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Characterization of novel regulatory components in the dynamic protein palmitoylation cycle Lin, David Tse Shen 2015

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CHARACTERIZATION OF NOVEL REGULATORY COMPONENTS IN THE DYNAMIC PROTEIN PALMITOYLATION CYCLE by  David Tse Shen Lin  M.Sc., McGill University, 2008 B.Sc., The University of British Columbia, 2004  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Medical Genetics)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  March 2015  © David Tse Shen Lin, 2015   ii Abstract  Protein palmitoylation represents the only reversible lipid modification in the cell. As a post-translational modification, it is highly dynamic and plays an important role in protein trafficking and localization. Two families of enzymes mediate dynamic palmitoylation: palmitoyl-acyl transferases (PATs) catalyze palmitate addition, and acyl-protein thioesterases (APTs) catalyze palmitate removal. In mammalian cells, twenty-three PATs have been identified; however, the mechanisms that regulate their enzymatic activity are largely unexplored. Only two APTs, APT1 and APT2, have been identified, but it is unclear if these enzymes act constitutively on all palmitoylated proteins, or if additional depalmitoylases exist. To determine if APT1 and APT2 are responsible for the depalmitoylation of all cytosolic substrates, in this dissertation, I first examined the roles of APT1 and APT2 in protein depalmitoylation. Using a dual pulse-chase strategy to compare protein and palmitate half-lives, I found that simultaneous knockdown or inhibition of APT1 and APT2 strongly blocked palmitate removal from the N-terminal domain of Huntingtin (N-HTT), but had no effect on the depalmitoylation of post-synaptic density-95 (PSD-95). By activity-based protein profiling (ABPP), I showed that the APT1/2 inhibitor Palmostatin B has additional serine hydrolase targets that may play a role in PSD-95 depalmitoylation. Moreover, Palmostatin B induced PSD-95-GFP re-distribution in COS-7 cells, a phenotype not observed with APT1- and APT2-selective inhibitors. These results demonstrate that serine hydrolases other than APT1 and APT2 mediate the substrate-specific removal of palmitate from cytosolic proteins. I also investigated a possible novel regulator of the PAT HTT-interacting protein 14 (HIP14), Optineurin (OPTN), a cargo adaptor known to interact with HTT to   iii mediate post-Golgi vesicle trafficking. I validated this interaction by co-immunoprecipitation and showed that OPTN is not a palmitoylated substrate. Furthermore, HIP14, OPTN, and HTT formed a trimeric complex. I mapped the binding of OPTN and HTT to the HIP14 ankyrin repeat domain, and identified mutations that selectively destabilized the HIP14/OPTN interaction. I hypothesize that OPTN transports HIP14 to distinct subcellular compartments to regulate its access to substrates. In summary, these results reveal potential novel regulatory components in the dynamic palmitoylation cycle.    iv Preface Chapter 1 Portions of Chapter 1 will be published in the journal Biochemical Society Transactions. Lin, D.T. and Conibear, E. (2015) Enzymatic Protein Depalmitoylation by Acyl Protein Thioesterases. Biochem. Soc. Trans. 43:193-198. As first author, I designed the outline of the review and wrote the manuscript.  Chapter 2 A version of Chapter 2 is being prepared for submission (Lin, D.T. and Conibear, E., “APT1- and APT2-indpendent depalmitoylation”). As first author, I designed and conducted all the experiments in this manuscript. I was responsible for making the figures and writing the paper.  Chapter 3 Part of Chapter 3 has been published (Figure 3.1) in Human Molecular Genetics. Butland, S.L., Sanders, S.S., Schmidt, M.E., Riechers, S.P., Lin, D.T., Martin, D.D.O., Vaid, K., Graham, R.K., Singaraja, R.R., Wanker, E.E., Conibear, E., and Hayden, M.R. (2014) The Palmitoyl Acyltransferase HIP14 Shares a High Proportion of Interactors with Huntingtin: Implications for a Role in the Pathogenesis of Huntington’s Disease. Hum. Mol. Genet. 23:4142-4160. As the fifth author, I was responsible for conducting the OPTN/HIP14 co-immunoprecipitation experiment and OPTN palmitoylation experiment, data analyses, and generating figures. Currently, the rest of Chapter 3 is based on the follow-up work from Figure 3.1 that I’m leading, which includes contributions from Matt Tinney, Mike Davey,   v Fuqiang Ban, and Jordan Shimell. I designed the project and, cloned constructs, carried out co-immunoprecipitation assays, and analyzed data. Matt Tinney and Mike Davey conducted the yeast-two-hybrid studies. Fuqiang Ban was responsible for identifying HIP14 protein-protein interaction sites and providing advice for structural-guided mutations. Jordan Shimell is currently analyzing the trafficking and localization of the HIP14 ANKRD mutants identified in this study.  In Chapters 2 and 3, I consistently use “we” to reflect the co-authors that contributed to the studies.   vi Table of Contents 	  Abstract ................................................................................................................................... ii	  Preface .................................................................................................................................... iv	  Table of Contents .................................................................................................................. vi	  List of Tables ........................................................................................................................ xii	  List of Figures ...................................................................................................................... xiii	  List of Abbreviations ........................................................................................................... xv	  Acknowledgements ........................................................................................................... xviii	  Dedication ............................................................................................................................. xx	  Chapter 1: Introduction ........................................................................................................ 1	  1.1	   Overview of Protein Palmitoylation ........................................................................... 1	  1.1.1	   General Introduction to Palmitoylation ............................................................... 1	  1.1.2	   Strategies to Assess Substrate Palmitoylation ..................................................... 3	   in silico Prediction of Palmitoylation Sites ................................................... 3	   Protein Localization Studies ......................................................................... 4	   Quantitative Measurement of Protein Palmitoylation ................................... 5	   Metabolic Labeling with Fatty Acid Analogues .................................... 5	   Acyl-Biotin Exchange (ABE) ................................................................ 7	   Inhibiting Palmitoylation with 2-Bromopalmitate (2-BP) ............................ 8	   Large-Scale & Proteomic Approaches .......................................................... 9	  1.1.3	   Basic Biological Functions of Palmitoylation ..................................................... 9	   Protein Sorting and Trafficking .................................................................. 10	    vii	   Association with Membrane Domains ........................................................ 11	   Protein-Protein Interactions ........................................................................ 11	   Membrane Tilting of Transmembrane Proteins .......................................... 12	  1.1.4	   Interplay with Other Post-Translational Modifications ..................................... 12	   Palmitoylation and Nitrosylation Compete to Regulate Protein Localization   ..................................................................................................................... 12	   Palmitoylation and Phosphorylation act as Cellular Pathway “Switches” . 13	   Palmitoylation Shields Substrate from Ubiquitin-Mediated Degradation .. 14	  1.1.5	   Dynamics of Protein Palmitoylation .................................................................. 15	  1.2	   Palmitoyl-Acyl Transferases: Enzymes that Mediate Palmitate Addition ............... 18	  1.2.1	   Identification of Palmitoyl-Acyl Transferases (PATs) ...................................... 18	  1.2.2	   Mechanism of PAT Activities ........................................................................... 20	  1.2.3	   Overlapping Substrate Specificity of PATs ....................................................... 21	  1.2.4	   Modulation of PAT Activities ........................................................................... 24	   Co-Factors can Modulate DHHC Activity ................................................. 24	   Post-Translational Modifications of DHHC Proteins ................................. 25	   Differential Endogenous Expression Levels of DHHC Proteins ................ 26	   Oligomerization as a Possible Mechanism of Regulating DHHC Activity 26	   Subcellular Trafficking of DHHC Proteins ................................................ 27	  1.2.5	   Involvement of DHHC Proteins in Neurodegenerative Diseases ...................... 28	   Cancer ......................................................................................................... 28	   Mental Retardation ...................................................................................... 29	   Schizophrenia .............................................................................................. 29	    viii	   Huntington’s Disease (HD) ......................................................................... 30	  1.3	   Enzymes that Mediate Palmitate Removal ............................................................... 32	  1.3.1	   Palmitoylthioesterases are Members of the Metabolic Serine Hydrolase (mSH) Family  ............................................................................................................................ 32	   Luminal Protein Depalmitoylation by Palmitoyl-Protein Thioesterase 1 (PPT1)  ..................................................................................................................... 34	   Cytosolic Protein Depalmitoylation by Acyl-Protein Thioesterases (APTs) .   ..................................................................................................................... 35	   Acyl-Protein Thioesterase 1 (APT1) ................................................... 35	   Acyl-Protein Thioesterase 2 (APT2) and APT1-Like (APT1L) .......... 37	  1.3.2	   Modulation of Protein Depalmitoylation ........................................................... 38	  1.3.3	   Evaluating APT Activity Using Genetic and Small Molecule Approaches ...... 39	   Using Genetic Manipulation to Study APT Activity .................................. 40	   Using Small Molecules and Chemical Biology to Study APT Activity ..... 40	   Verification of Small Molecule Targets by Activity-Based Protein Profiling (ABPP) .................................................................................................... 41	   Palmostatin B (PalmB) ........................................................................ 43	   Hexadecylfluorophosphonate (HDFP) ................................................. 43	   Piperazine Amide Compounds ............................................................ 44	  1.4	   Current Paradigm of PAT/APT-Directed Dynamic Cycling .................................... 45	  1.4.1	   Spatial Modulation of APT-Mediated Depalmitoylation .................................. 45	  1.4.2	   Current Issues with the Model ........................................................................... 47	   Evidence for Activity-Dependent Palmitoylation and Depalmitoylation ... 48	    ix	   Compartmentalized PAT/APT Activities ................................................... 49	  1.4.3	   Concluding Remarks .......................................................................................... 49	  1.5	   Thesis Goals and Objectives ..................................................................................... 50	  Chapter 2: APT1/APT2-Independent Protein Depalmitoylation .................................... 52	  2.1	   Introduction ............................................................................................................... 52	  2.2	   Materials and Methods .............................................................................................. 54	  2.2.1	   cDNAs & siRNAs .............................................................................................. 54	  2.2.2	   Chemicals ........................................................................................................... 57	  2.2.3	   Cell Culture Conditions ..................................................................................... 57	  2.2.4	   cDNA & siRNA Transfections .......................................................................... 57	  2.2.5	   Small Molecule Inhibitor Treatments ................................................................ 58	  2.2.6	   Pulse-Chase Metabolic Labeling with Inhibitors ............................................... 58	  2.2.7	   Immunoprecipitations ........................................................................................ 59	  2.2.8	   Sequential On-Bead CuAAC/Click Chemistry .................................................. 60	  2.2.9	   Competitive Activity-Based Protein Profiling ................................................... 60	  2.2.10	   In-Gel Fluorescence Analyses ........................................................................... 61	  2.2.11	   Western Blotting ................................................................................................ 61	  2.2.12	   PSD-95-GFP Localization by Microscopy & Cell Counting ............................ 62	  2.2.13	   Statistical Analyses ............................................................................................ 62	  2.3	   Results ....................................................................................................................... 63	  2.3.1	   Palmitate Turnover is Substrate-Specific ........................................................... 63	  2.3.2	   APT1 and APT2 Display Substrate Specificity ................................................. 67	  2.3.3	   Palmostatin B (PalmB) has Additional Serine Hydrolase Targets .................... 69	    x 2.3.4	   ABHD6, FASN, PNPLA6, and PNPLA8 do not Play Major Roles in PSD-95 Depalmitoylation ............................................................................................................ 73	  2.3.5	   Tetrahydrolipstatin (THL) Treatment Results in Partial Inhibition of PSD-95 Depalmitoylation ............................................................................................................ 75	  2.3.6	   Inhibition of PSD-95 Depalmitoylation Alters its Localization ........................ 78	  2.4	   Discussion ................................................................................................................. 79	  Chapter 3: The Huntingtin-Binding Partner Optineurin is a Novel Interactor of the PAT HIP14 ........................................................................................................................... 85	  3.1	   Introduction ............................................................................................................... 85	  3.2	   Materials and Methods .............................................................................................. 88	  3.2.1	   Mammalian cDNAs ........................................................................................... 88	  3.2.2	   Chemicals ........................................................................................................... 90	  3.2.3	   Cell Culture Conditions ..................................................................................... 90	  3.2.4	   cDNA Transfections .......................................................................................... 91	  3.2.5	   Co-Immunoprecipitations .................................................................................. 91	  3.2.6	   Yeast-Two-Hybrid ............................................................................................. 92	  3.2.7	   Metabolic Labeling and CuAAC/Click Chemistry ............................................ 92	  3.2.8	   HIP14 Ankyrin Repeat Domain Structure Analysis .......................................... 93	  3.2.9	   Statistical Analyses ............................................................................................ 94	  3.3	   Results ....................................................................................................................... 94	  3.3.1	   HIP14 Interacts with, but Does not Palmitoylate, OPTN .................................. 94	  3.3.2	   HIP14 Interaction with OPTN is Enhanced by the Presence of HTT ................ 96	  3.3.3	   OPTN Interacts with HTT via its C-Terminal Domain ..................................... 98	    xi 3.3.4	   Mutation of the HIP14 Ankyrin Repeat Domain (ANKRD) Causes Selective Loss of the OPTN Interaction ...................................................................................... 101	  3.4	   Discussion ............................................................................................................... 103	  Chapter 4: Discussion & Conclusions .............................................................................. 109	  4.1	   Summary of Major Findings ................................................................................... 109	  4.2	   Functional Redundancy in APTs ............................................................................ 110	  4.3	   The Role of OPTN in Regulating HIP14 Trafficking and Function ....................... 114	  4.4	   Conclusions ............................................................................................................. 119	  Bibliography ....................................................................................................................... 121	  Appendix ............................................................................................................................. 