating kinase 1 (ULK1). The former regulates mRNA translation initiation and the latter functions in the formation of autophagosomes, membrane-bound structures that enclose cellular constituents destined for lysosomal degradation (Thomas and Hall, 1997; 4

Mizushima et al., 2011). When activated by mTORC1-mediated phosphorylation, S6K, in turn, phosphorylates ribosomal protein S6 (rpS6) to initiate protein synthesis (Ma and Blenis, 2009). Conversely, phosphorylation of ULK1 by mTORC1 inhibits its kinase activity, suppressing autophagy (Jung et al., 2009). In addition to growth factor signalling, mTORC1 activity can also be modulated by nutrient availability. This mechanism is regulated by the Rag family of small guanosine triphosphatases (GTPases), which exist as heterodimers of either RagA/B with either RagC/D (Sancak et al., 2008). Amino acids stimulate Rag GTPase activity, leading to recruitment of mTORC1 to the lysosomal membrane where it encounters the small GTPase, Rheb, which, in turn, activates mTORC1 kinase activity (Sancak et al., 2008; Sancak et al., 2010).

1.5 Vesicle transport

Eukaryotic cells are subdivided into functionally-distinct membranous compartments with unique protein signatures, yet protein synthesis occurs only in the cytosol, across the endoplasmic reticulum (ER) membrane, and in mitochondria. The sorting and directed transport of lumenal and membrane cargo proteins to target membranes occurs by means of vesicular transport. Much of our understanding of the molecular mechanisms underlying vesicle trafficking comes from the work of three pioneers in this field who were collectively awarded the 2013 Nobel Prize in Physiology or Medicine: Drs Randy Schekman, James E. Rothman and Thomas C. Südhof.

It is widely recognised that vesicle transport, which can either happen constitutively or be tightly regulated, occurs in five general stages: vesicle budding, movement, tethering, docking and membrane fusion [Fig. 1.2].

Figure 1.2. Vesicular transport (Figure legend on next page).

Figure 1.2. Vesicular transport Schematic diagram showing the five general stages of vesicle transport. (1) Budding. Transmembrane and soluble protein cargo, including specific vesicular (v-) SNARE(s), become concentrated at the site of vesicle formation on the donor membrane. Coat proteins are recruited to the cytosolic face of the membrane via interactions with Arf GTPases and induce membrane curvature. Membrane scission at the bud neck, catalysed by dynamin GTPase activity, releases the vesicle from the donor membrane and the vesicle coat then disassembles. (2) Movement. The cargo-laden vesicle is guided along cytoskeletal elements to the acceptor membrane, a process powered by motor proteins which interact with vesicular cargo and/or receptors on the vesicle membrane. (3) Tethering. As the transport vesicle approaches the acceptor membrane, tethering factors interact with vesicular components in preparation for membrane fusion. (4) Docking. The v-SNARE and a target (t-) SNARE on the acceptor membrane zipper into a stable four-helix bundle. (5) Fusion. The trans-SNARE complex ("SNAREpin") promotes fusion of the vesicular and acceptor membranes, inserting transmembrane vesicular cargo into the target membrane and releasing soluble cargo into the acceptor compartment. The fully-zippered cis-SNARE complex components are then recycled. Figure adapted from Bonifacino and Glick (2004). Abbreviations: Arf, ADP-ribosylation factor; GTPase, guanosine triphosphatase; SNARE, soluble N-ethylmaleimide-sensitive fusion factor attachment protein receptor; t-, target; v-, vesicular.

Vesicle budding involves the selective sorting and packaging of intended protein cargo into nascent transport vesicles. This process is driven by the assembly of coat proteins on the cytosolic side of membranes. Coat proteins are recruited to forming vesicles through interactions with small GTP-binding proteins of the ADP-ribosylation factor (Arf) family and adaptor proteins, which themselves are associated with cargo on the donor membrane (Schekman and Orci, 1996). The coat complex serves both to shape the spherical transport vesicle by distortion of membrane conformation and to participate in cargo selection by recognising sorting signals in the cytoplasmic domains of transmembrane cargo (Bonifacino and Lippincott-Schwartz, 2003). Three distinct coats are associated with transport vesicles at the different stages of the protein-sorting pathways (Kirchhausen, 2000a). Clathrin coats are restricted to post-Golgi locations, including the trans-Golgi network (TGN), PM and endosomes (Kirchhausen, 2000b); Coatamer protein I (COP-I) coats are involved in intra-Golgi transport and retrograde transport from the cis-Golgi to the ER (Letoumeur et al., 1994); and COP-II coats mediate anterograde transport from the ER either to the ER-Golgi intermediate complex (ERGIC) or the Golgi complex (Barlowe et al., 1994). Membrane scission at the bud neck, catalysed by dynamin GTPase activity, releases the vesicle from the donor membrane (Hinshaw and Schmid, 1995). The vesicle coat components may then disassemble and return to the cytosol for

7 recycling. Alternatively, the coat may play a role in directing the vesicle to its correct destination (Cai et al., 2007).

Transit of cargo-laden vesicles throughout the cytosol occurs along cytoskeletal elements and is powered by motor proteins from the kinesin, dynein and myosin superfamilies (Kamal and Goldstein, 2000). Different motors are specialised for transporting certain types of cargo and are recruited to transport vesicles via specific interactions with vesicular cargo or other receptors present on the vesicular membrane, such as Rab GTPases (Hammer and Wu, 2002; Karcher et al., 2002) [see 1.7 Rab GTPases in membrane traffic]. When a transport vesicle approaches the acceptor membrane, it initially becomes physically linked to the membrane at a distance (‘tethering’). This process ensures that the vesicle will fuse only with the appropriate target membrane (Brown and Pfeffer, 2010). Tethering is mediated by two broad classes of molecules: extended, coiled-coil proteins, such as the golgin tethers; or large, multi-subunit complexes, such as the ER-localised Dsl1 complex and the exocyst complex at the cell surface (Brown and Pfeffer, 2010; Yu and Hughson, 2010). Tethering factors function by binding a component of the transport vesicle and a component of the opposing acceptor membrane to facilitate fusion. Tethers have been shown to interact with vesicle coat proteins, Rab GTPases and soluble N-ethylmaleimide-sensitive fusion factor (NSF)- attachment protein (SNAP) receptor (SNARE) proteins (Cai et al., 2007).

SNARE and Sec1/Munc18-like (SM) proteins comprise the minimal and universal machinery required for intracellular membrane fusion in eukaryotes (Südhof and Rothman, 2009). Vesicle fusion is mediated by cognate SNAREs on the vesicle and target membranes (v-SNAREs and t-SNAREs, respectively), which zipper into a stable, membrane-bridging four-helical bundle as a trans-SNARE complex (also known as a "SNAREpin") (Weber et al., 1998). The mechanical force of the protein folding involved in complex formation drives membrane fusion thermodynamically, inserting the vesicular cargo into the target membrane (Südhof and Rothman, 2009). NSF primes specific SNARE complexes for docking of vesicles with their target membranes prior to fusion, whereas clasp-shaped SM proteins organise SNAREpin assembly both spatially and temporally (Mayer and Wickner, 1997; Südhof and Rothman, 2009). Individual fusion reactions are fulfilled by different combinations of SNARE and SM proteins to ensure the compartmental specificity of transport processes (Pelham, 2001). In the post-fusion

state, the fully-zippered SNAREpin is termed the cis-SNARE complex (Chen and Scheller, 2001).

1.6 The GLUT4 lifecycle

In adipocytes, only approximately half of intracellular GLUT4 transporters translocate to the PM in response to insulin (Cushman and Wardzala, 1980; James et al., 1988; Holman et al., 1990; Slot et al., 1991a; Slot et al., 1991b), suggesting that GLUT4 occupies more than one cellular compartment. Indeed, electron microscopy studies indicate that GLUT4 is distributed across the endosomal system, TGN and the insulin-responsive GSV compartment in brown and white adipose tissue (Slot et al., 1991b; Malide et al., 2000). Under basal conditions, ~95% of total GLUT4 is retained intracellularly by continuous trafficking through two inter-related loops (Slot et al., 1991b; Yang and Holman, 1993; Blot and McGraw, 2008) [Fig. 1.3]. Surface GLUT4 is efficiently internalised to early endosomes (EE) by clathrin-mediated endocytosis and is sequestered away from the endosome-PM recycling pathway through sorting into GSVs and retrograde transport from endosomes to the TGN (Robinson et al., 1992; Chakrabarti et al., 1994; Martin et al. 2000b; Shewan et al., 2003; Shigematsu et al., 2003; Govers et al. 2004; Karylowski et al., 2004).

Figure 1.3. The GLUT4 lifecycle Schematic diagram of GLUT4 subcellular distribution and translocation between different membranous compartments. Nascent GLUT4 protein is synthesised in the endoplasmic reticulum (ER) and transported via the Golgi cisternae to the trans-Golgi network (TGN) (1). From here, GLUT4 is incorporated into budding GLUT4 storage vesicles (GSVs) (2), which fuse with the plasma membrane (PM) following insulin stimulation (3). When GLUT4 is positioned at the cell surface (4), glucose is transported into the cell along its concentration gradient. GLUT4 also undergoes endocytosis (5) and is trafficked to the endosomal system for recycling. If insulin levels remain elevated, GLUT4 is rapidly recycled back to the PM in vesicles of early endosomal (EE) origin (6). A slower recycling route involving recycling endosomes (REs) also operates (7, 8). In the absence of insulin, GLUT4 is sequestered away from the endosome-PM recycling pathway through retrograde transport from REs to the TGN (9) and sorting into (contd. on next page) 10

GSVs. Alternatively, GLUT4 may be targeted for lysosomal degradation (10). Figure adapted from Sadler et al. (2013). Abbreviations: EE, early endosome; ER, endoplasmic reticulum; GSV, GLUT4 storage vesicle; PM, plasma membrane; RE, recycling endosome; TGN, trans-Golgi network.

Newly synthesised GLUT4 is trafficked through the TGN to GSVs without first cycling to the PM (Watson et al., 2004). Insulin stimulation induces the redistribution of GLUT4 from intracellular membrane compartments to the PM, increasing steady-state surface GLUT4 levels 5 to 30-fold (Holman et al., 1990; Slot et al., 1991b; Malide et al., 2000; Govers et al., 2004). If insulin levels remain elevated, GLUT4 is internalised by endocytosis and recycled back to the PM in vesicles of endosomal origin (Coster et al., 2004; Govers et al., 2004). There are multiple potential endosomal recycling pathways: a direct route from EE to the PM, a slower route involving recycling endosomes (RE), and a third via the TGN (Stöckli et al., 2011). It is unclear which of these recycling routes dominates in the continuous presence of insulin. Once the insulin stimulus is withdrawn, GLUT4 traffics to the RE, from where it can either be targeted back to GSVs via the TGN or undergo lysosomal degradation (Shewan et al., 2003; Shi and Kandror, 2005). Unlike the nearly instantaneous release of neurotransmitter-containing vesicles at a stimulated neuronal synapse, the time-course of insulin action on GLUT4 translocation occurs more slowly (several minutes) (Pakti et al., 2001). This contrast is consistent with the notion that GSVs move over quite some distance prior to tethering, docking and fusion with the plasma membrane in response to insulin and, further, suggest the existence of additional regulatory mechanisms in this process.

1.7 Rab GTPases in membrane traffic

Studies in budding yeast (Saccharomyces cerevisiae) and mammalian systems indicate that successive steps in endomembrane trafficking are mediated by Rab and Arf proteins, subfamilies of the Ras superfamily of monomeric small (25-35 kDa) guanosine triphosphatases (GTPases) conserved throughout eukaryotes (Chavrier and Goud, 1999). Whereas Arfs function primarily in membrane budding (Jackson and Bouvet, 2014), Rab GTPases are master regulators of vesicular membrane trafficking events and constitute

11 the largest branch of the Ras-related GTP binding proteins, with 11 Rabs (Ypts) in yeast, 30 in the nematode worm, Caenorhabditis elegans, and at least 66 members in humans (Hutagalung and Novick, 2011; Klöpper et al., 2012). Rab proteins are distributed among distinct organelles and their associated transport vesicles by reversible membrane insertion (Ali and Seabra, 2005). Multiple structural regions of the Rab can contribute to membrane recruitment, including a divergent carboxy (C)-terminal hypervariable domain (Chavrier et al., 1991; Aivazian et al., 2006) and a double-cysteine prenylation motif in the carboxyl tail (Desnoyers et al. 1996). Since each Rab GTPase has a characteristic subcellular localisation, they are key determinants of organelle and vesicle membrane identity. For instance, in the endocytic pathway, Rab5 and Rab7 are markers of EE and late endosomes (LE), respectively (Rink et al., 2005). Furthermore, endosomal sub- compartments exhibit contiguous yet distinct and functionally diverse Rab domains (Zerial and McBride, 2001). Discrete Rab4, Rab5 and Rab11 membrane domains exist within sorting endosomes (SE) (Sönnichsen et al., 2000), and separate Rab7 and Rab9 domains within LE (Barbero et al., 2002). Zerial and colleagues suggest dynamic Rab conversion as the mechanism of cargo progression between endocytic compartments; that is, endosomes mature by recruiting the necessary Rabs required for recycling cargo to progress along the pathway whilst discarding those Rabs that define the previous endosomal compartment (Rink et al., 2005).

1.8 The Rab cycle

As for all Ras superfamily GTPases, Rab proteins utilise guanine nucleotide exchange and guanine triphosphate (GTP) hydrolysis to function as molecular on/off switches [Fig. 1.4]. The guanine nucleotide status of the Rab determines its cellular localisation and activity. GDP-bound Rabs are inactive and complexed with cytosolic chaperone proteins termed guanine nucleotide dissociation inhibitors (GDIs) (Sasaki et al., 1990; Ullrich et al., 1993; Pfeffer et al., 1995). GDI association masks the prenylated Rab C-terminus, thereby extracting the Rab from its target membrane, and blocks GDP dissociation (Pfeffer et al., 1995; Wu et al., 1996; Pylypenko et al., 2006). Conversely, GTP-bound Rabs are functionally active and distributed to the cytosolic face of membranous transport compartments (Zerial and McBride, 2001). Recently, however, it was demonstrated that

Rab13 traffics on endocytic vesicles regardless of its prenylation or nucleotide status, likely via protein-protein interactions (Ioannou et al., 2016).

Due to their low intrinsic rates of GDP dissociation and GTP hydrolysis, Rab GTPases can sluggishly interconvert between inactive and active forms spontaneously; however, additional upstream regulatory factors are necessary to accelerate their nucleotide cycling [Fig 1.4]. Rab GDP-GTP exchange is mediated by specific guanine nucleotide exchange factors (GEFs), which interact directly with the GDP-bound forms of their substrate Rab(s) and stimulate the removal of GDP, allowing GTP, 10 times more abundant in the cell than GDP, to bind (Stenmark, 2009). The GEF, having a low affinity for the GTP- bound Rab, then dissociates (Stenmark, 2009). There is some evidence that a GDI displacement factor (GDF) disrupts the Rab-GDI complex and helps to target and insert the Rab in the correct membrane prior to GEF action (Sivars et al., 2003). However, two recent studies show that Rab membrane targeting can be dictated by Rab-GEFs (Gerondopoulos et al., 2012; Blümer et al., 2013). Active, GTP-bound Rabs interact with additional downstream proteins, termed Rab effectors, which transduce their signal in their respective transport pathways [Fig. 1.4]. These interactions may involve recruitment of specific effectors to their site of action, protein complex assembly and/or direct stimulation of effector activity. A heterogeneous range of Rab effectors have been identified, including coat proteins, sorting adaptors, tethering factors, kinases, phosphatases, SNAREs, scaffolds and motor proteins (Hutagalung and Novick, 2011). Rab activity is terminated by a cognate GTPase activating protein (GAP) which binds to the Rab and accelerates its inherent GTPase activity, causing hydrolysis of GTP to GDP and consequent disengagement from its effector(s) (Stenmark, 2009). The inactive, GDP- bound Rab then becomes a substrate for membrane extraction by a GDI, returning the Rab to the cytosol where the Rab cycle is reset.

Figure 1.4. The Rab cycle (Figure legend on next page).

Figure 1.4. The Rab cycle Schematic diagram showing the molecular switch mechanism of a Rab GTPase. The inactive, GDP-bound Rab is anchored to the cytosolic face of a membranous donor compartment via its prenylated C-terminus (1). Rab GDP-GTP exchange is mediated by a GEF (2). The now active, GTP-bound Rab interacts with a downstream effector protein (3). In this instance, the effector is a cargo protein in a nascent transport vesicle (4). Fusion of the transport vesicle with the acceptor compartment deposits the Rab in the target membrane (5). Rab activity is then terminated by a GAP, causing hydrolysis of GTP to GDP (6) and consequent disengagement of the Rab from its effector. The inactive, GDP-bound Rab then becomes a substrate for membrane extraction by a GDI (7). A GDF may disrupt the Rab-GDI complex to allow Rab membrane insertion prior to GEF action and thus reset the cycle (8). Abbreviations: GAP, GTPase activating protein; GDF, GDI displacement factor; GDI, guanine nucleotide dissociation inhibitor; GEF, guanine nucleotide exchange factor; Pi, inorganic phosphate.

1.9 Rab cascades

A key question in the field of membrane traffic is how the directionality of transport is achieved. The emerging concept of Rab cascades, pathways in which multiple Rabs, Rab- GAPs and Rab-GEFs template the order of trafficking events by acting in series, provides a satisfying mechanistic answer (Pfeffer, 2013). In such models, an upstream Rab recruits, as an effector, the GEF for the next acting Rab in the transport pathway. To counterbalance the feed-forward GEF cascade, there is evidence for a countercurrent GAP cascade, whereby the downstream Rab recruits the GAP that terminates the activity of the previous Rab in the series (Hutagalung and Novick, 2011; Mizuno-Yamasaki et al., 2012; Pfeffer, 2013) [Fig. 1.5].

Several examples of such cascades are reported in the literature. The Rab5-to-Rab7 conversion of maturing endosomes involves the Rab5-dependent recruitment of Mon1/SAND-1, a GEF for Rab7, which in turn displaces Rabex-5, a GEF for Rab5, from endosomal membranes (Kinchen and Ravichandran, 2010; Poteryaev et al., 2010). In the yeast secretory pathway, Gyp1, a GAP for the early Golgi-localised Ypt1, is an effector of Ypt32 at the late Golgi (Rivera-Molina and Novick, 2009). Ypt32 also recruits Gyp6, a GAP for Ypt6 whose function is to regulate the fusion of endosomal-derived vesicles at the Golgi apparatus (Suda et al. 2013). Therefore, Ypt32 acts to terminate both intra- Golgi and endosome-Golgi traffic through a Rab-GAP cascade. GTP-bound Ypt32 proceeds to recruit Sec2, a GEF for the Rab8 homologue, Sec4, to initiate the budding of

15 post-Golgi, Sec4-containing secretory vesicles (Ortiz et al., 2002). Hence, it is apparent that countercurrent cascades of Rab-GAPs and Rab-GEFs serve to prevent the spatial and temporal overlap of sequentially acting Rabs in vesicular transport pathways, thus helping to define compartment boundaries.

Figure 1.5. Rab cascade model Schematic diagram of a cascade mechanism involving three sequentially-acting Rabs in a pathway: Rab A, Rab B and Rab C. Active, GTP-bound Rab A recruits, as an effector, GEF B, the GEF for the next acting Rab in the pathway, Rab B. In turn, activated Rab B recruits GEF C. To counter-balance the feed-forward GEF cascade and help define the boundaries between individual Rab domains, GTP-bound Rab B recruits the GAP for Rab A, GAP A; and, similarly, Rab C recruits GAP B. Figure adapted from Jean and Kiger (2012). Abbreviations: GAP, GTPase activating protein; GEF, guanine nucleotide exchange factor.

1.10 Rab GTPases in GLUT4 traffic

It is unsurprising, given that GLUT4 traverses multiple membrane compartments en route to the PM, that several Rab proteins have been implicated at some stage of GLUT4 traffic. However, in light of the complexity of the endomembrane system, the specific functions of individual Rabs in GLUT4 traffic have been challenging to define. Proteomic analyses of immunoisolated GLUT4 vesicles from cultured murine adipocytes (Larance et al., 2005; Mîinea et al., 2005) and primary rat adipocytes (Jedrychowski et al., 2010; Fazakerley et al., 2015) have detected Rab1b, Rab2a, Rab5b, Rab8a, Rab10, Rab11 and Rab14 on GSVs. Whereas, in skeletal muscle, Rabs 8a and 13 appear to participate in the

delivery of GSVs to the PM (Ishikura et al., 2007; Sun et al., 2010; Sun et al., 2016); evidence suggests that Rab10 regulates this process in adipocytes (Sano et al., 2007; Sano et al., 2008; Chen et al., 2012; Sadacca et al., 2013). Lienhard and colleagues have demonstrated that small interfering RNA (siRNA)-mediated depletion of endogenous Rab10 in 3T3-L1 adipocytes inhibits insulin-stimulated GLUT4 redistribution to the PM, and re-expression of wild-type Rab10 restores GLUT4 translocation to control levels (Sano et al., 2007; Sano et al., 2008). Knockdown of Rabs 8a, 8b and 14 had no significant effect on GLUT4 translocation in the same assay (Sano et al., 2007; Sano et al., 2008). Live imaging of adipocytes using total internal reflection fluorescence (TIRF) microscopy, a means for visualising cellular events occurring within ~200 nm of the PM (referred to as the TIRF zone) (Burchfield et al., 2010), reveals that Rab10 is the only Rab associated with GSVs that mobilise and fuse directly with the PM in response to insulin (Chen et al., 2012). A constitutively active Rab10 mutant that mimics the GTP-bound Rab conformation, Rab10-Q68L, associates with GSVs and causes their rapid mobilisation to the TIRF zone in the absence of insulin, suggesting that Rab10 activation alone is sufficient for GSV recruitment to the vicinity of the PM (Chen et al., 2012). Rab10-GTP has been shown to bind the actin-based myosin motor, MyoVa (Chen et al., 2012), and the homologous exocyst subunits, Exo6 and Exo6b (Sano et al., 2015), as effectors. Loss of either Exo6 or Exo6b, or both, inhibits GLUT4 translocation in adipocytes (Sano et al., 2015), as does overexpression of a truncated MyoVa construct which competes with the endogenous full-length protein (Chen et al., 2012). These data further implicate a role for Rab10 in the delivery and tethering of GSVs to the PM.

Rab14, on the other hand, resides on GLUT4-positive endosomal compartments and is thought to act upstream of Rab10 to regulate the endocytic trafficking of internalised GLUT4 and the sorting of GLUT4 into GSVs (Chen et al., 2012; Sadacca et al., 2013; Reed et al., 2013; Brewer et al., 2016). However, GLUT4 traffic through both the Rab10 and Rab14 compartments is necessary in insulin-stimulated adipocytes since the loss of Rab 10 and Rab14 additively inhibits GSV exocytosis, and re-introduction of either Rab only partially rescues the phenotype of the double-knockdown (Chen et al., 2012). Additional studies suggest the involvement of Rab4 (Vollenweider et al., 1997; Kaddai et al., 2009; Chen et al., 2012), Rab5 (Huang et al., 2001), Rab11 (Kessler et al., 2000) and Rab31 (Lodhi et al., 2007) in GLUT4 traffic; however, based on the literature, these 17

Rabs likely regulate the endosomal and/or TGN trafficking of GLUT4, as opposed to GLUT4 exocytosis.

1.11 TBC domain proteins: GAPs for Rab GTPases

Rab-GAPs in all eukaryotes share a conserved protein module of 180-200 amino acids, the Tre2/Bub2/Cdc16 (TBC) domain, that was originally identified as common to the tre- 2 oncogene and the yeast cell cycle regulators, Bub2 and Cdc16, and which harbours enzymatic GAP activity (Richardson and Zon, 1995; Fukuda, 2011). The only known Rab-GAP lacking a TBC domain is the heterodimeric Rab3GAP1/Rab3GAP2 complex that regulates Rab3 (Fukui et al., 1997; Nagano et al., 1998). To date, 44 TBC domain- containing proteins have been found in humans and mice, which is similar to the number of Rabs in mammals (Gabernet-Castello et al., 2013). One theory regarding this similarity is that each Rab-GAP has a specific Rab target. The Rab substrates for some TBC proteins have been identified; however, the majority of TBC domain proteins are poorly characterised. Crystallographic, mutational and functional analyses of the TBC domain of the yeast Rab-GAP, Gyp1, in complex with Rab GTPases revealed that the TBC domain uses an arginine/glutamine dual-finger mechanism to catalyse GTP hydrolysis (Albert et al., 1999; Rak et al., 2000; Du and Novick, 2001; Pan et al., 2006). However, since a substantial number of TBC proteins lack these conserved catalytic residues, alternative catalytic mechanisms may exist or such proteins may not function as Rab- GAPs at all (Fukuda, 2011). Several TBC proteins have been shown to interact with non- substrate Rabs via a domain other than their TBC domain, possibly as means of targeting GAP activity to the correct intracellular location and/or small GTPase cascade assembly (Itoh et al., 2006; Fukuda et al., 2008; Kanno et al., 2010).

The Rab10 GAP, TBC1 domain family member 4 (TBC1D4) (hereinafter referred to as Akt substrate of 160 kDa, AS160) has been strongly implicated in the regulation of GLUT4 translocation in adipocytes (Sano et al., 2003; Equez et al., 2005; Mîinea et al., 2005) [see 1.12 The Rab-GAP, AS160]; whereas the closely related GAP, TBC1D1, controls this process in skeletal muscle tissue (Peck et al., 2009). More recently, TBC1D13 has been identified as a suppressor of GLUT4 traffic in adipocytes (Davey et al., 2012). TBC1D13 is a GAP for Rab35, a PM-localised Rab that regulates a rapid 18

recycling pathway between early endosomes and the PM (Kouranti et al., 2006; Prior et al., 2011; Davey et al., 2012). Overexpression of TBC1D13 inhibits insulin-stimulated GLUT4 translocation in 3T3-L1 adipocytes and this effect is partially reversed by the constitutively active Rab35-Q69L mutant (Davey et al., 2012). These data implicate a novel role for Rab35 in the regulation of GLUT4 traffic. Furthermore, TBC1D13 interacts with Rab10 suggesting that this GAP may participate in a Rab cascade involving Rabs 10 and 35 (Davey et al., 2012).

1.12 The Rab-GAP, AS160

AS160 was first identified as a novel Akt substrate by immunoprecipitation of proteins from insulin-treated adipocytes using an antibody directed against the phospho-form of the Akt phosphorylation motif, RXRXXS/T (referred to as the phospho Akt substrate (PAS) antibody) (Kane et al., 2002). AS160 possesses a C-terminal TBC domain, a Calmodulin (CaM)-binding domain and two amino (N)-terminal PTB domains (Kane et al., 2002; Kane and Lienhard, 2005). A lipid-binding domain is contained within the second PTB domain (Tan et al., 2012). Six putative Akt phosphorylation sites have been identified in the human AS160 protein, five of which (Ser318, Ser570, Ser588, Thr642 and Ser751) are insulin-regulated (Kane et al., 2002; Sano et al., 2003).

There is strong experimental evidence to suggest a key role for AS160 in the regulation of GLUT4 traffic. Overexpression of an AS160 mutant in which four of the insulin- sensitive phosphorylation sites (Ser318, Ser588, Thr642 and Ser751) are mutated to alanine (AS160-4P) markedly inhibits insulin-stimulated GLUT4 translocation in 3T3-L1 adipocytes (Sano et al., 2003). Moreover, this inhibition does not occur when the GAP function of AS160-4P is abolished by mutation of the catalytic arginine residue of the AS160-4P TBC domain (Arg973) to lysine, indicating that the inhibitory effect of AS160- 4P on GLUT4 translocation requires Rab-GAP activity (Sano et al., 2003). These findings provided a key connection between the upstream PI3K/Akt insulin signalling pathway and the distal GLUT4 trafficking machinery, and prompted Lienhard and colleagues to propose a model, described here, whereby AS160 functions as a negative regulator of GLUT4 traffic in adipocytes (Sano et al., 2003). Under basal (serum-starved) conditions, 19 it is thought that the active GAP domain of dephosphorylated AS160 maintains its cognate Rab in an inactive, GDP-bound conformation thus retaining GLUT4 intracellularly. Insulin-stimulated, Akt-mediated AS160 phosphorylation presumably inhibits the GAP activity of AS160 (either directly or indirectly), allowing for the GTP- loading and activation of the Rab that regulates the fusion of GSVs with the PM.

Several candidates for the Rab substrate of AS160 are suggested in the literature. AS160 displays GAP activity towards Rabs 2a, 8a, 10 and 14 in vitro, all of which are present on GSVs purified from 3T3-L1 adipocytes (Larance et al., 2005; Mîinea et al., 2005); however, Rab10 is the most likely in vivo target of AS160 in these cells (Sano et al., 2007; Sano et al., 2008; Sadacca et al., 2013). siRNA-mediated knockdown of AS160 causes an increase in the surface GLUT4 level of basal adipocytes (Eguez et al., 2005; Larance et al., 2005) and Rab10 depletion in AS160 knockdown cells partially restores basal GLUT4 retention (Sano et al., 2007). Rab10 activity downstream of AS160 inhibition is thought to regulate a distal, pre-fusion step in GLUT4 traffic, involving the recruitment and/or docking of GSVs within the TIRF zone in response to insulin (Gonzalez and McGraw, 2006; Jiang et al., 2008; Fujita et al., 2010; Xiong et al., 2010; Brewer et al., 2011; Sadacca et al., 2013).

In the basal state, AS160 is localised to GSVs via its interaction with the cytosolic tails of the vesicle cargoes, insulin-regulated aminopeptidase (IRAP) and low density lipoprotein receptor-related protein 1 (LRP1) (Larance et al., 2005; Peck et al., 2006; Jedrychowski et al., 2010). Such membrane association is necessary for the intracellular retention of GLUT4 in the absence of insulin (Eguez et al., 2005; Stöckli et al., 2008). Insulin stimulation causes AS160 to dissociate from GSVs and bind to 14-3-3 protein isoforms [see Chapter 4, 4.1 Introduction] via its Akt phosphorylation site, Thr642 (Kane et al., 2002; Larance et al., 2005; Ramm et al., 2006). Insertion of a constitutive 14-3-3 binding peptide sequence into the aforementioned AS160-4P protein reverses the inhibitory effect of this dominant negative mutant on insulin-stimulated GLUT4 translocation (Ramm et al., 2006). However, since the fusion of AS160-4P to GLUT4 is sufficient to block insulin-stimulated GLUT4 translocation, the disengagement of AS160 from GSVs is not necessary to relay the insulin signal to GLUT4 (Stöckli et al., 2008). Rather, studies indicate that the insulin- and Akt-dependent phosphorylation/14-3-3 association of AS160 regulates its GAP activity in adipocytes (Ramm et al., 2006; Stöckli

et al., 2008). In contrast to Larance et al. (2005), Holman and colleagues observe an insulin-dependent recruitment of AS160 to GSVs in rat adipocytes (Koumanov et al., 2011). Furthermore, the γ isoform of 14-3-3 is highly enriched on GSVs in insulin- stimulated cells (Koumanov et al., 2011). Therefore it is possible that AS160 remains on GSVs until after fusion has occurred. Supporting this theory, a small pool of highly phosphorylated AS160 is located at the adipocyte PM following insulin treatment, suggesting an additional function for AS160 at this location (Ng et al., 2010). Indeed, a more recent study speculates that non-phosphorylated, GSV-associated AS160 encounters active Akt at the PM, where it becomes phosphorylated and proceeds to play an active role in the docking and/or fusion of GSVs (Tan et al., 2012). Hence, AS160 may behave as a regulatory switch capable of both inhibiting and facilitating GLUT4 translocation (Tan et al., 2012).

Despite the immense amount of research on the role of AS160 in GLUT4 traffic over the last decade, much less is known about the Rab-GEF(s) that positively co-regulate(s) Rab10 function in this process [see 1.14.8 DENND4A-C]. Since AS160 knockdown results in only a partial redistribution of GLUT4 to the PM (Eguez et al., 2005; Larance et al., 2005), one possibility is that simultaneous AS160 inhibition and GEF activation is necessary for insulin-stimulated GLUT4 exocytosis. Furthermore, in view of the complexity of the GLUT4 trafficking itinerary, additional, as-yet-unknown Akt substrates that participate in GLUT4 traffic are likely to exist.

1.13 Rab-GEFs

Unlike Rab-GAPs, unified by their TBC domains, Rab-GEFs comprise a diverse range of structurally unrelated, yet evolutionary conserved, proteins and protein complexes (Barr and Lambright, 2010). The most well-known Rab-GEF protein module is the vacuolar protein sorting 9 (Vps9) domain (Burd et al., 1996). The human genome encodes at least 7 Vps9 domain-containing proteins, the best characterised of which is Rabaptin- 5 associated exchange factor for Rab5 (Rabex-5) (Horiuchi et al., 1997; Delprato and Lambright, 2007). Rabex-5 and other Vps9 domain proteins are highly selective for members of the Rab5 subfamily (Rabs5a-c, 17, 21, 22a and 22b/31) (Delprato et al., 2004). 21

All of these Rabs are involved in endocytic trafficking (Zacchi et al., 1998; Zerial and McBride, 2001; Kauppi et al., 2002; Simpson et al., 2004; Magadán et al. 2006). Rabin proteins, the mammalian homologues of yeast Sec2, are GEFs for Rab8/Sec4 (Hattula et al., 2002; Hattula et al., 2005). Rab8 is implicated in cell migration, epithelial polarisation, neuronal structural plasticity and ciliary membrane formation (Peränen, 2011).

In several cases protein oligomerisation is necessary for GEF activity. Ric1 and Rgp1 form a dimeric complex that stimulates guanine nucleotide exchange on Rab6a/Ypt6 to control the recycling of SNAREs and other exocytic machinery from EE to the Golgi (Siniossoglou et al., 2000; Pusapati et al., 2012). Similarly, Mon1/SAND-1 and its binding partner Ccz1 form a GEF for Rab7/Ypt7 to regulate endosomal maturation (Nordmann et al., 2010; Poteryaev et al., 2010). Hermansky–Pudlak syndrome (HPS) proteins HPS1 and HPS4 associate to form the biogenesis of lysosome-related organelles complex 3 (BLOC-3) complex, a GEF for Rab32 and Rab38 in melanosome biogenesis (Nazarian et al., 2003; Gerondopoulos et al., 2012). HSP1 and HPS4 are homologues of Mon1 and Ccz1, respectively, and Rabs 32 and 38 are phylogenetic relatives of Rab7 (Cheli and Dell’Angellica, 2010; Klöpper et al., 2012). The multisubunit transport protein particle (TRAPP) complexes (TRAPP-I, TRAPP-II and TRAPP-III) use a common core of subunits to activate Rab1/Ypt1 (Barrowman et al., 2010). TRAPP-I is a heptameric vesicle tethering complex at the cis-Golgi (Sacher et al., 1998; Wang et al., 2000). The TRAPP-II tether contains three additional subunits which switch the GEF specificity of TRAPP to favour Ypt31/32, homologues of mammalian Rab11, in post-Golgi traffic (Morozova et al., 2006; Sacher et al., 2008; Zou et al., 2012; Levine et al., 2013a). TRAPP-III plays a role in autophagy and is directed to the phagophore assembly site (PAS) by its Trs85 and Trs120 subunits (Lynch-Day et al., 2010; Taussig et al., 2014).

A large proportion of the 66 human Rabs, however, lack a known cognate GEF (Barr and Lambright, 2010). Moreover, few mammalian homologues of yeast GEFs have been identified; therefore additional, mammal-specific Rab GEFs are likely to exist (Yoshimura et al., 2010).

1.14 DENN domain proteins: a novel Rab-GEF family

A major recent development in the search for Rab regulators came with the identification of the DENN (differentially expressed in normal versus neoplastic) domain as a novel conserved Rab-specific GEF module (Allaire et al., 2010; Yoshimura et al., 2010; Marat et al., 2011). Levivier et al. (2001) described the DENN domain architecture comprising three indissociable sub-regions: a core DENN module flanked by upstream (u- DENN/longin) and downstream (d-DENN) motifs. However, the current viewpoint is that the DENN domain is bi-lobed with the N-terminal lobe being a longin domain (LD) (Wu et al., 2011). LDs dimerise to form platforms for small GTPases and have emerged as structural features of several Rab-GEFs, including Mon1-Ccz1, BLOC-3 and the human TRAPP-II subunit, TRAPPC10 (Levine et al., 2013a). Like Rabs, DENN domain- containing proteins have diversified during evolution, with 1 found in the fission yeast, Schizosaccharomyces pombe, 5 in C. elegans and 26 members currently identified in humans (Marat et al., 2011; Levine et al. 2013a, Levine et al., 2013b; Zhang et al., 2012). They are notably absent in budding yeast (Yoshimura et al., 2010). The DENN domain was first indicated as a Rab-GEF domain when the DENN domain-containing protein, MADD (MAP kinase activating death domain protein), purified from rat brain tissue, displayed GEF activity towards members of the Rab3 subfamily (Wada et al., 1997; Brown and Howe, 1998). MADD and its worm homologue, AEX-3, were later shown to regulate Rab27, a close relative of Rab3 (Mahoney et al., 2006; Figueiredo et al. 2008; Klöpper et al., 2012). Rab3 and Rab27 have both been implicated in synaptic vesicle exocytosis (Pavlos et al., 2010).