146	  Appendix A Gene Fragments for cloning of HIP14 ANKRD mutants (5’ > 3’). ............ 146	     xii List of Tables  Table 2.1 List of Oligos Used for Cloning mSH Constructs. ................................................. 56	  Table 2.2 HDFP-targeting metabolic serine hydrolases. ........................................................ 72	  Table 3.1 Cloning Oligos and Methods Used to Generate the Constructs for This Study. .... 90	     xiii List of Figures  Figure 1.1 Schematic Illustration of the Dual Click Chemistry Assay to Simultaneously Monitor Protein and Palmitate Turnover. ................................................................................. 6	  Figure 1.2 Schematic Illustration of the Acyl biotin Exchange (ABE) Assay. ........................ 7	  Figure 1.3 Palmitoylation Regulates Synaptic Transmission. ................................................ 16	  Figure 1.4 Phylogentic Clustering of the Human and Saccharomyces cerevisiae DHHC Proteins Based on ClustalX Alignment of the 51 Amino Acid DHHC Core Sequence. ........ 20	  Figure 1.5 Dendrogram Showing all 128 Members of the Mouse Metabolic Serine Hydrolase Family with Branch Length Depicting Sequence Relatedness. .............................................. 33	  Figure 1.6 Schematic Illustration of Competitive Activity-Based Protein Profiling (ABPP). 42	  Figure 1.7 Spatiotemporal Model of Dynamic Palmitoylation Cycle for the Model Substrate Ras. .......................................................................................................................................... 46	  Figure 2.1 Palmitoylation Dynamics Differ between Substrates. ........................................... 65	  Figure 2.2 APT1 and APT2 Inhibition Inhibits N-HTT, but not PSD-95 Palmitate Turnover. ................................................................................................................................................. 68	  Figure 2.3 Palmostatin B and HDFP Inhibit PSD-95 and N-HTT Palmitate Turnover and Share Overlapping Serine Hydrolase Targets. ........................................................................ 71	  Figure 2.4 Treatment with WWL70, C75, BEL, and RHC-80267 Does not Affect PSD-95 Palmitate Turnover. ................................................................................................................. 75	  Figure 2.5 Tetrahydrolipstatin Inhibits BAT5 and ABHD6 and Causes Partial Inhibition of PSD-95 Depalmitoylation. ...................................................................................................... 77	    xiv Figure 2.6 Treatment with Inhibitors Causing PSD-95 Palmitoylation Changes Also Leads to Localization Shift of PSD-95-GFP. ........................................................................................ 79	  Figure 3.1 OPTN Interacts with HIP14 but is not a Palmitoylated Substrate. ........................ 96	  Figure 3.2 Expression of Exogenous HTT Enhances the Interaction between HIP14 and OPTN. ..................................................................................................................................... 98	  Figure 3.3 An Intact OPTN C-Terminus is Important for its Interaction with HTT and HIP14. ............................................................................................................................................... 100	  Figure 3.4 Mutations in HIP14 Ankyrin Repeats Region Selectively Destabilizes Interaction with OPTN but Maintains Interaction with HTT. ................................................................. 102	  Figure 3.5 Wild-Type HIP14 can be Found on Golgi Outposts in Cultured Primary Rat Neurons. ................................................................................................................................ 103	     xv List of Abbreviations 17-ODYA 17-octadecynoic acid 2-BP 2-Bromopalmitate AADACL1 Arylacetamide deacetylase-like 1 ABE Acyl biotin exchange ABHD Abhydrolase domain containing ABIN A20-binding inhibitor of nuclear factor-κB proteins  ABPP Activity-based protein profiling ACOT Acyl-CoA thioesterase  AF Alexa Fluor    ANKRD Ankyrin repeats domain AMPAR α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptors ALS Amyotrophic lateral sclerosis APEH Acylaminoacyl-peptide hydrolase APP Amyloid precursor APT Acyl Protein thioesterase APT1 Acyl protein thioesterase 1 (LYPLA1) APT2 Acyl protein thioesterase 2 (LYPLA2) APT1L Acyl Protein thioesterase-like 1 (LYPLAL1) ASO Antisense oligonucleotide BACE1 β-site amyloid precursor protein (APP) cleaving enzyme-1 BAT5 Human lymphocyte antigen B-associated transcript 5 BDNF Brain-derived neurotrophic factor BEL Bromoenol lactone BK Large conductance calcium-activated potassium cABPP Competitive ABPP Cbl Casitas B-lineage lymphoma CD-M6PR Cation-dependent mannose-6-phosphate receptor CHO Chinese hamster ovary COS Cell CV-1 (simian) in origin and carrying the SV40 genetic material CSP Cysteine string protein (DNAJC5) CUAAC Cu(I)-catalyzed azide-alkyne cycloaddition DHHC Asp-His-His-Cys DHHC-CRD DHHC cysteine-rich domain DIC Dynein intermediate chain DPP Dipeptidyl peptidase ER Endoplasmic reticulum FAAH Fatty acid amide hydrolase FAM108 Family with sequence similarity 108 FASN Fatty acid synthase (FAS) FKBP12 12kDa FK506-binding protein FP Fluorophosphonate FP-Rho Fluorophosphonate-rhodamine FRAP Fluorescence recovery after photobleaching   xvi GAD65 Glutamate decarboxylase 65 GAP43 Growth associated protein 43 GM130 Cis-Golgi matrix protein 130 (GOLGA2) GOLIM4 Golgi phosphoprotein 4 GRIP1b Glutamate receptor interacting protein-1b GWAS Genome-wide association study HAM Hydroxylamine HAP Huntingtin-associated protein HD Huntington Disease HIP14 Huntingtin interacting protein 14 (ZDHHC17) HTT Huntingtin interacting protein 14-like (ZDHHC13) INCL Infantile neuronal ceroid lipofuscinosis ITM2B Integral membrane protein 2B JNK3 c-Jun N-terminal kinase 3 L0 Liquid-ordered Lck Lymphocyte-specific protein tyrosine kinase L-AHA L-Azidohomoalanine LIPA Lipase A, lysosomal acid, cholesterol esterase LIPE Hormone sensing lipase (HSL) LRP6 Lipoprotein-related protein 6 LYPLA3 Phospholipase A2, Group XV (APT3 / PLA2G15) MAFP Methylarachnidonyl fluorophosphonate MAGL Monoacylglycerol lipase MBOAT Membrane-bound-O-acyltransferases mHTT Mutant HTT (128 CAG expansions used in this thesis) MS Mass spectrometry mSH Metabolic serine hydrolase NEM N-ethylmaleimide NEMO Nuclear factor-κB essential modulator N-HTT Huntingtin N-terminal domain (amino acids 1-548 in this study) NMDAR N-methyl-D-aspartate receptors NOS NO synthase enzymes OPTN Optineurin ORF Open reading frame PAFAH2 Platelet-activating factor acetylhydrolase type II (PAFAH2) PARL Presenilin-associated rhomboid-like protease (PSARL) PAT Palmitoyl acyl transferase PCR Polymerase chain reaction PDE6δ Phosphodiesterase 6 delta subunit PDE10A2 Phosphodiesterase 10A isoform 2 PDZ PSD-95/Discs-large/ZO-1 homology PGAP1 Post-GPI attachment protein 1 (PGAP1) PICK1 Protein interacting with C-kinase PKA Protein kinase A PKC Protein kinase C   xvii PM Plasma membrane PNPLA Patatin-like phospholipase PPT Protein palmitoyl thioesterase PREP Prolyl endopeptidase-like PSD Post synaptic density PSD-95 Post-synaptic density-95 (DLG4) PTM Post-translational modification PV Pervanadate R7BP R7 regulator of G protein signaling (RGS)-binding protein Rho Rhodamine RISC RNA-induced silencing complex RNAi RNA interference SEM Standard error of the mean SERHL2 Serine hydrolase-like 2 SILAC Stable isotope labeling by amino acids in culture STREX BK Stress axis regulated exon-containing BK Channel SNAP23 Synaptosomal-associated protein 23 SNAP25 Synaptosomal-associated protein 25 SNARE Soluble N-ethylmaleimide-sensitive-factor attachment protein receptor SNP Single nucleotide polymorphism SOD1 Cu/Zn superoxide dismutase (SOD1) SPRED2 Sprouty-related, EVH1 domain-containing protein 2 TBTA Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine  TCEP Tris(2-carboxyethyl)phosphine hydrochloride TEA Triethanolamine TEM8 Tumor endothelial marker 8 THL Tetrahydrolipstatin TIRFM Total internal reflection fluorescence microscopy TMD Transmembrane domain UBAN Ubiquitin binding in ABINs and NEMO XLMR X-linked mental retardation   xviii Acknowledgements  First and foremost, I would like to thank my research supervisor, Dr. Elizabeth Conibear. Thank you, Liz, for being so supportive and allowed me some independence on research. I know, sometimes it had been a crapshoot, but I appreciate you believing in me in general and letting me take control. This was certainly one of the most intense but rewarding learning experience for me. I would also like to thank my supervisory committee members: Dr. Lynn Raymond, Dr. Chris Loewen, and Dr. Stefan Taubert for the guidance and genuine support along the way. I could not ask for a better committee. A special thank you to Dr. Nicholas Davis for providing honest feedback and support on my research progress. Also thank you, to Dr. Michael Kobor, for acknowledging my work and just being a cool guy.  To dear members of the Conibear lab, thank you for keeping my sanity in check and not rejecting (and sometimes even accepting) my pessimism and cynicism. Thank you, Mike Davey, for keeping us hooligans in check and being the most resourceful and helpful person I know, especially with your expertise in yeast biology and molecular cloning. Thank you to the current members Bjorn Bean, Lauren Dalton, Shawn Whitfield, and my awesome bench buddy Kathleen Kolehmainen for all your friendship and support along the way. Thank you to past members Karen Lam, Cayetana Schluter, Helen Burston and Lindsay Rogers for teaching me a thing or two about protein trafficking and proteomics. Last but not least, to my graduation buddy Tejasvi Dharwada: yay girl, we did it! Thanks for being so very supportive during the home stretch and also congratulations on your degree!   xix I have made some incredible friends during my 6 years in CMMT and medical genetics. I’d especially like to thank Phoebe Lu, for being a supportive friend to me during hardships. I’d also like to thank Alexandre Lussier, who has also been nothing but awesome. As well I’d like to thank Shaun Sanders, Dr. Grace Leung, Dr. Alice Wang, Grace Thamarajah, Joanna Yeung, and Dr. Anna Poon for their friendship. Last but not least, I’d like to thank Seetha Kumaran for being a sweetheart who cheers me up when I am down.  I’d like to thank my family for always believing in me and putting up with me being grumpy. Thank you dad for always checking up on me and thank you mom for always caring. Thank you to my sibs as well, Jessica and Eden. I love you with all my heart.    xx Dedication ???????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????????  1 Chapter 1: Introduction1   1.1 Overview of Protein Palmitoylation  1.1.1 General Introduction to Palmitoylation Fatty acylation refers to the covalent modification of proteins with a variety of lipids (Resh, 2013). As this increases hydrophobicity of the protein, fatty acylation promotes association with the lipid bilayer and serves to modulate protein localization and function (Nadolski and Linder, 2007). The three most common lipid modifications that take place in the cytoplasm are prenylation, N-myristoylation, and palmitoylation (also referred to as S-acylation) (Resh, 1999). Whereas prenylation and the co-translational N-myristoylation are irreversible (Zha et al., 2000; Wright and Philips, 2006), palmitoylation is the only reversible post-translational modification (PTM) among lipid modifications (Resh, 1999; Smotrys and Linder, 2004). The reversible nature of palmitoylation means that it can be modulated by specific signaling pathways (Neve et al., 2003; Smotrys and Linder, 2004). Therefore, addition and removal of palmitate may provide an important tool for dynamically regulating responses to diverse cellular stimuli. Indeed, research within the last two decades has demonstrated the role of palmitoylation in a wide variety of biological functions.                                                  1 Portions of this chapter will be published in Biochemical Society Transactions. Lin, DT and Conibear, E. (2015) “Enzymatic Protein Depalmitoylation by Acyl Protein Thioesterases” Biochem. Soc. Trans. 43:193-198.   2 Protein S-palmitoylation occurs when a 16-carbon saturated fatty acid chain is added to cytosolic cysteine residues via a thioester bond (Greaves and Chamberlain, 2007). Although palmitate is the most common fatty acid chain to be added, this modification may also involve the addition of other fatty acids including palmitoyleate (16:1), stearate (18:0), oleate (18:1), arachidonate (20:4), and eicosapentaenoate (20:5) (Liang et al., 2001). Both soluble and transmembrane proteins can be palmitoylated. Although the lack of consensus motifs for this PTM has hampered the discovery of palmitoylation sites, these sites share some general features (Salaun et al., 2010). Cytosolic proteins that already possess a single lipid modification, such as N-terminal myristoylation (consensus MGXXX(S/T), where one of the X is a cysteine residue) or C-terminal prenylation (CAAX motif), are often palmitoylated on a proximal cysteine. Such dual lipidation provides a mechanism to stably tether these proteins to membranes (Hannoush and Sun, 2010). For other proteins lacking another lipid attachment, protein palmitoylation usually either occurs in pairs on adjacent cysteines or near basic sequences that facilitate interactions with the negatively charged acidic phospholipid groups (Heo et al., 2006; Gauthier-Campbell et al., 2004). Transmembrane proteins are often palmitoylated on a cysteine residue near the transmembrane domain (TMD) (Blaskovic et al., 2013).  Occasionally, two other types of palmitate modifications are found in cells: N-palmitoylation and O-palmitoylation (Shipston, 2011). In N-palmitoylation, the palmitate is attached via an amide linkage to the N-terminal cysteine by a family of enzymes referred to as the membrane-bound-O-acyltransferases (MBOATs) (Buglino and Resh, 2008). In O-palmitoylation, the palmitate is attached via an oxyester linkage to a Serine residue by the   3 enzyme ghrelin O-acyltransferase (Yang et al., 2008). Because these modifications are rare and irreversible, they will not be discussed in this dissertation.  Here, I will first discuss strategies that have been used to predict and assess substrate palmitoylation, then I will highlight some important physiological functions of protein palmitoylation and the interplay between palmitoylation and other PTMs. Finally, I will focus on established biological examples of dynamic palmitoylation and its importance in the central nervous system.  1.1.2 Strategies to Assess Substrate Palmitoylation in silico Prediction of Palmitoylation Sites Due to the lack of general consensus motifs, conventionally, palmitoylation sites are generally mapped by mutagenesis of candidate cysteine residues, which is labour-intensive and time-consuming. Therefore, in silico prediction tools have been developed to assist identification of potential palmitoylation sites. The first of these tools, termed CSS-Palm, uses a clustering and scoring strategy (CSS) algorithm based on a large training dataset of known palmitoylated proteins (Zhou et al., 2006; Ren et al., 2008). However, this approach may fail to detect sites that do not conform to motifs observed in previously identified substrates. Newer algorithms have been designed with this caveat in mind; for example, IFS-Palm uses the incremental feature selection method, which takes into account the amino acid physiochemical factors, conservation, and disorder (Hu et al., 2011). WAP-Palm further included evolutionary information and structural characteristics as prediction parameters (Shi   4 et al., 2013). Both IFS-Palm and WAP-Palm reported better sensitivity, specificity, and accuracy. Protein Localization Studies As palmitoylation increases the hydrophobicity of a protein, it is expected to increase the affinity of the protein to membranes. Indeed, palmitoylation often leads to change in subcellular localization as determined by microscopy or subcellular fraction techniques (Magee et al., 1987; Huang et al., 2004; Butland et al., 2014). This approach has been used in many studies to establish altered palmitoylation levels (Tomatis et al., 2010; Kong et al., 2013). Other studies focusing on the palmitoylation dynamics have used fluorescence recovery after photobleaching (FRAP), where fluorescently-tagged substrates are irreversibly bleached in a defined region of interest, and the exchange between bleached and unbleached molecules is monitored over time as a readout for palmitate turnover (Kenworthy, 2006). Notably, FRAP has been used to monitor shuttling of palmitoylated Ras isoforms between the Golgi and plasma membrane (Rocks et al., 2005), as well as the partitioning of GAD65 (Glutamate decarboxylase 65) between the Golgi and post-Golgi vesicles (Kanaani et al., 2008). However, because a single palmitate modification is not sufficient to confer stable membrane association (Conibear and Davis, 2010), trafficking and localization can be influenced by many factors and does not necessarily reflect changes in palmitoylation state. To truly measure protein palmitoylation, more quantitative approaches must be employed, as discussed in the next section.     5 Quantitative Measurement of Protein Palmitoylation Metabolic Labeling with Fatty Acid Analogues Historically, protein palmitoylation was quantified by metabolic labeling using the radioactive palmitate analogue [3H]-palmitate (or [125I]-palmitate) in cultured cells (Berthiaume et al., 1995; Resh, 2006). An added advantage of metabolic labeling is the ability to perform pulse-chase studies to determine palmitate cycling time on proteins, which has been done for a handful of substrates. Although this method provides a direct measure of palmitoylation, handling of radioactive analogues is cumbersome, the analogues are potentially toxic to cells during prolonged treatment, and signal detection time is usually long, making this technique insufficiently sensitive for most studies.  