In humans, a core family of 18 DENN proteins whose DENN domains are reasonably well conserved in sequence are grouped into 8 subfamilies, each activating a common Rab (Yoshimura et al., 2010; Marat et al., 2011). These are: DENND1A-1C; DENND2A- 2D; DENND3; DENND4A-4C; DENND5A/5B; DENND6A/6B; myotubularin-related proteins 5 and 13 (MTMR5/13) and MADD [Fig 1.7]. It is thought that having multiple GEFs for a single Rab within each subfamily may allow for activation of the Rab towards distinct cellular activities and/or at different subcellular locations (Yoshimura et al., 2010; Marat et al., 2011). The wider DENN family is more divergent and members, listed here, were identified by more sensitive sequence and fold-recognition analyses: AVL9,

C9orf72/DENNL72, FAM45A, folliculin (FLCN), folliculin-interacting proteins 1 and 2 (FNIP1/2), LCHN/KIAA1147, nitrogen permease regulator 2 and 3 (NPR2/3) and Smith- Magenis candidate region 8 protein (SMCR8) (Zhang et al., 2012; Levine et al. 2013a, Levine et al., 2013b). However, it is unclear whether these DENN-related proteins are Rab-GEFs. The functional roles of a small number of DENN proteins are known (reviewed in Marat et al., 2011); however the majority are poorly characterised. An overview of the current knowledge on each core DENN subfamily is provided in the following sections.

1.14.1 DENND1A-C (connecdenn1-3)

The best-described DENN subfamily is DENND1A-1C, also known as connecdenn1-3, whose members exhibit varying degrees of GEF activity towards Rab35 (Allaire et al., 2010; Marat and McPherson, 2010; Yoshimura et al., 2010). One study implicates DENND1C as a GEF for Rab13 (Yoshimura et al., 2010). DENND1A and DENND1B regulate Rab35 activity in endocytic trafficking, whereas DENND1C activates Rab35 to control actin dynamics (Allaire et al., 2006; Allaire et al., 2010; Yoshimura et al., 2010; Marat et al., 2012). In C. elegans oocytes, the single DENND1 orthologue, receptor- mediated endocytosis 4 (RME-4), functions as a GEF for Rab35/RME-5 in yolk protein endocytosis (Sato et al., 2008). The DENN domain of DENND1A/1B interacts directly with GDP-bound Rab35, though that of DENND1C does not, indicating that Rab35- binding and GEF activity are separable features of the DENN domain (Marat and McPherson, 2010). All three DENND1 family members bind to clathrin and to the clathrin adaptor, AP-2, via their divergent C-termini (Marat and McPherson, 2010). A unique motif in the DENND1C C-terminus confers actin binding (Marat et al., 2012). Genome-wide association studies (GWAS) have linked DENND1A to polycystic ovary syndrome (Welt et al., 2012) and DENND1B to childhood asthma and Crohn’s disease (Sleiman et al., 2010; Franke et al., 2010). DENND1B was also identified as a candidate gene for T2D (Chen et al., 2013).

Figure 1.6. Human DENN domain-containing proteins Schematic diagram showing the domain architecture and amino acid lengths of the 18 core DENN proteins encoded in the human genome. Proteins are grouped into 8 subfamilies based on sequence similarity. The DENN domain is located towards the N-terminus of each protein, besides in the DENND2 subfamily, where it positioned nearer the C-terminus. Outside of the DENN domain, there is little sequence homology between subfamilies; however each is characterised by unique motifs and protein modules. For DENND6 subfamily members, the DENN domain is the only recognisable feature. The protein isoform shown (if applicable) is that of the canonical sequence listed in the UniProtKB/Swiss-Prot database. (Contd. on next page).

Abbreviations: aa, amino acids; AP-2, AP-2 interaction motif; CB, clathrin-binding motif; CalB, calmodulin- binding motif; d, downstream DENN motif; DD, death domain; DENN, core DENN module; GRAM, glucosyltransferase/Rab-like GTPase activator/myotubularin domain; ISRE, interferon-stimulated response element; LD, longin domain; MABP, MVB12-associated β-prism; NLS, nuclear localisation signal; PH, pleckstrin homology domain; PLAT, polycystin-1/lipoxygenase/α-toxin domain; poly-D, poly-aspartic acid domain; poly-E, poly-glutamic acid domain; poly-Q, poly-glutamine domain; PPR, pentatricopeptide repeat; PRD, proline-rich domain; pseudo-phosph, pseudo-myotubularin phosphatase domain; RUN, RPIP8/UNC-14/NESCA domain; SRD, serine-rich domain; WD40, WD40 repeat motif.

1.14.2 Folliculin

The C-terminal lobe of the DENND1B DENN domain is structurally similar to the C- terminus of the tumour-suppressor protein, FLCN (Nookala et al., 2012). Mutations in the FLCN gene are associated with a rare autosomal dominant condition, Birt-Hogg-Dubé (BHD) syndrome, leading to renal cell carcinoma and other tumours, partly through deregulation of the mTOR signalling pathway (Nickerson et al., 2002; Baba et al., 2008). The most common FLCN mutations in BHD patients generate a truncated FLCN protein that lacks the DENN-like C-terminal half (Schmidt et al., 2005). The FLCN C-terminus is a GEF for Rab35 in vitro; however it is not known whether Rab35 is the physiological Rab substrate of the FLCN protein (Nookala et al., 2012). FLCN has two binding partners, FNIP1 and FNIP2, which interact independently with FLCN to form a FLCN/FNIP complex (Baba et al., 2006; Hasumi et al., 2006; Takagi et al., 2008). FNIP1 and FNIP2 are themselves DENN proteins, although large unstructured insertions exist within their lobular longin and DENN domains (Zhang et al., 2012). A FLCN/FNIP complex has been shown to interact with AMP-activated protein kinase (AMPK) to inhibit AMPK signalling (Baba et al., 2006; Possik et al., 2014; Yan et al., 2014). More recently, a role for the FLCN/FNIP1 complex in mTORC1 activation has emerged (Petit et al., 2013; Tsun et al., 2013). In HeLa cells lacking FLCN, mTORC1 fails to localise to the lysosome, its site of activation, in response to amino acid stimulation (Petit et al., 2013). In the same study, FLCN was shown to interact directly with RagA via its GTPase domain and be recruited, in a FNIP1-dependent manner, to lysosomal membranes under conditions of amino acid starvation, but rapidly dissociate in response to amino acid re-supplementation (Petit et al., 2013). These observations led the authors to speculate that FCLN may function as a GEF for RagA/B GTPases (Petit et al., 2013). In contrast to these findings,

Tsun et al. (2013) propose that FLCN is a GAP for RagC/D GTPases at the lysosome, and not a GEF. Therefore the precise role of FLCN is unclear.

1.14.3 DENND2A-D

Members of the DENND2A-D subfamily have been shown to function as Rab9 GEFs in vitro (Yoshimura et al., 2010). Rab9 controls the trafficking of the mannose-6-phosphate receptor (MPR) between the TGN and LE (Lombardi et al., 1993; Díaz et al., 1997). Several studies implicate DENND2D as a candidate tumour suppressor gene downregulated by promoter hypermethylation in cancer cells (Kanda et al., 2014; Hibino et al., 2014; Kanda et al., 2015). Recently, a role for DENND2B as a GEF for Rab13 in cancer cell metastasis has been described (Ioannou et al., 2015). In polarised epithelial cells, Rab13 regulates exocytic membrane traffic between the TGN and the PM via RE (Nokes et al., 2008). DENND2B activates Rab13 at the epithelial cell periphery and binds to the Rab13 effector, molecule interacting with CasL-like 2 (MICAL-L2), to induce the membrane remodelling necessary for cell migration and invasion (Ioannou et al., 2015).

1.14.4 DENND3

DENND3 is a GEF for Rab12, both in vitro and in vivo (Yoshimura et al., 2010; Matsui et al., 2014). The regulation of Rab12 activity by DENND3 has been implicated in mTORC1 signalling and autophagy (Matsui et al., 2014). In mouse embryonic fibroblasts (MEFs), DENND3 activates Rab12 to control the trafficking of proton/amino transporter 4 (PAT4) from RE to lysosomes for degradation (Matsui and Fukuda, 2013; Matsui et al., 2014). In the same study, knockdown of DENND3 was shown to reduce Akt activity (Matsui et al., 2014).

1.14.5 DENND5A/B

DENND5A and DENND5B (also known as Rab6-interacting protein 1 (Rab6IP1) and Rab6-interacting protein 1-like (Rab6IP1L), respectively) are GEFs for Rab39 in vitro (Yoshimura et al., 2010). Rab39a has been implicated in the trafficking of multivesicular bodies (MVBs) to bacterial inclusions, whereas Rab39b regulates post-Golgi vesicular traffic during neurogenesis (Giannandrea et al., 2010; Gambarte Tudela et al., 2015). DENND5A interacts with Rab6 via its first RPIP8/UNC-14/NESCA (RUN) domain, a module found in several proteins related to the functions of Rab GTPases (Yoshida et al., 2011). The second DENND5A RUN domain has been shown to bind the endosomal sorting protein, sorting nexin 1 (SNX1) (Fernandes et al., 2012). Additionally, DENND5A binds GTP-bound Rab11, possibly via its LD (Miserey-Lenkei et al., 2007). Rab11 acts upstream of Rab6 in a retrograde transport route from RE to the Golgi (Miserey-Lenkei et al., 2007). Hence, DENND5A juxtaposes Rab6 and Rab11, although the function of this and the precise relationship between Rab39, Rab6 and Rab11 remain uncertain.

1.14.6 DENND6A/B

Until recently, the DENND6 subfamily of proteins, which are composed exclusively of a DENN domain, were completely undescribed. However, DENND6A/B, also called FAM116A/B in the literature, are now known to function as GEFs for Rab14 in the endocytic recycling of cell surface disintegrin and metalloproteinase (ADAM) proteases and regulation of cell-cell adherens junctions (Linford et al., 2012).

1.14.7 MTMR5/13

In vitro, MTMR5 and MTMR13 display GEF activity towards Rab28, a distant member of the Rab superfamily (Yoshimura et al., 2010; Klöpper et al., 2012). Rab28 co-localises with Vps23, a component of the endosomal sorting complex required for transport (ESCRT) complex, in the endosomal pathway, where it regulates late endocytic traffic

(Lumb et al., 2011). The Drosophila melanogaster MTMR13 orthologue, Sbf, is a GEF for Rab21 in endolysosomal trafficking (Jean et al., 2012; Jean et al., 2015). Starvation induces Sbf/MTMR13 GEF activity towards Rab21 and their interaction with VAMP8, a SNARE effector required for autophagosome-lysosome fusion (Jean et al., 2015). Mammalian MTMR5 and MTMR13 may therefore likely play roles in endosomal function.

1.14.8 DENND4A-C

The DENND4 subfamily members are GEFs for Rab10 in vitro (Yoshimura et al. 2010). Besides the N-terminal DENN domain, DENND4A-C each possess an N-terminal MVB12-associated β-prism (MABP) domain, a membrane targeting module (Boura and Hurley, 2012); one or two pentatricopeptide repeats (PPR), regions thought to mediate RNA interactions (Manna, 2015); and a C-terminal interferon-stimulated response element (ISRE)-binding region (Marat et al., 2011) [Fig. 1.6]. In addition to these common structural elements, DENND4A, also known as IrlB/C-myc promoter-binding protein (MycPBP), has a central CaM-binding domain (CalB), and both DENND4A and DENND4C possess nuclear localisation signals (NLS). DENND4A, which is ubiquitously expressed, has been shown to interact with the ISRE in the promoter of human c-myc (Stasiv et al., 1994; Semova et al., 2003). However, it is unclear whether DENND4A and/or DENND4C localise to the nucleus. A role for these GEFs in this location is inconsistent with Rab10 function in membrane trafficking. Indeed, in HeLa cells, DENND4A exhibits a diffuse cytoplasmic localisation, whereas DENND4B and DENND4C are present on tubular structures emanating from the perinucleus (Yoshimura et al. 2010).

In Drosophila, calmodulin-binding protein related to a Rab3 GDP-GTP exchange protein (CRAG), the single fly orthologue of the human DENND4 family, regulates the Rab10- dependent polarised secretion of basement membrane proteins in epithelial cells (Denef et al., 2008; Lerner et al. 2013). CRAG displays GEF activity towards both Rab10 and Rab11 in vitro; however the kinetics of nucleotide exchange are much faster for Rab10 than Rab11 (Xiong et al., 2012). One study has shown that CRAG is a GEF for Rab11 in 29 the post-Golgi vesicular transport of Rhodopsin light sensors to the membrane stacks of photoreceptor cells in the Drosophila eye (Xiong et al., 2012). Notably, Rab10 is not expressed in these cells (Chan et al., 2011). Loss of CRAG, which leads to the accumulation of newly-synthesised Rhodopsin in the cytosol, can be rescued by human DENND4A, suggesting that these orthologous proteins have conserved functions (Xiong et al., 2012).

The discovery of DENND4A-C as putative physiological Rab10 GEFs is particularly intriguing given the key regulatory roles of Rab10 and its GAP, AS160, in GLUT4 translocation. Lienhard and colleagues have proposed that DENND4C is the primary Rab10 GEF required for insulin-regulated GLUT4 translocation in adipocytes (Sano et al., 2011). In insulin-stimulated 3T3-L1 adipocytes, siRNA-mediated knockdown of DENND4C reduces the amount of GLUT4 at the PM by 60% relative to control cells, whereas overexpression of DENND4C boosts surface GLUT4 levels by almost 30% (Sano et al., 2011). Neither knockdown nor overexpression of DENND4C has an effect on the level of PM GLUT4 under basal conditions (Sano et al., 2011). Three insulin- sensitive phosphorylation sites on murine DENND4C have been reported (Ser1043, Ser1096 and Ser1321) and each lies within the partial Akt phosphorylation motif, RXXS/T (Sano et al., 2011). It was hypothesised that insulin-stimulated, Akt-mediated phosphorylation at these sites might regulate the GEF activity of DENND4C. However, overexpression of the nonphosphorylatable DENND4C S1043A/S1096A/S1321A mutant in insulin-treated 3T3-L1 cells has a stimulatory effect on GLUT4 translocation comparable to its wild- type counterpart (Sano et al., 2011). These results imply that insulin-stimulated phosphorylation of DENND4C does not regulate its GEF activity. Consistent with this, there is evidence to suggest that the GEF activity of DENND4C is constitutively active (Sadacca et al., 2013). Knockdown of DENND4C reduces the increased level of surface GLUT4 observed in AS160 knockdown cells under basal conditions; that is, even in the absence of insulin, the DENND4C GEF is sufficiently active that its depletion can inhibit GLUT4 translocation to the PM (Sadacca et al., 2013).

Little is known, however, about the functional roles of DENND4A and DENND4B in mammals. Given the domain similarity between DENND4A and DENND4C, it is not unfathomable that DENND4A might also direct Rab10 activity in GLUT4 traffic. This possibility remains to be explored.

1.15 The present study

The James laboratory recently performed a mass spectrometry-based phosphoproteomic screen to identify novel insulin-regulated targets in the 3T3-L1 adipocyte proteome (Humphrey et al., 2013). Using this approach, 4,182 phosphorylation sites in 2,218 proteins that change greater than 2-fold with insulin stimulation were uncovered. Among the most insulin-responsive proteins identified in adipocytes were two members of the DENND4 subfamily of Rab-GEFs, DENND4A and DENND4C, which are thought to regulate Rab10 activity. These proteins exhibited 5- to 30-fold increases in phosphorylation at 1-2 sites following insulin treatment. Some of the phosphosites identified are high stringency consensus sites for Akt-mediated phosphorylation/14-3-3 binding. Notably, DENND4A has been identified independently as an insulin-responsive 14-3-3 binding partner in a previous study (Larance et al., 2010). A role for DENND4C in GLUT4 translocation is already implicated and three insulin-regulated phosphosites in the DENND4C protein are described (Sano et al., 2011). However, Humphrey et al. (2013) have identified novel insulin-sensitive phosphosites on DENND4C that have yet to be reported in the literature. Furthermore, DENND4A phosphorylation has not been described and the protein has yet to be linked to insulin signalling. DENND4C has also been shown to localise to GSVs, however it is not known whether DENND4A is similarly associated with this compartment (Sano et al., 2011).

The findings of Humphrey and colleagues (2013) are particularly intriguing given that the Rab10 GAP, AS160, which plays a major role in GLUT4 translocation, is regulated by Akt-mediated phosphorylation and 14-3-3 binding downstream of insulin signalling. I therefore hypothesise that DENND4A and/or DENND4C are regulated in a similar manner and, in concert with AS160 and Rab10, comprise the major regulatory hub in the insulin-stimulated GLUT4 trafficking network. Alternatively, DENND4A and/or DENND4C might direct Rab10 activity towards another insulin-regulated cellular process.

The specific aims of this study are to:

1. Map the insulin-regulated phosphorylation sites on DENND4A and DENND4C.

2. Identify the upstream kinase(s) responsible for DENND4 protein phosphorylation.

3. Determine whether DENND4A and DENND4C are 14-3-3 binding partners and, if so, identify the phosphosite(s) responsible for this interaction.

4. Characterise the functional role of DENND4A phosphorylation in insulin action.

5. Validate that DENND4C is localised to GSVs and probe the nature of this interaction.

Ultimately, understanding the underlying mechanisms of GLUT4 trafficking may uncover novel therapeutic targets for the treatment of insulin resistance and T2D.

Chapter 2: General Materials and Methods

This chapter contains all of the materials and methods related to this thesis which were used in multiple chapters. Those methods which are relevant to one chapter alone are found in the methods section of that particular chapter.

2.1 Materials and antibodies

All cell lines were obtained from the American Type Culture Collection (ATCC, Rockville, MD). Dulbecco’s Modified Eagle Medium (DMEM), Opti-Minimum Essential Medium (opti-MEM), fetal bovine serum (FBS) and GlutaMAX were purchased from Thermo Fisher Scientific (Waltham, MA). Cell culture dishes were from Corning (Corning, NY). Insulin was obtained from Calbiochem (San Diego, CA). MK- 2206 was from Sellekchem (Houston, TX). Torin-1 was from Tocris Bioscience (Bristol, UK). Bovine serum albumin (BSA) was from Bovogen Biologicals. Trypsin was from Promega (Madison, WI). Acrylamide/bis-acrylamide solution was from Bio-Rad (Hercules, CA). PVDF membrane and HRP chemiluminescent substrate solution was from Millipore (Billerica, MA). DNA polymerases, restriction endonucleases and competent bacteria were all from New England BioLabs (Ipswich, MA). All chemicals were sourced from Sigma-Aldrich (St. Louis, MO) unless stated otherwise in the text.

Antibodies were purchased from Abcam, Cambridge, UK (mouse anti-GFP); Cell Signalling Technology, Beverly, MA (anti-Akt, anti-phospho-Akt Ser473, anti-phospho- ERK1/2 Thr202/Tyr204, anti-GST, anti-phospho-mTOR Ser2448, anti-phospho-S6K Thr389, anti-phospho-ULK1 Ser757, anti-Rab10); Jackson ImmunoResearch, West Grove, PA (anti-mouse IgG; anti-rabbit IgG); Molecular Probes, Eugene, OR (rabbit anti-GFP); Novus Biologicals, Littleton, CO (anti-EDC3, anti-DENND4A); Santa Cruz Biotechnology, Santa Cruz, CA (anti-pan-14-3-3) and Sigma-Aldrich, St. Louis, MO (anti-DENND4C, anti-FLAG M2, anti-α-tubulin). The antibodies against GLUT4 (1F8) (James et al., 1988) and AS160 (Larance et al., 2005) were produced as previously

33 described. Horseradish peroxidase (HRP)-conjugated secondary antibodies were from GE Healthcare (Buckinghamshire, UK). Infrared dye 700- or 800-conjugated secondary antibodies were from Rockland Immunochemicals (Gilbertsville, PA).

2.2 Plasmids pcDNA™4/TO-eGFP-DENND4A (human) and pcDNA™4/TO-eGFP-DENND4C (mouse) were gifts from Prof. Francis Barr (University of Oxford, UK). pGEX-4T-3 plasmid vector expressing a thrombin-cleavable GST-14-3-3β (human) was obtained from Prof. John Hancock (Institute of Molecular Bioscience, Brisbane). pGEX-6P vector expressing the GST-tagged, C-terminal Rab-binding domain (amino acids 806-1009) of MICAL-L2 (mouse) (described by Sano et al., 2008) was a gift from Prof. Gustav Lienhard (Dartmouth College, Hanover, NH). pcDNA™3.1(+)-FLAG-Rab10 (mouse) (described by Sadacca et al., 2013) was obtained from Prof. Timothy McGraw (Cornell University, New York, NY). pGEM-T-GLUT4-TagRFP-T vector was obtained from Dougall Norris (James laboratory). p3XFLAG-CMV™-10-AS160 WT and p3XFLAG- CMV™-10-AS160 4P have previously been described (Ramm et al., 2006). pGEX-5X- 3 vector expressing the GST-tagged, N-terminal cytosolic domain (amino acids 2-109) of insulin-regulated aminopeptidase (IRAP/vp165) (described by Keller et al., 1995) was a gift from A/Prof. Susanna R. Keller (University of Virginia, Charlottesville, VA). pGEX- KG vector expressing the GST-tagged, N-terminal cytosolic domain (amino acids 1-94) of VAMP2 (described by Calakos et al., 1994) was a gift from Prof. Richard Scheller (Stanford University, Santa Clara, CA). pGEX-KG vector expressing the GST-tagged carboxyl tail (amino acids 466-509) of GLUT4 (described by Larance et al., 2005) was obtained from Dr. Mark Larance (James laboratory).

2.3 Methods

2.3.1 Molecular cloning

DENND4A cDNA (human) was amplified from pcDNA™4/TO-GFP-DENND4A using attB site-containing forward (5'-GGGGACAAGTTTGTACAAAAAAGCAGGCTTCA TTGAAGACAAGGGGCCTCGTG-3') and reverse (5'-GGGGACCACTTTGTACAAG AAAGCTGGGTCTTAAAGATAAGGTTCTCCAAAGGTTTTTCG-3') primers. DENND4C cDNA (mouse) was amplified from pcDNA™4/TO-GFP DENND4C using attB site-containing forward (5'-GGGGACAAGTTTGTACAAAAAAGCAGGCTTCA TAGAAGACAAAGGACCTAGAGTGAC-3') and reverse (5'-GGGGACCACTTTGT ACAAGAAAGCTGGGTCTTAAATGAGAGGCGCTCCAAAACAC-3') primers. Polymerase chain reaction (PCR) products were separated by agarose gel electrophoresis and the relevant DENND4A and DENND4C bands excised and purified using the Wizard® SV Gel and PCR Clean-Up System (Promega). attB-flanked DENND4A and DENND4C cDNA fragments were recombined into the pDONR™221 entry vector (Invitrogen) by performing BP reactions following Gateway® Cloning Technology (Invitrogen) guidelines. N-terminal FLAG-tagged DENND4A and DENND4C constructs were generated by performing a Gateway® LR reaction between the pDONR™221- DENND4A or pDONR™221-DENND4C entry vector and a p3XFLAG-CMV™-10 destination vector containing the Gateway® attL recombination cassette (Sigma-Aldrich). The pcDNA™-DEST53-eGFP Gateway® destination vector was created by replacing the cycle 3 GFP (N-terminal) sequence of pcDNA™-DEST53 (Invitrogen) with the eGFP sequence of pEGFP-C1 (Clontech) by enzymatic restriction digest using SnaBI and BspEI restriction endonucleases. Gateway® Cloning Technology was then used to shuttle DENND4A cDNA from the pDONR™221 entry vector into the pcDNA™-DEST53- eGFP destination vector to generate an N-terminal eGFP-tagged DENND4A construct.

2.3.2 Cell culture

2.3.2.1 3T3-L1 and HEK-293E cells 3T3-L1 fibroblasts and HEK293-E cells were maintained in DMEM supplemented with

10% FBS and 2 mM GlutaMAX (DMEM/FBS medium) at 37 °C with 10% CO2 and passaged at ~60% confluence. In the case of 3T3-L1 fibroblasts, confluent cells were differentiated into adipocytes by replacement of the growth medium with differentiation medium (DMEM/FBS medium supplemented with 4 μg/mL insulin, 100 ng/mL dexamethasone, 500 μM isobutyl methylxanthine (IBMX) and 100 ng/mL biotin). 72 h later, the differentiation medium was replaced with post-differentiation medium (DMEM/FBS medium supplemented with 4 μg/mL insulin). 6 days post-differentiation, the medium was replaced with DMEM/FBS medium and adipocytes were used in experiments at between 6-12 days post-differentiation. For serum starvation, cells were washed three times with phosphate-buffered saline (PBS) and incubated in DMEM containing 0.2% BSA and 2 mM GlutaMAX for 2 h, unless stated otherwise.

2.3.2.2 Transient transfection of HEK293-E cells HEK-293E cells were transiently transfected at 60-70% confluence with 10 μg of plasmid DNA per 10 cm culture dish. Unless stated otherwise, all transfections were performed using Lipofectamine 2000 transfection reagent (Invitrogen) according to the manufacturer’s instructions. Briefly, 10 μg DNA was added to 1.25 mL Opti-MEM medium (Gibco) and 25 μL Lipofectamine 2000, mixed and incubated at room temperature (RT) for 20 min. All media was then removed from the culture dish and replaced with 4 mL of fresh DMEM/FBS medium. The transfection mix was added dropwise to cells and the culture dish incubated for 5 h at 37 °C. Subsequently, the transfection mix was removed and replaced with 6 mL DMEM/FBS medium. Cells were returned to 37 °C and used in experiments at 48 h post-transfection.

2.3.3 SDS-polyacrylamide gel electrophoresis (PAGE) and Western blot analysis

All samples for SDS-PAGE were prepared in reducing Laemmli sample buffer (LSB; 50 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, 0.01% Bromophenol Blue, 50 mM DTT)

and heated at 65 °C for 10 min, followed by centrifugation at 16,000 x g for 5 min at RT. Equal amounts of protein (typically 15 μg per lane) were loaded for each sample in a single experiment on self-made SDS-polyacrylamide mini gels (7% acrylamide, unless stated otherwise). Proteins were separated by electrophoresis alongside Precision Plus Kaleidoscope pre-stained protein standards (Bio-Rad) at 120 V for ~75 min and then transferred to PVDF membrane (Millipore) at 120 V for 80 min using Criterion Blotter protein transfer apparatus (Bio-Rad). Membranes were blocked by incubation in 5% skim milk in Tris-buffered saline (TBS; 10 mM Tris-HCl, pH 7.6, 150 mM NaCl) for 1 h at RT with shaking, rinsed twice in TBS containing 0.1% Tween-20 (TBS-T), followed by overnight incubation with primary antibodies (indicated in the text) in 5% BSA in TBS-

T containing 0.02% NaN3 at 4 °C with rotation. The following day, membranes were rinsed three times (15 min each) in TBS-T and incubated with HRP-conjugated secondary antibodies in 5% skim milk in TBS-T for 1 h at RT with shaking. Membranes were then washed three times (15 min each) in TBS-T. Immunoreactive protein bands were detected by SuperSignal West Pico (Thermo Fisher Scientific) or Immobilon (Millipore) chemiluminescent substrate and either exposed to X-ray film (Fujifilm) or imaged using a ChemiDoc MP imaging system (Bio-Rad). In some cases, infrared dye 700- or 800- conjugated secondary antibodies were used, where antibody incubation and washes were performed in the dark. These membranes were imaged in the 700 or 800 nm channel on an Odyssey infrared imaging system (LI-COR). Band intensities were quantified using ImageJ software (McMaster University, Hamilton, ON).

2.3.4 Purification of recombinant GST fusion proteins

2.3.4.1 Bacterial cell culture, cell lysis and binding of GST fusion proteins to glutathione E.coli (BL21) containing the appropriate GST fusion expression vector (indicated in chapter text) was grown in 500 mL LB medium containing 100 μg/mL ampicillin at 37 °C with shaking until the culture reached an optical density at 600 nm (OD600) of 0.6-0.8. GST fusion protein expression was then induced by addition of 1 mM isopropyl thiogalactopyranoside (IPTG) and the culture incubated for a further 3-4 h at 37 °C with shaking. Bacterial pellets were collected by centrifugation at 5,250 x g for 15 min at 4 °C and stored overnight at -20 °C. The following day, bacterial pellets were thawed on ice and resuspended in 20 mL ice-cold PBS containing cOmplete™ protease inhibitor cocktail (Roche). Lysozyme was added to a final concentration of 1 mg/mL and the solution incubated on ice for 30 min. DNAase was then added to a final concentration of 10 μg/mL and the solution sonicated five times for 30 sec on ice using a Sonicator (Heat Systems Ultrasonics) at power setting 4. Triton X-100 was then added to a final concentration of 1% and the solution incubated on ice for 30 min to further solubilise proteins. The solution was then centrifuged at 10,000 x g for 30 min at 4 °C to remove insoluble material and the supernatant incubated overnight at 4 °C with rotation with 1.2 mL of an 80% glutathione sepharose slurry (GE Healthcare) that had been pre- equilibrated by washing three times with 20 mL volumes of ice-cold PBS. The following day, the resin was washed three times by centrifugation at 2,000 x g for 2 min at 4 °C followed by resuspension in 20 mL ice-cold PBS.

2.3.4.2 Elution of GST fusion proteins GST fusion proteins were eluted from the glutathione sepharose resin by competition with reduced glutathione. The resin was resuspended in 1 mL elution buffer (50 mM Tris-HCl, pH 8.0 containing 10 mM reduced glutathione) and incubated for 45 min at 4 °C with rotation. The resin was pelleted by centrifugation at 2,000 x g for 2 min at 4 °C and the supernatant retained. This step was repeated to obtain a second eluate from the resin. A further 1 mL of elution buffer was added to the resin and incubated for 12 h at 4 °C with

rotation. The resin was pelleted by centrifugation at 2,000 x g for 2 min at 4 °C and the supernatant retained as a third eluate. Eluates were combined and the purified protein buffer exchanged using an Amicon Ultra Centricon centrifugal filter device (Millipore). Concentrations of purified GST fusion proteins were quantified by BCA (Pierce) following the manufacturer’s protocol. Protein molecular weights and purities were confirmed by SDS-PAGE followed by Coomassie staining (see 2.3.5).

2.3.4.3 Thrombin cleavage to remove proteins from GST fusion For cleavage of proteins from the GST tag, the glutathione sepharose resin was transferred to a disposable 5 mL plastic column (Pierce) and washed with 30 mL PBS at RT. The resin was further washed with 20 mL TBS followed by 20 mL thrombin cleavage buffer

(50 mM Tris-HCl, pH 8.0, 10 mM CaCl2). 10 U thrombin was added to the column in 2 mL thrombin cleavage buffer containing 20 μL xylene cyanol and incubated overnight at RT. Thrombin cleavage buffer was added in small batches to elute the solution containing thrombin and the cleaved protein into a collection tube. Fractions were collected until the xylene cyanol dye was no longer visible in the column. The GST portion of the of the GST fusion protein remained bound to the resin in the column. To remove the thrombin from the solution, the eluate was incubated for 1 h at 4 °C with rotation with 40 μL of a 50% benzamidine sepharose slurry (GE Healthcare) that had been pre-equilibrated by washing three times with 10 mL volumes of ice-cold TBS. The resin was centrifuged at 2,000 x g for 2 min at 4 °C and the supernatant containing the purified protein retained.

2.3.5 Colloidal Coomassie brilliant blue staining

Colloidal Coomassie brilliant blue stain was prepared by first making a solution of 17% ammonium sulfate and 34% methanol in distilled water. Orthophosphoric acid was then added to a final concentration of 3%. Whilst stirring the solution, Brilliant Blue G was added to a final concentration of 0.1% and the solution left to stir overnight at RT. For staining of an SDS-PAGE mini gel, the gel was first rinsed in distilled water and then incubated with 100 mL stain in a covered petri dish overnight at RT with gentle rocking. 39

The gel was de-stained the following day by incubation in 1% acetic acid in distilled water for 3-4 h at RT with gentle rocking. Stained gels were imaged in the 700 nm channel on an Odyssey infrared imaging system (LI-COR).

2.3.6 Immunoprecipitation of FLAG-tagged proteins

HEK293-E cells expressing FLAG-tagged DENND4A (wild-type or phosphomutant species) were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Cells were then transferred to ice, washed twice in ice-cold PBS and harvested in IP buffer (1% IGEPAL CA-630, 10% glycerol, 50 mM Tris-HCl, pH 7.4, 150 mM NaCl) containing cOmplete™ protease inhibitor cocktail (Roche) and phosphatase inhibitors (2 mM sodium orthovanadate, 1 mM sodium pyrophosphate, 1 mM ammonium molybdate and 10 mM sodium fluoride). Cells were lysed by passing through a 22-gauge needle six times, followed by six times through a 27-gauge needle. Lysates were solubilised on ice for 20 min and then centrifuged at 18,000 x g for 20 min at 4 °C to remove insoluble material. The protein concentration of the supernatant was quantified by BCA assay (Pierce) following the manufacturer’s protocol. For each sample, 1.5 mg protein was combined with 1 μL anti-FLAG M2 mouse monoclonal antibody and 40 μL of a 50% Protein G sepharose slurry (GE Healthcare) that had been washed twice in IP buffer, and incubated overnight at 4 °C with rotation. The next day, the resin was washed three times with IP buffer, followed by twice with ice-cold PBS by repeated centrifugation at 2,000 x g for 2 min at 4 °C. Immunoprecipitated proteins were eluted by addition of 50 μL 2X LSB and incubation at 65 °C for 10 min. Samples were then centrifuged at 16,000 x g for 5 min at RT. The supernatant was transferred to a clean tube and stored at -20 °C.

Chapter 3: Insulin-regulated phosphorylation of DENN domain proteins

The supplementary material relating to this chapter is in the attached electronic files. Supplementary Table legends are given in the Appendix.

3.1 Introduction

Phosphorylation, the reversible, covalent attachment of negatively charged phosphoryl 2- (PO3 ) groups to the polar side-chains of specific amino acid residues, is a ubiquitous protein post-translation modification (PTM) that serves as a rapid molecular switch mechanism to regulate protein function in diverse biological processes. The addition and removal of phosphate groups, catalysed by protein kinases and phosphatases, respectively, can significantly alter the hydrophilicity/hydrophobicity of protein surface regions, inducing protein conformational change and/or intermolecular association. As a consequence, phosphorylation is capable of either directly or indirectly modulating the enzymatic activity, protein-protein interactions, subcellular localisation and turnover of target proteins, and thus plays a critical role in intracellular signal transduction. In eukaryotes, the most common site of phosphorylation is the serine hydroxyl group, followed by that of threonine and, rarer, the phenolic hydroxyl of tyrosine (Mann et al., 2002). More than 500 protein kinases are predicted in the human proteome and these are classified into subfamilies according to their sequence similarity and substrate specificity (Manning et al., 2002). Kinase substrate specificity is not only based on the target amino acid residue, but also on the sequences surrounding the modified site (referred to as the consensus phosphorylation motif) (Pearson and Kemp, 1991; Songyang et al., 1994). At present, it is estimated that at least two-thirds of a given cellular proteome is phosphorylated, with over 200,000 phosphosites reported in human cells (Olsen et al., 2010; Sharma et al., 2014; Hornbeck et al., 2015). Almost all insulin-regulated cellular processes are mediated by protein phosphorylation. Downstream of the IR, the serine/threonine kinase, Akt, is a key signalling hub that coordinates the majority of these signalling events. In the context of glucose metabolism, the best-characterised Akt substrate is the Rab-GAP, AS160. Insulin-stimulated, Akt-mediated phosphorylation of

AS160 at five sites (Ser318, Ser570, Ser588, Thr642 and Ser751) inhibits its intrinsic Rab-GAP activity (either directly, or indirectly through its enhanced interaction with 14-3-3 protein isoforms), which, in turn, relieves its inhibition on GLUT4 translocation to the adipocyte PM [see Chapter 1, 1.12 The Rab-GAP, AS160] (Kane et al., 2002; Sano et al., 2003; Ramm et al., 2006; Stöckli et al., 2008). A Rab-GEF involved in GLUT4 traffic and whose function is regulated by phosphorylation downstream of insulin signalling, however, has yet to be described.

Phosphorylation is becoming increasingly recognised as a mechanism to regulate Rab- GEF activity in the literature. During endolysosomal maturation, phosphorylation of the Mon1 subunit of the heterodimeric Ypt7/Rab7 GEF, Mon1-Ccz1, by the type I casein kinase, Yck3, is necessary for release of the complex from vacuoles following Ypt7 activation (Lawrence et al., 2014). Protein kinase D (PKD)-mediated phosphorylation of the Vps9 domain-containing Rab5 GEF, Ras and Rab interactor 1 (RIN1), at Ser292 inhibits epithelial cell migration by modulating RIN1-dependent Abl tyrosine kinase activation at the PM (Ziegler et al., 2011). In the yeast secretory pathway, Sec2, a GEF for the exocytic Sec4/Rab8, is recruited to Golgi-derived secretory vesicles by GTP- bound Ypt32/Rab11 (Ortiz et al., 2002). As secretory vesicles concentrate at exocytic sites, phosphorylation of Sec2 by an as-yet-unidentified kinase inhibits its interaction with Ypt32-GTP, and instead promotes the association of Sec2 with Sec15, a component of the exocyst complex and Sec4 effector (Stalder et al., 2013). This generates a Sec2/Sec4/Sec15 GEF/Rab/effector complex that initiates the tethering of secretory vesicles to the PM in preparation for fusion (Stalder et al., 2013). In parallel, the interaction of Rabin8, the mammalian homologue of Sec2, with its alternate binding partners, Rab11 and EXOC6, the mammalian Sec15, is regulated by nuclear Dbf2-related kinase 2 (NDR2)-mediated phosphorylation at Ser272 (Ultanir et al., 2012; Chiba et al., 2013). This site aligns with the phosphorylated region in Sec2; however, the Sec2 phosphosites, Ser181, Ser186 and Ser188, do not conform to the consensus sequence for phosphorylation by the yeast NDR kinase, Cbk1 (Stalder et al., 2013). In the human fungal pathogen, Candida albicans, recruitment of Sec2 to hyphal tips, the site of Sec4 activation during polarised hyphal growth, is reliant on Sec2 phosphorylation by the cyclin-dependent kinase, Cdc28 (Bishop et al., 2010). It is therefore conceivable that DENN domain-containing Rab-GEFs, too, are regulated by phosphorylation. Notably, one study has reported three insulin-sensitive phosphosites contained within partial Akt 42

consensus motifs (Ser1043, Ser1096 and Ser1321) on DENND4C, the primary Rab10 GEF required for insulin-stimulated GLUT4 translocation in adipocytes (Sano et al., 2011). However, none of these sites was found to regulate DENND4C GEF activity [see Chapter 1, 1.14.8 DENND4A-C] (Sano et al., 2011).