A more sensitive method exploits probes developed for use in bio-orthogonal reactions, including the Staudinger ligation and the copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC)/click chemistry (Sletten and Bertozzi, 2011). In this method, non-radioactive, bio-orthogonal alkyne- or azido-containing palmitate analogues are first added to label proteins in cultured cells (Hang et al., 2007; Kostiuk et al., 2008; Martin and Cravatt, 2009; Charron et al., 2009). Bio-orthogonal reactions are then performed to conjugate these analogues to alkyne- or azido-tagged reporters such as biotin or fluorophores to allow assessment of protein palmitoylation by Western blotting, mass spectrometry, in-gel fluorescence, or cellular imaging. The resulting signals from these labeled analogues are relatively strong compared to radioactive palmitate. In addition, simultaneous labeling with clickable palmitate and the methionine surrogate L-azidohomoalanine (L-AHA) has been used to control for the rate of protein degradation, which could otherwise confound measures of   6 palmitate turnover (Zhang et al., 2010). I have adapted this approach in this dissertation to study dynamic palmitate turnover of substrates (Figure 1.1). Taken together, these features make click chemistry an appealing approach for studying protein palmitate turnover.    Figure 1.1 Schematic Illustration of the Dual Click Chemistry Assay to Simultaneously Monitor Protein and Palmitate Turnover. In dual click chemistry, proteins in cultured cells are simultaneously metabolically labeled with the palmitic acid analogue 17-octadecynoic acid (17-ODYA) and the methionine analogue L-azidohomoalanine (L-AHA). Palmitate turnover dynamics can then be studied using a pulse/chase method. Following cell lysis, sequential click chemistry is performed. The first azide-containing compound (denoted by star) would only react with the alkyne group on 17-ODYA, thus tagging palmitate turnover; the second alkyne-containing compound (denoted by hexagon) will only react with the azide terminal of L-AHA, thus providing a readout for protein turnover.     However, metabolic labeling with fatty acid analogues is not the best approach under some circumstances. For instance, labeling is only feasible in live cells and less practical for studying animal tissues in vivo. Another potential caveat is that the labeling is limited by the rate of protein synthesis and/or palmitate turnover of substrate of interest. This means that proteins with slower palmitate turnover may not label well using this method unless an extended labeling period is used. Metabolic labeling is also not suitable when detection of the total pool of palmitoylated proteins is desired.  N317-ODYA (Palmitic acid analogue)L-AHA(Methionine analogue) MetCCCCProteinturnoverPalmitateturnover1. Pulse/ChaseN3+ N32. On-beadclick chem.+IN-GEL FLUORESCENCE  7 Acyl-Biotin Exchange (ABE) In 2004, Drisdel and Green developed a method that allows for the quantitation of the total pool of palmitoylated proteins present in cultured cells and animal tissue extracts (Drisdel and Green, 2004). In this method, now widely known as acyl-biotin exchange (ABE), cellular lysates or tissue extracts containing the protein of interest are treated in three sequential steps to assess substrate palmitoylation: (1) Addition of N-ethylmaleimide (NEM) which blocks free thiol-containing cysteines; (2) Treatment with hydroxylamine (HAM) to cleave thioester bonds leading to palmitate release from palmitoylated cysteines; and (3) Labeling of newly exposed thiols with a biotin-based, sulfhydryl-specific labeling compound. Following these reactions, the palmitoylated proteins are detected by Western blotting using streptavidin enrichment (Figure 1.2) (Drisdel and Green, 2004; Wan et al., 2007). Recently, a variation of the ABE assay has been described using resin-assisted capture (Forrester et al., 2011). The method, known as Acyl-RAC, follows the ABE procedure but uses thiol-reactive resin in place of the sulfhydryl-reactive biotinylated reagent in the last step to capture all palmitoylated proteins for downstream mass spectrometry or western blotting analyses.    Figure 1.2 Schematic Illustration of the Acyl biotin Exchange (ABE) Assay. In ABE, three chemical steps are carried out to switch a palmitate group for a biotin group to allow detection of palmitoylated proteins. This involves (1) Blockade of free thiols by N-ethylmaleimide (NEM); (2) Cleavage of thioester-bonded palmitate groups from palmitoylated cysteines by hydroxylamine (HAM); and (3) Labeling of newly exposed thiol groups with a sulfhydryl-reactive compound, such as Biotin-HPDP.  CysCysCysSHSHPalmitate+NEMCysCysCysPalmitateNEMNEM+HAMCysCysCysNEMNEMSH+Biotin-HPDPCysCysCysNEMNEMS SBiotin  8 Although ABE can be used to measure palmitoylation levels at distinct time points, the measurements represent a balance between palmitoylation and depalmitoylation activities and do not reflect the depalmitoylation kinetics of a substrate. Furthermore, results from ABE can sometimes be confounded by false positives that arise from other thioester modifications, such as nitrosylation. Hence, the data needs to be cautiously interpreted. Nevertheless, its high practicality for assessing substrate palmitoylation in animal tissues makes ABE a great complementary approach to metabolic labeling techniques. Inhibiting Palmitoylation with 2-Bromopalmitate (2-BP) A classic approach to demonstrate substrate palmitate turnover is to use the general palmitoylation inhibitor, 2-bromopalmitate (2-BP), when carrying out metabolic labeling, ABE, or cellular localization studies. 2-BP is a non-metabolizable, α-bromo-carboxyl-containing palmitate analogue that blocks palmitate incorporation into proteins (Webb et al., 2000). However, 2-BP is a highly toxic compound (Mikic et al., 2006) and blocks palmitoylation in a non-substrate-specific manner (Coleman et al., 1992). Interestingly, 2-BP was observed to decrease the depalmitoylation rate of lymphocyte-specific protein tyrosine kinase (Lck), suggesting that 2-BP may also inhibit depalmitoylation enzymes (Zhang et al., 2010). Subsequently, two elegant proteomic studies demonstrated that 2-BP has many off-targets. Importantly, 2-BP may directly compete for palmitoylation at select cysteine residues (Zheng et al., 2013; Davda et al., 2013). In addition, a recent report identified depalmitoylation enzymes as 2-BP targets. These findings raise caution against using 2-BP to study palmitoylation (Pedro et al., 2013).    9 Large-Scale & Proteomic Approaches Mass spectrometry (MS) has been used as a method to directly detect protein palmitoylation (Liang et al., 2002; Kordyukova et al., 2008). However, this approach is not quantitative, as palmitoyl loss may occur due the labile nature of thioester linkage, and the large difference in the hydrophobicity between palmitoylated and unpalmitoylated peptides (Ji et al., 2013).  More often, MS approaches are performed downstream of ABE or click chemistry with palmitate analogues (Kang et al., 2008b; Martin and Cravatt, 2009; Yount et al., 2010). To date, many large-scale studies focusing on the palmitoylome have generated potentially useful datasets of palmitoylated proteins in various cell types. Additionally, by altering the expression levels of select palmitoylation enzymes (Li et al., 2012), or inhibiting palmitate turnover (Martin et al., 2011), some large-scale studies have provided a global view of substrate selectivity and dynamics for distinct substrates. Many researchers now use these resources, together with palmitoylation site prediction software, as a starting point to first evaluate the likelihood that their protein of interested is palmitoylated.    1.1.3 Basic Biological Functions of Palmitoylation As a hydrophobic modification, palmitoylation facilitates protein insertion into membranes. However, functional studies in the past decade have provided compelling evidence that rather than simply serving as a membrane anchor signal, palmitoylation plays an especially important role in protein sorting and trafficking, associating proteins to lipid microdomains, and mediating protein-protein interactions.   10  Protein Sorting and Trafficking Perhaps the best-studied example of palmitoylation affecting sorting and trafficking of a substrate is the oncogenic N- and H-Ras proteins. Both N- and H-Ras are prenylated and this irreversible lipidation event allows N- and H-Ras to transiently associate with all cellular endomembranes with unstable kinetics (Conibear and Davis, 2010; Salaun et al., 2010). Post-translational palmitoylation, occurring exclusively on the Golgi where the enzymes responsible for Ras acylation are localized, acts as a “membrane trap” by increasing member avidity of both Ras isoforms, thereby providing a mechanism for stable membrane tethering (Shahinian and Silvius, 1995; Rocks et al., 2010). This Golgi-localized palmitoylation then further serves as a sorting signal, targeting Ras proteins from the Golgi to the plasma membranes (PM) via recycling endosomes (Misaki et al., 2010). The dynamic palmitoylation of Ras is important because it has distinct activity profiles on the PM versus the endomembranes, acting via different signaling pathways to regulate cell proliferation and differentiation (Omerovic and Prior, 2009; Misaki et al., 2010). To further control this signaling gradient, a rapid and ubiquitous “palmitate removal” event (depalmitoylation) has been shown to prevent N-Ras accumulation on the PM and subsequent “spillover” to all cellular membranes (Rocks et al., 2010). Depalmitoylation of Ras allows it to get repalmitoylated at the Golgi, creating a palmitoylation cycle that regulates its trafficking and signaling.  Aside from the Ras proteins, palmitoylation has also been shown to be critical for sorting and trafficking of other transmembrane proteins like the transferrin receptor and sortilin (Alvarez et al., 1990; McCormick et al., 2008).   11  Association with Membrane Domains Palmitoylation has been shown to influence the protein’s ability to associate with lipid rafts, which are functional membrane domains rich in cholesterol and glycosphingolipids (Moffett et al., 2000; Charollais and Van Der Goot, 2009). The lipids within these rafts promote the formation of a “liquid ordered” Lo state that segregates from the bulk lipids in the PM (Simons and Ikonen, 1997). With its unique chain length and its saturated nature, which shows a preference for ordered domain in vitro, palmitate serves as a driving force for proteins to associate with lipid rafts (Uittenbogaard and Smart, 2000; Resh, 2013). This is indeed true for many cytosolic proteins, such as the Src family kinases, which preferentially associate with and signal from rafts upon its palmitoylation (Resh, 2013); however, for transmembrane proteins, palmitoylation has been observed to either positively regulate raft association (i.e. the β-secretase BACE1) (Meckler et al., 2010), negatively regulate raft association (i.e. Tumor Endothelial Marker TEM8), or exert no effect on raft association (i.e. Transferrin receptor) (Blaskovic et al., 2013). Palmitoylation was also documented to target the endoplasmic reticulum (ER)-resident chaperon protein calnexin to the specialized ER-mitochondrial interaction sites and the nuclear envelope (Lynes et al., 2011; Lakkaraju et al., 2012). Protein-Protein Interactions Palmitoylation can also influence the ability of a protein to associate with other proteins. An example is the cation-dependent mannose-6-phosphate receptor (CD-M6PR), which sorts newly synthesized acid-hydrolases from the trans-Golgi to endosomes. CD-M6PR then is recycled following hydrolase delivery in a palmitoylation-dependent manner, as this   12 modification allows the interaction of the cytosolic tail of CD-M6PR with the retromer complex (McCormick et al., 2008). Palmitoylation has also been shown to mediate interactions between tetraspanins and integrins (Zevian et al., 2011). Membrane Tilting of Transmembrane Proteins Originally, it was proposed that palmitoylation of juxtamembrane cysteines induces TMD tilting (Joseph and Nagaraj, 1995). Since the ER bilayer membrane is thinner than the plasma membrane, TMD tilting minimizes hydrophobic mismatch and promotes substrate ER exit. This has been suggested for the yeast chitin synthase Chs3 (Lam et al., 2006) and demonstrated for the mammalian lipoprotein-related protein 6 (LRP6) (Abrami et al., 2008).   1.1.4 Interplay with Other Post-Translational Modifications Aside from the commonly observed myristoylation and prenylation modifications on soluble palmitoylated proteins, some studies have highlighted the potential interplay of palmitoylation with other PTMs. Here, I highlight recent examples where palmitoylation may affect protein phosphorylation, nitrosylation, ubiquitination, and vice versa, as a way to modulate substrate localization, function, and stability. Palmitoylation and Nitrosylation Compete to Regulate Protein Localization In addition to being palmitoylated, cysteine residues can also be oxidized, nitrosylated, or glutathionylated (Zhou et al., 2014). A notable example substrate in which identical cysteines are modified by two distinct PTMs is PSD-95 (post-synaptic density-95), where the two known sites of palmitoylation, Cys3 and Cys5, are also nitrosylated by neuronal nitric oxide   13 synthase (nNOS) (Ho et al., 2011). Nitrosylation was previously suggested to compete with palmitoylation (Hess et al., 1993), or displace palmitoyl moieties directly from palmitoylated cysteines (Baker et al., 2000). PSD-95 nitrosylation was demonstrated by a biotin-switch assay where ascorbate, which selectively reduces nitrosylated cysteines, was used to label these cysteines with biotin (Ho et al., 2011). Consistent with previous findings that glutamatergic transmission leads to PSD-95 depalmitoylation, glutamatergic activation via NMDAR (N-methyl-D-aspartate receptors) increased nitric oxide formation that inhibited PSD-95 palmitoylation and decreased the association of PSD-95 with synaptic clusters in cerebellar granule cells. Conversely, PSD-95 nitrosylation was also increased in Zdhhc8-/- mice where PSD-95 palmitoylation is decreased, showing that this regulation is reciprocal (Ho et al., 2011). Palmitoylation and Phosphorylation act as Cellular Pathway “Switches” Recently, interplay between palmitoylation and phosphorylation was shown for the large conductance calcium-activated potassium (BK) channel splice variant that contains the stress axis regulated exon (STREX BK) (Tian et al., 2008; Zhou et al., 2012). STREX BK channels contain two phosphorylation sites: a protein kinase A (PKA) motif at Ser636, and a protein kinase C (PKC) motif at Ser700. Phosphorylation at either site leads to inhibition of STREX BK activity (Zhou et al., 2012). STREX BK also contains conserved C-terminal cysteines which, when palmitoylated, target STREX BK to the PM. This palmitoylation also has a profound effect on channel conformation, rendering the channel insensitive to phosphorylation by PKC and thus to its inhibitory effect (Tian et al., 2008). However, the PKA consensus site is still exposed in this conformation, and its phosphorylation functions as   14 an electrostatic switch to promote STREX BK depalmitoylation, leading to STREX domain dissociation from the membrane to allow PKC phosphorylation and subsequent inhibition of STREX BK (Zhou et al., 2012).  Proximal phosphorylation and palmitoylation modifications can also compete with each other to affect peripheral protein subcellular localization, as demonstrated for phosphodiesterase 10A isoform 2 (PDE10A2), which has a PKA motif at Thr16 and a palmitoylation site at Cys11 (Charych et al., 2010). In primary mouse striatal neurons, Thr16 phosphorylation prevented PDE10A2 membrane localization by blocking N-palmitoylation at Cys11, which caused its translocation to the PM. Interestingly, however, if N-palmitoylation occurred on PDE10A2 first, PKA phosphorylation would no longer dissociate PDE10A2 from membranes (Charych et al., 2010). Palmitoylation Shields Substrate from Ubiquitin-Mediated Degradation Palmitoylation may shield substrates from ubiquitin modification and subsequent degradation (Blaskovic et al., 2013). For example, palmitoylation of the tumour endothelial marker 8 (TEM8), a type-I transmembrane anthrax receptor, prevents its association with lipid microdomains where modification by the E3 ubiquitin ligase Cbl (Casitas B-lineage Lymphoma) leads to its premature degradation (Abrami et al., 2006). Similarly, in yeast, palmitoylation of the SNARE (soluble N-ethylmaleimide-sensitive-factor attachment protein receptor) protein Tlg1 was found to prevent its ubiquitination and proteosomal degradation (Valdez-Taubas and Pelham, 2005). Interestingly, despite the general observation that palmitoylation serves as a stabilizing signal, the reverse has recently been reported for the E3   15 ubiquitin ER ligase Gp78 (Glycoprotein 78), where palmitoylation within its RING (Really Interesting New Gene) finger motif shifts its localization to peripheral ER to promote substrate degradation (Fairbank et al., 2012).  1.1.5 Dynamics of Protein Palmitoylation The highly reversible nature of protein palmitoylation provides the cells with a dynamic mechanism to respond to stimuli. Dynamic palmitate turnover was first reported in the 1980’s for the transferrin receptor, ankyrin, and N-Ras (Omary and Trowbridge, 1981; Staufenbiel, 1987; Magee et al., 1987). Subsequent studies demonstrated that palmitate turnover on stimulatory G-protein alpha subunit is accelerated by agonist treatment (Wedegaertner and Bourne, 1994), supporting the idea that dynamic palmitoylation is tightly regulated and can be mediated by enzymatic events. Indeed, studies in the last decade have revealed that dynamic palmitoylation is widespread. Aside from the dynamic cycling described for the ubiquitous Ras proteins, dynamic palmitoylation is especially important at the synapse, where it serves as a “switch” to control protein localization and function in response to neuronal stimuli that promote synaptic plasticity (El-Husseini and Bredt, 2002; Fukata and Fukata, 2010). Figure 1.3 briefly summarizes a subset of neuronal proteins where palmitoylation plays a role in their function and trafficking, often in a highly dynamic fashion. Note there are some exceptions here; for example, the presynaptic protein   synaptotagmin I was found to be stably palmitoylated (Heindel et al., 2003; Kang et al., 2004). It was proposed that palmitoylation likely functions as a structural signal for proper targeting and assembly of elements involved in neurotransmitter release.    16    Figure 1.3 Palmitoylation Regulates Synaptic Transmission. In the synapse, the palmitoylation of a variety of proteins regulate important aspects of synaptic function (with some examples highlighted in red). On the presynaptic site, palmitoylation of the SNARE proteins SNAP25 and Ykt6 regulates vesicle fusion and transmitter release; on the post-synaptic site, palmitoylation mediates the signaling pathway of many receptors by anchoring them to the PM. More importantly, palmitoylation also regulates the dynamic cycling of PSD-95 and glutaminergic receptors, where their palmitate turnover rates alter to mediate synaptic plasticity in response to stimuli. Adapted from (Huang and El-Husseini, 2005) with permission.    17 One of the most well-established dynamic neuronal substrates with rapid and regulatable palmitate turnover is the synaptic protein PSD-95 (El-Husseini et al., 2002). PSD-95 is a cytosolic scaffolding protein with a dual palmitoylation signal on its N-terminus on Cys3 and Cys5 (El-Husseini et al., 2000). Upon its palmitoylation, it translocates into the PSD to regulate assembly of complexes that contain cell adhesion molecules and glutamate receptors such as AMPARs (α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptors) and NMDARs at the post-synaptic PM to regulate synaptic plasticity (El-Husseini et al., 2000). PSD-95 shows a constitutive palmitate turnover as determined by metabolic labeling. Notably, palmitate cycling on PSD-95 is enhanced by excitatory stimulation of the synapse, leading to reduced PSD-95 synaptic localization and synaptic strength (El-Husseini et al., 2002).  On the other hand, synaptic activity blockade increases PSD-95 palmitoylation and localization to the postsynaptic membrane, a process that was demonstrated to be due to increased translocation of a palmitoylation enzyme, DHHC2, into postsynaptic density (Noritake et al., 2009) (discussed below in More recently, a conformational antibody that is specific to palmitoylated PSD-95 has been developed that can be used to track palmitoylation of PSD-95 in live cells (Fukata et al., 2013).   Other recent examples of neuronal substrates that show activity-dependent dynamic palmitoylation include some AMPAR and NMDAR subunits themselves (Hayashi et al., 2005, 2009), which will be discussed later in detail (Section 1.4.2); CDC42-Palm (Cell division cycle 42-Palm), which is a brain-specific splice variant that undergoes rapid depalmitoylation and dislocation from dendritic spine under high level glutamate stimulation (Kang et al., 2008b); and δ-catenin, whose palmitoylation is increased upon enhanced   18 synaptic activity, leading to its association with N-cadherin at synapses and eventual enlargement of postsynaptic spines and glutamate receptor insertions (Brigidi et al., 2014). Importantly, this “activity-dependence” of palmitate turnover has been demonstrated on a large-scale, where changes over the course of minutes in palmitoylation levels of several synaptic proteins were observed following excitatory and seizure-inducing stimuli administration in vitro and in vivo (Kang et al., 2008b). These findings further reinforce the complexity for the regulation of enzymes for palmitoylation and depalmitoylation to achieve such diversity of palmitoylation level changes between substrates under a same stimulus.  Efforts in the last two decades to find the enzymes responsible for palmitoylation and depalmitoylation have led to the identification of two separate enzyme families. I will devote the next two sections of my introduction to palmitoyl-acyl transferases (PATs), enzymes that mediate palmitate addition; and to acyl protein thioesterases (APTs), enzymes that mediate palmitate removal.   1.2 Palmitoyl-Acyl Transferases: Enzymes that Mediate Palmitate Addition 1.2.1 Identification of Palmitoyl-Acyl Transferases (PATs) Although protein palmitoylation was first reported in 1979 (Schmidt and Schlesinger, 1979), the enzymes responsible were not identified until two decades later. Palmitoylation is mediated by a family of highly conserved enzymes collectively referred to as palmitoyl-acyl transferases (PATs). The first PATs, the Erf2-Erf4 complex and Akr1, were identified in Saccharomyces cerevisiae. Erf2-Erf4 was initially identified in a genetic knockout screen for   19 mutants affecting Ras subcellular localization (Bartels et al., 1999), whereas casein kinase I was mislocalized in yeast lacking Akr1, suggesting that Akr1 has a role in its lipid modification (Feng and Davis, 2000). Both Erf2 and Akr1 are multispanning transmembrane domain proteins containing an Asp-His-His-Cys (DHHC) motif within a 51 residue cysteine-rich domain (CRD) that resembles a C2H2 zinc finger. DHHC-dependent palmitoyl-acyl transfer activity was later demonstrated for both Erf2-Erf4 and Akr1 (Putilina et al., 1999; Lobo et al., 2002; Roth et al., 2002), prompting the hypothesis that the family of >120 DHHC-CRD-containing polytopic proteins found across 7 species all have palmitoyl-acyl transfer activities (Roth et al., 2002; Linder and Deschenes, 2004). Subsequently, palmitoyl-acyl transferase activity of other DHHC-CRD-containing proteins was demonstrated in yeast (Valdez-Taubas and Pelham, 2005; Hou et al., 2005; Lam et al., 2006; Hou et al., 2009) and mammalian cells (Keller et al., 2004; Huang et al., 2004; Fukata et al., 2004).  To date, 7 yeast and 23 human DHHC-CRD proteins have been identified. All DHHC-CRD proteins contain four or six transmembrane domains with cytosolic-facing DHHC-CRDs (Fukata et al., 2013). Notably, a small subset contain variations of the classic DHHC motif; in particular, the human DHHC13 (HIP14L; Huntingtin-interacting protein 14-like) has a DQHC motif, and the yeast PATs Akr1, Akr2 and Pfa5 all sport a DHYC motif (Mitchell et al., 2006). Nevertheless, Akr1 and Pfa5 have established PAT activities in vivo (Hou et al., 2009) and HIP14L palmitoylates Huntingtin (Huang et al., 2011) and ClipR-59 (Ren et al., 2013) in mammalian cells and SNAP25 (Synaptosomal-associated Protein, 25kDa) in vivo (Sutton et al., 2013), although studies in other systems argue against HIP14L activity (Lemonidis et al., 2014). In contrast, activity has not been demonstrated for a subset of   20 DHHC-containing PATs, including DHHC1, 4, 16, 19, 23, and 24 (Ohno et al., 2012). Evolutionarily, PATs are highly conserved and may potentially be grouped into different subfamilies based on their amino acid identity within the DHHC-CRD domain (Figure 1.4) (Mitchell et al., 2006; Fukata and Fukata, 2010).    Figure 1.4 Phylogentic Clustering of the Human and Saccharomyces cerevisiae DHHC Proteins Based on ClustalX Alignment of the 51 Amino Acid DHHC Core Sequence. Human DHHC proteins are labeled in blue (brackets indicate their aliases) and yeast DHHC proteins are labeled in red. The brackets indicate six potential sub-families based on DHHC-CRD core sequence similarities. Adapted from (Mitchell et al., 2006) with permission.   1.2.2 Mechanism of PAT Activities Studies to dissect the mechanism of palmitate addition suggested that PATs first undergo autopalmitoylation on the active site cysteine in the DHHC motif using the “high energy” palmitoyl-CoA as a substrate (Roth et al., 2002; Fukata et al., 2004, 2006). This palmitoyl-Pfa3Pfa4Akr2Akr1Erf2Swf1Pfa5DHHC15DHHC2DHHC24DHHC12DHHC17 (HIP14)DHHC13 (HIP14L)DHHC18DHHC14DHHC9DHHC19DHHC8DHHC5DHHC4DHHC11DHHC1DHHC23DHHC21DHHC22DHHC16DHHC7DHHC3 (GODZ)DHHC6DHHC20  21 PAT intermediate subsequently transfers palmitate onto substrates (Mitchell et al., 2010). Indeed, mutations within the DHHC motif not only abolish palmitoylation of the substrate, but also the enzyme itself (Roth et al., 2002; Mitchell et al., 2010). Supporting this two-step ping-pong mechanism, it was further demonstrated that PAT autoacylation and acyltransfer to substrates displayed the same acyl-CoA specificity (Jennings and Linder, 2012). Interestingly, the same study found DHHC2 and DHHC3 have differing lipid substrate specificities (Jennings and Linder, 2012), although the basis of this selectivity remains to be examined.  A separate study on yeast Pfa3 also showed that palmitoyl-acyl transfer onto substrates is a two-step mechanism, whereby the PAT first binds its substrate independent of the DHHC domain, followed by palmitate transfer and lateral release of the palmitoylated substrate to the membrane (Hou et al., 2009). Although strong interactions between some substrate and enzyme pairs have been observed, other DHHC-substrate interactions are transient and more difficult to detect (Thomas et al., 2012; Lemonidis et al., 2014).  1.2.3 Overlapping Substrate Specificity of PATs The sheer number of PATs suggests that they may have different substrate specificities. Initial loss-of-function analyses in yeast that mapped the potential substrates for each DHHC enzyme support this hypothesis, and show PATs have both discrete and overlapping specificities (Roth et al., 2006). Subsequent studies in mammalian cells also demonstrated PAT overlapping and distinct substrate specificities; for example, while the closely-related DHHC3 and DHHC7 can both potentiate the palmitoylation of eNOS (epithelial nitric oxide   22 synthase), GAP43 (Growth associated protein 43), and SNAP25, only DHHC3 promoted the palmitoylation of the AMPAR subunits (Hayashi et al., 2005; Tsutsumi et al., 2008; Huang et al., 2009). Meanwhile, a number of studies suggested that PSD-95 can be targeted by numerous PATs including DHHC2, DHHC3, DHHC7, DHHC8, DHHC15, and DHHC17 (HIP14; Huntingtin-interacting protein 14) (Fukata et al., 2004; Mukai et al., 2008; Singaraja et al., 2011). Because DHHC overexpression may bypass DHHC regulators that would otherwise ensure palmitoylation by a selected DHHC protein (Hou et al., 2009), loss-of-function analyses are essential to determine specificity (Fukata and Fukata, 2010). However, results from loss-of-function studies should also be interpreted cautiously as this can too be confounded by the fact that other DHHCs may compensate and adopt a function that they would not normally perform.  What determines the substrate selectivity of PATs? Recent studies suggest elements of both substrates and enzymes determine preferential reaction pairs. Previous analyses in yeast suggest that substrates of a particular PAT tend to share common features: for example, Swf1 prefers transmembrane substrates with juxtamembranous cysteine sites, Erf2-Erf4 prefers substrates with another co-translational lipid modification, and Akr1 substrates tend to be hydrophilic proteins (Roth et al., 2006). However, the structural basis for these differences remains unknown. Meanwhile, extensive studies on the hydrophilic yeast kinase Yck2 revealed that a tripartite motif found on Yck2 functions as a recognition signal for Akr1 binding to allow its palmitoylation (Roth et al., 2011). A similar tripartite motif has also been noted for SNAP25 and may also be crucial for its palmitoylation since hydrophilic proteins   23 have otherwise no effective means of accessing membrane-localized PATs (Greaves et al., 2009, 2010; Roth et al., 2011).   Some PAT selectivity in mammalian cells may be attributed to structural motifs found in a subset of DHHC proteins that allow for stronger PAT-substrate interactions. For example, DHHC5 and DHHC8 are two related PATs that both contain a C-terminal PSD-95/Discs-large/ZO-1 homology (PDZ) binding site (Fukata and Fukata, 2010). This PDZ domain is required for both DHHC5 and DHHC8 to efficiently bind and palmitoylate the PDZ domain-containing neuronal substrates GRIP1b (glutamate receptor interacting protein-1b) and PICK1 (protein interacting with C-kinase) (Thomas et al., 2012, 2013). Another recent report uncovered the importance of the DHHC17 (HIP14) ankyrin-repeat domain (ANKRD) for the binding and palmitoylation of CSP (Cysteine String Protein) and SNAP25 (Lemonidis et al., 2014). Interestingly, however, the same study reported another ANKRD-containing PAT, DHHC13 (HIP14L), was unable to palmitoylate CSP and SNAP25 despite strong enzyme-substrate interactions. In addition, DHHC3 and DHHC7, both of which showed barely detectable interactions with CSP and SNAP25, palmitoylated CSP and SNAP25 more strongly than DHHC17 did (Lemonidis et al., 2014). These findings highlight the complexity of protein palmitoylation where interaction and enzymatic activity are not always in a linear relationship.   Differences in PAT subcellular localization may provide another explanation for the preferential reaction pairs. In yeast, the DHHC-CRD PAT activities are localized to distinct locations: while Akr1p is localized to the Golgi, Erf2p, Swf1p and Pfa4p are found on the   24 ER, Pfa3 is found on the vacuole, and Pfa5 is found on the PM (Ohno et al., 2006). Similarly, although a majority of DHHC proteins in mammalian cells are found on the Golgi (Ohno et al., 2006), others are reported to have ER (DHHC4, DHHC6) (Gorleku et al., 2011), endosomal (DHHC5) (Thomas et al., 2012) , vesicular (DHHC2) (Noritake et al., 2009), or PM (DHHC2) (Greaves et al., 2011; Fukata et al., 2013) localizations with localized enzymatic activities.  1.2.4 Modulation of PAT Activities Given the observation that palmitoylation levels can change rapidly in response to stimulatory or inhibition events, it is not surprising that a subset, if not all PATs, show evidence of regulation. Although a majority of studies to date have focused on identifying substrates and examining the biological consequences of palmitoylation, emerging evidence demonstrates PAT activities can be modulated in multiple ways.  Co-Factors can Modulate DHHC Activity One of the first PATs discovered in yeast, the Erf2-Erf4 complex, and its conserved mammalian counterpart, DHHC9-GCP16, remains the only multi-subunit PAT identified to date (Swarthout et al., 2005). Although only Erf2 contains a DHHC-CRD, both subunits are required for PAT activity (Lobo et al., 2002). The mystery as to why the auxiliary protein Erf4 is needed was not fully examined until 10 years following its initial discovery. Mitchell et al. reported that Erf4 stabilizes Erf2 and the palmitoyl-Erf2 intermediate by protecting Erf2 from ubiquitin-mediated degradation; furthermore, Erf4 is required for palmitate   25 transfer from Erf2 to substrates (Mitchell et al., 2012). Hence, Erf4 functions as an Erf2 co-factor to carry out transfer catalysis and/or substrate recognition.  A more recent example of a co-factor that stimulates PAT activity was described for the mammalian ANKRD-containing PAT HIP14. Huang and colleagues reported that the HIP14 substrate, HTT (Huntingtin), modulates the palmitoylation and activity of HIP14 through tight physical association with the HIP14 ANKRD (Huang et al., 2011). In the presence of HTT, but not mutant HTT (mHTT) which binds weakly to HIP14, HIP14 palmitoylation of its substrate SNAP25 was potentiated in vitro. In vivo, HTT knockdown using antisense oligonucleotides (ASOs) reduced palmitoylation of HIP14 substrates. Taken together, it was concluded that HTT functions as a co-factor for HIP14. However, the mechanistic details remain to be elucidated (Huang et al., 2011). Post-Translational Modifications of DHHC Proteins Recently, it was reported that DHHC8 is phosphorylated by the brain-specific protein kinase C variant, PKMζ, in primary mouse cortical neurons (Yoshii et al., 2011). This phosphorylation is selective as PKMζ did not phosphorylate 6 other DHHC proteins, and results in increased PSD-95 palmitoylation (Yoshii et al., 2011). However, the mechanism by which phosphorylation alters DHHC8 activity was not determined. Phosphorylation may directly regulate DHHC8 activity by changing its conformation, or affect DHHC8 localization or stability.      26 Differential Endogenous Expression Levels of DHHC Proteins Alteration in DHHC protein expression has also been implicated as a mode for cells to regulate protein palmitoylation. While examining protein palmitoylation in the fission yeast Schizosaccharomyces pombe, Zhang et al. observed global changes in the yeast palmitoylome during meiosis (Zhang et al., 2013). The authors found significant increases in the transcript and protein expression levels of Erf2 and Erf4, but not of other DHHC proteins, that corresponded to this transition in cell cycle (Zhang et al., 2013). Further studies showed that forced expression of Erf2 and Erf4 promoted meiotic entry whereas deletion of Erf2 delayed this process. Importantly, while changes in Erf2-Erf4 levels affected the palmitoylation status of Ras GTPase superfamily members Ras1 and Rho3, only the upregulation of Rho3 palmitoylation is required for the meiotic phenotype triggered by Erf2-Erf4 expression (Zhang et al., 2013) Oligomerization as a Possible Mechanism of Regulating DHHC Activity While homomultimer and heteromultimer formation between DHHC3 and DHHC7 was previously documented, the physiological significance of this finding remained unclear (Fang et al., 2006). More recently, through the use of bioluminescence resonance energy transfer (BRET) and co-immunoprecipitation methods, Lai and Linder demonstrated dynamic self-association of DHHC2 and DHHC3 in HEK293 cells (Lai and Linder, 2013). Interestingly, dimeric forms of DHHC3 were found to be less active than their monomeric counterparts, raising the hypothesis that dynamic oligomerization may provide one of the mechanisms to modulate PAT activity (Lai and Linder, 2013). Subsequently, self-interaction of another DHHC protein, HIP14, was also reported in a large-scale yeast-two-hybrid screen (Butland et   27 al., 2014). Given these findings, it will be important to examine the role of DHHC oligomer formation in vivo. Subcellular Trafficking of DHHC Proteins PSD-95 is a synaptic protein shown to have dynamic palmitate cycling that is regulated by synaptic activity. While synaptic excitation causes PSD-95 depalmitoylation via an unidentified depalmitoylase, Noritake et al. demonstrated that PSD-95 is actively palmitoylated and translocated to the synaptic densities upon activity blockade in cultured hippocampal neurons (Noritake et al., 2009). Although PSD-95 is palmitoylated by several PATs, the study focused on the two highly expressed PSD-95-palmitoylating PATs in hippocampal neurons, DHHC2 and DHHC3, and found that the activity-dependent palmitoylation of PSD-95 is specifically due to DHHC2. DHHC2 is localized to cytoplasmic vesicles throughout neurons and is constitutively active. Upon activity blockade, dendritic DHHC2-containing vesicles translocate to dendritic shafts to mediate palmitoylation (Noritake et al., 2009). A follow-up study demonstrated that DHHC2 is inserted into the PM, which is necessary for the formation of postsynaptic PSD-95 nanodomains that support AMPAR recruitment in response to changes in synaptic activity (Fukata et al., 2013). Taken together, these results demonstrate that post-synaptic density assembly is a highly regulated event modulated by local palmitoylation cycles. The C-terminus domain of DHHC2 was reported to be important for its distinct vesicular/endosomal localization (Greaves et al., 2011). Further studies will be needed to elucidate the pathway by which activity blockade promotes DHHC2 translocation and membrane insertion. In addition, it will be vital to   28 determine whether other DHHCs and APTs play a role in the global palmitoylation changes that occur in response to inhibitory and excitatory stimuli.  