With the advent of the field of mass spectrometry (MS)-based proteomics in systems biology research, the study of protein phosphorylation dynamics is now more accessible than ever before. Indeed, in recent years, phosphoproteomics has emerged as a flourishing branch of proteomics focused entirely on the detection, quantification and characterisation of phosphorylation sites on a global scale (Mann et al., 2002; Macek et al., 2009). Stable isotope labelling by amino acids in cell culture (SILAC), an in vivo labelling method whereby heavy isotopic forms of C, N and H are discriminated by MS to identify and quantify peptides from separate samples, has facilitated the quantitative phosphoproteomic comparison between different experimental conditions (Ong et al., 2002). Using a triple SILAC labelling strategy, Humphrey and colleagues (2013) conducted a comprehensive, quantitative MS-based proteomic screen to delineate the insulin-regulated phosphoproteome in 3T3-L1 adipocytes. This study uncovered 37,248 phosphorylation sites on 5,705 proteins in the adipocyte proteome, 15% of which were regulated by insulin (Humphrey et al., 2013). However, well-studied insulin-sensitive phosphosites/phosphoproteins represent only a small proportion of the reported insulin- regulated phosphoproteome in this analysis, suggesting that key regulatory nodes in the insulin signalling network remain unknown. In light of this, I hypothesise that other DENN domain-containing proteins besides DENND4C are phosphorylated downstream of insulin signalling and, moreover, that the GEF function of these additional DENN phosphoproteins might be regulated by insulin-stimulated phosphorylation.

Herein, I will perform an in silico analysis of one unpublished and two published (Humphrey et al., 2013; Humphrey et al., 2015a) datasets from large-scale phosphoproteomic studies of insulin-treated 3T3-L1 adipocytes (Humphrey et al., 2013), L6 myotubes (Hoffman et al., unpublished) and murine liver (Humphrey et al., 2015a) to identify novel insulin-regulated phosphosites on DENN domain-containing and DENN related proteins in these insulin-responsive cell types and tissue. Further, for those DENN proteins found to be highly phosphorylated with insulin stimulation, I intend to validate

43 the reported phosphosites by label-free quantitative phosphoproteomic analysis of the isolated protein(s).

3.2 Materials and methods

General methods other than those described below are located in Chapter 2.

3.2.1 Phosphoproteomic analysis of insulin-stimulated 3T3-L1 adipocytes (Humphrey et al., 2013)

The following is an adapted excerpt from the methods of Humphrey and colleagues (2013). For the full experimental procedure, please refer to the original journal paper.

3.2.1.1 SILAC 3T3-L1 cell culture and peptide preparation Triple-SILAC labelling (as described by Ong and Mann, 2006) of 3T3-L1 fibroblasts was performed by passaging cells in DMEM containing either L-arginine and L-lysine (light); 13 14 2 13 15 L-arginine- C6 N4 and L-lysine- H4 (medium); or L-arginine- C6 N4 and L-lysine- 13 15 C6 N2 (heavy) to generate three different isotopically-labelled cell populations. DMEM was supplemented with 10% FBS that was dialysed to remove unlabelled amino acids. Cells were cultured for at least five passages to allow sufficient SILAC amino acid incorporation (i.e. > 98%) prior to differentiation into adipocytes as described previously (see Chapter 2, 2.3.2.1). 3T3-L1 adipocytes were used in experiments on days 10-12 of differentiation. For all large-scale MS experiments, three biological replicates were performed, with switching of SILAC labels to account for any possible quantitation bias caused by the labelling process. For time-course experiments, triple-SILAC labelled adipocytes were serum-starved for 3 h prior to administration of insulin (100 nM) for 0.25, 0.5, 1, 2, 5, 10, 20 or 60 min. For the Akt inhibition study, triple-SILAC labelled adipocytes were serum-starved for 3 h, and then pre-treated with 10 μM MK-2206 for 30 min prior to insulin stimulation (100 nM, 20 min). Cells were then transferred to ice, washed twice in ice-cold TBS and harvested in 1 mL ice-cold sucrose buffer (250 mM sucrose, 10 mM HEPES, pH 8.0) containing cOmplete™ EDTA-free protease inhibitor cocktail (Roche) and PhosSTOP phosphatase inhibitor cocktail (Roche). Harvested cell lysates were disrupted using a Dounce glass homogeniser and cleared by centrifugation 44

(700 x g, 10 min). Pooling of SILAC-labelled cells for time-course experiments was performed as described by Humphrey et al. (2013). Proteins were precipitated overnight at -20 °C by the addition of 4 volumes of ice-cold acetone, pelleted by centrifugation (2,500 x g, 5 min), and then resuspended in 8 M urea containing 1% N-octylglucoside, 50 mM ammonium bicarbonate and phosphatase inhibitors. Proteins were reduced for 45 min with 10 mM DTT, alkylated with 55 mM iodoacetamide for 20 min in the dark at RT, and then digested with the endoproteinase, Lys-C, at a 1:150 enzyme:protein ratio for 4 h. Peptides were subsequently diluted to 2 M urea with 50 mM ammonium bicarbonate and digested with trypsin (1:100 enzyme:protein ratio) overnight. Digested peptides were acidified with trifluoroacetic acid (TFA) (0.1% final concentration) and centrifuged (5,000 x g, 30 min) prior to fractionation by strong anion exchange (SAX) (as described by Wiśniewski et al., 2009) for total-proteome analysis, or by strong cation exchange

(SCX) and titanium dioxide (TiO2) chromatography for phosphopeptide analysis (as described by Larsen et al., 2005; Olsen et al., 2006). Eluted peptides were dried in a vacuum concentrator and acidified with TFA (1% final concentration) prior to desalting on C18 stop and go extraction (STAGE) tips (see 3.2.6.4), LC-MS/MS analysis and MS data processing (as described by Humphrey et al., 2013).

3.2.2 Phosphoproteomic analysis of insulin signalling in murine liver (Humphrey et al., 2015a)

The following is an adapted excerpt from the methods of Humphrey and colleagues (2015a). For the full experimental procedure, please refer to the original journal paper.

3.2.2.1 In situ hepatic insulin stimulation Male C57BL/6J mice (10-12 weeks old) were fasted for 18 h prior to anesthetisation and perfusion of the hepatic portal vein with either insulin (1 mU/g of body weight) or vehicle control (PBS) for 5, 10, 15 or 30 sec (early time-course) or 0.5, 1, 2, 3, 4, 6 or 10 min (intermediate time-course). Murine livers were then rapidly excised and snap-frozen in liquid nitrogen. At least six (and up to 10) biological replicates (separate mice) were performed for each time-point.

3.2.2.2 Liver tissue lysis Frozen tissues were powdered using a mortar and pestle on liquid nitrogen, solubilised in Guanidinium chloride (GdmCl) lysis buffer (6 M GdmCl, 100 mM Tris, pH 8.5, 10 mM tris(2-carboxyethyl)phosphine (TCEP), 40 mM chloroacetamide), and heated for 5 min at 95 °C. Tissue lysates were cooled on ice for 15 min, sonicated, heated again (95 °C for 5 min) and then centrifuged for at 3,500 x g for 30 min at 4 °C. An aliquot was removed to a clean tube, diluted 50% with water, and precipitated overnight at -20 °C by the addition of 4 volumes of ice-cold acetone. Precipitated proteins were pelleted by centrifugation (2,000 x g, 15 min, 4 °C), pellets washed twice with ice-cold 80% acetone, and then air-dried at RT. Pellets were resuspended in 2 mL trifluoroethanol (TFE) digestion buffer (10% TFE, 100 mM ammonium bicarbonate) with sonication. Protein concentration was determined by BCA assay and samples diluted to equal concentration in 500 μL TFE digestion buffer.

3.2.2.3 Protein digestion and phosphopeptide enrichment (EasyPhos) Protein samples were digested overnight in 500 μL TFE digestion buffer with trypsin (1:100 protein:enzyme ratio) and LysC with rapid agitation at 37 °C. The following day,

150 μL 3.2 M KCl, 55 μL 150 mM KH2PO4, 800 μL acetonitrile and 95 μL TFA were added to the digested peptides. Peptides were mixed, cleared by centrifugation and transferred to clean 2 mL 96-well deep-well plates (96-DWPs). TiO2 beads were subsequently added to peptides at a 10:1 beads:protein ratio, suspended in 80% acetonitrile/6% TFA, and incubated at 40 °C for 5 min with rapid agitation. Beads were then pelleted by centrifugation (3,500 x g, 1 min) and the supernatant (containing non- phosphopeptides) was aspirated and discarded. Beads were suspended in wash buffer (60% acetonitrile, 1% TFA), transferred to a clean 2 mL 96-DWP, and washed a further four times with 1 mL wash buffer. After the final wash, beads were suspended in 100 μL transfer buffer (80% acetonitrile, 0.5% acetic acid), transferred onto a C8 STAGE tip, and centrifuged for 3–5 min at 500 x g (or until no liquid remained on the STAGE tip). Bound phosphopeptides were eluted with two 30 μL volumes of Elution buffer (40% acetonitrile, 15% ammonium hydroxide (25%, HPLC grade) and collected by centrifugation into clean tubes. Samples were vacuum-concentrated for 15 min at 45 °C and then acidified with 10

μL of 10% TFA prior to STAGE tip desalting, LC-MS/MS analysis and MS data processing as described by Humphrey et al. (2015a).

3.2.3 Phosphoproteomic analysis of insulin-stimulated L6 myotubes (Hoffman et al., unpublished)

The method is essentially that described by Hoffman et al. (2015).

3.2.3.1 SILAC L6 cell culture and peptide preparation Double-SILAC labelling of L6 myoblasts was performed by passaging cells in DMEM 13 15 13 15 containing either L-lysine and L-arginine- C6 N4 (light), or L-lysine- C6 N2 and L- 13 15 arginine- C6 N4 (heavy) to generate two different isotopically-labelled cell populations. DMEM was supplemented with 10% FBS that was dialysed to remove unlabelled amino acids. Cells were cultured for at least five passages to allow sufficient SILAC amino acid incorporation (i.e. > 98%) prior to differentiation into myotubes by reducing the serum content of the culture medium to 2% for 6 days. SILAC labelled myotubes were serum- starved for 2 h prior to insulin stimulation (100 nM, 20 min). Cells were then transferred to ice, washed twice in ice-cold PBS and harvested in 2% SDS lysis buffer containing cOmplete™ EDTA-free protease inhibitor cocktail (Roche) and phosphatase inhibitors (2 mM sodium orthovanadate, 1 mM sodium pyrophosphate, 1 mM ammonium molybdate and 10 mM sodium fluoride). Cells were lysed by passing through an 18-gauge needle ten times. Lysates were then centrifuged at 13,000 x g for 10 min at 4 °C to remove insoluble material. Protein concentrations were determined by BCA assay (Pierce) and equal protein amounts of light and heavy SILAC cell populations were mixed (1:1 ratio) and then trypsinised (as described in 3.2.1.1). Peptides were desalted using C18 SepPak cartridges (Waters) according to the manufacturer’s guidelines prior to fractionation by

SCX and TiO2 chromatography (as described by Larsen et al., 2005; Olsen et al., 2006). Four biological replicates were performed, and SILAC labels were switched between the control (basal) and insulin-treated groups in two replicates to account for any possible quantitation bias caused by the labelling process. LC-MS/MS analysis, MS data processing and statistics were performed as previously described (Hoffman et al. 2015).

3.2.4 In silico analysis of phosphoproteomic datasets

3.2.4.1 Ranked protein abundance of detected proteomes Relative protein amounts in each of the measured proteomes were estimated using the intensity based absolute quantification (iBAQ) algorithm (Schwanhäusser et al., 2011) integrated into the MaxQuant software environment (Cox and Mann, 2008). Briefly, protein intensities (summed peptide intensities) were normalised to the number of theoretically observable peptides (fully tryptic peptides, 6-30 amino acid residues in length). The resulting iBAQ values were then log2‐transformed and used to rank the abundance of detected proteins in each proteome studied [Supplementary Table S1].

3.2.4.2 Hierarchical cluster analysis Hierarchical clustering of phosphosites was performed in Perseus (Max Planck Institute of Biochemistry, Munich) on log2-transformed median values after Z-score normalization of the data, using Euclidean distances. Heat map colours are based on the log2- transformed median fold-change over basal values reported by Humphrey et al. (2013; 2015a) [Supplementary Tables S2 and S3].

3.2.5 DENN domain protein expression in murine tissues

Tissues were excised from the C57BL/6J mouse and homogenised in HES-SDS buffer (20 mM HEPES, pH 7.0, 0.5 mM EGTA, 250 mM sucrose, 2% SDS) containing cOmplete™ protease inhibitor cocktail (Roche) with a Dounce homogeniser, followed by sonication. Tissue lysates were cleared by centrifugation at 13,000 x g for 10 min at RT. The supernatant was retained and protein concentration determined by BCA assay (Pierce). Western blotting was performed as described in Chapter 2, 2.3.3.

3.2.6 Phosphosite mapping of DENN-domain proteins

3.2.6.1 Immunoprecipitation and gel electrophoresis of DENN domain proteins Label-free HEK-293E cells were transiently transfected with either FLAG-tagged DENND4A (human) or DENND4C (mouse) as described in Chapter 2, 2.3.2.2 and experiments were performed 48 h post-transfection. Cells were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). For PI3K inhibition, 100 nM wortmannin was administered 30 min prior to insulin treatment. Harvested cell lysates were subjected to immunoprecipitation using an anti-FLAG antibody as described in Chapter 2, 2.3.6. Four biological replicate experiments were performed in the case of DENND4A, and a single experiment was performed for DENND4C. Equal volumes of immunoprecipitated proteins were loaded on NuPAGE Novex 4-12% Bis-Tris pre-cast polyacrylamide gels (Invitrogen) and separated by electrophoresis alongside Novex Sharp unstained protein standards (Invitrogen). Protein bands were visualised by SYPRO Ruby (Invitrogen) staining [see 3.2.6.2].

3.2.6.2 SYPRO Ruby staining All steps for SYPRO Ruby staining of NuPAGE Novex Bis-Tris gels were carried out at RT with gentle shaking unless stated otherwise. Following electrophoresis, gels were rinsed in distilled water and fixed by two incubations (30 min each) in 100 mL 50% methanol and 7% acetic acid in water. Gels were then placed in 50 mL SYPRO Ruby gel stain and incubated in the dark overnight at 4 °C. The following day, gels were transferred to a clean container, rinsed in distilled water and destained by two incubations (30 min each) in 100 mL 10% methanol and 7% acetic acid in water. Gels were then washed twice in distilled water (5 min each) to remove residual destain solution before imaging on a FLA-5100 fluorescent image analyser (Fujifilm).

3.2.6.3 In-gel protein digestion Post-SYPRO Ruby staining, protein bands of interest were excised from polyacrylamide gels under UV light, transferred to clean LoBind microcentrifuge tubes (Eppendorf) and destained by incubation in 1 mL 50% acetonitrile and 250 mM ammonium bicarbonate,

49 pH 8.0 in water for 30 min at RT with gentle agitation. The solution was then removed and gel slices dehydrated by incubation in 1 mL acetonitrile for 10 min at RT with occasional agitation. All solution was then carefully removed prior to addition of 500 ng modified trypsin (Promega) in 100 mM ammonium bicarbonate, pH 8.0 and overnight incubation at 37 °C. The following day, tryptic digestion was stopped and peptides eluted from gel slices by addition of 100 μL formic acid and incubation at 37 °C for 1 h. Further extraction was performed by the addition of 100 μL acetonitrile and incubation at 37 °C for 1 h. Complete dehydration of gel slices was achieved by the addition of 500 μL acetonitrile and incubation for 10 min at RT. The supernatant containing eluted peptides was then transferred to a clean tube, vacuum-dried and stored at -20 °C. Peptides were thawed and resuspended in 50 μL C18 elution buffer (50% acetonitrile, 0.1% TFA) prior to desalting on C18 STAGE tips [see 3.2.6.4].

3.2.6.4 Peptide desalting using C18 STAGE tips Two stacked C18 membranes (3M Empore) (2-4 μg binding capacity each) were punched out and packed into the end of a 200 μL low-peptide binding (Maxymum recovery) pipette tip (Axygen) using a custom packing device. STAGE tips were equilibrated with 50 μL methanol, followed by 20 μL acetonitrile, 20 μL C18 elution buffer (50% acetonitrile, 0.1% TFA), and 20 μL C18 wash buffer (0.1% TFA in water), without allowing the membranes to dry out at any point. Acidified peptides were then loaded onto equilibrated C18 STAGE tips. Columns were washed with 50 μL C18 wash buffer and eluted with two 10 μL volumes of C18 elution buffer. Eluates were collected in LoBind 96-well microplates (Eppendorf) and vacuum-dried.

3.2.6.5 Quantitative mass spectrometry analysis Desalted peptides were resuspended in MS loading buffer (2% acetonitrile, 0.3% TFA) prior to LC-MS/MS analysis. Peptides were loaded onto a reverse-phase column (20 cm x 75 μM inner diameter) packed in-house with 3 μM C18 ReproSil particles (Dr Maisch GmbH HPLC). An Easy-nLC system was connected to the mass spectrometer using a 1.9-2.3 kV nano-ion spray and peptides were separated with a binary buffer system of

0.5% acetic acid (buffer A) and 80% acetonitrile/0.5% acetic acid (buffer B) using linear gradients of buffer B from 5% to 35% over 130 min at a flow rate of 250 nL/min. Peptides were analysed on a Q-Exactive benchtop Orbitrap mass spectrometer (Thermo Fisher Scientific).

3.2.6.6 Mass spectrometry data processing Raw mass spectrometry data were processed using MaxQuant software version 1.4.0.8 using the default settings with minor changes: Oxidised Methionine (M), Acetylation (Protein N-term) and Phospho (STY) were selected as variable modifications, and Carbamidomethyl of cysteines (C) as a fixed modification. A maximum of 2 missed cleavages was permitted and a minimum peptide length of 7 amino acids. The match between runs (MBR) algorithm was enabled with a matching time window of 1 min to transfer identifications between adjacent fractions. For protein and peptide identification, database searching was performed using the Andromeda search engine integrated into the MaxQuant environment (Cox et al., 2011) against the human (for DENND4A) or mouse (for DENND4C) UniProt FASTA database (July 2013). The protein, peptide, and site false discovery rates (FDRs) were controlled at a maximum of 1%. Label-free quantification was performed in MaxQuant as described by Luber et al. (2010). Phosphopeptide intensities [Supplementary Tables S5 and S6] were normalised to the summed MS1 intensity of the corresponding protein. For DENND4A phosphosites, statistical significance was determined using an unpaired t-test in Microsoft Excel. Orthologous phosphosites across mouse, rat and human species were obtained from the PhosphoSitePlus online database. For each site, the identity of the responsible kinase(s) and 14-3-3-binding capacity were predicted using the Scansite3 web tool at high and medium stringencies, and the NetworKIN 3.0 web tool with a minimum NetworKIN score of 1.5. For kinase prediction of DENND4C phosphosites, NetworKIN 3.0 did not yield any output when a minimum NetworKIN score of 0.0 was applied.

3.3 Results

Part I: Discovery

3.3.1 Rab, Rab-GEF and Rab-GAP protein abundance in insulin target cell types and tissue

MS-based proteomic analysis of cultured murine 3T3-L1 adipocytes (Humphrey et al., 2013), cultured rat skeletal L6 myotubes (Hoffman et al., unpublished) and murine liver tissue (James laboratory, unpublished dataset) identified a total of 7,105, 6,216 and 3,973 proteins in their respective proteomes [Supplementary Table S1]. I studied the Rab, Rab- GEF and Rab-GAP protein expression signature of each proteome to, firstly, investigate whether DENN domain proteins and their putative Rab substrates are expressed in insulin target cell types and tissue, and, secondly, to analyse the relative cellular abundance of Rab proteins and their regulators. To estimate absolute protein amounts in each of the measured proteomes, I used the intensity based absolute quantification (iBAQ) algorithm (Schwanhäusser et al., 2011), which essentially normalises the summed peptide intensities to the number of theoretically observable peptides of the protein. Plotting the calculated iBAQ values produced a typical S-shaped distribution where proteins in each proteome were ranked by abundance [Figs. 3.1-3.3]. DENN domain-containing and DENN-related proteins were present in all proteomes studied [Figs. 3.1b, 3.2b and 3.3b]. The 3T3-L1 adipocyte proteome was found to contain the richest diversity of DENN proteins, with representatives from 6 of the 8 subfamilies of core DENN domain proteins and 5 DENN-related proteins present [Fig. 3.1b]. Furthermore, when a member of one DENN subfamily was present, in all but one case, another protein in the same subfamily was also present [Fig. 3.1b]. The Rab10 GEF, DENND4C, and FAM45A, a DENN- related protein of unknown function, were present in all three proteomes studied, with the latter being the most abundant DENN protein in each [Figs. 3.1b, 3.2b and 3.3b]. In almost all cases, where a Rab-GEF or Rab-GAP was present in a given proteome, its putative Rab substrate was also present and, frequently, proportionally abundant [see Tables 3.1 and 3.2 for an exhaustive list of known Rab-GEFs and Rab-GAPs and their putative Rab substrates]. For example, the Rab-GAP, TBC1D15, is the most abundant Rab-GAP in L6 myotubes [Fig. 3.2c] and murine liver tissue [Fig. 3.3c], and its target, Rab7, is also ranked highly. However, in general, Rabs are more abundant than their

regulators, and this was particularly pronounced in the murine liver proteome [Fig. 3.3]. Notably, amongst the most abundant Rabs and Rab-GAPs in the adipocyte proteome, are those known to regulate GLUT4 traffic and/or previously identified on GSVs, namely, the Rab-GAPs, AS160 and TBC1D13, and Rabs 10, 11b, 14 and 35 [Fig. 3.1a and c].

Figure 3.1. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the 3T3-L1 adipocyte proteome (Figure legend on page 57).

Figure 3.2. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the L6 myotube proteome (Figure legend on page 57).

Figure 3.3. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the murine liver proteome (Figure legend on page 57).

Figure 3.1. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the 3T3-L1 adipocyte proteome (page 54). Absolute abundance of detected 3T3-L1 adipocyte proteome (Humphrey et al., 2013) was estimated using the log2-transformed summed peptide intensities (iBAQ) of each of the 7,105 proteins quantified. Proteins are ranked by abundance with a, Rab; b, Rab-GEF and/or DENN- related; and c, Rab-GAP proteins labelled. All DENN domain-containing and DENN-related proteins, as well as those Rab and Rab-GAP proteins identified on GSVs and/or implicated in GLUT4 traffic in adipocytes are highlighted in green text. Abbreviations: iBAQ, intensity-based absolute quantification.

Figure 3.2. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the L6 myotube proteome (page 55). Absolute abundance of detected L6 myotube proteome (Hoffman et al., unpublished) was estimated using the log2-transformed summed peptide intensities (iBAQ) of each of the 6,216 proteins quantified. Proteins are ranked by abundance with a, Rab; b, Rab-GEF and/or DENN- related; and c, Rab-GAP proteins labelled. All DENN domain-containing and DENN-related proteins, as well as those Rab and Rab-GAP proteins identified on GSVs and/or implicated in GLUT4 traffic in skeletal muscle are highlighted in green text. Abbreviations: iBAQ, intensity-based absolute quantification.

Figure 3.3. Ranked abundance of Rab, Rab-GEF and Rab-GAP proteins in the murine liver proteome (page 56). Absolute abundance of detected murine liver proteome (James laboratory, unpublished dataset) was estimated using the log2-transformed summed peptide intensities (iBAQ) of each of the 3,973 proteins quantified. Proteins are ranked by abundance with a, Rab; b, Rab-GEF and/or DENN-related; and c, Rab-GAP proteins labelled. All DENN domain-containing and DENN- related proteins are highlighted in green text. Abbreviations: iBAQ, intensity-based absolute quantification.

Protein name(s) Rab substrate(s) Reference(s) Allaire et al., 2010; Marat and McPherson, 2010; DENND1A (connecdenn1) Rab35 Yoshimura et al., 2010 DENND1B-L Marat and McPherson, 2010;

(connecdenn2) Rab35 Yoshimura et al., 2010

DENND2A Rab9a/b Yoshimura et al., 2010 DENND2B (ST5; HTS1) Rab9a/b; Rab34; Rab13 Yoshimura et al., 2010; Ioannou et al., 2015

DENND2D Rab9a/b Yoshimura et al., 2010 DENND4A (IrlB; MycPBP) Rab10 Yoshimura et al., 2010 DENND4B Rab10 Yoshimura et al., 2010

DENND4C Rab10 Yoshimura et al., 2010 DENND5A (Rab6IP1) Rab39 Yoshimura et al., 2010

DENND5B (Rab6IP-like) Rab39 Yoshimura et al., 2010 Core DENN domain protein family protein domain DENN Core DENND6A (FAM116A) Rab14 Linford et al., 2012 MTMR5 (SBF1) Rab28 Yoshimura et al., 2010 MTMR13 (SBF2) Rab28 Yoshimura et al., 2010

AVL9 unknown Linford et al., 2012

C9orf72 Rab8a; Rab39b Sellier et al., 2016

FAM45A unknown Linford et al., 2012 related - FLCN Rab35 Nookala et al., 2012

FNIP1/2 unknown DENN LCHN (KIAA1147) unknown

SMCR8 Rab8a; Rab39b Sellier et al., 2016 ALS2 (Alsin) Rab5 Otomo et al., 2003; Topp et al., 2004

GAPEX-5 (GAPVD1; Hunker et al., 2006; Su et al., 2006; Lodhi et al.,

RAP6) Rab31; Rab5 2007 Horiuchi et al., 2007; Delprato and Lambright, Rabex5 (RABGEF1) Rab5; Rab21; Rab17 2007; Mori et al., 2013

RIN1 Rab5 Tall et al., 2001 Vps9 Vps9 domain RIN2 Rab5 Saito et al., 2002 RIN3 Rab5 Kajiho et al., 2003

Hps1-Hps4 (BLOC-3) Rab32; Rab38 Gerondopoulos et al., 2012

Mon1a/b (SAND-1/2)-Ccz1 Rab7 Nordmann et al., 2010

Rab3GAP1-Rab3GAP2 Rab18 Gerondopoulos et al., 2014

GEFs - Rab3IL1 (GRAB) Rab3a; Rab8a/b Luo et al., 2001; Yoshimura et al., 2010 Rab3IP (Rabin3; Rabin8) Rab8a; Rab8b Hattula et al., 2002

Other Rab Other Moya et al., 1993; Burton et al., 1993; Miyazaki RABIF (MSS4) Rab3a et al., 1994; Burton et al., 1994 RIC1-RGP1 Rab6a Siniossoglou et al., 2000; Pusapati et al., 2012

TRAPPC1 (BET5; MUM2)

TRAPPC3 (BET3) Wang et al., 2000; Kim et al., 2006; Cai et al., Rab1 TRAPPC4 2008 (mTrs23; Synbindin)

TRAPPC5 (mTrs31)

TRAPP subunits TRAPP TRAPPC10 Morozova et al., 2006; Yamasaki et al., 2009; (mTrs130; TMEM1) Rab1; Rab11 Levine et al., 2013a

Table 3.1. Rab substrates of mammalian Rab-GEF and DENN domain proteins 58

Protein name(s) Rab substrate(s) Reference(s) Dabbeekeh et al., 2007; Fuchs et al., 2007; EVI5 Rab11; Rab35 Laflamme et al., 2012 EVI5L Rab4a; Rab7; Rab10; Rab23 Itoh et al., 2006; Yoshimura et al., 2007 Rab3GAP1-Rab3GAP2 Rab3a-d Fukui et al., 1997; Nagano et al., 1998 RUTBC1 (SGSM2) Rab32; Rab33b; Rab38 Nottingham et al., 2011; Marubashi et al., 2015 RUTBC3 (SGSM3; CIP85; RABGAP5) Rab5a-c; Rab39b Haas et al., 2005; Fuchs et al., 2007 Rab2a; Rab8a/b; Rab10; TBC1D1 Rab14 Roach et al., 2007 TBC1D2 (Armus; PARIS1) Rab7a Frasa et al., 2010 TBC1D2b unknown TBC1D4 (AS160) Rab2a; Rab8a; Rab10; Rab14 Mîinea et al., 2005 TBC1D5 Rab7 Seaman et al., 2009 TBC1D8b unknown TBC1D9b Rab11a Gallo et al., 2014 Itoh et al., 2006; Hsu et al., 2010; Imai et al., 2011; TBC1D10a (EPI64) Rab8a; Rab 27a; Rab35; Hokanson and Bretscher, 2012 TBC1D10b Rab3a; Rab22a; Rab27a/b; Itoh et al., 2006; Fuchs et al., 2007; Ishibashi et (EPI64B; FLJ13130) Rab31; Rab35 al., 2009; Hsu et al., 2010; Hou et al., 2013 TBC1D11 Rab2; Rab4; Rab6; Rab11; Cuif et al., 1999; Fuchs et al., 2007; (RABGAP1; GAPCenA) Rab36 Kanno et al., 2010 TBC1D12 unknown TBC1D13 Rab35 Davey et al., 2012 TBC1D14 unknown Longatti et al., 2012 TBC1D15 Rab7; Rab11a Zhang et al., 2005 TBC1D16 Rab4a Goueli et al., 2012 Fuchs et al., 2007; Vaibhava et al., 2012; TBC1D17 Rab8; Rab21; Rab35 Chalasani et al., 2014 TBC1D18 (RABGAP1L; FLJ38519) Rab22a; Rab34; Rab39b Itoh et al., 2006; Kanno et al., 2010 Haas et al., 2007; Sklan et al., 2007a; TBC1D20 Rab1; Rab2; Rab18 Sklan et al., 2007b; Handley et al., 2015 TBC1D22a Rab33 Pan et al., 2006 TBC1D22b Rab33 Pan et al., 2006 TBC1D23 unknown TBC1D24 Rab35 Uytterhoeven et al., 2011 Rab2a; Rab13; Rab34; TBC1D25 (OATL1) Rab33b Itoh et al., 2006; Itoh et al., 2011 TBC1D32 (BROMI) GAP inactive Pan et al., 2006; Itoh et al., 2006; Ko et al., 2010 TBCK unknown Rab1; Rab2; Rab3a; Rab5; Lanzetti et al., 2000; Lanzetti et al., 2004; USP6NL (RN-tre) Rab28; Rab43 (Rab41) Haas 2005; Fuchs et al., 2007; Frittoli et al., 2008

Table 3.2. Rab substrates of mammalian Rab-GAP proteins

3.3.2 Insulin-regulated DENN protein phosphorylation in 3T3-L1 adipocytes

A SILAC-based, global phosphoproteomic analysis of insulin-stimulated 3T3-L1 adipocytes (Humphrey et al., 2013) has revealed that 15% of the adipocyte phosphoproteome is regulated by insulin. I analysed the phosphoproteomic data from this study to determine whether any Class I phosphorylation sites (those defined by a localisation probability of ≥ 0.75) on DENN domain proteins [Supplementary Table S2] were present amongst the insulin-responsive phosphosites uncovered during the screen. I identified 78 Class I phosphosites on 12 DENN domain proteins that were either downregulated ≤ 0.67-fold or upregulated ≥ 1.5-fold following insulin treatment of 3T3- L1 adipocytes over a time-course of 0 to 60 min [Fig. 3.4a]. These 12 phosphoproteins were core DENN domain-containing proteins, DENND1A, DENND1B, DENND2A, DENND4A, DENND4C, DENND5A, DENND5B and DENND6A; and DENN-related proteins, AVL9, FLCN, FNIP1 and SMCR8. The Rab10 GEFs, DENND4A and DENND4C, were heavily phosphorylated, containing 16 and 17 insulin-sensitive sites, respectively [Fig. 3.4a]. The most highly upregulated phosphosites were DENND1A Ser536, DENND1B Ser632, DENND4A Ser1282 and DENND4C Ser1240, which displayed between 17- and 38-fold increases in phosphorylation during a time-course of 0 to 20 min post-insulin stimulation [Fig. 3.4b].

Figure 3.4. Insulin-regulated phosphorylation of DENN domain-containing and DENN-related proteins in 3T3-L1 adipocytes (contd. on next page).

Figure 3.4. Insulin-regulated phosphorylation of DENN domain-containing and DENN-related proteins in 3T3-L1 adipocytes a, (previous page) Heat map and hierarchical clustering based on the log2-transformed median fold change over basal (SILAC ratio) of 78 insulin-regulated Class I phosphosites in DENN domain- containing and DENN-related proteins expressed in 3T3-L1 adipocytes. SILAC-labelled adipocytes were serum-starved prior to insulin stimulation (100 nM) for 0.25, 0.5, 1, 2, 5, 10, 20 or 60 min. Only those sites downregulated ≤ 0.67-fold or ≥ 1.5-fold following insulin treatment are shown and data were normalised using Z-score transformation. Clusters discriminate between downregulated (blue text) and upregulated (red text) phosphosites. Heat map colours are based on the log2-transformed median fold-change over basal (SILAC ratio) values reported by Humphrey et al. (2013) [Supplementary Table S2]. Three biological replicate experiments were performed and only those sites where quantitative data was present in ≥ 50% of time- points are shown. b, Temporal profiles of four highly upregulated phosphosites following insulin stimulation of 3T3-L1 adipocytes. Plotted values indicate the median fold change in insulin- stimulated phosphorylation over basal as determined by SILAC-based MS proteomics (Humphrey et al., 2013). Dashed line indicates baseline phosphorylation (normalised to 1). Abbreviations: FC, fold change; min, minutes; S, serine; T, threonine.

Humphrey and colleagues (2013) proceeded to study the effect of pharmacological inhibition of Akt on the insulin-regulated adipocyte phosphoproteome to determine whether this kinase, a key signalling node activated by insulin, was responsible for phosphorylation of the reported upregulated, insulin-sensitive phosphosites. To this end, 3T3-L1 adipocytes were pre-treated with the potent Akt inhibitor, MK-2206, prior to insulin stimulation for 20 min. I investigated whether any phosphosites on DENN domain proteins upregulated ≥ 1.5-fold following insulin stimulation in this dataset were sensitive to Akt inhibiton. Here, I chose to focus on DENND4A and DENND4C as these related phosphoproteins were clearly highly insulin-sensitive. Moreover, experimental evidence

suggests DENND4C functions in insulin-regulated GLUT4 translocation (Sano et al., 2011), and both DENND4A and DENND4C are GEFs for Rab10, a Rab known to regulate GLUT4 traffic in adipocytes (Sano et al., 2007; Sano et al., 2008; Chen et al., 2012). At 20 min post-insulin stimulation of adipocytes, six phosphosites on each of DENND4A (Ser729, Ser1151, Ser1152, Ser1282, Ser1584 and Ser1586) [Fig 3.5a] and DENND4C (Thr966, Ser971, Ser1087, Ser1240, Ser1274, and Ser1321) [Fig. 3.5b] were upregulated ≥ 1.5- fold. As in Fig. 3.4b, the most insulin-sensitive sites were DENND4A Ser1282 and DENND4C Ser1240 [Fig. 3.5]. Akt inhibition, with MK-2206, had a dampening effect on all twelve phosphosites [Fig. 3.5]. However, only four sites achieved statistical significance: DENND4A Ser1152 and DENND4C Thr966, Ser971 and Ser1321 [Fig. 3.5]. Despite this observation, DENND4C Thr966 and Thr971 do not conform to the Akt consensus phosphorylation motif, RXRXXS/T [Table 3.4] and are therefore unlikely to be Akt phosphorylation sites. Both DENND4A Ser1152 and DENND4C Ser1321, however, display the partial Akt phosphorylation motif, RXXS [Tables 3.3 and 3.4]. Furthermore, Scansite3, a web-based tool for the prediction of kinase-specific phosphorylation sites and phospho-binding motifs, predicts DENND4C Ser1321 to be both an Akt target site (high stringency) and 14-3-3 docking site (medium stringency) [Table 3.4].

Figure 3.5. Effect of Akt inhibition on insulin-sensitive DENND4A and DENND4C phosphorylation sites SILAC-labelled adipocytes were serum-starved prior to insulin stimulation (100 nM, 20 min). Where indicated, the Akt inhibitor, MK-2206 (10 μM), was administered 30 min prior to insulin treatment. Plotted values indicate the log2-transformed median fold change in insulin-stimulated phosphorylation over basal at Class I a, DENND4A and b, (next page) DENND4C phosphosites determined by SILAC-based MS proteomics (Humphrey et al., 2013) [Supplementary Table S2]. Error bars indicate IQR. Three biological replicate experiments were performed and only those sites upregulated ≥ 1.5-fold following insulin stimulation in at least one replicate are shown. Note: DENND4A and DENND4C phosphosites correspond to the Mus musculus proteins, Uniprot accession #E9Q8V6 and #A6H8H2, respectively. Where permitted, statistical analysis employed an unpaired t-test. *, p < 0.05; **, p < 0.01; ***, p < 0.001. Abbreviations: FC, fold-change; MK, MK-2206; S, serine; T, threonine.

Figure 3.5. Effect of Akt inhibition on insulin-sensitive DENND4A and DENND4C phosphorylation sites (contd.) (Figure legend on previous page).