1.2.5 Involvement of DHHC Proteins in Neurodegenerative Diseases Given the importance of DHHC proteins for proper protein palmitoylation and function, it is not surprising that several reports have associated disease conditions with misregulated palmitoylation due to altered DHHC expression or activity. For example, the mutant form of superoxide dismutase 1 (SOD1) is aberrantly palmitoylated in some forms of amyotrophic lateral sclerosis (ALS) (Antinone et al., 2013). More recently, a link was suggested between diabetes and HIP14, which was shown to be predominantly expressed in β-cells and is required for β-cell survival and glucose-stimulated insulin secretion (Berchtold et al., 2011). Here, I will briefly discuss the strongest link between DHHC proteins and some diseases. Cancer Links between cancer and altered expression of different DHHC enzymes have been proposed. These include an increased copy number of a chromosome 5 region encompassing the ZDHHC11 gene in both lung and bladder cancers (Kang et al., 2008a; Yamamoto et al., 2007), reduced DHHC2 protein expression in colorectal cancers and gastric adenocarcinoma (Oyama et al., 2000; Yan et al., 2013), and increased DHHC9 protein expression in colorectal tumours (Mansilla et al., 2007). Interestingly, DHHC17 (HIP14) was proposed to have oncogenic functions in mice (Ducker et al., 2004), while a recent study reported that another DHHC protein, DHHC14, may function as a tumor suppressor (Yeste-Velasco et al., 2014).   29 Mental Retardation Two reports highlight the role of DHHC proteins in X-linked mental retardation (XLMR). In a case report, a 29-year-old female patient with severe, nonsyndromic XLMR carried a de novo balanced reciprocal translocation 46,XX,t(X;15)(q13.3;cen). The X chromosome breakpoint was mapped to within 3.9kb of exon1 of the ZDHHC15 gene, and expression studies showed this translocation disrupted ZDHHC15 transcription (Mansouri et al., 2005).   Another candidate gene implicated in XLMR is ZDHHC9. In a systematic screen for mutations in genes on the X chromosomes from a cohort of 250 families, each with multiple males suffering from mental retardation without known molecular diagnosis, mutations in ZDHHC9 at Xq26.1 were identified from 4 separate families. Among these, there was one frameshift mutation, one splice-site mutation, and two missense mutations in highly conserved residues DHHC-CRD domain, strongly suggesting that these individuals code for dysfunctional DHHC9 (Raymond et al., 2007). While the documented frameshift and splice-site mutations lead to deletion of the essential DHHC catalytic domains of DHHC9 and therefore nonfunctional enzymes, follow-up studies showed that the two missense mutations negatively affect the autopalmitoylation step of the reaction by lowering the steady state amount of the palmitoyl-zDHHC9 intermediate without altering protein expression levels, providing a molecular mechanism for the role of ZDHHC9 in XLMR (Mitchell et al., 2014) . Schizophrenia De novo microdeletion of 22q11.2, the locus that contains the ZDHHC8 gene, was associated with a higher risk of schizophrenia (Bassett et al., 2003). Of the 72 single nucleotide   30 polymorphisms (SNPs) found in the 1.5Mb microdeletion region, rs175174 showed the strongest association and was subsequently found to regulate the level of ZDHHC8 transcript (Mukai et al., 2004), suggesting a strong link between ZDHHC8 and schizophrenia.  In a mouse model of 22q11.2 the microdeletion, cognitive and behavioural phenotypes mimicking schizophrenic symptoms were reported (Mukai et al., 2008). In addition, primary hippocampal neurons from both the 22q11.2 deletion mouse model and zdhhc8-deficient mice showed decreased dendritic spine density and impaired dendritic growth that can be rescued by the introduction of DHHC8, suggesting that DHHC8 activity plays an important role in neuronal development (Mukai et al., 2008). Supporting this notion, zdhhc8-deficient mice have decreased PSD-95 palmitoylation (Mukai et al., 2008). Huntington’s Disease (HD) Huntington’s Disease (HD) is an autosomal dominant neurodegenerative disease that is typically adult-onset and characterized by progressive cognitive, psychiatric, and behavioural symptoms (Walker, 2007). HD is caused by mutations in the gene Huntingtin (HTT) (Schulte and Littleton, 2011). Wild type HTT is an ubiquitous, essential 3,144 amino acid protein that is highly expressed in the brain (Li and Li, 2004). HTT contains an N-terminal CAG (polyglutamine) repeat of variable length that, when expanded to greater than 35 repeats (mHTT), causes HD (Myers, 2004). The wild-type protein is reported to function in protein trafficking, vesicle transport, clathrin-mediated endocytosis, postsynaptic signaling, transcription regulation and to have anti-apoptotic activity (Gil and Rego, 2008; Imarisio et al., 2008).    31 A yeast-two-hybrid-based study first identified Huntingtin-Interacting protein (HIP14/DHHC17) as a HTT interacting protein (Singaraja et al., 2002). HIP14 binding to HTT was inversely correlated with CAG repeat length, suggesting a potential role in HD pathogenesis. Interestingly, HIP14 is a mammalian homologue of the yeast PAT Akr1, and was able to rescue the endocytic defects and temperature-sensitive lethality of the yeast akr1Δ strain (Singaraja et al., 2002). Subsequent studies confirmed the palmitoylacyl transfer activity of HIP14 and showed that HTT is a HIP14 substrate (Huang et al., 2004). Importantly, mutation of the mHTT Cys214 palmitoylation site to serine increased mHTT inclusion formation in COS cells and primary neuronal cultures, a finding that strongly suggests a role for faulty mHTT palmitoylation in the pathogenesis of HD (Yanai et al., 2006).  Transgenic mice lacking HIP14 were developed to further examine the role of HIP14 in HD pathogenesis in vivo (Singaraja et al., 2011). Hip14-/- mice recapitulated some of the phenotypes seen in mouse models of HD, including decreased brain weight and striatal volume, and motor deficits; however, HTT palmitoylation level was not altered (Singaraja et al., 2011). This may be due to compensation from other HTT PATs, such as the HIP14 homologue HIP14L. Interestingly, Hip14-/- animals showed decreased SNAP25 and PSD-95 palmitoylation (Singaraja et al., 2011). Additionally, Hip14-/- animals also displayed synaptic electrophysiological defects (Milnerwood et al., 2013).   The observation that palmitoylation of PSD-95 and SNAP25 was affected in Hip14-/- mice (Singaraja et al., 2011) suggests that HIP14 is one of the main PATs for PSD-95 and   32 SNAP25 in neuronal cells, and that redundant PATs are not sufficient to rescue palmitoylation of these particular substrates in in Hip14-/- mice. Alternatively, the synaptic changes in these mice could have somehow disrupted the palmitoylation cycle of proteins to shift them towards the depalmitoylated state.  1.3 Enzymes that Mediate Palmitate Removal 1.3.1 Palmitoylthioesterases are Members of the Metabolic Serine Hydrolase (mSH) Family To date, only three enzymes with depalmitoylation activity have been identified: the lysosomal PPT1 (Palmitoyl-Protein thioesterase 1), and the cytosolic APT1 (Acyl-protein thioesterase 1) and APT2 (Acyl-protein thioesterase 2) (Zeidman et al., 2009; Conibear and Davis, 2010; Tomatis et al., 2010). Sequence homology and structural studies show that all three of these thioesterases belong to the metabolic serine hydrolase (mSH) superfamily of proteins, which consists of >120 additional members (Long and Cravatt, 2011). A majority of these mSH proteins, including PPT1 and APT1/2, adopt an α/β hydrolase fold with a signature GXSXG motif, where the serine residue is usually the catalytic nucleophile embedded in a classic catalytic triad (Ser-His-Asp or Ser-Ser-Lys) or dyad (Ser-Lys or Ser-Asp) (Long and Cravatt, 2011). Despite these general similarities, distinct mSH members cleave different ester, amide, or thioester bonds in small molecules or proteins (Long and Cravatt, 2011). As depicted in the dendrogram shown below (Figure 1.5), despite PPT1, APT1 (LYPLA1), and APT2 (LYPLA2) all being reported to have palmitoylthioesterase activity, PPT1 is quite distant from APT1/2 in terms of sequence relatedness (Bachovchin et   33 al., 2010). Therefore, structural features local to the enzymatic active site are likely contributors to the distinct lipid substrate selectivity of the mSH proteins.   Figure 1.5 Dendrogram Showing all 128 Members of the Mouse Metabolic Serine Hydrolase Family with Branch Length Depicting Sequence Relatedness. FAAH2 and PNPLA4 are human mSHs that are included here as they lack mouse orthologues. SHs that can be labeled with activity-based probes are shown in red (105 enzymes or 82% of the metabolic serine hydrolase family). Blue arrow points to the lysosomal depalmitoylase PPT1; Green arrows point to the cytosolic depalmitoylases APT1 and APT2. Adapted from (Bachovchin et al., 2010) (Copyright © 2010, by Bachovchin and authors. Reprinted with permission).    PPT1 and APT1/2 carry out enzymatic depalmitoylation by the general mSH catalytic mechanism. This reaction first involves the formation of an acyl-enzyme intermediate at the active site serine by nucleophilic attack, followed by hydrolysis of the protein or lipid substrate and subsequent regeneration of a free active site for entry into the next reaction cycle (Long and Cravatt, 2011). Owing to the enhanced reactivity of the active site serine, the functional state of most mSHs can be assessed using active site-directed affinity labels,   34 namely fluorophosphonates (FPs), which provide a useful tool to study activity of serine hydrolases under different physiological states or manipulations (Long and Cravatt, 2011). Termed activity-based protein profiling (ABPP), this approach is a great asset for development of selective inhibitors (Liu et al., 1999), as will be discussed in section  Here, I will discuss the lysosomal and cytosolic palmitoylthioesterases and their substrate selectivity as well as structural features that make them unique. Luminal Protein Depalmitoylation by Palmitoyl-Protein Thioesterase 1 (PPT1) PPT1 (Palmitoyl-protein thioesterase 1) was isolated from bovine brain homogenates and identified in 1993 as the first enzyme with depalmitoylation activity. PPT1 removed palmitate from [3H]palmitate-labelled H-Ras and G(alpha) in vitro (Camp and Hofmann, 1993). Molecular cloning revealed that the enzyme contains the thioesterase signature that includes a His-Asp-Gly catalytic triad, and the GSXSG motif characteristic of mammalian lipases (Camp et al., 1994). The crystal structure at 2.25 Å showed PPT1 adopts an α/β-hydrolase fold, with Ser115 acting as the nucleophile, and a hydrophobic pocket that accommodates fatty acid chains (Bellizzi et al., 2000).  The enzymatic activity of PPT1 expressed in heterologous cells was initially reported to have an optimal pH of 7.0, and was largely found in cytosolic fractions (Camp and Hofmann, 1993). However, subsequent studies showed that PPT1 is a glycosylated lysosomal   35 depalmitoylase with an acidic pH optimum, whose primary role is to remove fatty acids from proteins undergoing breakdown in the lysosomal lumen (Verkruyse and Hofmann, 1996; Hellsten et al., 1996). Importantly, mutations in PPT1 that affect enzymatic activity lead to infantile neuronal ceroid lipofuscinosis (INCL), a neurodegenerative lysosomal storage disorder that results from the consequent accumulation of lipid-modified proteins (ceroid) in neurons (Vesa et al., 1995). PPT1 may be involved in other cellular functions (Kim et al., 2008). However, a role for PPT1 in cytosolic depalmitoylation has not been clearly demonstrated, where APT1 and APT2 are generally believed to mediate protein depalmitoylation in the cytosol. Cytosolic Protein Depalmitoylation by Acyl-Protein Thioesterases (APTs) Acyl-Protein Thioesterase 1 (APT1) In 1998, Duncan and Gilman reported the identification of the cytosolic acyl-protein thioesterase 1 (APT1) as the enzyme responsible for Gsα palmitate removal (Duncan and Gilman, 1998). APT1, which is originally designated as LYPLA1, was previously thought to be a lysophospholipase, but was found to have significantly higher enzymatic activity towards acyl thioester-containing proteins relative to lysophospholipid substrates (Sugimoto et al., 1996; Duncan and Gilman, 1998). Like PPT1, APT1 is a highly conserved α/β hydrolase that contains both the His-Asp-Gly catalytic triad and a GSXSG motif (Duncan and Gilman, 1998; Devedjiev et al., 2000). Importantly, APT1 is found mostly in cytosolic fractions in both Saccharomyces cerevisiae and mammalian cells (Duncan and Gilman, 1998, 2002; Hirano et al., 2009)    36 A number of candidate APT1 substrates have since been identified. Pulse-chase studies using [3H]palmitate showed that high levels of APT1 expression accelerated palmitate removal on eNOS and G(alpha) in vitro, but did not affect depalmitoylation of the raft protein caveolin (Yeh et al., 1999; Duncan and Gilman, 2002). APT1 was also able to depalmitoylate Synaptosomal-associated protein 23 (SNAP23) and viral proteins when these substrates were co-incubated with recombinant APT1 purified from Escherichia coli (Veit and Schmidt, 2001; Flaumenhaft et al., 2007). While these studies indicate that APT1 is capable of substrate-specific depalmitoylation in vitro, they do not exclude the possibility that other APTs contribute to the process in vivo. More recently, a functional role for APT1 was demonstrated by Siegel et al., who showed that miRNA-138 reduced APT1 levels in rat synaptosomes and increased the membrane association of its substrate, Gα13, thus altering dendritic spine size and synaptic transmission (Siegel et al., 2009). Furthermore, treatment of cells with Palmostatin B (PalmB, discussed below in section, a newly developed APT1 small molecule inhibitor, alters Ras localization (Dekker et al., 2010). The latest evidence of APT1-directed protein depalmitoylation is demonstrated in the parasitic protozoan Toxoplasma gondii, where TgASH1 (also known as TgPPT1), an APT1 orthologue, displayed depalmitoylating activity towards substrates important for its lytic cycle such as gliding-associated protein-45 (GAP45) in vitro (Kemp et al., 2013; Child et al., 2013). These lines of evidence substantiate the role of APT1 as a cytosolic depalmitoylase.   Thus, in vitro and in vivo evidence to date support the idea that APT1 depalmitoylates numerous cytosolic proteins, and that its activity, which can be regulated at the transcriptional level, is important for physiological processes. However, a comprehensive   37 study of APT1 substrate specificity is lacking, and because APT1 does not appear to indiscriminately depalmitoylate all proteins, additional proteins with cytosolic depalmitoylation activities may well exist. Acyl-Protein Thioesterase 2 (APT2) and APT1-Like (APT1L) Bioinformatics analyses revealed two related proteins in mammalian cells, APT2 and APT1-like (APT1L), that have 66% and 33% identity to APT1, respectively (Conibear and Davis, 2010). Like APT1, APT2 was initially identified as a lysophospholipase, and both APT2 and APT1L have a conserved catalytic triad (Toyoda et al., 1999; Zeidman et al., 2009). In Chinese Hamster ovary K1 (CHO-K1) cells, which lack endogenous APT1 expression, knockdown of APT2 altered the localization of GAP43 and H-Ras (Tomatis et al., 2010). However, changes in palmitoylation of both proteins were not measured directly. While a separate study revealed that APT2 can depalmitoylate semisynthetic N-Ras in vitro with higher kinetic parameters than APT1 (Rusch et al., 2011), direct in vivo evidence of APT2 depalmitoylating activity is still lacking.  APT1L, the second APT1 homologue, is highly expressed in adipose tissues and its variants have been associated with obesity in a number of genome-wide association studies (GWAS), suggesting that it may have lipase activity (Lindgren et al., 2009; Speliotes et al., 2011). Acyl-biotin exchange (ABE) studies in HEK293 cells showed that overexpression of wild type, but not mutant, APT1 or APT1L led to the robust deacylation of large conductance calcium-activated potassium (BK) channels, an effect not observed with APT2 (Tian et al., 2012). Intriguingly, structural studies revealed that, despite having the serine hydrolase   38 signature α/β hydrolase fold, the APT1L active site is shallower than that of APT1, and may not be able to accommodate palmitoyl chains (Burger et al., 2012). Correspondingly, in vitro analyses showed APT1L was unable to hydrolyse palmitoyl-N-Ras, suggesting this enzyme preferentially hydrolyses lipid substrates of shorter chain lengths (Burger et al., 2012). Based on these findings, it is currently inconclusive whether APT1L has depalmitoylating activity. It is possible that the observed BK channel depalmitoylation by APT1L in cells was through an indirect effect of APT1L overexpression. It is also possible that APT1L shows substrate selectivity and does not target N-Ras. Further studies will be imperative to reconcile these opposing findings.  While these initial analyses suggested roles for APT2 and APT1L in palmitate removal from selected proteins, the lack of direct in vivo evidence for APT2, and the contradictory reports on APT1L, suggest further examination into the enzymatic activity, substrate selectivity, and functional relevance of these APT1-like proteins is warranted.   1.3.2 Modulation of Protein Depalmitoylation The rate of depalmitoylation can also be influenced by factors that change protein conformation and accessibility to PATs or APTs. For example, the prolyl isomerase FK506 binding protein 12 (FKBP12) binds H-Ras in a palmitoylation-dependent manner, promoting its depalmitoylation and retrograde trafficking from the plasma membrane to the Golgi (Ahearn et al., 2011). FKBP12 targets a proline residue proximal to the palmitoylated cysteines of H-Ras, and by catalyzing cis-trans isomerization at this site, it may enhance the presentation of palmitoylated cysteines to the thioesterases (Ahearn et al., 2011).   39  Other proteins can also indirectly alter rates of palmitoylation or depalmitoylation. The phosphodiesterase 6 delta subunit (PDE6δ) interacts preferentially with depalmitoylated yet farnesylated H-Ras to extract it from membranes (Chandra et al., 2012). This enhances the cytosolic diffusion of H-Ras to promote its repalmitoylation by Golgi-localized PATs (Chandra et al., 2012). While FKBP12 and PDE6δ may be specific for Ras isoforms, distinct co-factors could influence depalmitoylation of other proteins by promoting their interaction with cytosolic APTs, thus adding another layer of complexity to the regulation of dynamic palmitoylation.  Recently, it has also been suggested that APT1 may be regulated at the translational level. In neurons, APT1 mRNA can be found locally in dendrites where they are bound to miRNA-138 that represses APT1 translation (Siegel et al., 2009; Banerjee et al., 2009). However, under NMDA stimulation, the RISC (RNA-induced silencing complex) protein MOV10 undergoes rapid degradation, which in turn disrupts the RISC function and results in rapid local APT1 synthesis through release of APT1 mRNA from miRNA-138 (Banerjee et al., 2009). Hence, this change in local APT1 translation may serve as a mechanism for activity-dependent depalmitoylation of some substrates.  1.3.3 Evaluating APT Activity Using Genetic and Small Molecule Approaches There is little evidence to date that APT1 and APT2 are the sole cytoplasmic depalmitoylases. Whereas APT1/2 are thought to act ubiquitously and constitutively, as-yet-undiscovered APTs may be specific for a particular substrate or cell type, or respond to   40 regulatory inputs. However, finding the complete set of relevant APTs will be challenging. The metabolic SH superfamily encompasses more than 100 members (Long and Cravatt, 2011). While many have known functions as lipases, a large number remain uncharacterized (Simon and Cravatt, 2010), making it a daunting task to evaluate each of these systematically. To date, studying APT activity has primarily used genetics and chemical biological approaches with small molecule inhibitors. Using Genetic Manipulation to Study APT Activity Depalmitoylation activity is often established using in vitro assays, or by overexpression of a candidate APT. While this indicates the APT is able to target a given substrate, it does not prove that it does so in vivo. Characterization of the functional role of protein depalmitoylation will require more detailed analyses. Knockdown studies can establish the requirement for a given APT, but are less effective if several APTs act redundantly. Both overexpression and knockdown are long-term manipulations that could give rise to indirect effects. For example, perturbation of a lipase activity could indirectly affect localization or palmitoylation of proteins by altering the membrane lipid environment. In contrast, small molecule inhibitors act quickly and may inhibit redundant, related enzymes. With the advent of small molecule inhibitors, many have now turned to using chemicals as an approach for studying the cellular role of protein depalmitoylation. Using Small Molecules and Chemical Biology to Study APT Activity The development of small molecules that inhibit APT activity has been more fruitful than for studies of their DHHC counterparts. This is due in part to the availability of the APT1 crystal   41 structure (Dekker et al., 2010; Davda and Martin, 2014). In the last decade, there have been many efforts to identify chemotypes that selectively inhibit APT1 and APT2. These APTs can be inhibited by compounds containing benzodiazepines, β-lactones, triazole ureas, chloroisocoumarins, boronic acid, and piperazine amides as backbones (Deck et al., 2005; Dekker et al., 2010; Adibekian et al., 2011, 2012; Zimmermann et al., 2013; Child et al., 2013). However, one caveat is that off-target effects often complicate their usage.  In this section, I will first discuss activity-based protein profiling (ABPP) as a method to verify the molecular targets of these compounds. Then, I will focus on four recently developed small molecule inhibitors that have gained wide popularity: Palmostatin B (PalmB), a commercially available β-lactone-core containing APT1/2 inhibitor (Dekker et al., 2010); hexadecylfluorophosphonate (HDFP), a lipase-selective inhibitor that blocks the depalmitoylation of a subset of proteins (Martin et al., 2011); and the piperazine amide inhibitors, C83 and C115, which are the first compounds to show selective inhibition of APT1 and APT2, respectively (Adibekian et al., 2012). Verification of Small Molecule Targets by Activity-Based Protein Profiling (ABPP) Because the mSH family contains >120 members in addition to the known palmitoylthioesterases, PPT1, APT1, and APT2 (Long and Cravatt, 2011), it is challenging to design molecules that selectively affect the activity of a single mSH protein. An activity-based protein profiling (ABPP) technology has been developed using the electrophilic fluorophosphonate (FP)-based probes, which directly and covalently modify the nucleophilic   42 serine of active mSHs, to facilitate screening of the enzymatic activity of mSH family members (Cravatt et al., 2008; Rajagopalan et al., 2014). In ABPP, nondenatured cell lysates are incubated with the FP-based probes to label enzymatically active mSHs prior to quenching the reactions with the denaturing agent SDS (Liu et al., 1999). These FP-probes are conjugated to either rhodamine to facilitate in-gel fluorescence as read-out, or biotin to facilitate downstream MS analyses (Barglow and Cravatt, 2007).   Variations of the ABPP technology are used to study inhibitor targets. In competitive ABPP (cABPP; Figure 1.6), cell lysates are first treated with the inhibitor in question for a short duration prior to addition of the FP probe. Any inhibitor that occupies the active site of its target will prevent labeling with the FP probe. With the advent of click chemistry, azide- and alkyne-containing ABPP probes have been introduced to allow in situ and in vivo labeling of the active mSH proteome to identify physiological targets of inhibitor in question (Kidd et al., 2001; Leung et al., 2003).   Figure 1.6 Schematic Illustration of Competitive Activity-Based Protein Profiling (ABPP). For simplification, two types of serine hydrolases are shown, one as the inhibitor target (Purple) and the other being a non-target (Green). Treating the non-denatured cellular pool of these enzymes with inhibitor leads to active site occupancy of targets. Subsequent activity profiling with FP-Rhodamine (FP-Rho) will only lead to covalent labeling of rhodamine on active non-targets. In-gel fluorescence can be used as readout to confirm inhibitor target and determine if the inhibitor in question has any additional off-targets. + Inhibitor (FP-Rho)+Activity ProbeIn-gel fluorescence+ Inhibitor+ DMSOTNTcABPP:Serine HydrolasesTARGETNON-TARGET  43 Palmostatin B (PalmB) The first effort to develop an APT1 inhibitor used a “protein structure similarity clustering” (PSSC) strategy, based on the assumption that the ligand-sensing core of APT1 may be inhibited by a compound backbone that blocks the activity of the structurally similar gastric lipase (Dekker et al., 2010). Subsequently, a collection of β-lactone core-containing compounds were synthesized and screened for APT1 binding. Palmostatin B (PalmB) was identified as a competitive inhibitor of APT1 through reversible covalent modification of the active site serine residue. In cells treated with PalmB, Ras depalmitoylation is potently inhibited, leading to disrupted Ras localization, and notably, phenotypic reversion in HRasG12V-transformed MDCK-F3 cells, demonstrating in situ biological activity of PalmB and further support of the role of APT1 in Ras depalmitoylation. Interestingly, direct knockdown of APT1 only led to a partial inhibition of Ras depalmitoylation when compared to PalmB treatment (Dekker et al., 2010). This initial report suggested that selectivity of PalmB for APT1 is high compared to other cellular lipid esterases (Dekker et al., 2010). This has subsequently attracted many groups to use PalmB to study the biological effects of APT1/2 inhibition. Hexadecylfluorophosphonate (HDFP) Hexadecyl fluorophosphonate (HDFP) is described as a non-selective lipid-like serine hydrolase inhibitor with a C16 backbone (Martin et al., 2011). In studying global depalmitoylation dynamics using a pulse-chase approach in T cell hybridomas, HDFP treatment robustly led to the inhibition of depalmitoylation of a subset of proteins, including G-proteins and Ras proteins. Using SILAC (Stable isotope labeling by amino acids in   44 culture)-ABPP, HDFP was found to potently inhibit the activities of ~20 serine hydrolases, some with unannotated functions, in addition to APT1 and APT2 (Martin et al., 2011). A recent study on R7BP (R7 regulator of G protein signaling (RGS)-binding protein) depalmitoylation in mouse neuro2a cells showed that HDFP treatment, but not selective APT1 and APT2 downregulation, inhibited R7BP palmitate turnover, giving rise to the hypothesis that some of the HDFP targets may also have palmitoylthioesterase activities (Jia et al., 2014). In addition, HDFP treatment could not prevent the palmitate turnover of some proteins in T cells, suggesting that there may also be additional depalmitoylases that are not inhibited by HDFP (Martin et al., 2011). Piperazine Amide Compounds Thus far, none of the molecules that block depalmitoylation showed isoform-selective inhibition, and, in the absence of an APT2 crystal structure, a rational design approach to create isoform-selective inhibitors has been challenging. Using a high throughput screening approach with an ABPP-based assay, Cravatt and colleagues identified distinct inhibitors selective for APT1 or APT2. These inhibitors, C83 (for APT1) and C115 (for APT2), were further validated in vivo by injection into mice, which subsequently verified selective APT inhibition in various tissues (Adibekian et al., 2012). Thus, C83 and C115 are the most selective and potent in vivo inhibitors of APT1 and APT2 identified to date, and represent great assets for future studies on protein depalmitoylation.    45 1.4 Current Paradigm of PAT/APT-Directed Dynamic Cycling  Despite the advances in the field of protein palmitoylation in the last decade summarized above, the mechanism by which cellular palmitoylation machineries balance the palmitoylated/depalmitoylated states of substrates is only now emerging. As discussed previously, changes in PAT activity/localization have been described as ways of tipping the balance; however, APT activity modulations have not been explored. For basal palmitate turnover, various studies on palmitoylated forms of Ras have suggested a spatiotemporal regulation by which PAT and APT activities keep substrate palmitoylation in check.  1.4.1 Spatial Modulation of APT-Mediated Depalmitoylation APTs are thought to act ubiquitously to remove mistargeted palmitoylated proteins from endomembranes, and promote their re-localization and re-palmitoylation at the Golgi (Rocks et al., 2010). For many substrates, this leads to their concentration at the Golgi, where their PATs reside (Rocks et al., 2010). This model suggests APTs must freely diffuse to target substrates at all membranes in cell. However, an important question remains as to how APTs, which are cytosolic proteins with no previously determined membrane-targeting signals, can access their substrates (Figure 1.7).        46  Figure 1.7 Spatiotemporal Model of Dynamic Palmitoylation Cycle for the Model Substrate Ras. Ras proteins are prenylated (maroon zigzag symbol), and this single lipid modification allows them to sample all endomembranes transiently. In the Golgi, the enzyme palmitoyl-acyl transferase (PAT) catalyzes Ras palmitoylation (green wavy symbol), which targets it to PM. APT1 (Acyl protein thioesterase 1) acts ubiquitously in the cytosol to depalmitoylate Ras to prevent Ras accumulation on endomembranes, and to promote Ras to return to the Golgi where the palmitoylation cycle starts again. This creates a continuous flux between the Golgi and the PM.    Two independent reports showed that both APT1 and APT2 were palmitoylated on a single cysteine (Cys2) near their N-termini (Kong et al., 2013; Vartak et al., 2014). While this single modification confers only transient membrane association it could, in principle, promote interaction with substrates. In fact, Kong et al. proposed that palmitoylation enriches APT1 and APT2 at the plasma membrane to enhance the depalmitoylation of substrates that are localized there (Kong et al., 2013). However, Vartak et al. used dynamic imaging to PALMITATEISOPRENYLRas RasGolgiPAT(DHHC9)RasRasRasAPT1 RasAPT1RasAPT1Endoplasmicreticulum (ER)Plasma membrane  47 analyze interactions between APTs and the model substrate Gαi(1-11) and found that unpalmitoylated APTs interacted with palmitoylated substrates on endomembranes (Vartak et al., 2014), consistent with previous studies that showed depalmitoylation is rapid and ubiquitous (Rocks et al., 2010).  In the absence of APT inhibitors, palmitoylated forms of APT1 and APT2 were partially localized to the Golgi, the site of bulk palmitoylation, where they interacted with and depalmitoylated each other in trans, leading to their release into the cytosol (Vartak et al., 2014). Vartak et al. proposed this cycle - in which palmitoylation drives local recruitment of a depalmitoylase - creates a negative feedback loop that establishes a balance between depalmitoylation and palmitoylation at the Golgi (Vartak et al., 2014). The idea that palmitoylation of APT1/2 serves a modulatory rather than essential role is supported by the observation that the palmitoylated cysteine is not conserved in APT1 homologs in lower organisms (Saccharomyces cerevisiae, Caenorhabditis elegans and Toxoplasma gondii) (D. Lin, unpublished observations). Whether such a feedback loop is dispensable, or another mechanism exists to balance rates of palmitoylation and depalmitoylation in these organisms, remains to be examined.   1.4.2 Current Issues with the Model The proposed model nicely describes the balance between Golgi-localized PATs and the cytosolic APT1/2 to modulate the known cytosolic APT1/2 substrates Ras and G(alpha) under basal conditions. However, this model does not fully explain the diversity in substrate turnover dynamics, PAT localization, and biological effects of palmitoylation for some   48 substrates. Furthermore, the model does not account for the rapid-onset activity-dependent changes in palmitoylation of some substrates, as outlined in the following biologically established scenarios. Evidence for Activity-Dependent Palmitoylation and Depalmitoylation Activity-dependent changes in palmitoylation levels of various substrates have been demonstrated for signaling proteins even in heterologous cells. Palmitate turnover on the stimulatory G-protein alpha subunit is accelerated by the treatment with its agonist isoproterenol (Wedegaertner and Bourne, 1994). It was also shown that stimulation of T cell activation by pervanadate (PV) treatment accelerates Lck palmitate turnover, and this is inhibited by the broad spectrum serine hydrolase inhibitor, methylarachidonyl fluorophosphonate (MAFP), suggesting that that Lck depalmitoylation is likely mediated by a serine hydrolase whose identity remains unknown (Zhang et al., 2010).  In neurons, activity-dependent palmitate turnover has been demonstrated on a large scale. Changes in palmitoylation levels of several synaptic proteins were observed over the course of minutes following excitatory and seizure-inducing stimuli administration in vitro and in vivo (Kang et al., 2008b). More recently, activity-induced palmitoylation of δ-catenin by DHHC5 was also reported (Brigidi et al., 2014). These observations, along with the activity-dependent DHHC2 palmitoylation of PSD-95 (Noritake et al., 2009; Fukata et al., 2013) as elaborated earlier, support the idea that both APT and PAT activities are regulated differently in response to external stimuli.    49 Compartmentalized PAT/APT Activities Transmembrane proteins are also often palmitoylated and their palmitoylation status may direct their recycling between compartments. In neurons, the glutaminergic receptors AMPAR and NMDAR translocate between the Golgi and the synaptic plasma membrane in an activity-dependent manner to modulate synaptic plasticity (Malinow and Malenka, 2002; Lau and Zukin, 2007). Palmitoylation regulates AMPAR and NMDAR cycling in a complex manner. NMDAR subunits NR2A and NR2B have two distinct clusters of palmitoylation sites in their C-terminal region (Hayashi et al., 2009). In mature neurons, palmitoylation of the first cluster upregulates NR2A tyrosine phosphorylation, which leads to stabilized surface expression of NMDAR, whereas palmitoylation of the second cluster causes NMDAR to accumulate in the Golgi (Hayashi et al., 2009). Therefore, specific activity-dependent depalmitoylation of cluster II cysteines must occur in the Golgi apparatus to allow timely delivery to the plasma membrane. Likewise, AMPAR subunits are also palmitoylated at two distinct sites, which results in surface expression or induced internalization, depending on the site that is palmitoylated (Hayashi et al., 2005). These activity-dependent events do not support the model that PATs are Golgi-localized, and APT1/2 activity is ubiquitous throughout the cell.  1.4.3 Concluding Remarks While the ubiquitous APT depalmitoylation model presented by Rocks et al. holds true to explain the constitutive palmitate cycling for cellular substrates such as Ras (Rocks et al., 2010), the major issues with this model are:   50 (1) The identity of the enzyme(s) responsible for depalmitoylating many of the substrates discussed above has not been established. In some cases, it is not even clear if these substrates undergo basal dynamic palmitate turnover. (2) Some substrates clearly exhibit altered rates and levels of protein palmitoylation/ depalmitoylation in the presence of extracellular stimuli, and the mechanism by which this phenomenon occurs is largely unexamined.  Thus, the model proposed by Rocks et al. (2010) cannot be applied to all palmitoylated proteins, and it will be important to identify other molecular players that regulate palmitoylation/depalmitoylation cycles to understand the mechanisms that control palmitate levels of different substrates.  