DENND4A phosphosite Kinase/14-3-3 prediction (species, Uniprot accession #)

Sequence window Mouse Rat Human Scansite NetworKIN (mouse) #E9Q8V6 #D4A518 #Q7Z401

GPRVADYFVVAGL Y12 Y12 Y12 MAPK1; LPSKPSSPGSPSM S729 S729 S728 Cdk1; Erk1 MAPK3 Erk1; GSK3; MAPK1; KPSSPGSPSPMFR S732 S732 S731 Cdk1; Cdc2 MAPK3 NESTKGSAECLPT S950 S950 S949 CK1 FRKRHKSDDGSHL S1016 S1016 S1015 Akt CaMKII SRNRNLSGGVLMG S1036 S1036 S1035 Akt; Aurora B CaMKII DSLEKESSDDDTP S1099 S1099 S1098 CK2 SLEKESSDDDTPF S1100 S1100 S1099 CK2 AMPK; PKA; CaMKII; RLQRRNSSFSVKP S1151 S1151 S1151 Aurora A; 14-3-3 Aurora B; LQRRNSSFSVKPS S1152 S1152 S1152 CaMKII PKCβ DEDDNKSVSTPSA S1194 S1194 S1194 CK1 DNKSVSTPSARRN T1197 T1197 T1197 Cdk1 LTSRTPSIDLQRA S1225 S1225 S1225 Akt CaMKII KLTNKKSPTLVKA S1241 S1241 S1240 Aurora A; KACRRSSLPPNSP S1252 S1252 S1251 Akt; PKA; PAK1; 14-3-3 14-3-3 MAPK1; SSLPPNSPRPVRL S1257 S1257 S1256 Cdc2; Cdk5 MAPK3 Akt; PRDRLWSSPAFSP S1282 S1282 S1281 14-3-3 14-3-3 IDVSRASLGSSAS S1405 S1405 S1404 Aurora A FLKSSSSTETMHF S1512 S1512 S1511 LFPMARSISTCGP S1584 S1588 S1587 CaMKII; PMARSISTCGPLD S1586 S1590 S1589 PAK1; 14-3-3 MARSISTCGPLDK T1587 T1591 T1590 LKB1

Table 3.3. Orthologous DENND4A phosphosites and protein kinase prediction Orthologous DENND4A phosphosites across mouse, rat and human species were obtained from the PhosphoSitePlus online database. Protein sequences shown correspond to the Mus musculus protein, Uniprot accession #E9Q8V6, and the modified residue is highlighted in red bold text. For each site, the identity of the responsible kinase(s) and 14-3-3-binding capacity were predicted using the Scansite3 web tool at high (underlined text) and medium stringencies, and the NetworKIN 3.0 web tool with a minimum NetworKIN score of 1.5.

DENND4C phosphosite (species, Uniprot accession #) Rat Scansite Sequence window Mouse Human #F1LTD7 kinase (mouse) #A6H8H2 #Q5VZ89 *#A0A0G2K089 prediction Cdk1; DLKLFDSPQKLKL S720 S719 R485 Cdk5 HAPEEHTPPELTT T938 M986 A704 SKHLQPTPEPQSP T966 T1014 S732 p38 MAPK; PTPEPQSPTEPPA S971 S1019 S737 Erk1 SIVKVPSGLFDTN S987 S1035 S753 FPERSCSFSSESR S1043 S1092 S810 Akt; 14-3-3 Cdc2; LTCPKTSPPHVTR S1087 S1336 S854 Cdk5 PPHVTRTHSFENV T1094 T1143 T861 HVTRTHSFENVNC S1096 S1145 S863 Akt SEDKLFSPVISRN S1221 S1270 S989 SYMNLKSPLGSKS S1240 S1289 S1008 SPLGSKSCSMELH S1246 S1295 S1014 LGSKSCSMELHGE S1248 S1297 S1016 AHPLERSSSLPSD S1273 S1322 S1041 PLERSSSLPSDRG S1274 S1323 S1042 Akt; 14-3-3 STETEKSSPAVSS S1292 S1336 S1060 Cdk1; Cdc2; GRFKPQSPYRAYK S1310 T1354 T1078 Cdk5 YKDRSTSLSALVR S1321 S1365 S1089 Akt; 14-3-3 QSVKMSSVPNSLS S1620 S1614* S1387 SLTRSHSVGGPLQ S1637 S1631* S1404 14-3-3 HGVSTVSLPSSLQ S1659 S1653* S1426 TVSLPSSLQEDVD S1663 S1657* S1430

Table 3.4. Orthologous DENND4C phosphosites and protein kinase prediction Orthologous DENND4C phosphosites across mouse, rat and human species were obtained from the PhosphoSitePlus online database. Protein sequences shown correspond to the Mus musculus protein, Uniprot accession #A6H8H2, and the modified residue is highlighted in red bold text. Red stars indicate sites absent from the short Rattus norvegicus DENND4C isoform, Uniprot accession #F1LTD7 (1601 amino acids), but present in the long isoform, Uniprot accession #A0A0G2K089 (1901 amino acids). For each site, the identity of the responsible kinase(s) and 14-3-3-binding capacity were predicted using the Scansite3 web tool at high (underlined text) and medium stringencies

3.3.3 Insulin-regulated DENN protein phosphorylation in murine liver tissue

More recently, Humphrey et al. (2015a) developed a novel platform for studying phosphoproteome dynamics in vivo, EasyPhos, and used this technology to generate a time-resolved atlas of insulin action in murine liver. This study revealed that approximately 10% of the 25,507 distinct phosphorylation sites that constitute the reported liver phosphoproteome are regulated by insulin (Humphrey et al., 2015a). Furthermore, it was observed that many of the insulin-regulated sites are modified at early (< 30 sec) time-points post-insulin delivery (Humphrey et al., 2015a). I analysed the phosphoproteomic data from the early (5-30 sec) and intermediate (0.5-10 min) time- courses of insulin delivery in murine liver presented in this study to determine whether any Class I sites on DENN domain proteins were present amongst the insulin-responsive phosphosites [Supplementary Table S3]. I identified 26 and 39 Class I phosphosites on a total of 7 DENN domain proteins that were either downregulated ≤ 0.67-fold or upregulated ≥ 1.5-fold in murine liver following in situ insulin administration over the early [Fig. 3.6a] and intermediate [Fig. 3.6b] time-courses, respectively. These 8 phosphoproteins were core DENN domain-containing proteins, DENND1A, DENND1B, DENND4A, DENND4C, DENND5A, DENND5B and DENND6A; and the DENN- related protein, FLCN. Remarkably, these DENN proteins overlapped completely with those found to be phosphorylated following insulin stimulation of 3T3-L1 adipocytes [Fig. 3.4a]. Again, the Rab10 GEFs, DENND4A and DENND4C, were heavily phosphorylated, with a total of 15 and 12 insulin-sensitive sites mapped, respectively [Fig. 3.6]. 11 of these sites, 9 in DENND4A and 2 in DENND4C, were unique to the liver (were not identified in the adipocyte phosphoproteome) [Figs 3.4a and 3.6]. Notably, neither of the most highly insulin-responsive sites in DENND4A and DENND4C in adipocytes (Ser1282 and Ser1240, respectively) [Fig. 3.4b] were identified in liver in either time-course. In general, those sites that were categorised in the down- or upregulated clusters in 3T3-L1 adipocytes [Fig. 3.4a] were present in the same cluster in murine liver [Fig 3.6], a notable exception being DENND4A S1151 and S1152, which were upregulated in adipocytes but downregulated in liver.

Figure 3.6. Insulin-regulated phosphorylation of DENN domain-containing and DENN-related proteins in murine liver tissue (contd. on next page).

Figure 3.6. Insulin-regulated phosphorylation of DENN domain-containing and DENN-related proteins in murine liver tissue (contd. Figure legend on next page).

Figure 3.6. Insulin-regulated phosphorylation of DENN domain-containing and DENN-related proteins in murine liver tissue Heat map and hierarchical clustering based on the log2-transformed median fold change over basal of a, (page 69) 26 and b, (previous page) 39 insulin-regulated Class I phosphosites in DENN domain-containing and DENN-related proteins expressed in murine liver tissue. Fasted mice received intravenous insulin infusion (1 mU/g) for a, 5, 10, 15 or 30 sec; or b, 0.5, 1, 2, 3, 4, 6 or 10 min. Only those sites downregulated ≤ 0.67-fold or ≥ 1.5-fold following insulin treatment are shown and data were normalised using Z-score transformation. Clusters discriminate between downregulated (blue text) and upregulated (red text) phosphosites. Heat map colours are based on the log2-transformed median fold-change over basal values reported by Humphrey et al. (2015a) [Supplementary Table S3]. At least six biological replicates (separate mice) were performed for each time-point. Only those sites where quantitative data was present in ≥ 2 biological replicates at all time-points, or ≥ 3 biological replicates in a, ≥ 3 of 4 time-points; or b, ≥ 6 of 7 time-points are reported. Abbreviations: FC, fold change; min, minutes; sec, seconds; S, serine; T, threonine.

3.3.4 Insulin-regulated DENN protein phosphorylation in L6 myotubes

A SILAC-based, global phosphoproteomic analysis of insulin-stimulated L6 myotubes was recently performed in the James laboratory (Hoffman et al., unpublished). I discovered 30 Class I phosphosites on 10 DENN domain proteins contained within this dataset [Supplementary Table S4] [Fig 3.7]. All DENN phosphoproteins reported (besides DENND2B) overlapped with those identified in adipocytes [Fig. 3.4a] and liver [Fig. 3.6]. Three phosphosites were significantly upregulated following insulin stimulation of cultured rat myotubes for 20 min: DENND4A Ser1036; and DENND4C Thr1014 and Ser1019 (corresponding to Thr966 and Ser971 in the mouse DENND4C protein) [see Tables 3.3 and 3.4 for a list of orthologous DENND4A and DENND4C phosphosites, respectively]. Both DENND4C Thr966 and Ser971 were previously identified as being significantly upregulated following insulin stimulation of 3T3-L1 adipocytes and, furthermore, sensitive to Akt inhibition with MK-2206 [Figs. 3.4a and 3.5a]. DENND4A Ser1036 was not identified in the adipocyte phosphoproteome, but was found to be downregulated in murine liver [Fig. 3.6b]. DENND2B (also known as ST5), which had not been previously been identified as a DENN phosphoprotein, here contained 10 phosphosites, three of which (Ser28, Ser30 and Ser32) were positioned in close proximity to each other and were significantly downregulated with insulin [Fig. 3.7].

Figure 3.7. Insulin-stimulated phosphorylation of DENN domain-containing and DENN-related proteins in L6 myotubes SILAC-labelled L6 myotubes were serum-starved prior to insulin stimulation (100 nM) for 20 min. Plotted values indicate the median fold change in insulin- stimulated phosphorylation over basal at Class I phosphosites determined by SILAC-based MS proteomics (Hoffman et al., unpublished). Error bars indicate IQR. Four biological replicate experiments were performed and only those sites with quantitative data present in ≥ 2 replicates are shown. Dashed lines indicate the 0.67-fold (blue) and 1.5-fold (red) cut-off values for down- and upregulated phosphosites, respectively. Note: DENND4A and DENND4C phosphosites correspond to the Rattus norvegicus proteins, Uniprot accession #D4A518 and #F1LTD7, respectively. Statistical analysis employed an unpaired t-test integrated into the LIMMA package in R statistical computing software. *, p < 0.05; **, p < 0.01; ***, p < 0.001. Abbreviations: FC, fold-change; min, minutes; S, serine; T, threonine.

Part II: Independent validation

3.3.5 DENND4A and DENND4C expression in murine tissues

In Part I, I discovered that DENND4A and DENND4C are insulin-sensitive phosphoproteins in cultured murine adipocytes and rat myoutubes, and in mouse liver tissue. To validate the expression of DENND4A and DENND4C in insulin-responsive tissues, and to examine whether their distribution profiles overlapped with that of Rab10 and GLUT4, a library of lysates from murine tissues was immunoblotted using anti- DENND4A, anti-DENND4C, anti-GLUT4 and anti-Rab10 antibodies [Fig. 3.8]. The Uniprot database lists a single murine DENND4A protein of 209 kDa (accession #E9Q8V6), and two murine DENND4C protein splicoforms of 211 kDa and 216 kDa (accession #A6H8H2-1 and #A6H8H2-2, respectively). In the 200 kDa region, the DENND4A antibody immunolabelled three faint bands, whereas the DENND4C antibody labelled a single strong band [Fig. 3.8]. Immunoblotting experiments performed simultaneously in 3T3-L1 adipocytes revealed that the middle band detected by the anti- DENND4A antibody corresponded to the murine protein [see Chapter 4]. DENND4A was expressed in white and brown adipose tissues, red and white skeletal muscle fibres, spleen and testes; and DENND4C was expressed in epididymal and perirenal white adipose tissue, brown adipose tissue, brain, testes and, to a lesser extent, in spleen. Curiously, DENND4C was not detectable in skeletal muscle, and neither DENND4A nor DENND4C was detectable in murine liver using commercial antibodies. GLUT4 expression was highest in epididymal white adipose tissue, heart, brain and testes. Varying levels of Rab10 were detected in all tissues examined. Notably, where Rab10 was most highly expressed, DENND4C tended to be present also [Fig 3.8].

Figure 3.8. DENND4A and DENND4C expression in murine tissues DENND4A, DENND4C, AS160, GLUT4 and Rab10 expression levels were assessed in a panel of murine tissues by Western blotting. Two replicates were performed and representative immunoblots are shown. Abbreviations: BAT, brown adipose tissue; s/c, subcutaneous; WAT, white adipose tissue; WB, Western blot.

3.3.6 Mapping insulin-sensitive DENND4A and DENND4C phosphosites in HEK-293E cells

Multiple phosphorylation sites on DENND4A and DENND4C have been detected in SILAC-based, global phosphoproteomic analyses of insulin responsive cell types and tissue [see Part I]. To validate the phosphosites reported in these studies, I performed label-free quantitative phosphoproteomic analyses of DENND4A and DENND4C proteins immunoprecipitated from insulin-stimulated HEK-293E cells. In this instance, label-free quantification was an attractive strategy since it simplified experimental workflow and in silico analysis. I had originally intended to overexpress FLAG-tagged DENND4A (human) and DENND4C (mouse) proteins in 3T3-L1 adipocytes using a retroviral expression system; however, the large sizes of DENN protein cDNA (~5.7 kb) exceed the packaging capacity of commercial retroviral vectors. Therefore, I chose instead to overexpress the FLAG-tagged DENN proteins in HEK-293E, a fast-growing, highly-transfectable and insulin-responsive cell line. Using this method, I identified a total of 10 Class I phosphosites on each of DENND4A [Fig. 3.9] and DENND4C [Fig. 3.10] [Supplementary Tables S5 and S6]. Three novel sites in DENND4A and 5 in DENND4C that had not previously been found in the large-scale phosphoproteomic

datasets [Part I] were identified. These were DENND4A Tyr12, Ser728 and Ser1240; and DENND4C Ser720, Thr738, Thr1094, Ser1221 and Ser1246. Two DENND4A sites, Ser1225 and Ser1251 [Fig. 3.9], and four DENND4C sites, Thr1094, Ser1096, Ser1221 and Ser1246/8 [Fig. 3.10] were upregulated ≥ 1.5-fold following 20 min insulin stimulation of HEK-293E cells. Upregulation of DENND4A Ser1251 was statistically significant. In experiments performed with DENND4A, HEK-293E cells were pre-treated with the PI3K inhibitor, wortmannin, prior to insulin stimulation. However, none of the DENND4A phosphosites identified were sensitive to PI3K inhibition [Fig. 3.9].

The PhosphositePlus database lists 22 DENND4A and 52 DENND4C phosphosites with greater than five MS references. In summary, here, from three global quantitative phosphoproteomic datasets and my own label-free quantitative MS experiments, I have identified 26 and 24 Class I insulin-sensitive phosphosites on DENND4A and DENND4C, respectively. The majority of these sites are positioned in the DENN protein C-terminus, downstream of the NLS and lying outside of any functional protein domains [Fig 3.11]. Fig. 3.12 summarises the dynamics of those DENND4A and DENND4C phosphosites upregulated ≥ 1.5-fold at early (0.5-1 min) and intermediate (10-20 min) time-points following insulin treatment across all datasets. Scansite3 and/or NetworKIN web tools predict many of the insulin-responsive sites in DENND4A and DENND4C to be targets of Akt and/or 14-3-3 docking sites [Tables 3.3 and 3.4]. Notably, the most highly insulin- regulated site in DENND4A, Ser1282, is contained within a perfect Akt phosphorylation consensus sequence, and several others (Ser1016, Ser1036, Ser1151, Ser1152, Ser1225, Ser1252, Ser1586) display full or partial Akt consensus motifs [Table 3.3]. Therefore, it will be necessary to investigate in the next chapter whether the Rab10 GEFs, DENND4A and DENND4C, are, like AS160, capable of binding 14-3-3 in an insulin-regulated manner.

Figure 3.9. Insulin-sensitive phosphorylation sites in DENND4A Label-free HEK-293E cells overexpressing FLAG-tagged DENND4A (human) were serum-starved for 2 hours prior to insulin stimulation (100 nM, 20 min). Where indicated, wortmannin (100 nM) was administered 30 min prior to insulin treatment. Harvested cell lysates were subjected to immunopreciptaion using an anti-FLAG antibody and immunoprecipitated proteins were subjected to SDS-PAGE. Protein bands were visualised by SYPRO Ruby staining and DENND4A protein excised for subsequent in-gel tryptic digestion and peptide desalting prior to quantitative LC-MS/MS analysis. Plotted values indicate the log2- transformed median (±range) fold change in insulin-stimulated phosphorylation over basal at Class I DENND4A phosphosites. Four biological replicate experiments were performed and only those sites with quantitative data present in ≥ 2 replicates are shown. Dashed lines indicate the log2-transformed 0.67- fold (blue) and 1.5-fold (red) cut-off values for down- and upregulated phosphosites, respectively. Note: DENND4A phosphosites correspond to the Homo sapiens protein, Uniprot accession #Q7Z401-1. Statistical analysis employed an unpaired t-test. Statistical significance was not achieved at any site. Abbreviations: FC, fold-change; S, serine; Y, tyrosine. 76

Figure 3.10. Insulin-sensitive phosphorylation sites in DENND4C Label-free HEK-293E cells overexpressing FLAG-tagged DENND4C (mouse) were serum-starved for 2 hours prior to insulin stimulation (100 nM, 20 min). Harvested cell lysates were subjected to immunopreciptaion using an anti-FLAG antibody and immunoprecipitated proteins were subjected to SDS-PAGE. Protein bands were visualised by SYPRO Ruby staining and DENND4C protein excised for subsequent in-gel tryptic digestion and peptide desalting prior to quantitative LC-MS/MS analysis. Plotted values indicate the fold change in insulin-stimulated phosphorylation over basal at all DENND4C phosphosites quantified from a single experiment. Dashed lines indicate the 0.67-fold (blue) and 1.5-fold (red) cut-off values for down- and upregulated phosphosites, respectively. Note: DENND4C phosphosites correspond to the Mus musculus protein, Uniprot accession #A6H8H2. It is unclear from the MS localisation probability score whether S1246 or S1248 is the modified site, hence they are represented as a single phosphosite. Abbreviations: FC, fold-change; S, serine; T, threonine.

Figure 3.11. DENND4A and DENND4C phosphorylation sites Schematic diagram of all Class I phosphosites identified in a, DENND4A (26 sites) and b, DENND4C (24 sites) across all MS experiments. Phosphosites listed correspond to the Mus musculus DENND4A and DENND4C proteins (Uniprot accession #E9Q8V6 and #A6H8H2, respectively). Proteins domains other than those shown are omitted for simplicity. Abbreviations: d, downstream DENN motif; DENN, core DENN module; IRSE, interferon-stimulated response element; LD, longin domain; MABP, MVB12-associated β-prism; NLS, nuclear localisation signal; S, serine; T, threonine; Y, tyrosine.

Figure 3.12. Insulin-regulated DENND4A and DENND4C phosphosite dynamics (Figure legend on next page).

Figure 3.12. Insulin-regulated DENND4A and DENND4C phosphosite dynamics (page 79). Schematic diagram of all Class I phosphosites in a, DENND4A and b, DENND4C that are upregulated ≥ 1.5-fold following insulin stimulation of 3T3-L1 adipocytes and/or L6 myoblasts and/or murine liver tissue and/or HEK-293E cells. Site responses ≥ 1.5-fold, but < 5-fold (small- sized phosphorylation symbol); ≥ 5-fold, but < 10-fold (medium symbol); and ≥ 10-fold (large symbol) between 0.5-1 min (01:00 on stopwatch) and 10-20 min (20:00 on stopwatch) post- stimulation are shown. Phosphosites listed correspond to the Mus musculus DENND4A and DENND4C proteins (Uniprot accession #E9Q8V6 and #A6H8H2, respectively). Proteins domains other than those shown are omitted for simplicity. Abbreviations: d, downstream DENN motif; DENN, core DENN module; IRSE, interferon-stimulated response element; LD, longin domain; MABP, MVB12- associated β-prism; NLS, nuclear localisation signal; S, serine; T, threonine.

3.4 Discussion

In this chapter, I have demonstrated that a diverse range of DENN domain-containing and DENN-related proteins are expressed in insulin target cell types and tissue. The proteome coverage depth of murine liver tissue (3,973 proteins) was considerably less than that observed for cultured 3T3-L1 adipocytes and L6 myotubes (7,105 and 6,216 proteins, respectively). This was expected, as starting material quantities are generally lower for harvested tissues than in cell-based studies. As a consequence, fewer Rabs, Rab- GEFs and Rab-GAPs were found in liver and, moreover, DENN domain proteins later identified in the liver phosphoproteomic data were absent from abundance estimates. Any "missing" proteins, if truly present in the liver proteome, however, will be at least less abundant than those already identified. The coverage of such low-abundance liver proteins is likely precluded by the very high abundance of metabolic enzymes in this tissue. Amongst the most abundant Rabs and Rab-GAPs in the 3T3-L1 adipocyte proteome were those involved in GLUT4 traffic and/or are present on GSVs, emphasising the fundamentality of GLUT4 translocation as a cellular process in these cells. Nevertheless, TBC1D1 and Rab13, known regulators of GLUT4 traffic in skeletal muscle (Peck et al., 2009; Sun et al., 2010; Sun et al., 2016), were not identified in the L6 myotube proteome. Again, this indicates that the measured proteomes are incomplete and that "missing" proteins are presumably expressed at very low levels. Rab7-mediated late endosome-lysosome trafficking is clearly a highly active process in insulin-responsive cell types also, as Rab7−TBC1D15 were the most abundant Rab−Rab-GAP pair in all proteomes studied. A general trend observed across proteomes was that Rabs are more

abundant than Rab GEFs and GAPs, which may reflect a higher turnover rate of Rab regulators relative to their substrates.

A distinct subset of DENN domain proteins found in insulin target cell types and tissue are phosphorylated following insulin treatment. The DENN-related protein, FAM45A, was the most abundant DENN protein found in all proteomes studied. Nevertheless, FAM45A did not appear in the phosphoproteomic data and, at present, it is unclear whether the protein functions as a Rab-GEF (Linford et al., 2012). In 3T3-L1 adipocytes, the most highly insulin-sensitive phosphosites were DENND1A Ser536, DENND1B Ser632, DENND4A Ser1282 and DENND4C Ser1240. All of four of these DENN domain proteins were previously identified as phosphoproteins in the toll-like receptor-activated macrophage phosphoproteome (Weintz et al., 2010) and in a global analysis of steady- state phosphorylation in nine murine tissues (Huttlin et al., 2010). However, DENND1A Ser536 was the only site amongst those listed above reported in these studies. DENND1A and DENND1B are both GEFs for Rab35, a PM-localised Rab newly implicated in the regulation of GLUT4 traffic (Marat and McPherson, 2010; Yoshimura et al., 2010; Davey et al., 2012). Notably, following insulin stimulation of adipocytes, dual phosphorylation of DENND1A at S536 and the nearby site, Ser538, by Akt kinase was recently found to regulate the binding of DENND1A to 14-3-3 proteins and, further, enhance its interaction with Rab35 (Kulasekaran et al., 2015). DENND1B and DENND4A have not been previously implicated as insulin-regulated phosphoproteins. One study, however, has described three insulin-sensitive sites on DENND4C in 3T3-L1 adipocytes (Sano et al., 2011) and these sites (Ser1043, Ser1096 and Ser1321) were upregulated by insulin in the adipocyte phosphoproteome studied here, too. Insulin-induced phosphorylation at DENND4C Ser1043 and Ser1096 also occurred in mouse liver, yet only the latter site (Ser1145 in the rat protein) was identified as being insulin-sensitive in L6 myotubes, where it was downregulated. DENND4A Ser1282 was not reported in either insulin-regulated liver or myotube phosphoproteomes. Such cell type/tissue-specific phosphosignatures were observed for several other DENN protein phosphosites, suggesting that insulin may activate discrete sets of protein kinases and phosphatases in target tissues to direct Rab- GEF activity towards distinct cellular processes. Label-free MS analysis failed to identify those sites corresponding to murine DENND4A Ser1282 and DENND4C Ser1240 in insulin- stimulated HEK-293E cells, a possible explanation being that insulin does not regulate 81 phosphorylation at these sites when the proteins are ectopically expressed in this renal cell line.

In addition to Ser1282 (DENND4A) and Ser1240 (DENND4C), more than 20 other insulin- sensitive phosphosites were identified in these two related Rab10 GEFs across the phosphoproteomes studied. MS-based phosphoproteomic analyses have uncovered that multisite phosphorylation such as this is the rule rather than the exception for regulatory proteins. Nearly all DENND4A/C phosphorylation sites are found in the DENN protein C-terminus, downstream of the N-terminal GEF domain. Indeed, regulated multisite phosphorylation has been shown to occur more commonly in disordered/unstructured regions and at protein interaction interfaces, than in functional, catalytic protein domains (Nishi et al., 2011; Tyanova et al., 2013). But why are DENND4A and DENND4C so highly phosphorylated following insulin stimulation? One hypothesis is that phosphorylation of the DENND4A/C C-terminus acts as a regulatory switch mechanism facilitating the association/dissociation of specific protein interaction partners. At least four of the insulin-regulated sites in each of DENND4A and DENND4C are predicted to be 14-3-3 docking sites. Hence, it will be crucial to determine next whether DENND4A and DENND4C are 14-3-3 binding partners, especially given that both DENND1A and the Rab10 GAP, AS160, are known to be regulated by Akt-mediated phosphorylation and 14-3-3-binding downstream of insulin signalling in adipocytes. 14-3-3-binding has also been implicated in the regulation of DENND3 GEF activity towards Rab12 in HeLa cells (Xu et al., 2015). The ULK1 phosphorylation sites in DENND3 that mediate the 14-3-3 interaction, Ser554 and Ser572, however, were not identified in the insulin-regulated phosphoproteomes studied here. Multisite phosphoproteins are typically substrates of several different kinases, which, depending on the cellular context, target both overlapping and discrete sets of phosphorylation sites within the same protein. Since particular combinations of phosphorylated sites may have antagonistic or synergistic effects on protein function, multisite phosphoproteins are often key metabolic signal integrators (Humphrey et al., 2015b). For example, in skeletal muscle, phosphorylation of the Rab-GAP, TBC1D1, at distinct phosphosites by Akt and AMPK kinases enables GLUT4 translocation in response to insulin and exercise, respectively (Roach et al., 2007; Chavez et al., 2008). NetworKIN and/or Scansite3 analysis of DENND4A and DENND4C indeed suggests that these phosphoproteins are the targets of multiple protein kinases, including Akt, CaMKII, and members of the Aurora and MAPK family of 82

kinases. The majority of the putative 14-3-3 docking sites in DENND4A/C, as well as several other insulin-sensitive phosphosites, including DENND4A Ser1282 and the three DENND4C sites previously reported by others (Sano et al., 2011), are predicted to be Akt phosphorylation sites. Further, Akt inhibition, with MK-2206, significantly diminished insulin-stimulated phosphorylation at four of the sites upregulated by insulin in adipocytes (Humphrey et al., 2013). Therefore, it is reasonable to predict that Akt is the dominant kinase responsible for DENND4A/C phosphorylation in insulin target cell types/tissue.

In summary, I have identified two related Rab10 GEFs, DENND4A and DENND4C, as highly insulin-sensitive phosphoproteins in insulin-responsive cell types and tissue. Several DENND4A/4C phosphorylation sites are predicted to be Akt sites and/or docking sites for 14-3-3 interaction, and therefore may be involved in a mode of regulation of the GEF activity of the DENN domain.

Chapter 4: Probing the insulin-regulated binding of 14-3-3 to DENND4A

The supplementary material relating to this chapter is located in the Appendix.

4.1 Introduction

14-3-3s are a family of highly conserved, ubiquitously expressed phosphoprotein-binding proteins that play diverse roles in kinase signalling pathways in eukaryotes (Aitken, 1996). 14-3-3-interacting proteins include regulators of transcription, cell cycle progression, cell division, cytoskeletal dynamics, metabolism and apoptosis, many of which are deregulated in cancer and Type II diabetes mellitus (T2D). Upon ligand binding, 14-3-3s are capable of modulating the conformation, activity, stability, subcellular localisation and protein interactions of their phosphoprotein targets (Mackintosh, 2004). In mammals, seven 14-3-3 isoforms (~30 kDa) are recognised: β, γ, ε, ζ, τ, η, and σ (Toker et al., 1992). 14-3-3α and δ are the phosphorylated forms of β and ζ, respectively (Aitken et al., 1995). Homo- or heterodimers of L-shaped 14-3-3 monomers form a central groove that docks onto tandem-phosphorylated serine and threonine residues in their target proteins (Yaffe et al., 1997; Yaffe, 2002). Muslin et al. (1996) showed that the phospho-form of the phosphoserine-containing motif, RSXpSXP, originally identified in Raf-1 kinase, was important for mediating high-affinity 14-3-3 binding. Two distinct sub-types of this consensus motif, RSX(pS/pT)XP (mode I) and RX(F/Y)XpSXP (mode II), were later described (Yaffe et al., 1997). There is also a third, C-terminal mode III motif, pS/pTX1-

2-COOH (where X is not proline), but it is more rare (Ganguly et al., 2005; Coblitz et al., 2006). Most experimentally validated 14-3-3-binding sites fit the mode I motif, possessing at least one basic residue in the -3 to -5 positions relative to the phosphorylated serine/threonine residue, which overlaps with the sequence preference of basophilic protein kinases of the protein kinase A/protein kinase G/protein kinase C (AGC) and calcium/calmodulin protein kinase (CAMK) subfamilies (Johnson et al., 2010). A proline in the +1 position disfavours 14-3-3 binding (Panni et al., 2011). Hence, proline-directed kinases, such as mitogen-activated protein (MAP) kinases, are thought not to create 14- 3-3 docking sites.

High-throughput (HTP) proteomic screens have identified over 2000 14-3-3-binding proteins in the human proteome (Benzinger et al., 2004; Jin et al., 2004; Meek et al., 2004; Pozuelo Rubio et al., 2004; Nishioka et al., 2012; Collins et al., 2013). Notably, among these, there are several proteins whose interaction with 14-3-3 is regulated by Akt- mediated phosphorylation downstream of insulin signalling (Chen et al., 2011; Collins et al., 2013). For example, phosphorylation of the Akt substrate, AS160, and its subsequent binding to 14-3-3 proteins inhibits AS160 Rab-GAP activity, relieving, in turn, its inhibition on GLUT4 translocation to the PM [see Chapter 1, 1.12 The Rab-GAP, AS160] (Kane et al., 2002; Sano et al., 2003; Ramm et al., 2006; Stöckli et al., 2008). 14-3-3 capture of Akt-phosphorylated Forkhead box O (FoxO) transcription factor family members both masks their nuclear localisation signal (NLS), excluding them from the nucleus, and physically prevents their binding to target DNA sequences in anti-apoptotic genes (Brunet et al., 1999; Brownawell et al., 2001; Cahill et al., 2001; Obsil et al., 2003; Zhao et al., 2004; Silhan et al., 2009). This is one mechanism by which Akt promotes cell survival. Another is through phosphorylation of the 14-3-3-interacting protein, Bcl- 2-associated antagonist of cell death (BAD), a pro-apoptotic member of the BCL-2 family of proteins (Farrow and Brown, 1996). Non-phosphorylated BAD promotes cell death by heterodimerising with the BCL-2 proteins, Bcl-2 and Bcl-XL, at the mitochondrial membrane, thereby obstructing their function as death antagonists (Yang et al., 1995; Kelekar et al., 1997; Ottilie et al., 1997; Zha et al., 1997). 14-3-3 sequesters Akt- phosphorylated BAD to the cytosol, releasing Bcl-2 and Bcl-XL, thus preventing apoptosis (Zha et al., 1996; Datta et al., 1997). Other insulin-regulated 14-3-3-binding proteins include the proline-directed serine/threonine kinase, glycogen synthase kinase 3 (GSK3) (Agarwal-Mawal et al., 2003); the Rheb-GAP and regulator of mTOR signalling, Tuberous sclerosis complex 2 (TSC2) (Liu et al., 2002; Yong et al., 2002; Shumway et al., 2003); proline-rich Akt substrate of 40 kDa (PRAS40), an inhibitory component and substrate of mTOR complex 1 (mTORC1) (Kovacina et al., 2003); the mTORC2 component, Rictor (Treins et al., 2010); Enhancer of mRNA decapping 3 (EDC3), a regulator of mRNA decapping (Larance et al., 2010) and the unconventional myosin motor protein, Myosin-1c (Myo1c) (Yip et al., 2008).

14-3-3 proteins have also been implicated in the regulation of GEF activity (Wang et al., 2002; Chahdi et al., 2008; O’Toole et al., 2011). 14-3-3ζ mediates integrin-induced Rac1 85 activation by recruiting the Rac1 GEF, Tiam1, to β1-integrin complexes at the leading edge of spreading cells, thus initiating cell migration (O’Toole et al., 2011). Protein kinase A (PKA)-dependent phosphorylation of the Pak-interacting exchange factor, β1Pix, a GEF for small GTPases Rac1 and Cdc42, enhances 14-3-3β binding to β1Pix homodimers, thereby inhibiting GTP loading of Rac1 (Chahdi et al., 2008). RIN1, a Ras effector and Vps9 domain-containing GEF for Rab5 on early endosomes, downregulates growth factor signalling by controlling the lysosomal degradation of endocytosed receptor tyrosine kinases (RTKs) (Han and Colicelli, 1995; Tall et al., 2001; Kong et al., 2007). The recruitment of RIN1 to membrane compartments is negatively regulated by protein kinase D (PKD)-mediated Ser351 phosphorylation, which promotes RIN1 association with cytosolic 14-3-3 proteins (Wang et al., 2002). Overexpression of a RIN1 S351A mutant, which exhibits reduced 14-3-3 binding and enhanced membrane localisation, markedly augments degradation of the epidermal growth factor receptor (EGFR) in HeLa cells (Wang et al., 2002; Balaji et al., 2012).

More recently, two publications have described the phosphorylation and 14-3-3 binding affinity of the DENN domain-containing proteins, DENND1A/connecdenn1 and DENND3 (Kulasekaran et al., 2015; Xu et al., 2015). Following insulin stimulation of 3T3-L1 adipocytes, Akt regulates the binding of DENND1A, a GEF for Rab35, to 14-3- 3 by phosphorylating residues Ser536 and Ser538 in the DENND1A C-terminus (Kulasekaran et al., 2015). These two phosphosites were initially reported in a phosphoproteomic study performed by Humphrey et al. (2013) [see Chapter 3]. Inhibition of Akt using the potent allosteric inhibitor, MK-2206, significantly reduces the insulin- regulated binding of DENND1A to 14-3-3, and also perturbs the interaction between DENND1A and its substrate, Rab35 (Kulasekaran et al., 2015). Curiously, the isolated DENN domain of DENND1A is twice as efficient as the full-length protein at catalysing the GTP loading of Rab35 (Kulasekaran et al., 2015). Furthermore, a peptide encompassing Ser536 and Ser538 binds the DENN domain, whereas a phosphomimetic version (S536E/S538E) does not, suggesting that the full-length protein is autoinhibited due to steric hindrance from the non-phosphorylated DENND1A C-terminus (Kulasekaran et al., 2015). The authors propose that the Akt-dependent phosphorylation of Ser536 and Ser538 disrupts this intramolecular interaction, and that consequent 14-3-3 binding to this region stabilises DENND1A in an open conformation, allowing the exposed DENN domain to bind Rab35 (Kulasekaran et al., 2015). Starvation-induced 86

activation of Unc-51-like kinase (ULK), the most upstream kinase in autophagy initiation, stimulates phosphorylation of the Rab12 GEF, DENND3, at Ser554 and Ser572 (Xu et al., 2015). The binding of phosphorylated DENND3 to 14-3-3 enhances its GEF activity towards Rab12, which triggers the membrane trafficking events required for autophagy (Xu et al., 2015). siRNA knockdown of DENND3 abolishes the starvation-induced increase in Rab12 activity (Xu et al., 2015). Rescue of the DENND3 knockdown phenotype can be achieved by reintroduction of wild-type DENND3 or the phosphomimetic mutant, DENND3 S554E/S572E, but not by the phospho-dead S544A/S572A mutant (Xu et al., 2015).

Given that several of the insulin-regulated phosphosites identified in the Rab10 GEFs, DENND4A and DENND4C, are putative 14-3-3 binding sites [see Chapter 3], I hypothesise that these proteins are insulin-regulated 14-3-3-interacting proteins in vivo. Further, it is tempting to speculate that 14-3-3 binding to DENND4 proteins, if verified, may, as for DENND1A and the Rab10 GAP, AS160, be regulated by Akt-mediated phosphorylation. Supporting my hypothesis, both DENND4A and DENND4C have been identified previously as 14-3-3β-binding proteins by affinity capture mass spectrometry of whole-cell extracts from L6 myotubes (Larance et al., 2010) and HEK-293 cells (Collins et al., 2013). However, insulin treatment did not significantly alter the association of either protein with 14-3-3 (Larance et al., 2010). Herein, I intend to probe the 14-3-3 binding capacity of DENND4A and DENND4C in HEK-293E cells and 3T3- L1 adipocytes. Specifically, I aim to determine whether any observed interactions between 14-3-3 and DENND4 proteins are regulated by insulin. Further, where relevant, I will investigate the identity of the kinase(s) responsible for phosphorylating insulin- sensitive sites and characterise which of these phosphosites are responsible for mediating 14-3-3 binding. To this end, I will employ two established methods for assaying 14-3-3 protein interactions, affinity purification ("pull-down") of 14-3-3-binding partners using immobilised 14-3-3β and GST-14-3-3β protein overlay (Far Western blotting) analysis.

4.2 Methods

General methods other than those described below are located in Chapter 2.

4.2.1 Molecular cloning

4.2.1.1 Site-directed mutagenesis (SDM) of DENND4A pDONR™221-DENND4A was used as a template to generate pDONR™221- DENND4A S1281A (4A-1P); S1251A/S1281A (4A-2P); DENND4A S1015A/S1035A/ S1281A/S1251A (4A-4P); and DENND4A S1015A/S1035A/S1281A/S1251A/S1511A/ T1512A (4A-6P) using the QuikChange II XL site-directed mutagenesis kit (Stratagene). Mutagenic primers are listed in Supplementary Table S7. All point mutations were confirmed by DNA sequencing. Gateway® Cloning Technology (Invitrogen) was used to shuttle DENND4A phosphomutant cDNA species from the pDONR™221 entry vector into the p3XFLAG-CMV™-10 destination vector.

4.2.1.2 35-site DENND4A phosphomutant generation A 3717 bp nucleotide sequence spanning the 1819-5536 bp region in the human DENND4A gene (NCBI RefSeq accession XM_005254121) and containing 35 serine/threonine to alanine point mutations at the candidate 14-3-3 binding phosphorylation sites listed in Table 4.1 [page 103] was generated by commercial gene synthesis (GenScript, Piscataway, NJ). This mutant cDNA fragment was then subcloned into pDONR™221-DENND4A by enzymatic restriction digest using AgeI and AleI restriction endonucleases, which cut at positions 1826 bp and 5525 bp in the human DENND4A gene, respectively. Gateway® Cloning Technology (Invitrogen) was then used to shuttle the 35-site DENND4A phosphomutant cDNA from the pDONR™221 entry vector into p3XFLAG-CMV™-10 and pcDNA™-DEST53-eGFP destination vectors.

4.2.2 14-3-3 interactions studies

4.2.2.1 14-3-3β-coupled cyanogen bromide (CNBr)-activated sepharose resin production

E.coli (BCL21) containing the pGEX-GST-14-3-3β plasmid was grown overnight in 5 mL LB medium containing 100 μg/mL ampicillin at 37 °C with shaking. The following morning, this culture was used to inoculate 500 mL LB medium containing 100 μg/mL ampicillin which was incubated at 37 °C with shaking until the culture reached an OD600 of 0.6-0.8. IPTG-mediated protein expression induction, bacterial cell lysis, affinity purification of GST-14-3-3β, and thrombin cleavage of 14-3-3β from the GST tag were performed as described in Chapter 2, 2.3.4.1 and 2.3.4.3. Purified 14-3-3β protein was then buffer exchanged in CNBr coupling buffer (100 mM NaHCO3, 500 mM NaCl) and concentrated to a volume of 2 mL using an Amicon Ultra Centricon centrifugal filter device (Millipore) with a 10 kDa cut-off. The concentration of the eluted 14-3-3β protein was quantified by BCA assay. The average yield was ~12 mg of pure 14-3-3β protein. The molecular weight and purity of the protein were confirmed by SDS-PAGE (15% gel), followed by Coomassie staining [see Chapter 2, 2.3.5]. The 14-3-3β protein was then coupled to CNBr-activated sepharose (GE Healthcare) according to the manufacturer’s instructions at a ratio of 5 mg of 14-3-3β/1 mL of resin. For preservation, NaN3 was added to a final concentration of 0.01% and the resin stored at 4 °C.

4.2.2.2 GST-14-3-3β protein purification For purification of GST-tagged 14-3-3β, all steps described in 4.2.2.1 were performed, except that, instead of thrombin cleavage of the GST tag, the GST-14-3-3β fusion protein was eluted from the glutathione sepharose resin by competition with reduced glutathione as described in Chapter 2, 2.3.4.2. The molecular weight and purity of the fusion protein were confirmed by SDS-PAGE (15% gel) and Coomassie staining [see Chapter 2, 2.3.5]. The concentration of pure GST-14-3-3β was determined by BCA assay and adjusted to 5 mg/mL with 50 mM Tris-HCl, pH 8.0. For preservation, NaN3 was added to a final concentration of 0.01% and the solution stored at 4 °C.

4.2.2.3 14-3-3 pull-down from 3T3-L1 adipocytes and HEK293-E cells DENND4A or DENND4C constructs were expressed in HEK-293E cells by transient transfection as described in Chapter 2, 2.3.2.2. For experiments with 3T3-L1 adipocytes, endogenous DENN domain proteins were studied. Cells were serum-starved for 2 h prior to insulin stimulation (100 nM) for 20 min (unless stated otherwise in the text). For kinase inhibitor studies, adipocytes were administered with either wortmannin (100 nM), MK- 2206 (10 μM), rapamycin (50 nM), Torin-1 (250 nM) or U0126 (10 μM) 30 min prior to insulin treatment. Following insulin stimulation, cells were transferred to ice, washed twice in ice-cold TBS and harvested in 700 μL IP buffer (1% IGEPAL CA-630, 10% glycerol, 50 mM Tris-HCl, pH 7.4, 150 mM NaCl) containing cOmplete™ protease inhibitor cocktail (Roche) and phosphatase inhibitors (2 mM sodium orthovanadate, 1 mM sodium pyrophosphate, 1 mM ammonium molybdate and 10 mM sodium fluoride). Cells were lysed by passing through a 22-gauge needle six times, followed by six times through a 27-gauge needle. Lysates were solubilised on ice for 20 min and then centrifuged at 18,000 x g for 20 min at 4 °C to remove insoluble material. The protein concentration of the supernatant was quantified by BCA assay following the manufacturer’s protocol. For each sample, equal amounts of protein (typically 1-1.5 mg) were combined with 14-3-3β-coupled CNBr-sepharose resin (50 μg 14-3-3β/mg of lysate) that had been washed twice in IP buffer, and then incubated overnight at 4 °C with rotation. The next day, the resin was washed once with IP buffer, once with high-salt (0.5 M NaCl) IP buffer, and then twice again in IP buffer by repeated centrifugation at 2,000 x g for 2 min at 4 °C. The resin was dried by aspiration of the supernatant through a 30- gauge needle, resuspended in 200 μL of ice-cold TBS, and then transferred to a spin column (Pierce) and centrifuged dry at 500 x g for 1 min at 4 °C. 50 μL of 2X LSB (pre- heated to 65 °C) was then added to the spin column and the column incubated at 65 °C for 5 min. The eluate was harvested into a collection tube by centrifugation at 500 x g for 1 min at RT and stored at -20 °C. Western blotting was performed as described previously [see Chapter 2, 2.3.3].

4.2.2.4 14-3-3 overlay assay (far Western analysis) AS160 or DENND4A constructs were expressed in HEK-293E cells by transient transfection as described in Chapter 2, 2.3.2.2. Cells were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). For PI3K and mTORC1 inhibition, wortmannin 90

(100 nM) and rapamycin (50 nM) were administered 30 min prior to insulin treatment, respectively. Harvested cell lysates were subjected to immunoprecipitation (IP) with an anti-FLAG antibody [Chapter 2, 2.3.6] and IP eluates separated by SDS-PAGE as previously described [Chapter 2, 2.3.3]. Following transfer of proteins to PVDF membrane, membranes were blocked by incubation in 5% skim milk in TBS for 1 h at RT with shaking, rinsed twice in TBS-T and then incubated overnight with 5 ug/mL GST-

14-3-3β in 5% BSA in TBS-T containing 0.02% NaN3 at 4 °C with rotation. The next day, membranes were washed three times (15 min each) in TBS-T and incubated with a mouse anti-GST primary antibody in 5% BSA in TBS-T containing 0.02% NaN3 for 1 h at RT with rotation. Membranes were rinsed three times (15 min each) in TBS-T and incubated with an anti-mouse HRP-conjugated secondary antibody in 5% skim milk in TBS-T for 1 h at RT with shaking. Membranes were then washed three times (15 min each) in TBS- T and proteins detected by ECL as described in Chapter 2, 2.3.3.

4.2.2.5 Statistical analyses Statistical analyses were performed using the statistical software package, GraphPad Prism 6 (Graph Pad Software Inc.). Data are presented as the average ±SEM. Comparisons between groups were performed using the appropriate statistical tests as indicated in the figure legends.

4.3 Results

4.3.1 14-3-3 binding to DENND4A is enhanced by insulin

Large-scale proteomic analyses of the 14-3-3β interactome have uncovered DENND4A and DENND4C as 14-3-3 binding partners. To validate these findings, I performed 14-3- 3 pull-down assays in HEK-293E cells overexpressing either eGFP-tagged human DENND4A, eGFP-DENND4C (mouse) or an empty GFP vector (negative control) [Fig. 4.1a and b]. Endogenous EDC3 in HEK-293E was used as positive control for insulin- regulated 14-3-3 binding. Both DENND4A and DENND4C were found to interact with 91

14-3-3. However, whereas 14-3-3 binding to DENND4C was unchanged following insulin treatment, the 14-3-3 capture of DENND4A, as for EDC3, was enhanced by insulin stimulation and reduced to near-basal levels when cells were pre-treated with the PI3K inhibitor, wortmannin. Endogenous DENND4A in 3T3-L1 adipocytes exhibited a similar 14-3-3 interaction profile [Fig. 4.1c]. In these cells, endogenous AS160 was used as the positive control for insulin-regulated 14-3-3 binding. Since phosphorylation of DENND4C in response to insulin did not alter its 14-3-3-binding capacity, the protein was not studied further in this chapter. Rather, subsequent experiments were focused on characterising the insulin-regulated interaction between 14-3-3 and DENND4A.

Figure 4.1. 14-3-3 binding to DENND4A and DENND4C (Figure legend on next page).

Figure 4.1. 14-3-3 binding to DENND4A and DENND4C. The interaction between 14-3-3 and DENND4 proteins was determined by 14-3-3 pull-down assay. a, eGFP-tagged DENND4A or b, DENND4C were overexpressed in HEK-293E cells. c, 14-3-3 pull-down of endogenous DENND4A in 3T3-L1 adipocytes. In all panels (a-c), cells were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Where indicated, wortmannin (100 nM) was administered 30 min prior to insulin treatment. Harvested cell lysates were incubated with 14-3-3-conjugated sepharose beads overnight. Starting material and pull-down eluates were subjected to SDS- PAGE for Western blot analysis with the indicated antibodies. Black arrowhead indicates the endogenous DENND4A protein in 3T3-L1 adipocytes. EDC3 and AS160 are positive controls for insulin-regulated 14-3-3 binding. Three biological replicates were performed for each experiment and representative immunoblots are shown. Abbreviations: EV, empty vector; WB, Western blot.

4.3.2 14-3-3 binding to DENND4A occurs slowly following insulin stimulation

The insulin-regulated phosphoproteome, more than half of which in adipocytes is accounted for by the PI3K-Akt-mTOR signalling axis, is temporally clustered, with Akt substrates phosphorylated rapidly following insulin stimulation, and mTORC1 targets phosphorylated substantially slower (Humphrey et al., 2013). Hence, to probe the identity of the kinase(s) responsible for phosphorylating the insulin-regulated 14-3-3-interacting site(s) in DENND4A, I first investigated the time course of 14-3-3 binding to endogenous DENND4A in 3T3-L1 adipocytes by 14-3-3 pull-down assay [Fig. 4.2]. Again, here, endogenous AS160 was used as the positive control for insulin-regulated 14-3-3 binding. Phosphorylation of Akt on Ser473 occurred almost immediately following insulin stimulation and was sustained over the entire 30 min time course [Fig. 4.2b]. 14-3-3 binding to AS160, one of the best-characterised Akt substrates in adipocytes, mirrored the Akt activation profile. mTORC1, however, was activated more slowly, with its substrate, S6K, exhibiting peak Thr389 phosphorylation at 20 min post-insulin stimulation. Compared to AS160, 14-3-3 binding to DENND4A occurred more slowly [Fig. 4.2a]. Furthermore, the DENND4A−14-3-3 interaction kinetics were similar to that of mTORC1-dependent S6K phosphorylation. This suggests that insulin-regulated 14-3-3 binding to DENND4A is controlled by a slower kinase downstream of Akt, possibly mTORC1 or one of its substrates. Unlike the AGC kinases, Akt and mTORC1, MAPK/ERK kinase (MEK), a member of the MAPK family of proline-directed kinases, is thought not to create 14-3-3 interaction sites. Phosphorylation of the MEK substrates,

extracellular signal-regulated kinases (ERKs), ERK1/2, at Thr202 and Tyr204 (a read-out of MEK activity) was greatest at 5 min post-insulin treatment and slowly diminished thereafter. ERK1/2 phosphorylation kinetics did not exhibit substantial overlap with the 14-3-3 binding profile of either AS160 or DENND4A, indicating that MEK is not responsible for mediating 14-3-3 binding to these proteins.

Figure 4.2. Time course of AS160−14-3-3 and DENND4A−14-3-3 interactions (next page) The interaction between 14-3-3 and endogenous AS160 or DENND4A in 3T3-L1 adipocytes was determined by 14-3-3 pull-down assay. Adipocytes were serum-starved for 2 h prior to insulin stimulation (100 nM) for 0, 0.5, 1, 2, 5, 10, 20 or 30 min. Harvested cell lysates were incubated with 14-3-3-conjugated sepharose beads overnight. Starting material and pull-down eluates were subjected to SDS-PAGE for Western blot analysis with the indicated antibodies. a, Time course of 14-3-3 binding to AS160 and DENND4A. Plotted values indicate the average fold- change in 14-3-3 binding from three biological replicates. Immunoblot densitometries were quantified using ImageJ software (Schneider et al., 2012). Data are normalised to the 20 min insulin condition. Error bars indicate SEM. Statistical analysis employed a paired t-test in GraphPad Prism 6 software. Statistical significance (*, p < 0.05) was achieved at 0.5 and 2 min time-points. b, Immunoblot from a representative experiment. Black arrowheads indicate the endogenous AS160 and DENND4A proteins in 3T3-L1 adipocytes. Abbreviations: FC, fold change; WB, Western blot.

Figure 4.2. Time course of AS160−14-3-3 and DENND4A−14-3-3 interactions (Figure legend on previous page).

4.3.3 The DENND4A−14-3-3 interaction is sensitive to inhibitors of mTORC1

To complement the time course data, I performed further 14-3-3 pull-down experiments using a panel of kinase inhibitors [Fig. 4.3]. The insulin-induced increase in 14-3-3 binding to endogenous AS160 in 3T3-L1 adipocytes was abolished when cells were pre- incubated with either wortmannin or the selective Akt inhibitor, MK-2206 [Fig. 4.3a]. This result validates the findings of a previous study in which the AS160−14-3-3 interaction was shown to be regulated by Akt (Ramm et al., 2006). In adipocytes pre- treated with the PI3K inhibitor, wortmannin, the insulin-regulated binding of 14-3-3 to endogenous DENND4A was perturbed [Fig. 4.3b and c]. Despite this, inhibition of Akt, with MK-2206, had inconsistent effects on the DENND4A−14-3-3 interaction in the same cells [Fig. 4.3b]. Inhibition of mTORC1, with either Torin-1 or rapamycin, however, reproducibly inhibited 14-3-3 binding to DENND4A following insulin stimulation [Fig. 4.3b and c]. MEK inhibition, using U0126, had no effect on the DENND4A−14-3-3 interaction at 5 min post-insulin [Fig. 4.3d]. Together, these data suggest that mTORC1 is the primary kinase responsible for phosphorylating the insulin-sensitive 14-3-3 binding sites in the DENND4A protein.

Figure 4.3. Kinase inhibitor sensitivity of AS160−14-3-3 and DENND4A−14-3-3 interactions (next page). The interaction between 14-3-3 and endogenous AS160 or DENND4A in 3T3-L1 adipocytes in the presence of various kinase inhibitors was determined by 14-3-3 pull-down assay. Adipocytes were serum-starved for 2 h prior to insulin stimulation (100 nM) for a-c, 20 min or d, 5 min. Where indicated, either wortmannin (100 nM), MK-2206 (10 μM), rapamycin (50 nM), Torin-1 (250 nM) or U0126 (10 μM) was administered 30 min prior to insulin treatment. Harvested cell lysates were incubated with 14-3-3-conjugated sepharose beads overnight. Starting material and pull-down eluates were subjected to SDS-PAGE for Western blot analysis with the indicated antibodies. a, Immunoblot showing kinase inhibitor sensitivity of AS160−14- 3-3 interaction. b, Kinase inhibitor sensitivity of DENND4A−14-3-3 interaction. Plotted values indicate the average fold-change in 14-3-3 binding from three biological replicates. Immunoblot densitometries were quantified using ImageJ software. Data are normalised to the basal condition. Statistical analysis employed an unpaired t-test in GraphPad Prism 6 software. **, p < 0.01; ***, p < 0.001; ****, p < 0.0001; ns, non-significant. Error bars indicate SEM. c-d, Immunoblots from representative experiments. Black arrowhead indicates the endogenous DENND4A protein in 3T3-L1 adipocytes. Abbreviations: avg., average; Bas, basal; FC, fold change; Ins, insulin; MK, MK-2206; rap, rapamycin; Thr, threonine; tor, Torin-1; Tyr, tyrosine; WB, Western blot; wrt, wortmannin.

Figure 4.3. Kinase inhibitor sensitivity of AS160−14-3-3 and DENND4A−14-3-3 interactions (Figure legend on previous page).

4.3.4 14-3-3 binding to a 6-site DENND4A phosphomutant is comparable to the wild- type protein

The DENND4A protein is phosphorylated at multiple sites in response to insulin [see Chapter 3]. To probe which of the DENND4A phosphosite(s) are responsible for mediating insulin-regulated 14-3-3 interactions, I proceeded to test the 14-3-3-binding capability of various DENND4A phospho-dead mutant species. The precipitation of a protein in a pull-down assay does not necessarily indicate a specific interaction with the bait (in this case, 14-3-3β). That is, there is a possibility that the interaction is mediated by an intervening protein that binds both the bait and the precipitated target protein. Therefore, to eliminate any non-specific interactions, 14-3-3 binding to DENND4A species was determined by GST-14-3-3β overlay (Far Western blotting) herein.

Firstly, to verify the performance of the overlay assay and to establish suitable controls, I tested the 14-3-3-binding capacity of FLAG-tagged AS160 immunoprecipitated from HEK-293E cell lysates [Fig. 4.4a]. Insulin stimulation caused a marked enhancement of GST-14-3-3 binding to wild-type AS160, and this binding was reduced to basal levels when cells were pre-incubated with wortmannin. An AS160 mutant in which the insulin- sensitive 14-3-3 phosphorylation site, Ser341, was mutated to alanine (AS160 S341A) exhibited negligible 14-3-3 binding under non-stimulated conditions. Therefore, in subsequent experiments, wild-type AS160 (isolated from insulin-stimulated cells) and AS160 S341A (from non-stimulated cells) were employed as positive and negative controls for GST-14-3-3 binding, respectively.

Figure 4.4. Direct binding of 14-3-3 to AS160 and DENND4A phosphomutants (Figure legend on next page).

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Figure 4.4. Direct binding of 14-3-3 to AS160 and DENND4A phosphomutants (previous page). The ability of wild-type and phosphomutant(s) of AS160 and DENND4A to directly bind 14-3-3 was determined by GST-tagged 14-3-3β overlay assay. a-b, FLAG-tagged wild-type AS160 or AS160 S341A mutant; b-c, wild-type DENND4A (4A-WT); b, DENND4A S1281A (4A-1P); c, DENND4A S1281A/S1251A (4A-2P) or DENND4A S1015A/S1035A/S1281A/S1251A (4A-4P) were overexpressed in HEK-293E cells. In all panels (a-c), cells were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Where indicated, wortmannin (100 nM) was administered 30 min prior to insulin treatment. Harvested cell lysates were subjected to immunopreciptaion using an anti-FLAG antibody. Starting material and immunoprecipitated proteins were subjected to SDS-PAGE for Western blot analysis with the indicated antibodies, and for far-Western blot analysis (immunoprecipitates only) with GST-tagged 14-3-3β and an anti-GST antibody. Two biological replicates were performed for each experiment and representative immunoblots are shown. Abbreviations: ø, non-transfected; far-WB; far-Western blot; IP, immunoprecipitation; WB, Western blot.

The first amino acid residue I chose to mutate in human DENND4A was S1281. Phosphorylation of the corresponding site in the mouse protein (S1282) was upregulated almost 20-fold at 10 min post-insulin stimulation of 3T3-L1 adipocytes [see Chapter 3, Fig. 3.4b]. This is the time-point at which DENND4A exhibits maximal 14-3-3 binding [Figs. 4.2a and b]. Furthermore, S1281 lies within a medium stringency mode I consensus 14-3-3-binding motif (RXXpSXP) [see Chapter 3, Table 3.3]. In spite of this, the binding of GST-14-3-3 to a DENND4A mutant in which the S1281 phosphosite was mutated to alanine (referred to as the 4A-1P mutant) was no less than that observed for the wild-type DENND4A protein in a pilot overlay assay [Fig. 4.4b and Fig. 4.5]. Consequently, I used consecutive rounds of site-directed mutagenesis to generate further DENND4A phosphomutants for testing. These were DENND4A S1251A/S1281A (4A-2P); DENND4A S1015A/S1035A/S1281A/S1251A (4A-4P); and DENND4A S1015A/S1035A/S1281A/S1251A/S1511A/T1512A (4A-6P). S1251 (corresponding to S1252 in the murine DENND4A protein) exhibited robust 1.5 to 2.5-fold increases in phosphorylation following insulin stimulation of 3T3-L1 and HEK-293E cells [Supplementary Tables S2 and S5] and is the only DENND4A phosphosite to be contained within a high stringency mode I consensus 14-3-3-binding motif (RRSpSXP) [see Chapter 3, Table 3.3]. Nevertheless, unexpectedly, the binding of GST-14-3-3 to 4A- 2P was comparable to that observed for the wild-type protein [Fig. 4.4c]. Further, mutation of S1015 and S1035, both of which lie within the minimum stringency mode I consensus 14-3-3-binding motif, RXXpSXX, and exhibit ~1.3-fold increases in insulin- induced phosphorylation in HEK-293E [see Chapter 3, Table 3.3 and Supplementary 101

Table S5], did not significantly alter the insulin sensitivity of 14-3-3 binding to DENND4A either [Fig. 4.4c and Fig. 4.5]. Phosphorylation at S1511/T1512 was upregulated 12-fold in insulin-stimulated HEK-293E cells [Supplementary Table S5]. It is unclear from the mass spectrometry data whether one or both of these residues are modified. Despite the highly insulin-responsive nature of the site(s) in this region, 14-3- 3 binding to 4A-6P, as assessed by 14-3-3 pull-down and GST-14-3-3 overlay methods, although reduced overall relative to the wild-type protein, was still enhanced by insulin [Fig. 4.5]. This indicated that the insulin-regulated 14-3-3-binding site(s) in the DENND4A protein remained unidentified.

Figure 4.5. 14-3-3 binding to multi-site DENND4A phosphomutants Indirect and direct interactions between 14-3-3 and multi-site DENND4A phosphomutants were determined by 14-3-3 pull-down and GST-14-3-3β overlay assays, respectively. FLAG-tagged wild-type DENND4A, DENND4A S1281A (4A-1P); DENND4A S1015A/S1035A/S1281A/S1251A (4A-4P) or DENND4A S1015A/S1035A/S1281A/S1251A/S1511A/T1512A (4A-6P) were overexpressed in HEK-293E cells. Cells were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Harvested cell lysates were either incubated with 14-3-3-conjugated sepharose beads overnight or subjected to immunopreciptaion using an anti-FLAG antibody. Starting material, pull-down eluates and immunoprecipitated proteins were subjected to SDS- PAGE for Western blot analysis with the indicated antibodies, and for far-Western blot analysis (immunoprecipitates only) with GST-tagged 14-3-3β and an anti-GST antibody. Three biological replicates were performed and a representative immunoblot is shown. Abbreviations: ø, non- transfected; far-WB; far-Western blot; IP, immunoprecipitation; WB, Western blot.

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4.3.5 Insulin-enhanced 14-3-3 binding to DENND4A is abolished in a 35-site phosphomutant

Since the mutation of several top candidate 14-3-3 binding phosphosites in the DENND4A protein had little to no effect on the insulin-regulated binding of 14-3-3 to DENND4A, I next adopted a broader approach for interrogating the DENND4A−14-3-3 interaction, in which I systematically analysed the entire catalogue of DENND4A modification sites and selected all candidate 14-3-3 binding sites for subsequent mutation. More than 60 putative serine/threonine phosphorylation sites in the human DENND4A protein are described in the PhosphositePlus database. A sub-set of these sites was selected for mutation by application of scrupulous inclusion and exclusion criteria. Only those sites in DENND4A which were identified by Humphrey et al. (2013) and/or myself [see Chapter 3], and/or sites contained within a consensus motif for 14-3-3 binding (minimum stringency) were selected. All proline-directed sites, that is (pS/T)P, and all sites that were not conserved between mouse and human proteins were eliminated. Additionally, only those sites positioned outside of the DENND4A DENN domain were included, as this region, which exhibits considerable sequence homology with the DENN domain of the other DENND4A subfamily members, is thought not to be a regulatory portion of the protein.

Using this strategy, 35 sites spanning the DENND4A C-terminus were selected [Table 4.1] and a 35-site DENND4A phosphomutant in which all of these sites were mutated to a phospho-dead alanine (4A-35P) was generated by a combination of commercial gene synthesis and in-house molecular cloning [Fig. 4.6].

Figure 4.6. Schematic diagram of 35-site DENND4A phosphomutant (4A-35P) The human DENND4A protein (Uniprot accession #Q7Z401) was mutated at the indicated 35 phosphosites to generate a 4A-35P phosphomutant. Abbreviations: CalB, calmodulin-binding domain; d, downstream DENN motif; DENN, core DENN module; IRSE, interferon-stimulated response element; LD, longin domain; MABP, MVB12-associated β-prism; NLS, nuclear localisation signal; PPRs, pentatricopeptide repeats; S, serine; T, threonine. Note: Diagram is not to scale.

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DENND4A phosphosite Sequence window (Human, UniProt accession #Q7Z401) S728 LPSKSSSPNSPLP S731 KSSSPNSPLPMFR S755 IAKRYSSIPQMWS S866 WPSRSRSGYFLWT T921 EVRRGDTSTEDIQ S922 VRRGDTSTEDIQE T923 RRGDTSTEDIQEE S1015 FRKRHKSDNETNL S1035 RNRNLSGGVLMG T1079 KSTRPNTLDIGKP S1093 LRSKRDSLEKESS S1151 RIQRMNSSFSVKP S1152 IQRMNSSFSVKPF S1154 RMNSSFSVKPFE S1196 DDSKSISTPSARR S1225 LTSRTPSIDLQRA S1240 DKLNKKSPPLVKA S1251 KACRRSSLPPNSP S1256 SSLPPNSPKPVRL S1267 RLTKSKSYTKSEE T1269 TKSKSYTKSEEKP S1281 PRDRLWSSPAFSP S1282 RDRLWSSPAFSPT T1288 SPAFSPTCPFREE S1302 QDTLTHSSPSFNL S1339 KASKWYSRFTMYT T1427 PGKSEVTSSFNAS T1455 SCSRCRTCDCLVH S1508 GRYFLKSSPSTEN S1511 FLKSSPSTENMHF T1512 LKSSPSTENMHFP T1574 PANRSKTAMSKCP S1587 IFPMARSISTSGP S1589 PMARSISTSGPLD S1798 HFKRQRSLYREIL

Table 4.1. Candidate 14-3-3-binding sites spanning the DENND4A C-terminus Thirty-five candidate 14-3-3 binding sites in the human DENND4A (UniProt accession #Q7Z401) C-terminus (listed) were selected for mutation from the PhosphoSitePlus online database using the selection criteria outlined in the text.

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I proceeded to investigate the 14-3-3-binding capacity of the 35-site mutant by performing 14-3-3 pull-down [Fig. 4.7] and GST-14-3-3β overlay [Fig. 4.8] assays in HEK-293E cells overexpressing either wild-type DENND4A or the 4A-35P mutant. In these cells, insulin stimulated phosphorylation of Akt at Ser473, mTORC1 at Ser2448, and mTORC1 substrates, S6K and ULK1, at Thr389 and Ser757, respectively. All sites besides Akt Ser473 were sensitive to treatment with the mTORC1 inhibitor, rapamycin. Whereas, the binding of 14-3-3 to wild-type DENND4A was enhanced with insulin and reduced to basal levels when cells were pre-incubated with rapamycin, the precipitation of 4A-35P with 14-3-3 was negligible across all conditions in the pull-down assay [Fig. 4.7]. Similarly, GST-14-3-3 did not bind to immunoprecipitated 4A-35P [Fig. 4.8].These results confirm that all DENND4A phosphorylation sites responsible for insulin- enhanced 14-3-3 binding are absent in the 35-site mutant. Hence, the 4A-35P construct will serve as a valuable tool for probing the biological function of the DENND4A−14-3- 3 interaction in subsequent studies. Curiously, in HEK-293E cells overexpressing DENND4A species, the phosphorylation of the mTORC1 substrates, S6K and ULK1, was slightly blunted relative to non-transfected cells [Figs. 4.7 and 4.8]. However, these effects were not reproducible across biological replicates and pursuing their significance is beyond the scope of this chapter.

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Figure 4.7. 14-3-3 binding to DENND4A 35-site phosphmutant The interaction between 14-3-3 and a 35-site DENND4A phosphomutant (4A-35P) was determined by 14-3-3 pull-down assay. eGFP-tagged wild-type DENND4A (4A-WT) or 4A-35P were overexpressed in HEK-293E cells. Cells were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Where indicated, rapamycin (50 nM) was administered 30 min prior to insulin treatment. Harvested cell lysates were incubated with 14-3-3-conjugated sepharose beads overnight. Starting material and pull-down eluates were subjected to SDS- PAGE for Western blot analysis with the indicated antibodies. Black arrowheads indicate endogenous and overexpressed DENND4A species. AS160 is a positive control for insulin- regulated, rapamycin-insensitive 14-3-3 binding. Three biological replicates were performed and a representative immunoblot is shown. Abbreviations: ø, non-transfected; WB, Western blot.

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Figure 4.8. Direct binding of 14-3-3 to DENND4A 35-site phosphomutant The ability of a DENND4A 35-site phosphomutant (4A-35P) to directly bind 14-3-3 was determined by GST-tagged 14-3-3β overlay assay. FLAG-tagged wild-type DENND4A (4A-WT) or 4A-35P were overexpressed in HEK-293E cells. Cells were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Where indicated, rapamycin (50 nM) was administered 30 min prior to insulin treatment. Harvested cell lysates were subjected to immunopreciptaion using an anti-FLAG antibody. Starting material and immunoprecipitated proteins were subjected to SDS- PAGE for Western blot analysis with the indicated antibodies, and for far-Western blot analysis (immunoprecipitates only) with GST-tagged 14-3-3β and an anti-GST antibody. Three biological replicates were performed and a representative immunoblot is shown. Abbreviations: ø, non- transfected; far-WB; far-Western blot; IP, immunoprecipitation; WB, Western blot.

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4.4 Discussion

In this chapter, I have independently identified DENND4A and DENND4C as 14-3-3 binding partners in vitro. Furthermore, I have expanded on the work of others (Larance et al., 2010; Collins et al., 2013) by demonstrating that the DENND4A−14-3-3 interaction, in adipocytes and HEK-293E cells, is regulated by insulin. The binding of 14-3-3 to DENND4C was unchanged following insulin stimulation; however, this was not entirely unexpected given that evidence in the literature suggests the GEF activity of DENND4C is constitutively active (Sadacca et al., 2013). Therefore, a mechanism for regulation of DENND4C GEF activity in response to insulin, for instance, through 14-3- 3 binding, would not be required. Given that the insulin-regulated binding of 14-3-3 to DENND4A was inhibited by the PI3K inhibitor, wortmannin, I initially hypothesised that this interaction was dependent on Akt. However, this was found not to be the case. Rather, multiple lines of evidence indicated that mTORC1 was the primary kinase responsible for phosphorylating the insulin-regulated sites which mediate 14-3-3 binding to DENND4A. Following insulin stimulation, the binding of 14-3-3 to DENND4A occurred much more slowly than to the well-characterised Akt substrate, AS160. The kinetics of 14-3-3 binding to DENND4A notably resembled the temporal profile of mTORC1 activation. Furthermore, when adipocytes were pre-incubated with an mTORC1 inhibitor, either Torin-1 or rapamycin, the insulin-induced binding of 14-3-3 to DENND4A was abolished. Akt is classically positioned upstream of mTORC1, yet inhibition of Akt, with MK-2206, had inconsistent effects on the DENND4A−14-3-3 interaction. My results therefore suggest the existence of an alternative pathway between PI3K and mTORC1, independent of Akt. However, given the irreproducibility of the MK-2206 data, I cannot exclude the possibility that DENND4A is an Akt substrate. It is likely that multiple members of the AGC kinase subfamily phosphorylate DENND4A at sites that have the capacity to bind 14-3-3, with mTORC1 being the dominant kinase in the insulin- stimulated state. Indeed, multiple signalling pathways converge on several 14-3-3- binding sites (Lee et al., 2011), and, naturally, 14-3-3 dimers can engage two tandem sites that are phosphorylated by different kinases. In light of a newly published study demonstrating that ULK-dependent phosphorylation of DENND3 enhances its interaction with 14-3-3 proteins (Xu et al., 2015), it will be interesting to test whether DENND4A is a substrate for ULK, especially considering that ULK activity is modulated downstream of mTORC1. 108

Mapping the insulin-regulated 14-3-3 binding sites in DENND4A proved more challenging than first anticipated. Serine-to-alanine mutation of the most highly insulin- sensitive phosphosite in DENND4A, S1281, which is contained within a mode I consensus 14-3-3-binding motif, did nothing to disrupt the DENND4A−14-3-3 interaction. This result was not discouraging, however, as the S1281 site is predicted to be a substrate for Akt [see Chapter 3, Table 3.3] and I had already demonstrated that Akt was not the primary kinase responsible for mediating the 14-3-3 binding to DENND4A. Consecutive mutation of a further 5 top candidate 14-3-3 -binding sites did not perturb the insulin-regulated binding of 14-3-3 to DENND4A either. Given the large size of the human DENND4A protein (209 kDa), I reasoned that multiple phosphosites, certainly more than 6, were likely capable of mediating 14-3-3 interactions in response to insulin. Only when a set of 35 candidate 14-3-3-binding sites spanning the DENND4A C- terminus were mutated to the phospho-dead alanine was the insulin-enhanced binding of 14-3-3 to DENND4A abolished. It is noteworthy that these 35 sites appear segregated into distinct clusters [Fig. 4.6]. One possibility is that unique clusters mediate the basal and the insulin-regulated 14-3-3 binding to DENND4A. Future work, therefore, could involve using further DENND4A species (for instance, truncated phosphomutants) to tease apart the DENND4A−14-3-3 interaction more rigorously. Further, armed with the new knowledge of the autoinhibitory mode of DENND1A regulation (Kulasekaran et al., 2015), one could investigate whether the isolated DENND4A DENN domain has greater GEF activity than the full-length protein and, if so, assess whether peptides encompassing the distinct phosphosite clusters in the DENND4A C-terminus interact with the DENN domain.

14-3-3 binding to phosphorylated proteins is a well-established mechanism for the regulation of target protein function. Hence, it will be important to next address the functional relevance of the insulin-stimulated binding of 14-3-3 to DENND4A. Specifically, it will be interesting to test whether 14-3-3 binding regulates the GEF activity of DENND4A in response to insulin. The majority of known insulin-regulated 14-3-3 interactions inhibit the function of the target phosphoprotein. However, it was recently reported that 14-3-3 binding to phosphorylated DENND1A and DENND3 enhances their GEF activity towards Rab35 and Rab12, respectively (Kulasekaran et al., 2015; Xu et al., 2015). Therefore, it is possible that insulin-regulated 14-3-3 binding to 109

DENND4A augments its Rab10 GEF activity. Rab10 and its GAP, AS160, are known to play major roles in insulin-stimulated GLUT4 translocation; however, this process is regulated by Akt and is unaffected by rapamycin-mediated mTORC1 inhibition. Therefore, given that 14-3-3 binding to DENND4A downstream of insulin signalling is dependent on mTORC1, but not Akt, this functional attribute is likely to regulate DENND4A GEF activity in an alternative insulin-regulated, Rab10-dependent cellular process. This notion is consistent with evidence in the literature suggesting that DENND4C, not DENND4A, is the primary Rab10 GEF regulating insulin-regulated GLUT4 traffic in adipocytes (Sano et al., 2011).

In summary, I have identified DENND4A as a novel, insulin-regulated 14-3-3 binding protein in adipocytes. Phosphomutant studies have led the generation of a 35-site DENND4A phospho-dead mutant which lacks 14-3-3-binding capacity and which will now serve as a tool for investigating the functional relevance of the insulin-regulated DENND4A−14-3-3 interaction in the next chapter.