1.5 Thesis Goals and Objectives The primary goal of this thesis is to discover novel factors that play a role in dynamic protein palmitoylation, and to define how they modulate the palmitoylation or depalmitoylation of selected substrates. Thus far, studies of protein depalmitoylation have focused on the activity of APT1 towards a subset of substrates; however, the existence of additional cytosolic palmitoylthioesterases remains largely unaddressed. Meanwhile, changes in the localization or activity of specific DHHC-CRD-containing PATs can also and have been shown to modulate palmitoylation, but whether this is a general property of all PATs, and whether there are additional regulators of PAT activity, remains to be examined. I was involved in a recent DHHC17 (HIP14) interactome study, where we identified the known HTT-interactor, Optineurin (OPTN), as a novel HIP14 binding partner (Butland et al., 2014). We speculated   51 that the binding of OPTN to HIP14 may affect HIP14 function. The overall hypothesis of this thesis is that there are some yet undiscovered cytosolic APTs besides APT1 and APT2 that mediate the depalmitoylation of a subset of proteins, and that there are novel regulators of PATs that can mediate dynamic palmitoylation through its trafficking and/or a co-factor. Since OPTN is a known cargo adaptor protein, it may bind HIP14 to mediate its trafficking (del Toro et al., 2009). Thus, the specific objectives were to: 1. Determine if APT1 and APT2 are responsible for bulk depalmitoylation in the cell 2. Determine the identity of additional SH(s) that play a role in protein depalmitoylation 3. To determine whether the interaction between OPTN and HIP14 is important for HIP14 enzymatic activity.    52 Chapter 2: APT1/APT2-Independent Protein Depalmitoylation2   2.1 Introduction Protein S-palmitoylation refers to the post-translational attachment of the 16-carbon fatty acid, palmitate, to cysteine residues (Salaun et al., 2010). Protein palmitoylation can be highly dynamic and represents the only reversible lipid modification (Magee et al., 1987; Conibear and Davis, 2010). Dynamic palmitoylation is implicated in the modulation of protein localization, trafficking, and protein-protein interactions (Rocks et al., 2010; McCormick et al., 2008; Conibear and Davis, 2010; Greaves and Chamberlain, 2011b) and involves two enzyme families. Protein acyl-transferases (PATs) are conserved cysteine-rich DHHC (Asp-His-His-Cys) domain-containing transmembrane proteins that mediate palmitate addition (Greaves and Chamberlain, 2011a), whereas acyl palmitoyl thioesterases (APTs) belong to the metabolic serine hydrolase (mSH) superfamily, and remove palmitate from proteins (Long and Cravatt, 2011).   Palmitate turnover has been demonstrated in several cellular contexts (Wedegaertner and Bourne, 1994; El-Husseini et al., 2002; Baker et al., 2003; Zhang et al., 2010; Brigidi et al., 2014). In particular, stimulation or blockade of synaptic activity leads to dramatic alterations in the palmitoylation of many neuronal proteins, which accompany changes in synaptic                                                 2 A version of this chapter is being prepared for submission. Lin, DT and Conibear, E. “APT1- and APT2-indpendent depalmitoylation.” DTL designed and conducted all the experiments. DTL and EC analyzed the data and wrote the manuscript for publication.    53 plasticity (Kang et al., 2008b). One such protein, post synaptic density 95 (PSD-95), is a neuronal scaffolding protein whose synaptic targeting is dependent on its palmitoylation status (El-Husseini et al., 2002; Craven et al., 1999). Activity-dependent changes in PSD-95 palmitoylation result from local cycling of its PAT, DHHC2, in the postsynaptic density to mediate its repalmitoylation (Noritake et al., 2009; Fukata et al., 2013). However, the APT(s) that depalmitoylate PSD-95 to allow its re-entry into the dynamic palmitoylation cycle are not yet known.  To date, only two cytosolic APTs have been found to mediate the enzymatic removal of palmitate from cytosolic cysteine residues in vitro and in vivo (Duncan and Gilman, 1998; Tomatis et al., 2010; Siegel et al., 2009). APT1 and APT2 are closely related α/β hydrolases that contain a signature GxSxG motif and Ser-His-Asp catalytic triad (Long and Cravatt, 2011). Whereas APT2 is only expressed in mammals (Tomatis et al., 2010; Rusch et al., 2011), APT1 is a highly conserved enzyme that is also found in yeast and Toxoplasma (Duncan and Gilman, 2002; Child et al., 2013), and regulates dendritic spine size in cultured rat hippocampal neurons through the depalmitoylation of Gα13 (Siegel et al., 2009). Both APT1 and APT2 have been shown to target other proteins, including N- and H-Ras, when expressed in cultured cells (Tomatis et al., 2010; Siegel et al., 2009; Tian et al., 2012), and in vitro (Yeh et al., 1999; Veit and Schmidt, 2001; Flaumenhaft et al., 2007).    Efforts to determine the role of these two APTs were facilitated by the recent development of small molecule inhibitors. Palmostatin B (PalmB), which targets APT1 and 2, potently inhibited Ras depalmitoylation in heterologous cells (Dekker et al., 2010; Rusch et al., 2011).   54 Hexadecyl fluorophosphonate (HDFP), which inhibits a subset of serine hydrolases including APT1 and APT2, suppressed the palmitate turnover of numerous proteins in immune T cell hybridomas (Martin et al., 2011). Although a number of studies have shown that manipulating APT1 and APT2 expression level or activity can cause quantitative changes in palmitate level (Yeh et al., 1999; Dekker et al., 2010; Tian et al., 2010), it is unclear if APT1 and APT2 are the only palmitoylthioesterases responsible for the depalmitoylation of cytosolic proteins.   Here, we examine the substrate specificity of APT1 and APT2 using a dual pulse-chase strategy that directly compares the time course of palmitate loss and protein turnover under controlled conditions. We show that APT1 and APT2 are redundantly responsible for the depalmitoylation of huntingtin protein. In contrast, neither APT1 nor APT2 mediate palmitate removal from PSD-95, despite the robust inhibition of PSD-95 depalmitoylation by the APT1/2 inhibitor PalmB. We show that PalmB has targets other than APT1 and APT2, and define a small number of uncharacterized serine hydrolases, including BAT5 and the FAM108 family of proteins, as potential palmitoylthioesterases. These results reinforce the substrate selectivity of APTs and highlight the role of novel APTs in protein depalmitoylation.  2.2 Materials and Methods 2.2.1 cDNAs & siRNAs pcDNA Myc-hAPT1 was a generous gift of Dr. Tetsuro Izumi (Gunma University). GFP-N-Ras, PSD-95-GFP, N-HTT-GFP, SNAP25-GFP were provided by Dr. Michael Hayden   55 (University of British Columbia), and GAD65-GFP was a gift from the late Dr. Alaa El-Husseini (University of British Columbia). GOLIM4-GFP was a gift from Dr. Adam Linstedt (Carnegie Mellon University). FLAG-SPRED2 was a gift from Dr. Akihiko Yoshimura (Keio University). GFP-ITM2B was cloned by polymerase chain reaction (PCR) amplification of the ITM2B ORF from MGC Fully sequenced Human ITM2B cDNA, clone ID 3163436 (OpenBiosystems;  http://www.ncbi.nlm.nih.gov/nuccore/BC000554), using the forward primer ATTTAACCCGGGATGGTGAAGATTAGCTTCCAGCC (including an XmaI site) and the reverse primer ATTTAAGGTACCTCACACCACCCCGCAGAT (including a KpnI site), followed by restriction digest and ligating the flanking sites to pEGFP-C3 vector (Clontech) linearized with BspEI-KpnI digestion.   For cloning of mSHs, plasmids containing corresponding human ORFs (open reading frames) were purchased from DNASU and OpenBiosystems, or obtained as clones from the hORFeome v8.1 Collection (Yang et al., 2011). Genes of interest were amplified by PCR with flanking restriction sites. mSH-encoding PCR products were subcloned into the vector of interest (FLAG-CT or FLAG-NT, generously gifted by Dr. Stefan Taubert, University of British Columbia; or pCINeo, Promega). See Table 2.1 for oligos used for PCR and restriction sites used for cloning into their respective plasmids.   For knockdown studies, shRNA pSUPER-APT1, pSUPER-scrambleAPT1, and pSUPER-control plasmids were generous gifts from Dr. Gerhard Schratt (University of Marburg). Smartpool siGENOME siRNAs to APT2, was purchased from Dharmacon (Thermo).    56 Table 2.1 List of Oligos Used for Cloning mSH Constructs.      57 2.2.2 Chemicals Lipofectamine 2000, sodium dedocyl sulfate (SDS) solution, L-azidohomoalanine (L-AHA), Alexa Fluor 488-azide (AF488-az), and Alexa Fluor 647-alkyne (AF647-alk) were purchased from Life technologies. X-tremeGENE 9 was purchased from Roche. Palmostatin B was purchased from Merck Scientific. Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA), Tris(2-carboxyethyl)phosphine hydrochloride (TCEP), Triton-X 100 (TX-100), sodium deoxycholate, CuSO4, palmitic acid, and 2-bromopalmitate (2-BP) were obtained from Sigma-Aldrich.17-ODYA (17-octadecynoic acid), MAFP (Methylarachidonyl fluorophosphonate), THL (Tetrahydrolipstatin), BEL (Bromoenol lactone), C75, WWL70, and RHC 80267 were purchased from Cayman Chemical. HDFP (Hexadecylfluorophosphonate), C83, and C115 were generously gifted by Dr. Brent Martin (University of Michigan), and FP-rhodamine was generously provided by Dr. Benjamin Cravatt (Scripps Institute).   2.2.3 Cell Culture Conditions  COS-7 cells were maintained and propagated in high glucose Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS; Life Technologies), 4mM L-Glutamine and 1mM sodium pyruvate, in a humidified incubator at 37°C, 5% CO2.  2.2.4 cDNA & siRNA Transfections For Pulse-chase metabolic studies and activity-based protein profiling studies, COS-7 cells were transfected with cDNAs as indicated in each experiment using Lipofectamine 2000 as per manufacturer’s instructions. Cells were grown in 6-well plates (CoStar) and transfected at   58 90% confluence with 1µg of cDNA per well. For immunofluorescence studies, COS-7 cells were grown on glass coverslips (Fisher) in 24-well plates (CoStar) and transfected at 60-90% confluence with 0.5µg of cDNA per well using Xtreme-GENE 9 according to product instructions. All experiments were carried out 20-24 hours following transfections as described below.  For knockdown studies with siRNA, a double knockdown approach was used (Bond et al., 2011) where COS-7 cells were transfected with siRNA (100 nM final concentration per transfection) on days 1 and 3 with 5µL of Lipofectamine 2000 per transfection. 1µg of cDNA was co-transfected with the siRNA on day 3, and pulse-chase studies were carried out on day 4, at 20 hours following the co-transfection.  2.2.5 Small Molecule Inhibitor Treatments Unless otherwise indicated, for all experiments where treatments with small molecule inhibitors were carried out, the chemicals were used at the following concentrations: PalmB (10µM), C83 (10µM), C115 (10µM), HDFP (20µM), THL (20µM), MAFP (20µM), RHC 80267 (20µM), WWL70 (10µM), C75 (10µM), BEL (20µM), and 2-BP (100µM). All chemicals were solubilized in DMSO to make 1,000x stock solutions that are kept at -20°C until use. Freeze and thaw cycles were minimized.  2.2.6 Pulse-Chase Metabolic Labeling with Inhibitors Twenty hours following transfection, COS-7 cells were washed twice in phosphate-buffered saline (PBS) and then starved in cysteine- and methionine-free DMEM containing 5%   59 charcoal-filtered FBS (Invitrogen) for 1 hour. Cells were then labeled with 30µM 17-ODYA and 50µM L-AHA for 1.5 hours in this media. For chase, cells were removed of the labeling media, and briefly washed twice in PBS. Cells were then incubated in complete DMEM supplemented with 10% FBS and 300µM palmitic acid. Small molecule inhibitors or DMSO (vehicle) were added at time 0. At indicated time points, cells were washed twice in PBS then lysed with 500µL triethanolamine (TEA) lysis buffer [1% TX-100, 150 mM NaCl, 50 mM TEA pH 7.4, 2x EDTA-free HaltTM Protease Inhibitor (Thermo)]. The lysates were transferred to 1.5mL Eppendorf tubes (CoStar), vigorously shaken (3 x 20s) while placed on ice in between each agitation. Lysates were cleared by centrifugation at 16,000 x g for 15 minutes at 4°C. Solubilized proteins in the supernatant were quantified using Bicinchoninic acid (BCA) assay (Pierce) and subsequently used for immunoprecipitations as described below.  2.2.7 Immunoprecipitations For immunoprecipitations, Protein A or Protein G sepharose beads (GE Healthcare) were washed thrice in TEA lysis buffer. Protein A beads were pre-incubated with rabbit anti-GFP antibodies (Life Technologies) and Protein G beads were pre-incubated with FLAG M2 antibodies (Sigma) for 2 hours at 4°C, before the addition 500µg – 1mg of transfected COS-7 cell lysates containing indicated proteins. Immunopreciptations were carried out for 12-16 hours on an end-to-end rotator at 4°C. Following immunoprecipitation, sepharose beads were washed thrice in modified RIPA buffer (150mM NaCl, 1% Sodium deoxycholate (w/v), 1% TX 100, 0.1% SDS, 50mM TEA pH7.4) before proceeding to sequential on-bead CuAAC/click chemistry.   60 2.2.8 Sequential On-Bead CuAAC/Click Chemistry Sequential on-Bead click chemistry of immunoprecipitated 17-ODYA/L-AHA labeled proteins was carried out as previously described (Zhang et al., 2010), with minor modifications. After immunoprecipitation, sepharose beads were washed three times in RIPA buffer, and resuspended in 50µL PBS containing 3.5µL of freshly premixed click chemistry reaction mixture (Final concentrations in reaction: 1mM TCEP in dH2O, 1mM CuSO4·5H2O in dH2O, 100µM TBTA in DMSO, and 100µM AF488-az in DMSO). The reaction was carried out for 1 hour at room temperature, followed by three washes in 500µL RIPA buffer. The subsequent reaction was carried out in 50µL RIPA buffer containing 3.5µL of freshly premixed click chemistry reagents (AF647-alk in place of AF488-az) for 1 hour at room temperature. Beads were washed thrice with RIPA buffer then resuspended in 10µL SDS buffer (150mM NaCl, 4% SDS, 50mM TEA pH7.4), 4.35µL 4x SDS-loading buffer (8% SDS, 4% Bromophenol Blue, 200mM Tris-HCl pH 6.8, 40% Glycerol), and 0.65µL 2-mercaptoethanol. Samples were heated for 5 min at 95°C, and separated on 10% tris-glycine SDS-PAGE gels for subsequent in-gel fluorescence analyses as described below.  2.2.9 Competitive Activity-Based Protein Profiling  Twenty-four hours following transfection with corresponding mSH constructs, COS-7 cells were washed twice in PBS and transferred to a new vial by scraping in PBS. Cells were lysed by gentle sonication on ice (18 Amps, 4 seconds). Protein amounts were quantified by BCA assay. 30µg of total protein were treated either with DMSO or small molecule inhibitors at indicated concentrations at room temperature for 30 minutes, prior to the addition of FP-Rho (10µM final concentration). Labeling reactions were carried out at room temperature for 1   61 hour. Reactions were quenched with 4x SDS sample buffer and heated to 95°C for 5 minutes. Samples were separated on SDS-PAGE and in-gel fluorescence was analyzed with Typhoon Trio scanner (GE Healthcare) as described. Gels were transferred onto nitrocellulose membrane for Western blotting analysis, as described below.  2.2.10 In-Gel Fluorescence Analyses SDS-PAGE gels containing Alexa Fluor- or rhodamine-tagged proteins were immediately used for in-gel fluorescence analysis using the Typhoon Trio scanner (GE). For pulse-chase studies, AF488 signals were acquired using the blue laser (excitation 488nm) with selected emission filter of 520BP40; AF647 signals were acquired using the red laser (excitation 633nm) with selected emission filter of 670BP30; and rhodamine signals were acquired with the green laser (excitation 532nm), with emission filter set at 580BP30. ImageQuant TL 7.0 software (GE) was used for signal quantification.  2.2.11 Western Blotting   Nitrocellulose membranes with transferred proteins were blocked with PBS with 0.1% Tween-20 (PBST) containing 3% bovine serum albumin (BSA, Sigma) for 1 hour. Membranes were incubated with primary antibodies (anti-rabbit GFP, 1:1,000; or anti-mouse FLAG M2, 1:1,000) in PBST + 3% BSA for 2 hours, followed by 3x 15 minute washes with PBST + 0.3% BSA. Membranes were further incubated with secondary antibodies (IRDye® 800CW goat anti-mouse IgG,1:10,000; or IRDye® 680RD goat anti-rabbit IgG,1:10,000) in PBST + 0.3% BSA for 1 hour. After three final washes in PBST, membranes were imaged using the Li-COR Odessey Scanner (Li-COR). Images were acquired using the 680nm and   62 800nm lasers. Signals were acquired in the linear range and quantified using the Image StudioTM software (Li-COR).  2.2.12 PSD-95-GFP Localization by Microscopy & Cell Counting Twenty hours post-transfection, cells were briefly washed twice with PBS, then re-incubated in fresh DMEM medium containing inhibitors or DMSO (vehicle) for 4 hours. Cells were washed briefly with PBS twice then fixed in 4% paraformaldehyde (PFA) solution (4% PFA, 4% sucrose in PBS) for 30 minutes, then washed three more times in PBS. Coverslips were mounted onto glass slides with ProLong® Gold Antifade Mountant with DAPI (Life Technologies). Images were visualized with an Axioplan 2 fluorescence microscope (Carl Zeiss) using a Plan-Apochromat 100x 1.40 NA oil immersion objective lens. Images were taken with a CoolSNAP camera (Roper Scientific) using MetaMorph 7.7 software (MDS analytical Technologies), and adjusted using Metamorph 7.7. Images were acquired with the GFP and DAPI filters. In each experiment, 100 cells were counted to determine the ratio of cells showing perinuclear PSD-95-GFP localization (n = 3).  2.2.13 Statistical Analyses  Statistical analyses were carried out by performing Student’s two-tailed t-tests using Prism 6  (GraphPad Software, Inc., La Jolla, CA).  A p < 0.05 was considered to indicate a statistically significant difference.      63 2.3 Results 2.3.1 Palmitate Turnover is Substrate-Specific APT1 was proposed to act constitutively to remove palmitate from a variety of substrates (Rocks et al., 2010). The physiological rate of depalmitoylation varies widely between proteins, which may reflect substrate-specific differences in enzyme activity. However, even for a single protein, palmitate half-life measurements can vary based on cell type, expression level, and the method used to detect palmitoylation (Lane and Liu, 1997; Heindel et al., 2003; Kang et al., 2004; Greaves and Chamberlain, 2011b). Failure to accurately measure protein turnover rate can also be an important confounding factor (Qanbar and Bouvier, 2004).  