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Chapter 5: Exploring the function of DENND4A Rab10-GEF activity in insulin-regulated cellular processes

The supplementary material relating to this chapter is in the attached electronic files. Supplementary Table legends are given in the Appendix.

5.1 Introduction

The small GTPase, Rab10, is distributed between the trans-Golgi network (TGN) and endosomes where it functions in endosomal sorting and polarised membrane trafficking (Babbey et al., 2006; Chen et al., 2006; Glodowski et al., 2007; Schuck et al., 2007; Babbey et al., 2010; Shi et al., 2010; Lerner et al., 2013; Chen et al., 2014a; Deen et al., 2014; Xu et al., 2014; Wang et al., 2016; Isabella and Horne-Badovinac, 2016). In the previous chapter, I demonstrated that the insulin-regulated interaction between the Rab10 GEF, DENND4A, and 14-3-3 is sensitive to rapamycin, indicating that mTORC1 (and/or a kinase activated downstream of mTORC1) is the dominant kinase responsible for DENND4A phosphorylation following insulin stimulation. Rab10 has previously been implicated in insulin-regulated GLUT4 exocytosis in adipocytes [see Chapter 1, 1.10 Rab GTPases in GLUT4 traffic] (Sano et al., 2007; Sano et al., 2008; Chen et al., 2012). Yet, given that direct activation of Akt kinase, positioned upstream of mTORC1, fully recapitulates insulin-stimulated GLUT4 translocation in adipocytes (Ng et al., 2008), it is unlikely that the mTORC1-dependent, and seemingly Akt-independent, binding of 14- 3-3 to DENND4A modulates Rab10 activity within the context of GLUT4 exocytosis. Rather, it is more convincing that this functional attribute is dedicated to an alternate insulin-regulated membrane trafficking event that is downstream of mTORC1, such as autophagy (outlined in the following sections). Importantly, however, there are insulin- regulated phosphorylation sites in DENND4A that display full or partial Akt consensus motifs [see Chapter 3] but do not confer 14-3-3 binding [see Chapter 4]. For example, phosphorylation of DENND4A Ser1282, a predicted Akt site [Chapter 3, Table 3.3], was upregulated almost 20-fold with insulin [Chapter 3, Fig. 3.4b]; yet mutation of this residue to a phospho-dead alanine had no effect on the 14-3-3-binding capacity of DENND4A [Chapter 4, Fig. 4.4b and Fig. 4.5]. Therefore, a role for DENND4A in insulin-regulated GLUT4 traffic cannot be excluded and requires further examination. 111

5.1.1 Autophagy

Autophagy ("self-eating") is an evolutionary conserved catabolic process in which cytoplasmic cargoes, including proteins and organelles, are targeted for lysosomal degradation (de Duve, 1963). In almost all eukaryotic cells, autophagy occurs constitutively at low basal levels to maintain normal cellular homeostasis and integrity. However, autophagy can also be triggered in response to environmental stress stimuli, such as starvation and hypoxia, to promote cell survival (Kroemer et al., 2010). Impaired autophagy is a hallmark of numerous human pathologies, including ageing, cancer, bacterial infection, inflammatory bowel disease and neurodegeneration (Shintani and Klionsky, 2004; Yano and Kurata, 2009). Three modes of autophagy, defined by distinct mechanisms of cargo selection and delivery to lysosomes, have been described: macroautophagy, microautophagy and chaperone-mediated autophagy (Klionsky, 2005). The best-characterised and most prevalent of these is macroautophagy (herein referred to as autophagy). Autophagy begins with the formation of a membranous structure (termed a phagophore or isolation membrane) from a membrane source known as the phagophore assembly site (PAS) (Suzuki et al., 2001; Kim et al., 2002). The phagophore elongates and engulfs cytoplasmic material in a double-membraned vesicle termed an autophagosome. In mammalian cells, autophagosomes fuse with lysosomes to form autolysosomes, in which the autophagosome inner membrane and luminal contents are degraded by lysosomal hydrolases (Klionsky, 2005). In most circumstances, the products of autophagy liberated upon digestion of the autophagosomal cargo are recycled into biosynthetic pathways. However, under starved conditions, these macromolecules become substrates for ATP production (Lum et al., 2005). Genetic screens in yeast have identified over 30 autophagy-related (ATG) genes whose protein products participate directly in the molecular mechanism of autophagy (Xie and Klionsky, 2007). Many of these Atg proteins have known mammalian orthologues.

5.1.2 mTORC1 regulation of autophagy

In the fed state, insulin signalling inhibits cellular autophagy via activation of the nutrient- sensing kinase, mTORC1, a central hub for the integration of diverse environmental and nutritional signals (including growth factors, cellular energy status and amino acids) and

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a master regulator of cell growth (Laplante and Sabatini, 2009). Autophagy initiation in mammals is mediated by a cytosolic protein complex comprising the serine/threonine kinase, UNC51-like kinase 1 or 2 (ULK1/2) (Atg1 in yeast), Atg13, Atg101 and focal adhesion kinase (FAK) family interacting protein of 200 kDa (FIP200) (Ganley et al., 2009; Hosokawa et al., 2009; Mercer et al., 2009). mTORC1 activation disrupts formation of this complex through phosphorylation of Atg13 and ULK1, and thereby inhibits autophagosome biogenesis (Kamada et al., 2000; Jung et al., 2009; Kamada et al., 2010; Kim et al., 2011). Inhibition of mTORC1 during nutrient deprivation or following treatment with rapamycin causes dephosphorylation of Atg13 and ULK1, which, in turn, enhances both complex assembly and ULK1 kinase activity, thus stimulating autophagy induction (Kamada et al., 2000; Ganley et al., 2009; Shang et al., 2011). A second pathway of starvation-induced regulation of autophagy exists through the AMP-activated protein kinase (AMPK), another cellular energy sensor. In response to glucose deprivation (low ATP:ADP ratio), AMPK interacts with, and directly phosphorylates, ULK1 on at least two serine residues to trigger autophagy (Lee et al., 2010; Bach et al., 2011; Egan et al., 2011; Kim et al., 2011). Conversely, when nutrients are plentiful, mTORC1-mediated ULK1 phosphorylation disrupts the AMPK−ULK1 interaction, thus preventing autophagy induction (Egan et al., 2011; Shang et al., 2011). AMPK can also promote autophagy by phosphorylating the Rheb GTPase activating protein, tuberous sclerosis complex 2 (TSC2), and the mTORC1 component, Raptor, leading to mTORC1 inhibition (Inoki et al., 2003; Gwinn et al., 2008).

5.1.3 Autophagic roles of Rab GTPases and DENN domain proteins

The involvement of a DENND4A−Rab10 pathway in autophagy is attractive for several reasons. Firstly, autophagy is a process heavily reliant on regulated membrane trafficking and numerous endosomal Rab GTPases are already known to participate in autophagosome formation and maturation. For example, during starvation-induced autophagy in HEK-293A cells, Rab11 mediates autophagosome formation by regulating vesicular transport from ULK1-positive recycling endosomes (REs) to the nascent phagophore (Longatti et al., 2012). The Rab11 effector and ULK1 interaction partner, TBC1D14, negatively regulates this process in the fed state by tubulation of RE, which 113 disrupts their function and inhibits autophagosome biogenesis (Longatti et al., 2012). Others, however, have suggested that Rab11 participates in autophagosome maturation (Fader et al., 2008; Richards et al., 2011; Szatmári et al., 2014). In Drosophila melanogaster, Rab11 translocates from RE to autophagosomes in response to autophagy induction, and interacts with the mictrotubule-binding protein, Hook, a negative regulator of endosome maturation. By removing Hook from mature late endosomes (LE), Rab11 facilitates endosome−autophagosome fusion (Szatmári et al., 2014). Loss of Rab11 causes accumulation of abnormal autophagosomes and acidic LE (Szatmári et al., 2014). Many studies have described the involvement of the late endosome-associated Rab7 in the maturation of autophagosomes and their fusion with lysosomes (Harrison et al., 2003; Gutierrez et al., 2004; Jäger et al., 2004; Bains et al., 2011; Toyofuku et al., 2015). It was recently shown that Rab7 promotes microtubule plus end-directed transport of endosomes and autophagic vesicles through a novel effector, the kinesin motor adaptor FYVE and coiled-coil domain-containing protein 1 (FYCO1) (Pankiv et al., 2010). FYCO1 is recruited to endosomal or autophagosomal compartments via interactions with Rab7, PIP3 and the autophagosomal marker, microtubule-associated protein 1 light chain 3 alpha (LC3) (Pankiv et al., 2010). siRNA-mediated knockdown of FYCO1 in HeLa cells results in the accumulation of LC3-positive early autophagosomal and phagophoric clusters, suggesting an inhibition of their maturation (Pankiv et al., 2010). In macrophages, Rab8b, a paralogue of Rab10 (Klöpper et al., 2012), facilitates autophagic elimination of Mycobacterium tuberculosis var. bovis BCG by controlling autophagosome maturation via its downstream effector, innate immunity regulator TANK-binding kinase 1 (TBK-1) (Pilli et al., 2012). Rab8b and TBK-1 co-localise in LC3-positive autophagic organelles and knockdown of Rab8b inhibits the conversion of BCG phagosomes into autophagolysosomes following autophagy induction (Pilli et al., 2012). In contrast, active Rab14 prevents the fusion of phagosomes containing live BCG with lysosomes (Kyei et al., 2006). Furthermore, during Salmonella typhimurium infection, the bacterial effector protein, SopB, maintains a phagosome maturation block via activation of host Akt1 and downstream inhibition of the Rab14 GAP, AS160, thereby promoting intracellular survival (Kuiji et al., 2007). This is noteworthy because, in 3T3- L1 adipocytes, AS160 is a GAP for Rab10 (Mîinea et al., 2005). In the nematode worm, Caenorhabditis elegans, dietary restriction stimulates autophagy partly through inhibition of Rab10 (Hansen et al., 2008); however, at present, it is not understood

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whether Rab10 plays a role in autophagy in mammalian cells. Other Rab proteins involved in autophagic processes include Rab1 (Zoppino et al., 2010; Pinar et al., 2013; Wang et al., 2015), Rab5 (Ravikumar et al., 2008; Su et al., 2011; Dou et al., 2013; Chen et al., 2014b), Rab8a (Dupont et al., 2011), Rab9a (Nozawa et al., 2012), Rab12 (Matsui et al., 2014; Xu et al., 2015), Rab21 (Jean et al., 2015), Rab23 (Nozawa et al., 2012), Rab24 (Munafó and Colombo, 2002; Egami et al., 2005), Rab25 (Liu et al., 2012), Rab26 (Binotti et al., 2015), Rab32 (Hirota and Tanaka, 2009) and Rab33b (Fukuda and Itoh, 2008; Itoh et al., 2008; Itoh et al., 2011).

Secondly, there are many key examples of DENN domain and DENN-related proteins with functions related to cellular autophagy. Both DENND3 and its substrate, Rab12, have been shown to interact with LC3 (Behrends et al., 2010; Xu et al., 2015). Further, starvation-induced, ULK1-mediated phosphorylation of DENND3 enhances its GEF activity towards Rab12, promoting Rab12 integration into forming autophagosome membranes from where it facilitates autophagosomal transport (Xu et al., 2015). In another study, Rab12, upon activation by DENND3, indirectly induces autophagy by targeting proton/amino transporter 4 (PAT4) to lysosomes for degradation, leading to downregulation of cellular amino acid transport, thus mTORC1 inhibition (Matsui et al., 2014). Sbf, the fly orthologue of the mammalian DENN domain-containing pseudophosphatase, myotubularin-related protein 13 (MTMR13), functions as a Rab21 GEF in endolysosomal traffic (Jean et al., 2012). In Drosophila melanogaster larval fat bodies and human HeLa cells, starvation stimulates Sbf/MTMR13 GEF activity towards Rab21 to upregulate endolysosomal flux of Vamp7/VAMP8, a Rab21 effector required for SNARE-mediated autophagosome-lysosome fusion, to accommodate enhanced levels of autophagy (Jean et al., 2015). siRNA-mediated depletion of Sbf or Rab21 causes autophagosomes to accumulate in the cytoplasm of larval fat bodies, consistent with a defect in starvation-induced autophagy at the level of autophagosome clearance (Jean et al., 2015).

The DENN-related protein, folliculin (FLCN), considered a tumour suppressor, is becoming well established as an autophagy regulator in the literature. Mutations in FLCN are responsible for Birt-Hogg-Dubé (BHD) syndrome, an autosomal dominant disorder characterised by renal cell carcinoma, cysts and benign skin tumours (Nickerson et al.,

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2002). Luijten et al. (2013) find FLCN localised to primary cilia in human renal proximal tubule (HK-2) cells and propose that BHD is a ciliopathy, that is, its symptoms are partly due to abnormal ciliogenesis. Curiously, cilia formation is compromised or lost in multiple cancer types (Cao and Zhong, 2016) and two landmark studies have uncovered a functional relationship between autophagy and ciliogenesis (Pampliega et al., 2013; Tang et al., 2013). In the wake of these findings, Dunlop and colleagues (2014) present evidence that FLCN is a positive regulator of autophagy and, further, demonstrate that impaired autophagy is indeed a hallmark of BHD syndrome, as indicated by the elevated levels of LC3, GABA(A) receptor-associated protein (GABARAP) and sequestome 1 (SQSTM1) proteins, established autophagy markers, expressed in tumour tissue harvested from BHD patients. Knockdown of FLCN increases SQSTM1 expression in HK-2 cells, together with reduced autophagosome maturation, whereas re-expression of FLCN restores normal basal autophagic flux (Dunlop et al., 2014). FLCN communicates with the autophagic machinery through association with GABARAP, and the FLCN−GABARAP interaction is enhanced when FLCN is complexed with either folliculin-interacting protein 1 or 2 (FNIP1/2) (Dunlop et al., 2014). Further, ULK1 negatively regulates the GABARAP binding capacity of FLCN, partly through phosphorylation of three serine residues (Ser406, Ser537, and Ser542) located in the DENN- like FLCN C-terminus (Dunlop et al., 2014). Notably, the most common FLCN mutations in BHD patients generate a truncated FLCN protein that lacks the C-terminal half (Schmidt et al., 2005). In this study, the C-terminal FLCN truncation mutants, Y463X and H429X, exhibit reduced binding to GABARAP, yet enhanced ULK1 interaction, which could explain the dysregulation of autophagy observed in BHD-associated renal tumours (Dunlop et al., 2014). Supporting a role for FLCN in autophagy, FLCN is recruited to lysosomal membranes under nutrient stress or in response to Torin1-mediated mTORC1 inhibition (Martina et al., 2014) and, moreover, modulates mTORC1 kinase activity through association with Rag GTPase heterodimers (Petit et al., 2013; Tsun et al., 2013) [see Chapter 1, 1.14.2 Folliculin]. FLCN has also been shown to bind to AMPK in a phosphorylation-sensitive manner (Piao et al., 2009; Wang et al., 2010; Baba et al., 2006; Possik et al., 2014; Yan et al., 2014). However, in C. elegans and mouse embryonic fibroblasts (MEFs), loss of FLCN causes constitutive activation of AMPK, which induces autophagy (Possik et al., 2014; Yan et al., 2014). These data are in contrast to the findings of Dunlop et al. (2014) where FLCN is found to positively regulate autophagy. Further,

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siRNA-mediated FLCN depletion dampens AMPK signalling in human bronchial and murine alveolar epithelial cells, but has no effect on AMPK in human small airway epithelial cells (Goncharova et al., 2014; Khabibullin et al., 2014). It is therefore apparent that FLCN function is highly cell type-specific and hence the precise role of FLCN in autophagy remains unclear.

5.1.4 Rab7 regulation of lipophagy

Another recent study (Schroeder et al., 2015) has implicated Rab7, the most abundant Rab GTPase in insulin responsive cell types and tissues [see Chapter 3, Figs. 3.1-3.3], as a key regulator of lipophagy. Lipophagy is the autophagic degradation of lipid droplets (LDs), dynamic lipid storage organelles, by lysosomal acid lipases to mobilise fatty acids for mitochondrial β-oxidation and ATP production during periods of nutrient deprivation (Singh et al., 2009). Lipophagy occurs in almost every cell type investigated to date and, in many of them, is a constitutive process (Singh et al., 2009). In hepatocellular carcinoma (Hep3B) cells, starvation induces marked Rab7 activation and concomitant recruitment to LD membranes (Schroeder et al., 2015). In this manner, Rab7 "primes" LDs for autophagic degradation and targets them for docking/fusion with lysosomal compartments, thus promoting lipophagy (Schroeder et al., 2015). siRNA-mediated depletion of Rab7 significantly impairs starvation-induced LD breakdown, and this phenotype can be rescued by re-expression of wild-type (WT) and constitutively active (Q67L) forms of Rab7, but not by dominant-negative (T22N) or prenylation-defective Rab7 mutants, indicating that Rab7-mediated lipophagy is dependent on both Rab7 GTPase activity and Rab7 membrane association (Schroeder et al., 2015). Furthermore, a Rab7 mutant unable to bind Rab7-interacting lysosomal protein (RILP), a Rab7 effector and microtubule-based motor adaptor involved in minus end-directed lysosomal transport (Jordens et al., 2001), but otherwise displays normal GTPase activity and membrane distribution, cannot reverse the inhibitory effect of Rab7 knockdown on lipophagy. Considering Rab7 simultaneously regulates plus end-directed transport of autophagosomes through FYCO1 (Pankiv et al., 2010 - see above text), this observation implies that Rab7 directly orchestrates the union of degradative compartments with those LDs primed for lipophagy through regulation of microtubule dynamics (Schroeder et al., 117

2015). Intriguingly, Rab10, the Rab substrate of DENND4A, has been found to reside on LDs isolated from human squamous epithelial carcinoma (A431) cells (Umlauf et al., 2004), Chinese hamster ovarian (CHO) K2 cells (Liu et al., 2004; Bartz et al., 2007), human hepatoma (Hepswx) cells (Sato et al., 2006), human monocytes (U937) (Wan et al., 2007), mouse skeletal muscle (C2C12) cells (Zhang et al., 2011) and murine primary adipocytes (Ding et al., 2011). However, a role for Rab10 in lipophagy has yet to be described.

5.1.5 Chapter aims

Herein, I aim to establish a function for DENND4A phosphorylation and/or 14-3-3 binding in an mTORC1- and/or insulin-regulated cellular process that involves Rab10. I will first assess whether DENND4A phosphorylation modulates its GEF activity towards Rab10 in HEK-293E cells using a Rab effector pull-down strategy. I will then proceed to investigate whether overexpression of wild-type and phosphomutant DENND4A species influence insulin-regulated GLUT4 translocation in 3T3-L1 adipocytes using live-cell total internal reflection fluorescence (TIRF) microscopy. Lastly, I intend to examine a Rab10 knockout (Rab10 −/−) mouse embryonic fibroblast (MEF) cell line previously established in the James laboratory for autophagic defects and, if any are found, probe whether a DENND4A−Rab10 pathway is involved.

5.2 Methods

General methods other than those described below are located in Chapter 2.

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5.2.1 In vivo GEF assay

5.2.1.1 GST-MICAL-L2 domain (GST-ML2) protein purification E.coli (BL21) containing the pGEX-GST-ML2 plasmid was grown overnight in 5 mL LB medium containing 100 μg/mL ampicillin at 37 °C with shaking. The following morning, this culture was used to inoculate 500 mL LB medium containing 100 μg/mL ampicillin which was incubated at 37 °C with shaking until the culture reached an OD600 of 0.6. IPTG-mediated protein expression induction, bacterial cell lysis and affinity purification of GST fusion proteins were performed as described in Chapter 2, 2.3.4.1 and 2.3.4.2, except that 2 mM DTT was added to the elution buffer. The purified GST-ML2 protein was then buffer exchanged in 50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 5 mM MgCl2 and concentrated using an Amicon Ultra Centricon centrifugal filter device (Millipore) with a 10 kDa cut-off. The protein concentration was quantified by BCA assay. The molecular weight and purity of the protein were confirmed by SDS-PAGE (10% gel), followed by Coomassie staining [see Chapter 2, 2.3.5].

5.2.1.2 GST-ML2 pull-down from HEK293-E cells The Rab effector pull-down method was performed as described by Sano et al. (2008). HEK-293E cells were transiently transfected as described in Chapter 2, 2.3.2.2 with FLAG-tagged Rab10 and either eGFP-tagged wild-type DENND4A, eGFP-DENND4A- 35P or empty pcDNA™-DEST53-eGFP vector at a 1:2 molar ratio (15 μg total) to ensure cells expressing Rab10 also expressed the GEF or empty plasmid. Where endogenous Rab10 was studied (indicated in text), cells were transfected with 10 μg FLAG-tagged wild-type DENND4A or p3XFLAG-CMV™-10 empty vector only. Cells were serum- starved for 2 h prior to insulin stimulation (100 nM) for 10 min. Following insulin stimulation, cells were transferred to ice, washed twice in ice-cold PBS and harvested in 1 mL lysis buffer (3% IGEPAL CA-630, 50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 5 mM

MgCl2) containing 100 μg/mL GST-ML2 (or, where indicated, 50 μg/mL GST tag only), cOmplete™ protease inhibitor cocktail without EDTA (Roche) and phosphatase inhibitors (2 mM sodium orthovanadate, 1 mM sodium pyrophosphate, 1 mM ammonium molybdate and 10 mM sodium fluoride). Cells were lysed by passing through a 22-gauge needle six times, followed by six times through a 27-gauge needle, and then centrifuged

119 at 16,000 x g for 10 min at 4 °C to remove insoluble material. For each sample, the supernatant was combined with 40 μL of a 50% glutathione sepharose slurry (GE Healthcare) that had been washed twice in wash buffer (0.5% IGEPAL CA-630, 50 mM

Tris-HCl, pH 8.0, 100 mM NaCl, 5 mM MgCl2), and incubated for 1 h at 4 °C with rotation. The resin was then washed 3 times with wash buffer by repeated centrifugation at 2,000 x g for 2 min at 4 °C, and dried by aspiration of the supernatant through a 30- gauge needle. Precipitated proteins were eluted by addition of 40 μL 2X LSB and incubation at 65 °C for 10 min. Samples were then centrifuged at 16,000 x g for 5 min at RT and stored at -20 °C. Western blotting was performed as described in Chapter 2, 2.3.3. Immunoblot densitometries were quantified using ImageJ software.

5.2.2 Live-cell GLUT4 translocation assay

5.2.2.1 Preparation of Matrigel-coated imaging dishes 35 mm glass-bottom imaging μ-dishes (ibidi) were incubated at RT for 3 h with a 1:50 dilution of Matrigel (Corning) in ice-cold PBS. Dishes were washed twice with PBS prior to use.

5.2.2.2 Electroporation of 3T3-L1 adipocytes Adipocytes at day 7 post-differentiation were washed twice with PBS, trypsinised with 5X trypsin-EDTA (Invitrogen), resuspended in 30 mL of DMEM/FBS medium and then centrifuged at 500 x g for 2 min. The supernatant was removed and pelleted cells were resuspended in 30 mL PBS and centrifuged at 500 x g for 2 min. This step was repeated, followed by aspiration of the medium and resuspension of cells in electroporation buffer

(20 mM HEPES, 135 mM KCl, 2 mM MgCl2, 0.5% Ficol 400, 1% DMSO, 2 mM ATP, 5 mM reduced L-glutathione, pH 7.6) that was filter-sterilised using a 0.2 μM filter. Cells were then transferred to electroporation cuvettes (Bio-Rad) containing 4 μg pGEM-T- GLUT4-TagRFP-T and either 50 ug pDEST53-eGFP-DENND4A WT; 50 μg pDEST53- eGFP-DENND4A 35P; 20μg p3XFLAG-CMV™-10-AS160 WT; or 20 μg p3XFLAG- CMV™-10-AS160 4P. The contents of each cuvette were mixed and cuvettes were electroporated with a single square wave pulse (200 V, 40 ms) using an ECM 830 Square Wave Electroporator (BTX-Harvard Apparatus). Post-electroporation, 2 mL of 120

DMEM/FBS medium was quickly added to the cells before seeding into 2-well silicone inserts (ibidi) placed on Matrigel-coated 35 mm glass-bottom imaging μ-dishes (ibidi). Cells were then returned to 37 °C and the culture medium replaced with fresh DMEM/FBS after cell attachment. Adipocytes were used for microscopy 24-48 h post- electroporation. 5.2.2.3 Confocal microscopy Adipocytes on μ-dishes expressing eGFP-DENND4A (WT or 35P mutant) and GLUT4- tagRFP-T were imaged with a Dragonfly multimodal imaging system using confocal functionality (Andor) on a Nikon TiE inverted microscope equipped with a 60x NA 1.49 objective lens and a iXon3 888 EMCCD camera (Andor). NIS Elements software (Nikon) was used for all microscope control and image acquisition.

5.2.2.4 Live single-cell total internal reflection fluorescence (TIRF) microscopy Adipocytes on μ-dishes were serum-starved in FluoroBrite DMEM medium (Thermo

Fisher Scientific) for 2 h at 37 °C with 10% CO2. Cells were then placed on a Nikon TiE inverted microscope equipped with an Okolab incubator maintained at 37 °C with 10%

CO2 and allowed to equilibrate for a further 1 h. Healthy and suitably transfected adipocytes were identified by bright-field and fluorescence microscopy using a 60x NA 1.49 objective (NA 1.49) and imaged with a Dragonfly multimodal imaging system using TIRF functionality (Andor). Cells were imaged over a time-course of 40 min with 1 nM insulin injection at 10 min. The Nikon Perfect Focus System was used for maintenance of focus over time. eGFP and tagRFP-T fluorescence was detected using an iXon3 888 EMCCD camera (Andor). NIS Elements software (Nikon) was used for all microscope control and image acquisition.

5.2.2.5 Image analysis Image analysis was performed using a custom image analysis pipeline in Fiji, a distribution of ImageJ (Schindelin et al., 2012).

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5.2.2.6 Statistical analyses Statistical analyses were performed using the statistical software package, GraphPad Prism 6 (Graph Pad Software Inc.). Data are presented as the average ±SEM. Comparisons between groups were performed using the appropriate statistical tests as indicated in the figure legends.

5.2.3 Lipid droplet studies

5.2.3.1 Isolation and immortalisation of mouse embryonic fibroblasts (MEFs) Rab10 knockout mice were generated previously in the James laboratory from mouse embryonic stem (ES) cell line XC820 (BayGenomics). MEFs were isolated from embryonic day 13.5 mouse embryos and immortalised by serial passaging following the guidelines of Xu (2005). All animal work was performed by Jacqueline Stöckli (James laboratory). MEFs were maintained in DMEM/FBS medium at 37 °C with 10% CO2 and passaged at ~60% confluence. Rab10 knockout was confirmed by Western blotting [see Fig. 5.5a].

5.2.3.2 Electron microscopy MEFs grown on Matrigel (Corning)-coated 13mm glass coverslips were either serum- starved for 2 h or left in DMEM/FBS (full-serum) medium. Cells were then transferred to ice, washed twice in ice-cold PBS and fixed in 2.5% glutaraldehyde in 0.1M phosphate buffer, pH 7.4 at RT. Following post-fixation in 1% osmium tetroxide/1.5% potassium ferricyanide in 0.065 M sodium cacodylate buffer, pH 7.4 for 2 h at 4 °C, cells were washed with distilled water and incubated in 0.5% uranyl acetate for 1 h at 4 °C. Samples were then rinsed in distilled water, dehydrated in ethanol and embedded in Epon 812 resin (Electron Microscopy Sciences). Electron microscopy was performed by Dr. Georg Ramm (Bio Electron Microscopy Facility, Monash University) and he gave permission for the work to be included in this thesis.

5.2.3.3 Preparation of oleate-supplemented growth medium DMEM/FBS medium containing 200 μM oleate was prepared following the guidelines of Listenberger and Brown (2007). Briefly, 100 μL oleic acid was added to a solution containing 200 μL 1 M NaOH in 15.7 mL distilled water that had been pre-warmed to 122

70 °C in a glass conical flask and incubated at 70 °C for 30 min with constant stirring. 5 μL 5 M NaOH was then added and the solution incubated for a further 5 min at 70 °C. This step was repeated until micelles were no longer visible. 3.5 mL of this 20 mM sodium oleate solution was then added dropwise to 11.6 mL 5% BSA (fatty acid-free) in PBS that had been filter-sterilised and pre-warmed to 37 °C, and mixed thoroughly. In a sterile glass vial, 750 μL of the BSA/oleate mix was added to 16.75 mL DMEM/FBS medium (pre-warmed to 37 °C) to yield a final concentration of 200 μM oleate. BSA- conjugated oleate was stored at -20 °C and oleate-supplemented DMEM was prepared fresh and filter-sterilised prior to addition to cells.

5.2.3.4 Lipophagy assay in MEFs Wild-type (WT) or Rab10 knockout (Rab10 −/−) MEFs grown on Matrigel-coated 13mm glass coverslips were incubated in DMEM/FBS containing 200 μM oleate overnight at

37 °C with 10% CO2. The following morning, cells were either immediately fixed [see 5.2.3.6] to assess intracellular lipid levels prior to starvation ('loaded'); or the oleate- supplemented medium was removed and cells were washed with PBS prior to the addition of starvation medium (substrate-free DMEM (Sigma D5030) containing 0.1% FBS, 5 mM D-glucose and 3.7 g/L sodium bicarbonate). Cells undergoing starvation were then returned to 37 °C for a further 30 h.

5.2.3.5 Preparation of Oil Red-O (ORO) stock and working solutions ORO stock solution was prepared by dissolving 2.5 g ORO powder in 400 mL 100% isopropanol and mixed by magnetic stirring for 2 h at RT. ORO working solution was prepared by adding 1.5 parts ORO stock solution to 1 part distilled water, allowing the solution to stand for 10 min at 4 °C prior to filtration through a 0.45 μM filter (Millipore). The ORO working solution was made fresh and used with 2 h of preparation.

5.2.3.6 ORO staining Cells were washed twice in PBS, fixed in 10% formalin in PBS for 10 min at RT and then rinsed with 60% isopropanol to facilitate the staining of neutral lipids. Cells were stained with ORO working solution for 10 min at RT, destained with 60% isopropanol and then counter-stained with Mayer’s haemotoxylin solution for 1 min at RT. Coverslips were 123 then washed 3 times with distilled water prior to mounting onto glass slides with Immuno- Fluore mounting medium (MP Biomedicals).

5.2.3.7 Differential interference contrast (DIC) microscopy and image analysis Images were acquired using a Leica DM6000 upright microscope (Leica Microsystems) using a x40 DIC objective and a DFC365-FX camera. Images were pre-processed using Fiji software (ImageJ). Semi-supervised image segmentation and classification was performed as described herein. Probability maps of cellular 'objects' (cell perimeter, cell area, cytoplasm, nuclei and lipid droplets) were generated using the machine-learning application, Ilastik (Sommer et al., 2011). Probability maps were further analysed using a custom image analysis pipeline in CellProfiler (Carpenter et al., 2006). Data were exported to Microsoft Excel and analysed using R statistical software (R Development Core Team, 2010).

5.3 Results

5.3.1 DENND4A 35-site phosphomutant exhibits enhanced Rab10-directed GEF activity

Phosphorylation and consequent 14-3-3 binding of the DENN domain-containing proteins, DENND1A and DENND3, has been shown to enhance their GEF catalytic activity towards Rab35 and Rab12, respectively [see Chapter 4, 4.1 Introduction] (Kulasekaran et al., 2015; Xu et al., 2015). I therefore questioned whether the GEF activity of DENND4A towards its substrate, Rab10, is also augmented by phosphorylation and 14-3-3 association. Sano et al. (2008) previously demonstrated that a 204 amino acid domain contained within the effector protein, molecule interacting with CasL-like 2 (MICAL-L2), binds selectively to the active, GTP-bound form of Rab10. The authors developed a method to measure in vivo GTP loading of Rab10 by pull-down of Rab10-GTP in complex with the Rab-binding domain of MICAL-L2 fused to GST (Sano et al., 2008). I used this same Rab effector pull-down strategy to test whether wild-type (WT) DENND4A, or a 35-site phosphomutant which does not bind 14-3-3 (DENND4A- 124

35P) [see Chapter 4], modulates Rab10 GTP loading in HEK-293E cells following 10 minutes† of insulin stimulation [Fig. 5.1]. Cells were lysed in non-ionic detergent containing GST-MICAL-L2, and Rab10-GTP complexed to GST-MICAL-L2 in the lysate was adsorbed to glutathione sepharose resin. Overexpression of DENND4A WT in HEK-293E cells had no effect on the level of endogenous Rab10-GTP [Fig. 5.1a and b]. Since the transfection efficiency of DENND4A was low (approximately 20%), I co- transfected FLAG-Rab10 with eGFP-tagged DENND4A WT or DENND4A-35P (or empty vector control) into HEK-293E cells at a 1:2 molar ratio to ensure that cells expressing FLAG-Rab10 also expressed DENND4A. GTP loading of Rab10 was enhanced by 50% in cells overexpressing DENND4A-35P versus the WT protein [Fig 5.1c and d]. This result suggests that phosphorylation and/or 14-3-3 binding of DENND4A downstream of insulin signalling impairs its GEF activity towards Rab10.

†10 minutes is the time-point at which DENND4A exhibits maximal 14-3-3 binding [see Chapter 4, Fig. 4.2a and b].

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Figure 5.1. In vivo GEF assay The GEF activity of DENND4A was determined by Rab effector pull-down assay. HEK-293E cells expressing a,b, FLAG-tagged DENND4A (WT) or empty vector or c,d, eGFP-tagged DENND4A (WT or 35P mutant) or empty vector and FLAG-Rab10 were serum-starved prior to insulin stimulation (100 nM, 10 min). Cells were lysed with non-ionic detergent in the presence of GST-MICAL L2 or GST tag alone. Cleared lysates were subjected to GST pull-down using glutathione sepharose beads. Input and pull-down eluates were immunoblotted with anti-FLAG, anti-GFP, anti-GST and anti-Rab10 antibodies as indicated. Three biological replicates were performed for each experiment. Data are presented as the average fold-change in a, endogenous Rab10-GTP over that observed in the empty vector condition or c, FLAG-tagged Rab10-GTP over that observed cells in expressing DENND4A WT, and representative immunoblots (b and d, respectively) are shown. Quantifications of FLAG-Rab10 in pull-down eluates were normalised to input. In each experiment, Rab10-GTP amounts in pull-down eluates were normalised to the average of all eluates. Error bars indicate SD. Black arrowhead indicates eGFP-tagged DENND4A species. Statistical analysis employed an unpaired t-test. **, p < 0.01; ns, non-significant. Abbreviations: EV, empty vector; WB, Western blot.

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5.3.2 Wild-type DENND4A inhibits insulin-stimulated GLUT4 translocation when overexpressed in 3T3-L1 adipocytes

A role for the Rab10 GEF, DENND4C, in insulin-stimulated GLUT4 translocation has previously been described [see Chapter 1, 1.14.8 DENND4A-C] (Sano et al., 2011; Sadacca et al., 2013). However, it is unclear whether DENND4A, which belongs to the same DENN subfamily as DENND4C, also regulates this process. TIRF microscopy allows for the selective visualisation of dynamic cellular events occurring at or within ~200 nm of the PM (the TIRF zone) and is an established method for the detection of GLUT4 exocytic events in living cells (Burchfield et al., 2010; Burchfield et al., 2013). To assess whether DENND4A participates in insulin-regulated GLUT4 translocation, I used single live-cell TIRF microscopy to monitor real-time GLUT4 exocytosis in 3T3- L1 adipocytes co-expressing eGFP-tagged DENND4A (WT or 35P mutant) and GLUT4- tagRFP-T over a 30 min time-course of insulin stimulation [Fig. 5.2]. Sano et al. (2003) demonstrated that overexpression of the non-phosphorylatable "4P" mutant of the Rab10 GAP, AS160 (AS160-4P), inhibits insulin-stimulated GLUT4 exocytosis in adipocytes, whereas overexpression of the wild-type protein has no effect [see Chapter 1, 1.12 The Rab GAP, AS160]. Therefore, I employed AS160 WT and AS160-4P as negative and positive controls for inhibition of insulin-stimulated GLUT4 translocation, respectively, in my study. In adipocytes overexpressing AS160 WT, insulin treatment caused a ~2.2- fold increase in GLUT4 exocytosis, as assessed by tag-RFP-T signal in the TIRF zone [Fig. 5.2a]. Cells expressing AS160-4P exhibited the expected inhibition of insulin- stimulated GLUT4 translocation [Fig. 5.2]. Remarkably, overexpression of DENND4A WT blunted insulin-stimulated GLUT4 exocytosis to 65% of that observed in the AS160 WT control [Fig. 5.2]. Furthermore, in adipocytes overexpressing the DENND4A 35P mutant, the GLUT4 response following insulin treatment resembled that of control cells [Fig. 5.2], suggesting that the inhibitory effect of the DENND4A WT protein is dependent on its phosphorylation downstream of insulin signalling. DENND4A WT and the 35P mutant display similar cytoplasmic and nuclear localisation patterns when overexpressed in 3T3-L1 adipocytes [Fig. 5.3a]. The amount of DENND4A in the TIRF zone was reduced, to an equal extent for both species, following insulin stimulation [Fig. 5.3b and c]. However, this is unlikely to reflect a loss of DENND4A localisation at the PM as no obvious membrane association of DENND4A was observed in these cells and, further, there was no detectable change in DENND4A localisation following insulin stimulation. 127

Rather, this observation can be explained by insulin-stimulated changes in cell morphology (cell spreading), which would be expected to reduce detectable eGFP fluorescence in the TIRF zone in a given region of interest.

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Figure 5.2. Live-cell GLUT4 translocation assay in 3T3-L1 adipocytes (Figure legend on next page).

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Figure 5.2. Live-cell GLUT4 translocation assay in 3T3-L1 adipocytes (previous page) Adipocytes co-expressing either eGFP-DENND4A (WT or 35P mutant) or AS160 (WT or 4P mutant) and GLUT4-tagRFP-T were serum-starved for 3 hr prior to insulin stimulation (1 nM, 30 min). Cells were imaged by TIRF microscopy over a time-course of 40 min where insulin was administered at 10 min. a, Time-lapse TIRF measurements of GLUT4-tagRFP-T. Data are presented as the average fold response over basal from two experiments [Supplementary Table S8]. n = 36 cells (AS160 WT); n = 38 cells (AS160 4P); n = 35 cells (DENND4A WT); n = 32 cells (DENND4A 35P). b, GLUT4 exocytosis during the 40 min time-course. Area under curve in a was calculated using the average Y-value of the first nine time-points as the baseline. Error bars indicate SEM. Statistical analysis employed a one-way ANOVA in GraphPad Prism 6 software. *, p < 0.05; ****, p < 0.0001; ns, non-significant. c, Representative cells before (basal) and after 20 min of insulin stimulation. Scale bar, 10 μm. Abbreviations: AUC, area under curve; FC, fold change; WT, wild-type.

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Figure 5.3. DENND4A localisation in 3T3-L1 adipocytes Adipocytes co-expressing eGFP-DENND4A (WT or 35P mutant) and GLUT4-tagRFP-T were serum-starved for 3 hr prior to insulin stimulation (1 nM, 30 min). Cells were imaged by TIRF microscopy over a time-course of 40 min where insulin was administered at 10 min. a, Representative adipocytes co-expressing eGFP-DENND4A WT or 35P mutant and GLUT4-tagRFP- T imaged by confocal microscopy at an arbitrary time post-insulin stimulation. Scale bar, 10 μm. b, Time-lapse TIRF measurements of eGFP-DENND4A species. Data are presented as the average fold response over basal from two experiments [Supplementary Table S9]. n = 35 cells (DENND4A WT); n = 32 cells (DENND4A 35P). c, Presence of DENND4A species in the TIRF zone during the 40 min time-course. Area under curve in b was calculated using the average Y-value of the last two time-points as the baseline. Error bars indicate SEM. Statistical analysis employed an unpaired t-test in GraphPad Prism 6 software. ns, non-significant. Abbreviations: AUC, area under curve; FC, fold change; WT, wild-type.

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5.3.3 Rab10 −/− mouse embryonic fibroblasts exhibit hallmarks of compromised autophagy under nutrient stress

A mouse embryonic fibroblast (MEF) cell line deficient in Rab10, the Rab substrate of DENND4A, has previously been established in the James laboratory. To investigate whether Rab10 plays a role in cellular autophagy, WT and Rab10 knockout (Rab10 −/−) MEFs were either maintained in full serum medium or serum-starved and subcellular structures examined by transmission electron microscopy (TEM) [Fig. 5.4]. Under fed conditions, WT MEFs displayed normal Golgi, ER and mitochondrial morphology; the presence of intracellular lipid droplets; and few degradative organelles (late endosomes, lysosomes and autophagic structures) [Fig. 5.4a]. Following a 2 h period of serum starvation, the intracellular lipid content of WT MEFs was reduced, and this was accompanied by an increase in degradative organelles [Fig. 5.4a]. Rab10 −/− MEFs maintained in full serum medium had disorganised Golgi stacks, irregular mitochondrial morphology, and the ER was markedly shortened [Fig. 5.4b]. As in WT cells, lipid droplets were present and degradative organelles were sparse [Fig. 5.4b]. Under nutrient deprivation conditions, however, Rab10 −/− MEFs exhibited a less severe starvation response than their WT counterparts. Degradative organelles increased in numbers, but to a lesser extent than in WT cells, and lipid droplets were maintained [Fig. 5.4b]. These observations imply that cellular autophagy is compromised in Rab10-deficient cells and, moreover, suggest that Rab10 may function specifically in lipid droplet autophagy (lipophagy). To gain further insight into the potential role of Rab10 in lipophagy, I performed a pilot, fixed-cell microscopy-based experiment to quantitatively measure intracellular lipid droplet area in MEFs following nutrient stress. WT or Rab10 −/− MEFs were incubated overnight in full serum medium supplemented with oleate, a monounsaturated fatty acid ester, to promote intracellular lipid accumulation, and then starved for 30 h in low serum medium to induce lipophagy [Fig. 5.5]. However, using this methodology, there was no significant difference in intracellular lipid droplet area between cells imaged pre- and post-starvation [Fig. 5.5b] and therefore I could not assess whether Rab10 deficiency altered lipophagy in this setting. The lipid content of MEF cells was highly variable [Fig. 5.5b and Supplementary Table S10]. Furthermore, some cells failed to accumulate lipid during the loading phase [Fig. 5.5c]. Collectively, these observations emphasise the limitations of population-based assays due to cellular heterogeneity. Therefore, at this stage, further work is required to delineate the

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compromised autophagic phenotype observed in Rab10 −/− MEFs by TEM and these studies should focus on single-cell approaches.

Figure 5.4. Transmission electron microscopy (TEM) of wild-type and Rab10 knockout MEFs under fed and starved conditions a, Wild-type (WT) or b, (next page) Rab10 knockout (Rab10 −/−) mouse embryonic fibroblasts were either maintained in DMEM/FBS medium (full-serum) or subjected to serum-starvation (serum-free) for 2 hours. Subcellular structures are labelled as follows: D, degradative organelle (i.e. late endosome/lysosome/autophagic structure); ER, endoplasmic reticulum; G, Golgi; gsv, vesicles surrounding the Golgi apparatus; L, lipid droplet; M, mitochondrion; N, nucleus.

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Figure 5.4. Transmission electron microscopy (TEM) of wild-type and Rab10 knockout MEFs under fed and starved conditions (contd.).

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5.4 Discussion

Here, I have demonstrated that the GEF catalytic activity of DENND4A towards Rab10 is directly inhibited by phosphorylation downstream of insulin signalling. Moreover, DENND4A impairs insulin-regulated GLUT4 translocation when overexpressed in adipocytes, and this inhibitory effect is phosphorylation-dependent. Based upon my findings in Chapter 4, DENND4A is an insulin-regulated 14-3-3-binding protein and that mTORC1 is the primary kinase responsible for phosphorylating the insulin-sensitive 14- 3-3-interacting sites on DENND4A. Since insulin-regulated GLUT4 exocytosis is dependent on Akt activity (and unaffected by mTORC1 inhibition), collectively, these data imply that the phosphorylation site(s) in DENND4A that confer inhibition on GLUT4 translocation are independent of those that create docking sites for 14-3-3 and, further, are likely Akt phosphorylation sites. DENND4A Ser1282, Ser1252, Ser1036 and Ser1016 (murine) phosphorylation sites are attractive candidates as each was upregulated following insulin stimulation [Supplementary Tables S2 and S5] and they lie within a full/partial Akt consensus sequence [see Chapter 3, Table 3.3], yet mutation at these sites had no effect on the insulin sensitivity of 14-3-3 binding to DENND4A. The DENND4A- 35P mutant has served as a useful, albeit blunt, tool for studying the behaviour of DENND4A when its 14-3-3 binding capacity is absent. However, it is now clear that in order to define the phosphorylation sites that regulate DENND4A GEF activity and its inhibitory effect on GLUT4 exocytosis (which may be independent of one another or overlapping) future studies will require the use of DENND4A mutants with fewer mutated phosphosites. It will be especially important to test whether the GEF activity of DENND4A is enhanced and/or inhibition of GLUT4 translocation is lost in those mutants whose 14-3-3 interaction profiles resembled the WT protein (4A-1P, 4A-2P, 4A-4P and 4A-6P) [see Chapter 4, 4.3.4]. Moreover, it will be interesting to assess the kinase sensitivity of DENND4A GEF activity towards Rab10 by Rab effector pull-down. That is, whether the GEF activity of DENND4A WT is comparable to that of the 35P mutant in the presence of rapamycin or MK-2206, inhibitors of mTORC1 and Akt, respectively. This will provide an indication of whether DENND4A GEF activity is regulated by 14- 3-3 association versus phosphorylation per se; and, further, reveal whether DENND4A inhibition on GLUT4 traffic (presumably involving the Akt pathway) is a direct reflection of a change in DENND4A GEF activity towards Rab10. One could complement these 135 experiments by demonstrating that the in vivo GEF activity of DENND4A is specific to Rab10, for instance by employing other Rab species that bind to MICAL-L2, such as Rab13 (Terai et al., 2006), in the Rab effector pull-down assay.

The embryonic lethality of Rab10 knockout in mice (Lv et al., 2015) suggests that Rab10 is likely to function in other processes besides endosomal trafficking and GLUT4 exocytosis. Rab10 −/− MEFs examined by TEM displayed signs of compromised lipophagy following serum starvation. Furthermore, others (Lv et al., 2015) and I have observed abnormal ER morphology in Rab10 −/− embryonic cells, which is consistent with the proposed role of Rab10 in the regulation ER structure and dynamics (English and Voeltz, 2013). This is noteworthy since evidence suggests that, during autophagy initiation, the phagophore/autophagosome membrane is supplied by the ER (Hayashi- Nishino et al., 2009; Ylä-Anttila et al., 2009). In contrast, Vazirani et al. (2016) did not detect any gross alterations in ER function in the adipocytes of adipose-specific Rab10 knockout mice. Evidently, therefore, further studies are required to characterise the autophagic defects observed in Rab10 −/− MEFs and investigate whether a disrupted DENND4A−Rab10 pathway is causative.

DENND4C has been described as the primary Rab10 GEF responsible for insulin- regulated GLUT4 translocation in adipocytes [see Chapter 1, 1.14.8 DENND4A-C] (Sano et al., 2011; Sadacca et al., 2013). My findings clearly indicate that DENND4A also participates in this process. According to our current understanding, Rab10 regulates a single, prefusion step in GLUT4 traffic, presumably involving the delivery and/or tethering of GLUT4 vesicles to the PM (Gonzalez and McGraw, 2006; Bai et al., 2007; Sano et al., 2007; Sadacca et al., 2013). In fact, the latest viewpoint is that Rab10 accelerates the biogenesis of GSVs from a perinuclear recycling endosome/TGN compartment during insulin stimulation via the novel effector, SEC16A (Bruno et al., 2016). However, this simplistic one-step model cannot fully explain my observations and it is becoming obvious that Rab10 functions at multiple prefusion steps in the GLUT4 trafficking itinerary. The inhibition of DENND4A GEF activity by phosphorylation downstream of insulin signalling is inconsistent with a role for DENND4A in GSV biogenesis, as this process would require active Rab10. Rather, it is more likely that this is the step in GLUT4 traffic at which DENND4C functions and, as others (Sadacca et al., 2013) have suggested, it may be constitutively active. Where then in the GLUT4 lifecycle

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might DENND4A be controlling Rab10 activity? Perhaps DENND4A functions in the endosomal sorting of GLUT4 or, given the mounting evidence supporting a role for DENN domain proteins as regulators of Rab GTPases in autophagic trafficking events, it may be that DENND4A/Rab10 functions in the lysosomal degradation of endocytosed GLUT4 transporters. Either way, this would create competition between discrete functions of Rab10 and, further, indicate that DENND4A and DENND4C might play antagonising roles in GLUT4 traffic. GLUT4 traversing the endocytic system presumably has either one of three fates: to enter a rapid recycling route that leads directly to the PM; to be directed along the autophagosome-lysosome pathway for degradation; or, alternatively, be steered towards the TGN for eventual reincorporation into GSVs. One can imagine that, in the absence of insulin, the active, non-phosphorylated form of DENND4A might regulate Rab10-dependent retrograde transport of GLUT4 from the endosomal system to the TGN so that it can re-enter the GSV pathway. DENND4A phosphorylation following insulin stimulation would inhibit GTP loading of Rab10 at this location, allowing GLUT4 to enter the rapid recycling circuit, as would occur if insulin levels remained elevated. This newer model aligns with my observation that overexpression of DENND4A WT inhibits insulin-stimulated GLUT4 translocation, as one can picture here that an increase in the phosphorylated pool of DENND4A would inhibit the reincorporation of GLUT4 into GSVs and, further, promote the movement of GLUT4 circulating the endosomal recycling system towards a degradative pathway. This would then inhibit GLUT4 translocation simply due to a decrease in the number of transporters present in the insulin-responsive compartment (GSVs). In the case of the DENND4A 35P mutant, the block on endosome to TGN traffic would not occur, and so the magnitude of the GSV pool would be unaffected. However, the GLUT4 trafficking itinerary is undoubtedly very complex and the precise details of DENND4A/Rab10 involvement are not clear. Notably, Sano and colleagues (2011) have demonstrated that siRNA-mediated DENND4A knockdown reduces PM-localised GLUT4 by 24% in the insulin-stimulated state. In the model described above, this makes sense, as less GLUT4 would enter the GSV pathway if DENND4A levels were depleted. It is also possible that knockdown of DENND4A might have a profound effect on autophagy and this could inhibit GLUT4 traffic via an indirect mechanism.

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In summary, I have uncovered that the Rab10 GEF, DENND4A, inhibits insulin- stimulated GLUT4 exocytosis when overexpressed in 3T3-L1 adipocytes. Further, phosphorylation of DENND4A following insulin treatment inhibits its GEF activity towards Rab10. Curiously, Rab10 knockout cells exhibit defects in autophagy. These collective findings may be explained by an updated model of GLUT4 traffic, in which DENND4C and DENND4A regulate Rab10 function in competing GLUT4 secretory and sorting transport pathways, respectively.

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Chapter 6: Investigating the nature of DENND4C association with GLUT4 vesicles

The supplementary material relating to this chapter is located in the Appendix.

6.1 Introduction

GLUT4 storage vesicles (GSVs) constitute a specialised insulin-sensitive, GLUT4- containing compartment derived from the trans-Golgi network (TGN) and/or endosomes (Martin et al. 2000a; Stöckli et al., 2011) [see Chapter 1, 1.2 GLUT4]. Since GLUT4 traverses multiple intracellular membrane compartments during its lifecycle [see Chapter 1, Fig. 1.3], the purification and molecular characterisation of GSVs has proven biochemically challenging. Several studies have applied immunoisolation procedures to purify intracellular GLUT4-enriched membranes from cultured adipocytes and fat tissue to interrogate the protein composition of GSVs, a methodology later complemented by mass spectrometry (MS)-based proteomics. These analyses have led to the identification of insulin-regulated amino peptidase (IRAP, also known as gp160/vp165) (Kandror and Pilch, 1994), the v-SNARE, VAMP2/synaptobrevin-II (Cain et al., 1992; Volchuck et al., 1995; Martin et al., 1996), sortilin (Lin et al., 1997; Morris et al., 1998), low density lipoprotein receptor-related protein 1 (LRP1) (Jedrychowski et al., 2010) and tumour suppressor candidate 5 (TUSC5) (Fazakerley et al., 2015) as major integral membrane constituents of GSVs. Among these, IRAP (Mastick et al., 1994; Ross et al., 1996; Malide et al., 1997; Garza and Birnbaum, 2000) and TUSC5 (Fazakerley et al., 2015) both display a similar insulin-regulated trafficking pattern to GLUT4. Other proteins associated with GLUT4-containing membranes include mannose-6-phosphate receptors (Tanner and Lienhard, 1989; Kandror and Pilch, 1996; Ramm et al., 2000); the transferrin receptor (TfR) (Tanner and Lienhard, 1989; Livingstone et al., 1996); GLUT1 (Calderhead et al., 1990; Piper et al., 1991); secretory carrier-associated membrane proteins (SCAMPs) (Laurie et al., 1993; Thoidis et al., 1993); VAMP3/cellubrevin (Volchuck et al., 1995); the t-SNARE, syntaxin 4 (Volchuck et al., 1996); α-tubulin (Guilherme et al., 2000); the intermediate filament protein, vimentin (Guilherme et al., 2000); the Rab-GAP, AS160 (Larance et al., 2005); and Rabs 1b, 2a, 5b, 8a, 10, 11 and 139

14 (Larance et al., 2005; Mîinea et al., 2005; Jedrychowski et al., 2010; Fazakerley et al., 2015). However, immunoisolated GLUT4-enriched membranes likely represent a mixture of bona fide GSVs and GLUT4-containing endosomal compartments, as well as other compartments that GLUT4 might traverse en route to GSVs. As such, many of the proteins identified in these studies may not reside in insulin-sensitive GLUT4 vesicles. For instance, evidence suggests that the constitutive recycling proteins, TfR and GLUT1, are both excluded from GSVs (Kandror et al., 1995; Livingstone et al., 1996).

Recent studies implicate a role for the DENN domain-containing Rab10 GEF, DENND4C, in insulin-regulated GLUT4 traffic (Sano et al., 2011; Sadacca et al., 2013) [see Chapter 1, 1.14.8 DENND4A-C]. Notably, the Lienhard laboratory (Sano et al., 2011) has shown that DENND4C is present on IRAP-positive GLUT4 vesicles purified from 3T3-L1 adipocytes and, further, the amount of DENND4C localised to GSVs is unchanged following insulin stimulation. To date, DENND4C has not been detected in proteomic analyses of GLUT4-containing membranes (Larance et al., 2005; Jedrychowski et al., 2010; Fazakerley et al., 2015); however, given that each analysis identified proteins not found in either of the others, none are fully comprehensive. Herein, I will use an immunoisolation technique to purify GLUT4 vesicles from cultured 3T3-L1 adipocytes to validate the observations of Sano and colleagues (2011) described above. If DENND4C is indeed found to be a resident protein of GLUT4 vesicles, then I intend to probe further the nature of this association by determining the DENND4C interaction partner(s) and/or the DENND4C protein domain(s) responsible for its GSV localisation.

6.2 Methods

General methods other than those described below are located in Chapter 2.

6.2.1 Subcellular distribution and protein interactions of DENND4C

6.2.1.1 Subcellular fractionation of 3T3-L1 adipocytes 3T3-L1 adipocyte fractionation was performed as described previously by Fazakerley et al. (2015). Briefly, adipocytes on day 10-12 post-differentiation were serum-starved for

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2 h prior to insulin stimulation (100 nM, 20 min). Cells were then transferred to ice, washed three times in ice-cold PBS and harvested in HES buffer (10 mM HEPES, pH 7.4, 1 mM EDTA, 250 mM sucrose) containing cOmplete™ protease inhibitor cocktail (Roche) and phosphatase inhibitors (2 mM sodium orthovanadate, 1 mM sodium pyrophosphate, 1 mM ammonium molybdate and 10 mM sodium fluoride). Cells were lysed by passing through a 22-gauge needle six times, followed by six times through a 27-gauge needle. Lysates were solubilised on ice for 20 min and then centrifuged at 500 x g for 10 min at 4 °C to pellet unbroken cells. The pellet was discarded and the supernatant was centrifuged at 13,550 x g for 12 min at 4 °C to yield two fractions: the pellet fraction consisting of plasma membranes (PM) and mitochondria/nuclei (mito/nuc); and the supernatant fraction consisting of cytosol, low-density microsomes (LDM) and high-density microsomes (HDM). To isolate HDM, the supernatant fraction was transferred to a clean tube and centrifuged at 21,170 x g for 17 min at 4 °C. The resulting HDM pellet was resuspended in HES buffer containing protease and phosphatase inhibitors. The supernatant from this spin was transferred to a clean tube and centrifuged at 235,200 x g for 75 min at 4 °C. The supernatant (cytosol fraction) was transferred to a clean collection tube and the pellet (LDM fraction) was resuspended in HES buffer containing protease and phosphatase inhibitors. The protein concentration of each fraction was quantified by BCA assay (Pierce) following the manufacturer’s protocol. Western blotting was performed as described in Chapter 2, 2.3.3.

6.2.1.2 Immunoisolation of GLUT4-containing vesicles from 3T3-L1 adipocytes 3T3-L1 adipocytes on day 6 or 7 post-differentiation were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Subcellular fractionation was performed as described in 6.2.1.1 above. The supernatant from the centrifugation spin used to pellet the HDM (post-HDM supernatant, pHS) containing the LDM and cytosol fractions was retained and used as the starting material for immunoprecipitations. Control (non- specific) mouse IgG or anti-GLUT4 monoclonal 1F8 antibody was covalently coupled to Protein G sepharose resin (GE Healthcare) at a concentration of 2 μg IgG per 50 μL of resin (50% slurry) in PBS containing 2% BSA for 1 h at 4 °C with rotation. The resin was washed three times with ice-cold PBS by repeated centrifugation at 2,000 x g for 2 min at 4 °C, and then incubated overnight with 1.5-2.5 mg of pHS in HES containing 0.1% 141

BSA and 150 mM NaCl in a final volume of 800 μL at 4 °C with rotation. The following day, the resin was pelleted by centrifugation at 2,000 x g for 2 min at 4 °C and (where indicated) an aliquot of the supernatant retained as the unbound fraction. The resin was washed twice with HES, then twice in ice-cold PBS by repeated centrifugation at 2,000 x g for 2 min at 4 °C. The resin was then dried by aspiration of the supernatant through a 30-gauge needle. Immunoprecipitated proteins were eluted by addition of 50 μL 2X LSB and incubation at 65 °C for 10 min. Samples were then centrifuged at 16,000 x g for 5 min at RT. The supernatant was transferred to a clean tube and stored at -20 °C. Western blotting was performed as described in Chapter 2, 2.3.3. In some cases (indicated in the text), solid magnetic Dynabeads (Invitrogen) were used in place of the Protein G sepharose resin, in which case all washes were performed by magnetic capture and resuspension.

6.2.1.3 Immunoisolation of DENND4C- and AS160-associated vesicles For immunoisolation of DENND4C- and AS160-associated vesicles, vesicle immunoprecipitation was performed as described above [see 6.2.1.2], except that either an anti-DENND4C or an affinity purified anti-AS160 polyclonal antibody was covalently coupled to the Protein G sepharose resin or Dynabeads. A control (non-specific) rabbit IgG antibody was used in these instances.

6.2.1.4 GST fusion protein-coupled CNBr-activated sepharose resin production E.coli (BL21) cultures containing either the pGEX-KG empty vector, pGEX-GST-

IRAP2-109, pGEX-GST-VAMP21-94, or pGEX-GST-GLUT4466-509 plasmids were grown overnight in 5 mL LB medium containing 100 μg/mL ampicillin at 37 °C with shaking. The following morning, each culture was used to inoculate 500 mL LB medium containing 100 μg/mL ampicillin which was incubated at 37 °C with shaking until the culture reached an OD600 of 0.6-0.8. IPTG-mediated protein expression induction, bacterial cell lysis and affinity purification of GST fusion proteins were performed as described in Chapter 2, 2.3.4.1 and 2.3.4.2. Purified GST fusion proteins were then buffer exchanged in CNBr coupling buffer (100 mM NaHCO3, 500 mM NaCl) and concentrated to a volume of 2 mL using Amicon Ultra Centricon centrifugal filter devices (Millipore)

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with a 3 kDa cut-off. Protein concentrations were quantified by BCA assay and the molecular weight and purity of the proteins were confirmed by SDS-PAGE (15% gel), followed by Coomassie staining [see Chapter 2, 2.3.5]. GST fusion proteins were then coupled to CNBr-activated sepharose (GE Healthcare) according to the manufacturer’s instructions at a ratio of 5 mg of GST fusion protein/1 mL of resin. For preservation,

NaN3 was added to a final concentration of 0.01% and the resin stored at 4 °C.

6.2.1.5 GST pull-down of FLAG-tagged AS160 and DENND4C FLAG-tagged AS160 or DENND4C constructs were expressed in HEK-293E cells by transient transfection as described in Chapter 2, 2.3.2.2. Cells were serum-starved for 2 h, transferred to ice, washed twice in ice-cold PBS and harvested in 700 μL IP buffer (1% IGEPAL CA-630, 10% glycerol, 50 mM Tris-HCl, pH 7.4, 150 mM NaCl) containing cOmplete™ protease inhibitor cocktail (Roche) and phosphatase inhibitors (2 mM sodium orthovanadate, 1 mM sodium pyrophosphate, 1 mM ammonium molybdate and 10 mM sodium fluoride). Cells were lysed by passing through a 22-gauge needle six times, followed by six times through a 27-gauge needle. Lysates were solubilised on ice for 20 min and then centrifuged at 18,000 x g for 20 min at 4 °C to remove insoluble material. The protein concentration of the supernatant was quantified by BCA assay following the manufacturer’s protocol. GST pull-down method was essentially that of Larance et al. (2005). For each sample, 1 mg of lysate was incubated with either GST tag alone, GST-

IRAP1–109, GST-GLUT4466–509, or GST-VAMP21–94 coupled to CNBr-activated sepharose resin (50 μg GST-tagged protein/mg of lysate) overnight at 4 °C with rotation. The next day, the resin was washed twice with IP buffer, followed by twice with ice-cold PBS by repeated centrifugation at 2,000 x g for 2 min at 4 °C. The resin was dried by aspiration of the supernatant through a 30-gauge needle. Precipitated proteins were eluted by addition of 50 μL 2X LSB and incubation at 65 °C for 10 min. Samples were then centrifuged at 16,000 x g for 5 min at RT. The supernatant was transferred to a clean tube and stored at -20 °C. Western blotting was performed as described in Chapter 2, 2.3.3.

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6.2.2 Molecular cloning

6.2.2.1 Truncated DENND4C mutant species generation pcDNA™4/TO-eGFP DENND4C was used as a template to generate the following truncated murine DENND4C species using PCR primers introducing a double stop codon after the desired nucleotide positions: pcDNA™4/TO-eGFP DENND4C−MABP (corresponding to amino acids 1-191 of the full-length protein) and DENND4C−ΔC- terminus (1-631 aa). Mutagenic primers are listed in Supplementary Table S7. All deletion mutations were confirmed by DNA sequencing and Western blot analysis [Supplementary Fig. S1]. To generate the DENND4C−ΔMABP (192-1906 aa) truncation mutant, DENND4C (192-1906 aa) cDNA was amplified from pcDNA™4/TO-eGFP- DENND4C using attB site-containing forward (5'-GGGGACAAGTTTGTACAAAAAA GCAGGCTTCTTTCTGTGTTACAAGAAGTCTGTGCC-3') and reverse (5'-GGGGAC CACTTTGTACAAGAAAGCTGGGTCTTAAATGAGAGGCGCTCCAAAACAC-3') primers. The attB-flanked DENND4C (192-1906 aa) cDNA fragment was then recombined into pDONR™221 by performing a Gateway® BP reaction, and the eGFP- tagged DENND4C−ΔMABP construct made by performing a Gateway® LR reaction between pDONR™221-DENND4C and pcDNA™-DEST53-eGFP [see Chapter 2, 2.3.1].

6.2.3 DENND4C truncation mutant localisation

6.2.3.1 Transient transfection of HeLa cells

HeLa cells were maintained in DMEM/FBS medium at 37 °C with 10% CO2 and passaged at ~60% confluence. Cells were transiently transfected at 60-70% confluence using TransIT-X2 Dynamic Delivery System (Mirus) according to the manufacturer’s instructions. Briefly, per well of a 6-well culture dish, 2.5 μg DENND4C plasmid DNA (wild-type or mutant) was added to 250 μL Opti-MEM medium (Gibco) and 7.5 μL TransIT-X2, mixed and incubated at RT for 20 min. All media was then removed from the culture well and replaced with 2.5 mL of fresh DMEM/FBS medium. The transfection mix was added drop-wise to cells and the culture plate incubated overnight at 37 °C. Subsequently, the transfection mix was removed and replaced with 2 mL DMEM/FBS

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medium. Cells were returned to 37 °C and re-seeded onto Matrigel-coated [see Chapter 5, 5.2.2.1] 13mm glass coverslips at 24 h post-transfection.

6.2.3.2 Confocal microscopy HeLa cells expressing eGFP-tagged DENND4C species seeded on glass coverslips were serum-starved for 2 h at 48 h post-transfection. Cells were then washed twice in ice-cold PBS and fixed in 3% paraformaldehyde (PFA, ProSciTech) in PBS for 15 min at RT. Subsequently, cells were washed 3 times with PBS and quenched with 50 mM glycine in PBS containing 2% BSA. Coverslips were then washed three times in PBS and once in distilled water prior to mounting onto glass slides with Immuno-Fluore mounting medium (MP Biomedicals). eGFP fluorescence was imaged with a Dragonfly multimodal imaging system using confocal functionality (Andor) on a Nikon TiE inverted microscope equipped with a 60x NA 1.49 objective lens and a iXon3 888 EMCCD camera (Andor). NIS Elements software (Nikon) was used for all microscope control and image acquisition. Images are presented as maximum intensity Z-projections created using Fiji (ImageJ) software.

6.3 Results

6.3.1 DENND4C is enriched in the low density microsome fraction of 3T3-L1 adipocytes

Insulin-sensitive GLUT4 vesicles are enriched in the low density microsomal (LDM) fraction of non-stimulated (basal) 3T3-L1 adipocytes (Simpson et al., 1983; James and Pilch, 1988; Piper et al., 1991). To examine the cellular distribution of endogenous DENND4C in 3T3-L1 adipocytes and hence determine whether DENND4C, like GSVs, resides in the LDM, I performed subcellular fractionation of non-stimulated and insulin- stimulated cells by differential centrifugation, followed by Western blotting using polyclonal anti-DENND4C and monoclonal anti-GLUT4 (1F8) antibodies [Fig. 6.1]. The

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Uniprot database lists two murine DENND4C isoforms of 211 kDa and 216 kDa (accession #A6H8H2-1 and #A6H8H2-2, respectively). Both DENND4C isoforms were enriched in the LDM and the amount of DENND4C in this fraction was unchanged following insulin treatment [Fig. 6.1]. The DENND4C antibody also labelled a single faint band in the cytosolic fraction with an apparent heavier molecular weight [Fig. 6.1]. DENND4C was absent from both high density microsomes (HDM), which consist primarily of membranes derived from the ER and Golgi apparatus (Simpson et al., 1983), and a single fraction† containing plasma membranes (PM), nuclei and mitochondria [Fig. 6.1]. GLUT4 was highly enriched in HDM and LDM [Fig. 6.1]. Insulin treatment of adipocytes led to a loss of GLUT4 from these fractions, and this was accompanied by an increase in PM-localised† GLUT4 [Fig. 6.1].

†The PM was not separated from the PM/nuclear/mitochondrial fraction as the purpose of the experiment was to determine whether DENND4C is enriched in the LDM, the fraction containing GLUT4 vesicles. The GLUT4 detected in the PM/nuclear/mitochondrial fraction represents PM-localised GLUT4 [refer to Simpson et al., 1983].

Figure 6.1. Subcellular distribution of DENND4C and GLUT4 in 3T3-L1 adipocytes DENND4C and GLUT4 expression in subcellular fractions of cultured murine adipocytes was assessed by Western blotting with anti-DENND4C and anti-GLUT4 (1F8) antibodies. Black arrowheads indicate DENND4C isoforms of different molecular mass. Three biological replicates were performed and a representative immunoblot is shown. Abbreviations: HDM, high density microsomes; LDM, low density microsomes; mito, mitochondria; nuc, nuclei; PM, plasma membrane; WB, Western blot.

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6.3.2 DENND4C associates with GLUT4 vesicles in the non-stimulated state and dissociates following insulin stimulation in 3T3-L1 adipocytes

The Rab10 GAP, AS160, is a well-established GSV resident protein (Larance et al., 2005; Jedrychowski et al., 2010; Fazakerley et al., 2015). Sano et al. (2011) demonstrated that DENND4C, a GEF for Rab10, is also localised to GLUT4-containing vesicles purified from murine adipocytes. To verify this novel observation, I immunoisolated GSVs from non-stimulated and insulin-stimulated 3T3-L1 adipocytes using an anti-GLUT4 (1F8) antibody coupled to Protein G sepharose resin. A non-specific mouse IgG antibody was used as a negative control. To avoid disrupting fragile protein-membrane interactions by pelleting and resuspending the LDM fraction, the post-HDM supernatant (pHS), a mixture of cytosol and small vesicular membranes, was used as the starting material for immunoprecipitations (IPs). In accordance with the findings of Larance et al. (2005), AS160 was present on GLUT4 vesicles in the non-stimulated (basal) state and dissociated from GLUT4-containing membranes in response to insulin stimulation [Fig. 6.2a]. DENND4C was also detected on GSVs and, moreover, the behaviour of DENND4C association with GLUT4 vesicles under basal and insulin-stimulated conditions was analogous to that of AS160 [Fig. 6.2b]. Immunoblotting of the IP supernatant, however, revealed that a fraction of DENND4C in the pHS was not bound to GSVs in the non- stimulated state [Fig. 6.2b].

6.3.3 AS160 and DENND4C occupy distinct GLUT4 vesicle populations

Since AS160 and DENND4C were both found to reside on GLUT4 vesicles in the absence of insulin, I questioned whether these two Rab10 regulatory proteins could be co-localised on the same vesicles. To investigate this possibility, I immunoisolated DENND4C-containing membranes from the adipocyte pHS using a polyclonal anti- DENND4C antibody coupled to Protein G sepharose resin, and then immunoblotted the IP eluates and supernatants for the presence of AS160 [Fig. 6.2c]. In this instance, a non- specific rabbit IgG antibody was used as the negative control. AS160 was enriched in the anti-DENND4C IP supernatant in non-stimulated and insulin-stimulated cells [Fig. 6.2c]. However, it was unclear whether a small amount of AS160 was also associated with

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DENND4C-containing vesicles in the basal state due to the high level of non-specific binding of AS160 to the rabbit IgG-coupled resin [Fig. 6.2c]. To address this issue, I replaced the Protein G sepharose resin with solid magnetic Dynabeads and repeated the anti-DENND4C IP using pHS purified from non-stimulated adipocytes alone [Fig. 6.2d]. Non-specific binding of AS160 to the affinity support was suitably eliminated, revealing AS160 to be absent from the anti-DENND4C IP eluate [Fig. 6.2d]. To confirm that AS160 and DENND4C did not occupy the same membranes, I simultaneously performed a reciprocal IP, using a polyclonal anti-AS160 antibody, to isolate AS160-containing vesicles [Fig. 6.2e]. Immunoblotting revealed that DENND4C was indeed absent from the anti-AS160 IP eluate [Fig. 6.2e]. Interestingly, GLUT4 was present on AS160- containing membranes purified from non-stimulated adipocytes [Supplementary Fig. S2], yet was undetectable in the anti-DENND4C IP eluates [n=3; data not shown].

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Figure 6.2. Immunoisolation of GLUT4-, DENND4C- and AS160-containing vesicles from 3T3-L1 adipocytes (previous page) Adipocytes were serum-starved for 2 h prior to a-c, insulin stimulation (100 nM, 20 min), or d-e, were left unstimulated. Harvested cell lysates underwent differential centrifugation to purify the post-HDM supernatant (pHS) containing the LDM and cytosol fractions. The pHS was subjected to immunoprecipitation using a-b, anti-GLUT4 (1F8); c-d, anti- DENND4C; or e, anti-AS160 antibodies to isolate GLUT4-, DENND4C- and AS160-containing vesicles, respectively; and either a-b, mouse or c-e, rabbit IgG antibodies (non-specific controls). Starting material, immunoprecipitation eluates and b-e, immunoprecipitation supernatants were immunoblotted for the presence of endogenous GLUT4, DENND4C and AS160 as indicated. Immunoblots are from representative experiments (a, n=4; b, n=3; c, n=2; d, n=2; e, n=3). Black arrowhead indicates endogenous AS160 protein in 3T3-L1 adipocytes. Abbreviations: D4C, DENND4C; IgG, immunoglobulin G; IP, immunoprecipitation; pHS, post-HDM supernatant; sup., supernatant; WB, Western blot.

6.3.4 DENND4C does not interact with a panel of established GLUT4 vesicle integral membrane proteins

In the basal state, AS160 is localised to GLUT4 vesicles through association with the cytosolic tails of GSV resident integral membrane proteins, IRAP (Larance et al., 2005; Peck et al., 2006) and LRP1 (Jedrychowski et al., 2010). It is therefore conceivable that DENND4C may also interact with the intracellular domain(s) of established GSV cargo protein(s). To investigate this possibility, lysates from non-stimulated HEK-293E cells overexpressing either FLAG-tagged DENND4C or AS160 (positive control) were subjected to GST pull-down assay using either GST tag alone (negative control), GST fused to the GLUT4 C-terminus (GLUT4466-509), or GST fused to the N-termini of IRAP

(GST-IRAP1-109) and VAMP2 (GST-VAMP21–94) [Fig. 6.3]. Whereas AS160 associated strongly with the IRAP N-terminus, DENND4C did not bind to any of the GST fusion proteins in the panel of candidate GSV resident binding partners [Fig. 6.3].

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Figure 6.3. Binding of AS160 and DENND4C to GSV resident proteins Interactions between AS160 or DENND4C and established GLUT4 vesicle proteins in non- stimulated HEK-293E cells were determined by GST pull-down assay. Cells overexpressing FLAG- tagged AS160 or DENND4C were serum-starved for 2 hours. Harvested cell lysates were incubated with either GST tag alone, GST-IRAP1-109, GST-VAMP21–94, or GST-GLUT4466-509 coupled to CNBr-activated sepharose beads overnight. Starting material and pull-down eluates were immunoblotted with anti-FLAG and anti-GST antibodies. Black arrowheads indicate overexpressed FLAG-tagged AS160 and DENND4C. Two biological replicates were performed and a representative immunoblot is shown. Abbreviations: ø, non-transfected; D4C, DENND4C; G4, GLUT4; V2, VAMP2; WB, Western blot.

6.3.5 Localisation of DENND4C truncation mutants in HeLa cells

Since DENND4C did not directly interact with any of the established GSV integral membrane proteins tested in 6.3.4, I proceeded to investigate which DENND4C protein domain(s) are responsible for its membrane association. Besides the DENN module, which harbours Rab GEF catalytic activity, DENND4C contains multiple proteins domains, namely an N-terminal MVB12-associated β-prism (MABP) domain, a central nuclear localisation signal (NLS) and C-terminal interferon-stimulated response element (ISRE)-binding region (Marat et al., 2011) [see Chapter 1, Fig. 1.6]. A recent study has shown that the MABP domains of MVB12A and MVB12B, subunits of the human ESCRT-I complex, bind to acidic lipids and facilitate ESCRT-I function at the PM and late endosomes (Boura and Hurley, 2012). It is therefore possible that the MABP domain of DENND4C mediates its localisation to GLUT4-containing membranes, which are likely to incorporate GLUT4-stabilising anionic lipid species (Hresko et al., 2016). To 151 determine the intracellular localisation of the DENND4C MABP domain and, further, assess the contribution of the DENND4C C-terminus to its membrane association, I generated three eGFP-tagged truncated DENND4C species [Fig 6.4] and expressed them ectopically in HeLa cells [Fig. 6.5]. Wild-type (full-length) DENND4C and a truncation mutant lacking the entire region C-terminal to the DENN domain (DENND4C−ΔC- terminus) showed cytosolic staining as well as a punctate distribution consistent with endosomes [Fig. 6.5]. The localisation of the DENND4C mutant missing the N-terminal MABP domain (DENND4C−ΔMABP) also displayed a punctate distribution similar to the wild-type protein, albeit with a slightly more diffuse pattern [Fig. 6.5]. The DENND4C MABP domain alone localised to large punctate structures and displayed a much reduced level of diffuse cytosolic localisation relative to the other DENND4C species examined [Fig. 6.5]. This is consistent with the formation of insoluble aggregates suggesting that the MABP alone is unstable.

Figure 6.4. DENND4C truncation mutant species Schematic diagram showing the domain architecture and amino acid length of wild-type murine DENND4C, as well as three truncated DENND4C species. Abbreviations: aa, amino acids; C-term, carboxy-terminus; d, downstream DENN motif; Δ, deletion; DENN, core DENN module; ISRE, interferon- stimulated response element; LD, longin domain; MABP, MVB12-associated β-prism; NLS, nuclear localisation signal; PPR, pentatricopeptide repeat.

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Figure 6.5. Localisation of DENND4C truncation mutants in HeLa cells HeLa cells were transiently transfected with eGFP-tagged wild-type DENND4C (WT), DENND4C MABP domain alone (MABP), or truncated DENND4C species lacking either the MABP domain (ΔMABP) or the entire C-terminus downstream of the DENN domain (ΔC-terminus). Cells were serum-starved for 2 hr at 48 hr post-transfection prior to fixation. eGFP fluorescence was visualised directly by spinning disk confocal microscopy. For each DENND4C construct, images shown are representative of cells examined from three biological replicates. Scale bar, 5 μm.

6.4 Discussion

In this chapter, I have demonstrated that the Rab10 GEF, DENND4C, is localised to intracellular membranes and the cytoplasm. In 3T3-L1 adipocytes, the membranous fraction coincides with a subpopulation of GLUT4 vesicles that does not contain the Rab10 GAP, AS160. There is some disagreement in the literature over whether the Rab10 GAP, AS160, remains associated with, or dissociates from, GLUT4 vesicles in response to insulin [see Chapter 1, 1.12 The Rab GAP, AS160] (Larance et al., 2005; Koumanov et al., 2011). My data support the former claim, as AS160 was present in the anti-GLUT4 IP eluate in the basal state, yet the amount of AS160 precipitated with insulin resembled that of non-specific binding in the IgG control IP. Further, GLUT4 was barely detectable on AS160-containing membranes purified from insulin-stimulated adipocytes. Sano and colleagues (2011) have demonstrated that the amount of DENND4C localised to GSVs is unaffected by insulin treatment; however, I found that this was not the case. Rather, 153 akin to AS160, DENND4C disengaged from GLUT4-containing membranes in response to insulin. Notably, in the basal state, a portion of total DENND4C was present in the IP supernatant. Further, the fractionation data showed that DENND4C, despite dissociating from GSVs, remained in the LDM following insulin stimulation. These observations could imply that, in non-stimulated cells, DENND4C resides on different membranes that either include or exclude GLUT4, and, moreover, insulin causes DENND4C to redistribute to those membranes which lack GLUT4. Alternatively, since the LDM fraction comprises both intracellular membranes and large polypeptide complexes (Clark et al., 1998), it may just be that DENND4C is contained within the latter when not present on GLUT4 vesicles. However, given that only approximately one fifth of Rab10 in the LDM fraction is localised to GLUT4 vesicles (Sano et al., 2008), it is tempting to speculate that DENND4C may be colocalised with Rab10 at membranes besides those which contain GLUT4, most likely endosomal membranes, in both insulin-stimulated and non-stimulated cells. As AS160 was absent from DENND4C-containing membranes and vice versa, AS160 and DENND4C presumably occupy separate GLUT4 vesicle pools. Considering that the GAP domain of dephosphorylated AS160 is active under basal conditions (Sano et al., 2003) and that the GEF activity of DENND4C is thought to be constitutively active (Sadacca et al., 2013), it seems intuitive that these two counteracting regulators of Rab10 GTPase activity are not juxtaposed. Additionally, this implies that AS160 and DENND4C may function at distinct Rab10-regulated steps of GLUT4 traffic. In future, it will be interesting to determine, for instance by immunoelectron microscopic analysis of the LDM, the proportion of AS160-positive versus DENND4C-positive GLUT4 vesicle populations and/or their colocalisation with Rab10 to probe this matter further.

In support of published data (Larance et al., 2005), AS160 was found to interact with the N-terminal, cytosolic region of IRAP. DENND4C did not bind to the intracellular domain of IRAP, VAMP2 or GLUT4; however, it remains unclear whether DENND4C can associate with GSVs through direct protein-protein interaction(s), as other candidate GSV resident binding partners (namely LRP1, sortilin and TUSC5) were not tested in this study. Yoshimura et al. (2010) have previously shown that DENND4C is localised to a tubular Golgi-proximal membrane compartment when ectopically expressed in HeLa cells, an observation that is congruent with DENND4C enrichment in the LDM and association with GLUT4-containing membranes in vivo. Consistent with these findings, I also 154

observe localisation of DENND4C to punctate structures in HeLa cells. A similar distribution was observed using DENND4C truncation mutants missing either the N- terminal MABP domain (ΔMABP) or the entire region C-terminal to the DENN domain (ΔC-terminus). These data suggest that the DENN domain of DENND4C, not the MABP domain as originally hypothesised, may mediate its association with membranes, as this region was common to all DENND4C species that displayed a punctate distribution. It will therefore be important in the future to assess whether the membrane localisation of DENND4C is perturbed by removal of the DENN domain. Furthermore, given that HeLa cells have no endogenous GLUT4, it will now be necessary to validate the localisation of the DENND4C constructs in 3T3-L1 adipocytes and, moreover, assess their colocalisation with GLUT4 and/or markers of intracellular membranes under basal and insulin-stimulated conditions. Notably, the DENN module, the defining feature of the DENN family of proteins, contains a longin domain (Wu et al., 2011). Several GEFs are known to use longin domain dimerisation to create platforms for small GTPases (Levine et al. 2013a). Hence, one could envisage the DENN domain of DENND4C conferring membrane association via specific interaction(s) with substrate and/or non-substrate Rab(s) which, further, could represent a means of establishing a Rab cascade mechanism. In fact, the transient nature of Rab interactions could explain why DENND4C was not detected in proteomic analyses of GLUT4-containing membranes. The DENN domains of DENND1A-C (connecdenn1-3), GEFs for Rab35, have been shown to bind lipids (Allaire et al., 2010), and it will be interesting to test whether the DENN domains of DENND4C and other DENN domain proteins share this property. Indeed, there may be multiple modes of interaction between DENND4C and intracellular membranes involving Rab GTPases, membrane lipids and/or transmembrane proteins. Curiously, DENND4C contains an NLS, yet others (Yoshimura et al., 2010; Sano et al., 2011; Sadacca et al., 2013) and I provide no evidence for DENND4C nuclear localisation and/or function. Therefore, it remains unknown whether DENND4C plays a role in the nucleus.

In summary, I have demonstrated that DENND4C is localised to a distinct subpopulation of AS160-negative GLUT4 vesicles in non-stimulated 3T3-L1 adipocytes. Thus DENND4C is placed in a cellular location where it can access its substrate, Rab10. The physical nature of DENND4C association with GLUT4-containing membranes, however, remains unclear. The DENND4C DENN domain is an attractive candidate for mediating 155

DENND4C-Rab GTPases interactions. In future, it will be of interest to determine whether this domain and/or others contribute to DENND4C targeting in 3T3-L1 adipocytes.

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Chapter 7: General Discussion

One of the major outcomes of insulin signalling in adipocytes is the translocation of GLUT4 to the plasma membrane to permit glucose uptake. However, the precise points of intersection between upstream signalling via Akt and the distal GLUT4 trafficking machinery are not fully resolved. The inhibitory role of the Rab10-GAP, AS160, in GLUT4 traffic has been recognised for quite some time; yet the identity of a Rab-GEF that counter-regulates Rab10 activity in GLUT4 translocation has remained elusive. The recent emergence of DENN domain-containing proteins as a novel family of Rab-GEFs, therefore, has caused a ripple of excitement in GLUT4 and broader fields of membrane trafficking research. Sano and colleagues (2011) were the first to describe a role for the DENN protein, DENND4C, a GEF for Rab10, in insulin-regulated GLUT4 translocation. Given the complexity of the GLUT4 itinerary and the assortment of Rab GTPases identified on GLUT4 membranes, however, other DENN domain proteins besides DENND4C are probably involved in GLUT4 traffic. To interrogate this possibility, I studied the datasets from three large-scale phosphoproteomic studies of insulin stimulation and discovered that, not only are a diverse range of DENN proteins expressed in insulin target cell types/tissues, but many are insulin-regulated phosphoproteins. Of these, the Rab10 GEF, DENND4A, a close relative of DENND4C, was highly insulin- responsive. Further, many DENND4A phosphorylation sites exhibited predicted 14-3-3- binding motifs which suggested that, as for several known Akt substrates, 14-3-3 binding may be a mode of regulating the cellular function of DENND4A. Complementary 14-3- 3 pull-down and overlay assays in 3T3-L1 adipocytes and/or HEK-293E cells demonstrated that DENND4A was indeed a binding partner for 14-3-3. However, despite my original prediction, kinase inhibitor studies revealed that mTORC1 (and/or a downstream kinase) was the dominant kinase responsible for DENND4A phosphorylation. A multitude of phosphosites in the DENND4A C-terminus are candidates for 14-3-3 binding, and mutation of each of these to a phospho-dead alanine residue eliminated the interaction between DENND4A and 14-3-3. Phosphorylation of DENND4A downstream of insulin was found to impair its GEF catalytic activity towards Rab10 in vivo, as assessed by Rab effector pull-down. Furthermore, DENND4A inhibited real-time insulin-stimulated GLUT4 exocytosis when overexpressed in adipocytes, whereas the phospho-dead DENND4A mutant that does not bind 14-3-3 had no effect. 157

DENN proteins are becoming increasingly recognised as key players in cellular autophagy [see Chapter 5, 5.1.3 Autophagic roles of Rab GTPases and DENN domain proteins]. Rab10 knockout MEFs examined by TEM displayed hallmarks of defective lipophagy, suggesting that DENND4A, positioned downstream of mTORC1, may also function in this process. DENND4C is also an insulin-sensitive phosphoprotein and associates with 14-3-3. However, unlike DENND4A, the DENND4C−14-3-3 interaction is not regulated by insulin. DENND4C has previously been found to reside on GSVs purified from 3T3-L1 adipocytes (Sano et al., 2011). Here, I have expanded on the work of others by demonstrating that, in the absence of insulin, DENND4C localises to a distinct subpopulation of GLUT4 vesicles that exclude AS160 and dissociates from GLUT4-containing membranes following insulin stimulation.

The current opinion in the literature is that Rab10 acts at a single, prefusion step in the GLUT4 trafficking itinerary, possibly functioning in GSV biogenesis (Bruno et al., 2016). However, the present findings lead me to propose an updated model of GLUT4 traffic, in which there are multiple Rab10-dependent transport steps and, further, where DENND4C and DENND4A may regulate Rab10 function in competing exocytic and endosomal sorting pathways, respectively (described herein). In the absence of insulin, GLUT4 is sequestered in perinuclear GSVs and the active GAP domain of AS160 maintains its cognate Rab (presumably Rab10 in adipocytes) in an inactive, GDP-bound conformation, thus preventing docking and/or tethering of GSVs to the PM. The biogenesis of GSVs from the TGN, on the other hand, is presumably a constitutively active process and may involve DENND4C control of Rab10 activity. Meanwhile, at recycling endosomes (RE), I hypothesise that the active GEF domain of DENND4A maintains Rab10 in an active, GTP-bound conformation to regulate the retrograde transport of circulating GLUT4 to the TGN for eventual reincorporation into GSVs. Following insulin stimulation, Akt- mediated phosphorylation of AS160 inhibits its GAP activity, allowing the Rab10- dependent steps in GLUT4 exocytosis to proceed. DENND4A GEF activity is also inhibited by phosphorylation and/or 14-3-3 binding downstream of insulin and this allows GLUT4 to enter to a rapid recycling route for direct return to the PM. Once insulin is withdrawn, however, the GEF activity of DENND4A is restored due to its dephosphorylation and internalised GLUT4 traffics to REs, from where it can re-enter the GSV pathway [Fig 7.1].

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Since AS160 and DENND4C are found to occupy distinct GLUT4-containing membranes, they presumably regulate Rab10 activity at discrete locations. As indicated above, GSV biogenesis from the TGN may involve DENND4C and would require active, GTP-bound Rab10. As nascent GSVs bud off, one can picture they abandon DENND4C at the TGN but acquire AS160. In the absence of insulin, AS160 stimulates GDP hydrolysis of Rab10-GTP, thereby converting Rab10 to its inactive, GDP-bound conformation. One would expect this process to be highly dynamic, with GSVs constantly being formed and consumed as Rab10 cycles between active/inactive forms. In this context, whilst the GAP (AS160) and the GEF (DENND4C) may be simultaneously active, it is likely that the equilibrium favours the GDP-bound conformation of Rab10 leading to inhibition of GLUT4 exocytosis. With insulin, the inactivation of AS160 by phosphorylation would therefore achieve two goals: firstly, to stabilise the Rab10 packaged in GSVs in its active, GTP-bound conformation so that GLUT4 translocation can proceed and, secondly, to allow AS160 to facilitate the docking and/or fusion of GSVs at the PM, as described by Tan et al. (2012) [see Chapter 1, 1.12 The Rab-GAP, AS160]. It is unclear at present whether insulin-stimulated phosphorylation of DENND4C might further enhance its GEF activity towards Rab10 (discussed below). Therefore, in the newer model, there may be as many as three Rab10-regulated steps in the GLUT4 lifecycle: endosomal sorting of GLUT4 (DENND4A/Rab10); GSV biogenesis (DENND4C/Rab10); and GSV docking and/or tethering (AS160/Rab10) [Fig. 7.1].

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Figure 7.1. Model for the role of DENND4A and DENND4C in GLUT4 trafficking (Figure legend on next page). 160

Figure 7.1. Model for the role of DENND4A and DENND4C in GLUT4 trafficking (previous page) Schematic diagram showing Rab10 activity at the various steps in GLUT4 traffic under the regulation of GEFs, DENND4A and DENND4C, and the GAP, AS160. a, In the absence of insulin, active DENND4A/Rab10 control the retrograde transport of GLUT4 from recycling endosomes (RE) to the trans-Golgi network (TGN) (1). DENND4A/Rab10 may also regulate degradation of endocytosed GLUT4 transporters via an autophagosome−lysosome pathway (not shown). At the TGN, DENND4C/Rab10 function in GLUT4 storage vesicle (GSV) biogenesis, presumably a constitutively active process (2). As nascent GSVs bud off, they abandon DENND4C at the TGN but acquire AS160, which binds to the GSV resident protein, IRAP and inactivates Rab10. GSVs are constantly being formed and consumed; however the equilibrium favours Rab10-GDP thus inhibiting GLUT4 translocation (3). b, In the presence of insulin (4), DENND4A GEF activity is inhibited by phosphorylation and/or 14-3-3 binding (5). This allows GLUT4 circulating the endocytic system to enter a rapid recycling route for direct return to the plasma membrane (PM) (6). It is not understood whether insulin-stimulated phosphorylation of DENND4C might further enhance its GEF activity towards Rab10 in GSV biogenesis (7). Akt- mediated phosphorylation of AS160 enhances its binding to 14-3-3 and these events inhibit its GAP activity (8), allowing for GTP-loading of Rab10 and the Rab10-dependent step(s) in GLUT4 exocytosis to proceed (9). A pool of phosphorylated AS160 remains on GSVs to facilitate the docking and/or fusion of GSVs at the PM (10). Abbreviations: D, degradative compartment; EE, early endosome; GSV, GLUT4 storage vesicle; IR, insulin receptor; PM, plasma membrane; RE, recycling endosome; TGN, trans-Golgi network.

Further studies should now focus on testing the hypothetical model outlined above. It will be of highest priority to investigate which insulin-sensitive phosphorylation site(s) in DENND4A regulate(s) a) its GEF activity; and b) its inhibitory function in GLUT4 exocytosis, and whether these site(s) are unique or overlapping. In Chapter 4, using various DENND4A phosphomutant species, I demonstrated that some of the most highly upregulated phosphorylation sites in DENND4A following insulin treatment do not create docking sites for 14-3-3. It may be that those DENND4A phosphosites that confer 14-3-3 binding are independent of those that regulate DENND4A GEF activity; however, in the 35P mutant, all regulatory sites are mutated and so I cannot distinguish between them. DENND4A S1282 (murine) is an attractive candidate site for regulating DENND4A GEF activity in the context of GLUT4 traffic, as it conforms to the Akt consensus motif and phosphorylation at this position occurred rapidly following insulin treatment. Therefore, further in vivo GEF and live-cell GLUT4 trafficking assays [Chapter 5] should be performed using DENND4A mutants with fewer mutated phosphosites, especially those mutants whose 14-3-3 binding profiles resembled the WT protein. One of the most remarkable findings of this study was the large number of phosphosites confined to the 161

DENND4A C-terminus and, moreover, their segregation into distinct clusters [see Fig. 4.6]. Here, it is tempting to speculate that different kinases might target discrete clusters of DENND4A phosphorylation sites to regulate separate protein functions, such as 14-3- 3 binding and GEF activity. The insulin-regulated DENND4A–14-3-3 interaction was sensitive to rapamycin treatment, indicating that mTORC1 (and/or a kinase activated downstream of mTORC1) is the dominant kinase responsible for DENND4A phosphorylation in this context. Notably, DENND3 (Xu et al., 2015) and the DENN- related protein, C9orf72 (Sullivan et al., 2016; Webster et al., 2016), are now recognised as a substrate and interaction partner, respectively, of the autophagy-initiating kinase, ULK1, which is positioned downstream of mTORC1 and is itself a 14-3-3 binding protein (Bach et al., 2011). Since DENND4A/Rab10 may play a role in autophagy, it could be that ULK1 is in fact the major kinase acting on DENND4A. It will be interesting, therefore, to examine whether the association between DENND4A and 14-3-3 is sensitive to pharmacological inhibition of ULK1. Also, given the existence of candidate Akt phosphorylation sites in DENND4A and its proposed role in GLUT4 traffic, it will be necessary to test whether DENND4A is a direct substrate of recombinant Akt in vitro. The above model proposes that the inhibitory effect of DENND4A WT overexpression on GLUT4 exocytosis in adipocytes is due to a defect in the endosomal sorting of GLUT4, possibly involving enhanced GLUT4 degradation and/or a reduced GSV pool. If an overactive autophagosome-lysosome pathway is causative, then one would expect that by blocking autophagic processes (using rapamycin, for instance) the inhibitory effect of the WT protein would be abolished, and this warrants further investigation. Further, in this scenario, one would expect cells overexpressing DENND4A WT to have less total GLUT4 protein. In the live-cell GLUT4 trafficking assay, DENND4A WT- overexpressing adipocytes did indeed appear to contain less GLUT4-tag-RFP-T than surrounding non-expressing cells (observations of Murrow et al.); however, it will now be necessary to confirm this observation by accurately quantifying total GLUT4 protein in these cells. One could complement these experiments by examining whether the numbers of GFP-LC3 puncta (autophagosomes), a widely used method to assess autophagy, are increased in cells co-expressing GFP-LC3 and DENND4A WT. Lastly, it will be important to decipher if AS160 counter-regulates Rab10 at the same step(s) in GLUT4 traffic as DENND4A, or whether the control of DENND4A GEF function by

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phosphorylation and/or 14-3-3 binding alone is sufficient to generate appropriate Rab10 activity/inactivity at this discrete location.

Despite the knowledge gained from this study, my findings raise several outstanding questions discussed herein. Firstly, what is the functional significance of DENND4C phosphorylation? Sano and colleagues (2011) previously described three insulin- sensitive DENND4C phosphosites with partial Akt consensus sequences (Ser1043, Ser1096 and Ser1321); yet, overexpression of a non-phosphorylatable S1043A/S1096A/S1321A DENND4C mutant did not inhibit GLUT4 translocation in 3T3-L1 adipocytes. This implies that insulin-stimulated phosphorylation of DENND4C at these sites does not regulate its GEF activity. However, I have identified further insulin-sensitive sites in DENND4C: Thr966, Ser971, Ser1087, Ser1240 and Ser1274 [see Chapter 3]. Therefore, it will now be necessary to assess whether phosphorylation at these novel sites enhances DENND4C GEF activity towards Rab10, thus GLUT4 exocytosis, following insulin stimulation. Intriguingly, an unpublished study from the James laboratory (Hoffman et al.) has uncovered an additional insulin-sensitive site in DENND4C in L6 rat skeletal muscle cells, Ser1035 (corresponding to Ser987 in the murine protein), that is independently upregulated by 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR), an AMPK agonist. Since activation of AMPK mediates glucose transport during exercise (Hayashi et al., 1998; Kurth-Kraczek et al., 1999), this phosphorylation site could represent an important point of convergence between insulin and contraction signalling pathways on GLUT4 in muscle. Whereas Rab10 regulates GLUT4 exocytosis in adipocytes, Rab8a and Rab13 are implicated in this process in skeletal muscle (Ishikura et al., 2007; Sun et al., 2010; Sun et al., 2016). Presumably DENND4A and/or DENND4C do not exhibit GEF activity towards Rabs 8a and 13, and hence it is currently unclear whether the roles of DENND4A and/or DENND4C in GLUT4 traffic are conserved in muscle.

Secondly, what is the mode of interaction between DENN proteins and their substrate Rabs? Wu et al. (2011) describe the DENN domain as bi-lobed with the N-terminal lobe being a longin domain (LD). LDs are found in several known Rab GEFs and dimerise to create platforms for small GTPases (Levine et al. 2013a). Therefore, one might speculate that DENN domain proteins also use LD dimerisation to associate with their Rab substrates. This would presumably involve the formation of DENN protein homo- or

163 heterodimers. Notably, there are examples of heterodimerisation between DENN proteins in the literature: FLCN interacts with either of FNIP1 or FNIP2 to form a FLCN/complex (Baba et al., 2006; Hasumi et al., 2006; Takagi et al., 2008), and the DENN-related proteins, C9orf72 and SMCR8, associate in a complex that has GEF activity for Rab8a and Rab39b (Sellier et al., 2016; Sullivan et al., 2016). It remains to be tested whether DENND4A and DENND4C use homo- or heterodimerisation as a means for regulating substrate interactions and/or GEF catalytic activity. One could investigate this possibility, to begin with, by assessing whether DENND4A and/or DENND4C constructs fused to different epitope tags co-immunoprecipitate when expressed in the same cells. In Chapter 6, my data suggest that the DENND4C DENN domain may mediate its association with membranes. If this is correct, then it could indicate that an interaction between DENND4C and Rab10 is responsible for the membrane targeting of DENND4C. Therefore, it will be interesting in future to examine whether DENND4C localisation is perturbed when overexpressed in Rab10 knockout cells. In a similar vein, given the evidence supporting a role for Rab GEFs in Rab membrane targeting (Gerondopoulos et al., 2012; Blümer et al., 2013), one could assess whether Rab10 localisation is disturbed in DENND4A and/or DENND4C knockdown adipocytes.

What is the role of DENND4A in the adipocyte nucleus? DENND4A and DENND4C both possess a nuclear localisation signal (NLS). In 3T3-L1 adipocytes, I discovered that ectopic DENND4A was localised to two very distinct subcellular locations – the nucleus and bulk cytoplasm. Curiously, neither DENND4A (Yoshimura et al., 2010) nor DENND4C is found in the nucleus when overexpressed in HeLa cells. I wonder whether the purpose of targeting DENND4A to the nucleus in adipocytes is somehow linked to a speculative role in autophagy? It is unlikely that DENND4A functions as a GEF in the nucleus as this is inconsistent with Rab10 function in membrane trafficking. However, DENND4A has been shown to interact with the promoter of the human c-myc gene, which encodes the c-myc transcription factor (Stasiv et al., 1994; Semova et al., 2003). Intriguingly, overexpression of c-myc in rat 3Y1 fibroblasts induces autophagy (Tsuneoka et al., 2003), whereas knockdown of c-myc in HeLa cells inhibits autophagy by impairing autophagosome formation (Toh et al., 2013). Therefore, it is tempting to guess that DENND4A could not only regulate a Rab10-dependent autophagy pathway via its GEF function, but also control the expression of autophagy genes via c-myc.

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Lastly, do other DENN domain proteins function in the GLUT4 trafficking itinerary? In this study, I chose to focus my attention on DENND4A and DENND4C as these insulin- sensitive phosphoproteins are GEFs for Rab10. However, several other Rabs besides Rab10 are implicated in GLUT4 traffic [see Chapter 1, 1.10 Rab GTPases in GLUT4 traffic] and, further, a diverse range of DENN proteins are expressed in adipocytes [Fig. 3.1]. Davey et al. (2012) have described a novel role for the Rab-GAP, TBC1D13, and its substrate, Rab35, in GLUT4 traffic in 3T3-L1 adipocytes. Rab35 regulates a rapid recycling route between early endosomes and the PM (Kouranti et al., 2006). Further to this, DENND1A, a GEF for Rab35, was recently shown to be an Akt substrate in adipocytes (Kulasekeran et al., 2015). Therefore, it may be that DENND1A and TBC1D13 counter-regulate Rab35 activity in a rapid GLUT4 recycling pathway and this will be an important avenue of future research.

In conclusion, I have demonstrated that insulin targets two related GEFs, DENND4A and DENND4C, that act on the same Rab, but are likely to function in distinct processes at different cellular locations and be regulated via different signalling mechanisms. The concept that DENND4A may sit at the nexus between GLUT4 translocation and autophagy suggests that these processes are mutually exclusive. Moreover, the decision of the cell to employ a single Rab for several functions using different GEFs implies that the separate functions may share common Rab effectors. Insulin-regulated GLUT4 trafficking has become a paradigm for the precise yet profoundly intricate regulation of a system in response to the external environment. My research has added another piece to the GLUT4 puzzle by demonstrating that insulin clearly targets a highly organised series of molecules in order to orchestrate the movement of GLUT4 to the PM. These include Rab-GAPs and motor proteins from previous studies, but also Rab-GEFs, as shown here. In future, these proteins may serve as novel therapeutic targets for the treatment or prevention of insulin resistance and Type II diabetes mellitus. It is hardly surprising that the insulin regulation of glucose transport has captured the imagination of numerous researchers since the early 1960s. While my findings have evidently provided an advance, they also expose the need to forge ahead if we are to completely understand the intricacies of this extraordinary biochemical phemonenon.

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Appendix: Supplementary Material

Supplementary Figures

Figure S1. Immunoblot of DENND4C truncation mutants overexpressed in HEK-293E cells Predicted molecular weights of wild-type DENND4C, DENND4C MABP domain, and ΔMABP and ΔC-terminus mutants (inclusive of eGFP epitope tag) are approximately 240 kDa, 47 kDa, 218 kDa and 97 kDa, respectively. Abbreviations: ø, non-transfected; Ct, carboxy-terminus; Δ, deletion; Da, Dalton; k, kilo; MABP, MVB12-associated β-prism; WB, Western blot; WT, wild-type.

Figure S2. Immunoisolation of AS160-containing vesicles from 3T3-L1 adipocytes. Adipocytes were serum-starved for 2 h prior to insulin stimulation (100 nM, 20 min). Harvested cell lysates underwent differential centrifugation to purify the post-HDM supernatant (pHS) containing the LDM and cytosol fractions. The pHS was subjected to immunoprecipitation using an anti-AS160 antibody to isolate AS160-containing vesicles, or a rabbit IgG antibody (non- specific control). Starting material, immunoprecipitation eluates and immunoprecipitation supernatants were immunoblotted for the presence of AS160 and GLUT4. Two biological replicates were performed and a representative immunoblot is shown. Abbreviations: IgG, immunoglobulin G; IP, immunoprecipitation; pHS, post-HDM supernatant; sup., supernatant; WB, Western blot.

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Supplementary Tables

Table S1. Ranked abundance of proteins in 3T3-L1 adipocyte, L6 myotube and murine liver proteomes (electronic file). Absolute abundance of detected 3T3-L1 adipocyte [Sheet 1], L6 myotube [Sheet 2] and murine liver tissue [Sheet 3] proteomes were estimated using the log2- transformed summed peptide intensities (iBAQ) of each of the 7,105, 6,216 and 3,973 proteins quantified, respectively. Abbreviations: iBAQ, intensity-based absolute quantification.

Table S2. Class I phosphorylation sites in DENN domain-containing and DENN-related proteins expressed in 3T3-L1 adipocytes (electronic file). Phosphorylation status of 135 Class I phosphosites in 14 DENN proteins following insulin stimulation (100 nM) of 3T3-L1 adipocytes over a time-course of 0-60 min, or following pre-treatment with the Akt inhibitor, MK-2206, prior to insulin stimulation (100 nm, 20 min). Three biological replicate experiments were performed and the log2-transformed median fold change in insulin-stimulated phosphorylation over baseline (starved) at each site determined. The data are excerpted from Humphrey et al., 2013. Abbreviations: Exp, experiment; min, minutes; MK, MK-2206; S, serine; T, threonine; TC, time- course; Y, tyrosine.

Table S3. Class I phosphorylation sites in DENN domain-containing and DENN-related proteins expressed in murine liver tissue (electronic file). Phosphorylation status of 103 Class I phosphosites in 10 DENN proteins [Sheet 1] following in situ insulin delivery to murine liver (1 mU/g) over early (5-30 sec) [Sheet 2] and intermediate (0.5-10 min) [Sheet 3] time-courses. At least six biological replicates (separate mice) were performed for each time-point and the median fold change in insulin-stimulated phosphorylation over baseline at each site determined. Only those sites where quantitative data was present in ≥ 2 biological replicates at all time- points, or ≥ 3 biological replicates in ≥ 3 of 4 time-points (early time-course); or ≥ 6 of 7 time- points (intermediate time-course) are reported. The data are excerpted from Humphrey et al., 2015a. Abbreviations: Bio, biological replicate; FC, fold change; quant., quantified; s, seconds; S, serine; T, threonine; TC, time-course; Y, tyrosine.

Table S4. Class I phosphorylation sites in DENN domain-containing and DENN-related proteins expressed in L6 myotubes (electronic file). Phosphorylation status of 56 Class I phosphosites in 11 DENN proteins following insulin stimulation (100 nM, 20 min) of L6 myotubes. Four biological replicate experiments were performed and the median fold change in insulin-stimulated phosphorylation over baseline at each site determined. The data (unpublished) were provided by Nolan Hoffman. Abbreviations: Exp, experiment; FC, fold change; S, serine; T, threonine; Y, tyrosine.

Table S5. Phosphosite mapping of human DENND4A in HEK-293E cells (electronic file). Label- free HEK-293E cells overexpressing human DENND4A were serum-starved prior to insulin stimulation (100 nM, 20 min) with or without pre-treatment with the PI3K inhibitor, wortmannin (100 nM, 30 min). 23 DENND4A phosphorylation sites were mapped [Sheet 1]. Four biological replicate experiments were performed [Sheets 2-5] and the log2-transformed median fold change in insulin-stimulated phosphorylation over baseline at each site determined. Abbreviations: Exp, experiment; FC, fold change; S, serine; T, threonine; Y, tyrosine. 211

Table S6. Phosphosite mapping of murine DENND4C in HEK-293E cells (electronic file). Label- free HEK-293E cells overexpressing murine DENND4C were serum-starved prior to insulin stimulation (100 nM, 20 min). 14 DENND4C phosphorylation sites were mapped from a single experiment and the fold change in insulin-stimulated phosphorylation over baseline at each site determined. Abbreviations: FC, fold change; S, serine; T, threonine; Y, tyrosine.

Table S7. Mutagenic primer sequences (next page).

Table S8. Time-lapse TIRF measurements of eGFP-DENND4A species in insulin-stimulated 3T3- L1 adipocytes (electronic file). Adipocytes co-expressing eGFP-DENND4A WT (Sheet1) or 35P mutant (Sheet2) and GLUT4-tagRFP-T were serum-starved for 3 hr prior to insulin stimulation (1 nM, 30 min). Cells were imaged by TIRF microscopy over a time-course of 40 min where insulin was administered at 10 min (grey shaded row). Data are presented as fold response in TIR fluorescence (eGFP) over basal for 35 (DENND4A WT) and 32 (DENND4A 35P) individual cells imaged from two experiments.

Table S9. Time-lapse TIRF measurements of GLUT4-tagRFP-T in insulin-stimulated 3T3-L1 adipocytes (electronic file). Adipocytes co-expressing FLAG-AS160 WT (Sheet1) or 4P mutant (Sheet2) or eGFP-DENND4A WT (Sheet3) or 35P mutant (Sheet4) and GLUT4-tagRFP-T were serum-starved for 3 hr prior to insulin stimulation (1 nM, 30 min). Cells were imaged by TIRF microscopy over a time-course of 40 min where insulin was administered at 10 min (grey shaded row). Data are presented as fold response in TIR fluorescence (tagRFP-T) over basal for 36 (AS160 WT), 38 (AS160 4P), 35 (DENND4A WT) and 32 (DENND4A 35P) individual cells imaged from two experiments. Abbreviations: Exp, experiment.

Table S10. Lipid droplet content in wild-type and Rab10 knockout MEFs (electronic file). Wild- type (WT) or Rab10 knockout (Rab10 −/−) mouse embryonic fibroblasts (MEFs) were incubated in the presence of 200 μM oleate overnight and then subjected to 30 hr starvation in low serum medium. Lipid droplets were visualised by Oil Red-O staining and lipid droplet area in WT and Rab10 −/− cells post-oleate loading ('Loaded') and post-starvation ('Starved30hr') was quantified as described in 5.3: Methods. 189 (WT, Loaded), 155 (WT, Starved30hr), 107 (Rab10 −/−, Loaded) and 123 (Rab10 −/−, Starved30hr) cells were examined from 4-8 coverslips per condition.

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Primer name Primer sequence

hsDENND4A_S1015A.F 5'-GCTTCAGGAAAAGACATAAAGCTGACAATGAAACTAATTTGCAGCAGCAAGTGG-3' hsDENND4A_S1015A.R 5'-CCACTTGCTGCTGCAAATTAGTTTCATTGTCAGCTTTATGTCTTTTCCTGAAGC-3'

hsDENND4A_S1035A.F 5'-GGAAATAGAAACCGTAATCTTGCTGGAGGGGTACTGATGG-3' hsDENND4A_S1035A.R 5'-CCATCAGTACCCCTCCAGCAAGATTACGGTTTCTATTTCC-3' hsDENND4A_S1251A.F 5'-GGCATGCAGAAGATCTGCTCTGCCTCCTAATTCTCC-3'

hsDENND4A_S1251A.R 5'-GGAGAATTAGGAGGCAGAGCAGATCTTCTGCATGCC-3’

directed mutagenesis directed - hsDENND4A_S1281A.F 5'-GGGATAGACTTTGGGCTTCTCCAGCCTTCTC-3' hsDENND4A_S1281A.R 5'-GAGAAGGCTGGAGAAGCCCAAAGTCTATCCC-3'

hsDENND4A_S1511A/T151A.F 5'-GTCAAGCCCAGCAGCAGAAAATATGCAC-3'

DENND4A site DENND4A hsDENND4A_S1511A/T1512A.R 5'-GTGCATATTTTCTGCTGCTGGGCTTGAC-3'

mmDENND4C−MABP.F 5'-GGCATGTGGGGTTCCAACGTGTAATAGTGTTACAAGAAGTCTGTGCC-3' mmDENND4C−MABP.R 5'-GGCACAGACTTCTTGTAACACTATTACACGTTGGAACCCCACATGCC-3'

mmDENND4C−ΔC-terminus.F 5'-GATTTTTATTCGTTTCATTGAAGAATGATGATTTGTAAGTGATAAAGATACTGGATTGG-3'

DENND4C DENND4C truncation truncation mmDENND4C−ΔC-terminus.R 5'-CCAATCCAGTATCTTTATCACTTACAAATCATCATTCTTCAATGAAACGAATAAAAATC-3'

Table S7. Mutagenic primer sequences Primers listed were used for site-directed mutagenesis of human DENND4A and truncation of murine DENND4C. Abbreviations: A, alanine; C, carboxy; Δ, deletion; .F, forward primer; hs, Homo sapiens; MABP, MVB12-associated β-prism; mm, Mus musculus; NLS, nuclear localisation signal; .R, reverse primer; S, serine.

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