To examine if different substrates have inherently different rates of depalmitoylation when expressed in the same cell type, we surveyed several established palmitoyl-proteins proteins using a dual pulse-chase scheme that simultaneously measures protein and palmitate turnover as adopted from Zhang et al. (Zhang et al., 2010). GFP- or FLAG-tagged versions of each protein were expressed in COS-7 cells and subjected to pulse-chase metabolic labeling with the palmitate analogue 17-octadecynoic acid (17-ODYA) and the methionine surrogate L-azidohomoalanine (L-AHA) (Martin and Cravatt, 2009; Zhang et al., 2010). Following cell lysis and immunoprecipitation of the protein of interest, analogues were conjugated with distinct fluorophores by CuAAC/click chemistry (Kolb et al., 2001). The level of palmitoylation remaining at each time point was adjusted for protein degradation by calculating the ratio of 17-ODYA and L-AHA signals (Figure 2.1A).    64      0 1 2 3 4010.51.520 2 4 010.51.5260 1 2 3 4010.51.522.50 1 2 3 4010.51.52CHASE (Hours)PALM B (10µM)0 0.5GFP-N-RAS0.5+17-ODYAL-AHAACHASE (Hours)PALM B (10µM)PSD95-GFP17-ODYAL-AHA0 2 2+4 4+CHASE (Hours)PALM B (10µM)17-ODYAL-AHA0 2GAD65-GFP2+CHASE (Hours)PALM B (10µM)0 4SNAP25-GFP4+17-ODYAL-AHABCHASE (Hours)PALM B (10µM)0 3N-HTT-GFP3+6 6+17-ODYAL-AHAPalmitoylated N-HTTCHASE (Hours)PALM B (10µM)0 2SPRED2-GFP4+4 No ODYA17-ODYAL-AHAPalmitoylated SPRED20 2GOLIM4-GFP2+4 4+CHASE (Hours)PALM B (10µM)17-ODYAL-AHAPalmitoylated GOLIM40 2GFP-ITM2B2+4 4+CHASE (Hours)PALM B (10µM)17-ODYAL-AHAPalmitoylated ITM2BChase (Hours)Palm BDMSOPalm BDMSOChase (Hours)Chase (Hours) Chase (Hours)Palm BDMSOPalm BDMSO  65 Figure 2.1 Palmitoylation Dynamics Differ between Substrates. Representative in-gel fluorescence images showing palmitate (“17-ODYA”) and protein (“L-AHA”) turnover of each test substrate. COS-7 cells expressing indicated proteins were metabolically labeled with the palmitate analogue 17-ODYA and methionine analogue L-AHA for 90 minutes, followed by chase in excess palmitate and methionine in the presence of PalmB or DMSO (vehicle). Proteins were immunoprecipitated at indicated time points post-labeling for on-bead click chemistry to fluorescently tag the analogues for in-gel fluorescence. The amount of palmitoylated substrate remaining at each time point was quantified as the ratio of (17-ODYA signal/L-AHA signal at chase time):(17-ODYA signal/L-AHA signal at time 0)  (A) The established palmitoylated substrates N-Ras, SNAP25, PSD-95, and GAD65 all display distinctive palmitate turnover rates and their palmitate turnovers are inhibited by the APT1/APT2 inhibitor PalmB (n = 2 for SNAP25 and PSD-95; n = 3 for N-Ras and GAD65). The PSD95-GFP image is a composite of multiple lanes from in-gel fluorescence images of a single polyacrylamide gel scan. (B) N-HTT is dynamically palmitoylated and its palmitate turnover is affected by PalmB, whereas SPRED2, GOLIM4, and ITM2B are stably palmitoylated (n = 2 each). Error bars indicate SEM from two independent experiments.  We found the well-established substrate N-Ras (Rocks et al., 2005) had a rapid palmitate turnover (~24% remaining after 0.5 hours of chase), in agreement with previous estimates under similar experimental conditions (Magee et al., 1987) (Figure 2.1A, upper left panel, n = 3). In contrast, SNAP25 turned over slowly (~75% remaining 4 hours post-chase), consistent with past reports (Lane and Liu, 1997; Greaves and Chamberlain, 2011b) (Figure 2.1A, upper right panel, n = 2). Intermediate rates of depalmitoylation were found for the synaptic proteins PSD-95 and the glutamate decarboxylase subunit GAD65. The 1 hour palmitate half-life of PSD-95 (Figure 2.1A, lower left panel, n = 3) was faster than the 2 hours as previously determined by pulse-chase labeling in rat hippocampal neurons with [125I]Palmitate (El-Husseini et al., 2002). Palmitate turnover on GAD65 (~70% remaining 2 hours post-chase) was slower than on PSD-95 (Figure 2.1A, lower right panel, n = 3). For each of these proteins, treatment with the APT1/2 inhibitor Palmostatin B (PalmB) inhibited palmitate turnover without affecting protein turnover (Figure 2.1A). This suggests that the rate of palmitate turnover is a characteristic of the substrate, and that palmitate removal from SNAP25, PSD-95 and GAD65 may be mediated by APT1 and/or APT2, as previously suggested for N-Ras (Dekker et al., 2010).   66 Next, we examined proteins whose palmitate turnover is less well characterized. Palmitoylation of the Huntingtin (HTT) N-terminal domain has been implicated in the pathogenesis of Huntington’s disease (Singaraja et al., 2002; Huang et al., 2004; Yanai et al., 2006). The disease-causing CAG expansion results in less efficient HTT palmitoylation due to decreased interaction with its PAT (Singaraja et al., 2002); however, the APT responsible for HTT depalmitoylation is not known. The huntingtin N-terminal domain (N-HTT; amino acids 1-548) (Hermel et al., 2004) showed dynamic palmitate turnover that was blocked in the presence of PalmB (Figure 2.1B, upper left panel, n = 3), suggesting N-HTT could be a substrate of APT1 or APT2.   A recent global proteomic analysis of dynamic palmitoylation (Martin et al., 2011) identified several proteins whose palmitate turnover did not appear to be blocked by the APT inhibitor HDFP. We selected three proteins that were identified as minor hits in this study, including the Shiga toxin receptor GOLIM4 (Golgi integral membrane protein 4; GPP130) (Mukhopadhyay and Linstedt, 2012), which was not previously known to be palmitoylated; ITM2B (Integral membrane protein 2B), which is implicated in the pathogenesis of familial dementias (Vidal et al., 1999); and SPRED2 (Sprouty-related, EVH1 comain-containing protein 2), the disruption of which has been shown to cause dwarfism (Bundschu et al., 2005). We confirmed the palmitoylation status of each protein, but did not observe significant palmitate turnover, suggesting the apparent decline in palmitate labeling, which was PalmB-resistant, was due to protein instability (Figure 2.1B, n = 2). Taken together, these results show that palmitate turnover occurs at varying rate for each protein, and that not   67 all modified proteins are subject to rapid depalmitoylation. In all cases examined, PalmB inhibited the palmitate turnover of dynamically palmitoylated proteins.   2.3.2 APT1 and APT2 Display Substrate Specificity PalmB is thought to disrupt palmitate cycling via potent inhibition of both APT1 and APT2 (Dekker et al., 2010; Rusch et al., 2011). Because COS-7 cells express both APT1 and APT2, as determined by qPCR and Western blotting analyses (data not shown), one or both of these enzymes may be responsible for the observed depalmitoylation. In the presence of the selective small molecular inhibitors C83 and C115, which are specific for APT1 and APT2, respectively (Adibekian et al., 2012), we observed minimal inhibition of N-HTT depalmitoylation after a 3 hours chase when either inhibitor was used alone (Figure 2.2A,B, n = 3). Interestingly, simultaneous treatment with C83 and C115 resulted in a synergistic effect that translated to significant inhibition of N-HTT palmitate turnover (p=0.033), which represented 73.6% of the inhibition seen with PalmB (p=0.002). A similar synergistic effect was observed for GAD65 depalmitoylation, whereas C83 and C115 had no effect on PSD-95 depalmitoylation when these inhibitors were used alone or together (data not shown).    To verify these observations, we used RNAi (RNA interference) to perform a double knockdown of APT1 and APT2. We observed a significant inhibition of N-HTT depalmitoylation, comparable to that observed for PalmB (Figure 2.2C,D, n = 3). A similar effect of APT1/APT2 knockdown was also seen for GAD65 (data not shown), further confirming GAD65 as a cellular target of both APT1 and APT2. In contrast, APT1/APT2 double knockdown using RNAi produced no inhibition of PSD-95 depalmitoylation (Figure   68 2.2E,F, n = 3). The results of the inhibitor and knockdown experiments indicate that while APT1 and APT2 are responsible for the depalmitoylation of a subset of palmitoylated proteins (i.e. N-HTT, GAD65), depalmitoylation of other cellular substrates, including PSD-95, does not require APT1 and APT2, and that the specificity of PalmB is different that previously thought in that it may also inhibit APTs other than APT1 and APT2.        Figure 2.2 APT1 and APT2 Inhibition Inhibits N-HTT, but not PSD-95 Palmitate Turnover. Pulse-chase metabolic labeling in COS-7 cells expressing indicated substrates, immunoprecipitation and click chemistry, in gel-fluorescence and image quantification were performed as described in Figure2.1. (A,B) N-HTT palmitate turnover is significantly inhibited by PalmB or co-treatment with the APT1 selective inhibitor C83 and the APT2 selective inhibitor C115, but not when C83 or C115 was used alone (n = 3, Two-tailed Student’s t-test). (C,D) N-HTT in cells with APT1 and APT2 knocked down show significant palmitate turnover inhibition comparable to PalmB or C83 and C115 co-treatment (n = 3, Two-tailed Student’s t-test). (E,F) PSD-95 depalmitoylation is inhibited by PalmB but unaffected by APT1 and APT2 knockdown or C83 and C115 co-treatment (n = 3, Two-tailed Student’s t-test). Error bars indicate SEM from three independent experiments. Palmitoylated N-HTT(relative to Time 0)T=0 3 Hrs 6 HrsChase Time:DMSO APT1/2 RNAi C83+C115 Palm BL-AHA17-ODYACHASE (3 Hours) APT1/2 KNOCKDOWNSC83&C115 6HrsC83&CHASE (6 Hours)3HrsCHASENOHTTNOHTTT=0 DMSO PalmBDMSO PalmB T=0N-HTT-GFPC115C Palmitoylated PSD95(relative to Time 0) 2 Hrs 4 HrsChase Time:L-AHA17-ODYACHASE (2 Hours) APT1/2 KNOCKDOWNSC83&C115 4HrsC83&CHASE (4 Hours)2HrsCHASENOPSDT=0 DMSO PalmBDMSO PalmB T=0PSD95-GFPC115NOPSDDMSO APT1/2 RNAi C83+C115 Palm B00. hours chaseDMSO PalmB C83 C115 C83&C115Palmitoylated N-HTT p=0.033p=0.002T=0AL-AHA17-ODYACHASE (3 Hours)C83&C115DMSOT=0N-HTT-GFPPalmB C115C83p=0.003p<0.001p=0.011 p<0.001p<0.001p<0.001BDEF  69 2.3.3 Palmostatin B (PalmB) has Additional Serine Hydrolase Targets Previous approaches suggested APT1, APT2, and the luminal palmitoylthioesterase PPT1 were the sole serine hydrolases targeted by PalmB (Rusch et al., 2011). However, another depalmitoylation inhibitor, HDFP (hexadecylfluorophosphonate), has its targets determined by SILAC-ABPP and the target list included several serine hydrolases other than APT1 and APT2 (Martin et al., 2011). In pulse-chase experiments, we found HDFP robustly inhibited the depalmitoylation of both PSD-95 and N-HTT (Figure 2.3A-D, n = 3 for PalmB and n = 1 for HDFP). Because palmitate removal on PSD-95 does not require APT1 or APT2, PSD-95 depalmitoylation is likely mediated by a distinct serine hydrolase that is a common target of both PalmB and HDFP.   To identify overlapping targets of HDFP and PalmB, we defined an initial list of 28 candidate SHs that showed >25% inhibition by HDFP (Martin et al., 2011). We added to this list LYPLAL1 (also known as APT1L), which was previously implicated in the depalmitoylation of the big calcium-activated potassium (BK) channel (Tian et al., 2012), and excluded 3 known proteases (DPP8, PREPL, and PARL) and 7 SHs with established luminal activity (PPT1, ABHD12, PGAP1, AADACL1, LIPA, and LYPLA3, SERHL2), leaving 20 SHs for our analysis (Table 2.2).    70  FP-RhoWBPNPLA6 ABHD6FAM108A1 FAM108B1FAM108C1 BAT5FASNPalmB + + + + + + +FAAH PAFAH2 ABHD4ACOT1 ACOT2ABHD10(NO TAG)APT1L(NO TAG)FP-RhoWBPalmB + + + + + + +APT1 APT2FP-RhoWBPalmB + +(NO TAG)ACHASE (2 Hrs)PSD95-GFPT=0 DMSOPalmBHDFPCHASE (4 Hrs)17-ODYADMSOPalmB HDFPL-AHADMSO PalmB HDFP0. % Palmitoylated PSD9501.01.21.4T=0 2 Hrs 4 Hrs(relative to Time 0)Chase Time:DMSO PalmB HDFPCHASE (3 Hrs)N-HTT-GFPT=0 DMSOPalmB HDFPCHASE (6 Hrs)17-ODYADMSOPalmBHDFPL-AHA0. % Palmitoylated N-HTT(relative to Time 0)T=0 3 Hrs 6 HrsChase Time:BCDEF% Activity Blockade by PalmB(Relative to DMSO)02080100120FAAHABHD10APT1LACOT1ACOT2PAFAH2ABHD4FAM108C1FASNPNPLA6FAM108A1ABHD6FAM108B1BAT5APT2APT16040*  71 Figure 2.3 Palmostatin B and HDFP Inhibit PSD-95 and N-HTT Palmitate Turnover and Share Overlapping Serine Hydrolase Targets.  (A-D) Pulse-chase metabolic labeling in COS-7 cells expressing indicated substrates, immunoprecipitation and click chemistry, in gel-fluorescence and image quantification were performed as described in Figure2.1. (A,B) PalmB and HDFP show similar extent of inhibition of PSD-95 palmitate turnover (n = 3 for PalmB; n = 1 for HDFP). (C,D) HDFP inhibits N-HTT palmitate turnover more strongly than PalmB (n = 3 for PalmB;  n = 1 for HDFP). Note that (A) and (C) are composites of multiple lanes from in-gel fluorescence images of a single polyacrylamide gel scan. (E,F) Targeted competitive activity-based protein profiling reveals multiple novel targets of PalmB. COS-7 cells expressing indicated serine hydrolases were lysed by sonication under non-denaturing conditions, and the lysates were treated with DMSO (-) or 10µM PalmB (+) for 30minutes prior to labeling with Fluorophosphonate-rhodamine (FP-Rho) for 1 hour. The extent of PalmB activity inhibition of each enzyme were determined by in-gel fluorescence as shown in (E), and quantified as 1- (PalmB:DMSO) signal ratio in a graph (F) (n = 3 for each mSH). * indicates endogenous reactivity. Error bars indicate SEM from three independent experiments.    The activity of each candidate was determined in the presence or absence of PalmB using a competitive ABPP (cABPP) approach, in which enzymes are expressed in COS-7 cells and treated with either PalmB or DMSO in the presence of the activity probe fluorophosphonate-rhodamine (FP-rho) (Kidd et al., 2001) (Table 2.2). Binding of PalmB occludes the active site and prevents labeling by FP-rho, leading to a reduced signal. As expected, we observed significantly diminished FP-rho labeling of both APT1 and APT2, the two known cytosolic PalmB targets, in the presence of the inhibitor (Figure 2.3E,F, n = 3). PalmB treatment did not affect the labeling of 7 candidates, including FAAH, ABHD10, LYPLAL1, ACOT1/2, PAFAH2, and ABHD4 (Figure 2.3E,F, n = 3). 4 of the 22 SHs (ABHD13, PNPLA7, PNPLA8, and LIPE) did not label with the FP probe due to low activity and could not be assessed (Table 2.2). Strikingly, PalmB potently inhibited 7 of the candidates including FASN, PNPLA6, ABHD6, BAT5 and FAM108A/B/C family members (Figure 2.3E,F, n = 3). Thus, PalmB has additional cellular targets, some of which may function as protein depalmitoylases. These results also show that PalmB and HDFP have different substrate specificities.  72 Table 2.2 HDFP-targeting metabolic serine hydrolases.    73  2.3.4 ABHD6, FASN, PNPLA6, and PNPLA8 do not Play Major Roles in PSD-95 Depalmitoylation This analysis identified at least 7 additional targets of PalmB that represent potential depalmitoylation enzymes (Table 2.2, Figure 2.3E,F). ABHD6 was recently shown to associate with PSD-95-containing α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPAR) complexes at synapses (Schwenk et al., 2012, 2014), whereas FASN functions in the palmitoyl-CoA synthesis pathway (Wakil, 1989) and thus may indirectly affect depalmitoylation. Treatment with 10µM of the established ABHD6 inhibitor WWL70 (Li et al., 2007) gave a highly selective and potent inhibition of ABHD6 with no effect on the remaining 8 PalmB-sensitive serine hydrolases, including APT1 and APT2 (Figure 2.4A, n = 2). However, we saw no change in the palmitate turnover of PSD-95 (Figure 2.4B,C, n = 1).   Next, we targeted FASN, using the established FASN inhibitor C75 (Kuhajda et al., 2000). As C75 irreversibly inhibits the β-ketoacyl-ACP (Acyl Carrier Protein) synthase domain, but not the thioesterase domain of FASN (Nasheri et al., 2013), FP-Rho labeling of FASN remained strong in the presence of C75, and C75 did not inhibit the activity of other candidates (data not shown). Importantly, C75 had no effect on PSD-95 depalmitoylation (Figure 2.4B,C, n = 1). PSD-95 palmitate turnover was also unaffected by Bromoenol lactone (BEL) (Figure 2.4E,F, n = 1), a known PNPLA8 (iPLA2 gamma) inhibitor (Hazen et al., 1991) that did not alter the activity of other candidates (data not shown). Finally, the diacylglycerol lipase inhibitor RHC80267 (Hoover et al., 2008) showed moderate inhibition   74 of ABHD6 and PNPLA6 at 20µM, but exerted little effect on PSD-95 palmitate turnover (Figure 2.4D-F, n = 2).     FAM108A1APT1ABHD6FASNACOT1BAT5PNPLA6DMSOPalm B RHC80267FP-RhoAWWL C75 BEL17-ODYAL-AHACHASE (2 Hrs)PSD95-GFPT=0 DMSOPalmB WWL C75 BELCHASE (4 Hrs)DMSOPalmBDMSO Palm B WWL70 C75 BEL0. Palmitoylated PSD95(relative to Time 0)T=0 2 Hrs 4 HrsChase Time: Palmitoylated PSD95(relative to Time 0)T=0 2 Hrs 4 HrsChase Time:RHC80267 MAFPDMSO Palm B17-ODYAL-AHACHASE (2 Hrs)PSD95-GFPT=0 DMSOPalmB MAFPCHASE (4 Hrs)DMSOPalmBRHC RHC MAFPDMSOPalm B WWL70FAM108A1APT1ABHD6FASNACOT1BAT5PNPLA6FP-RhoDBCEF  75  Figure 2.4 Treatment with WWL70, C75, BEL, and RHC-80267 Does not Affect PSD-95 Palmitate Turnover. (A) WWL70 is highly selective for ABHD6. cABPP assay was carried out as described in Figure 2.3(E) (n = 2). (B-C) The ABHD6 inhibitor WWL70, FASN inhibitor C75, and PNPLA8 inhibitor BEL do not affect PSD-95 palmitate turnover. Pulse-chase metabolic labeling in COS-7 cells expressing PSD-95-GFP, immunoprecipitation and click chemistry, in gel-fluorescence and image quantification were performed as described in Figure2.1 (n = 1). (D) RHC80627 moderately inhibits both ABHD6 and PNPLA6. cABPP assay was carried out as described in Figure 2.3(E) (n = 2) (E-F) RHC80267 treatment does not affect PSD-95 dynamic palmitate turnover while MAFP shows a partial effect.  Pulse-chase metabolic labeling in COS-7 cells expressing PSD-95-GFP, immunoprecipitation and click chemistry, in gel-fluorescence and image quantification were performed as described in Figure2.1 (n = 2). Note that (B) and (E) are composites of multiple lanes from in-gel fluorescence images of a single polyacrylamide gel scan.    These observations suggest that ABHD6, PNPLA6, PNPLA8, and FASN are unlikely to play a key role in PSD-95 depalmitoylation, as their inhibition did not affect palmitate turnover on PSD-95. However, given our finding that APT1 and APT2 act redundantly on N-HTT (Figure 2.2 C,D), we cannot exclude the possibility that one or more of these mSHs can mediate palmitate removal from PSD-95, but their activity is masked by other, more potent, depalmitoylases.   2.3.5 Tetrahydrolipstatin (THL) Treatment Results in Partial Inhibition of PSD-95 Depalmitoylation There are no established specific inhibitors for the remaining 4 candidates, the unannotated BAT5 and FAM108A/B/C family members. However, the lipase inhibitor Tetrahydrolipstatin (THL) was previously shown to inhibit several of these candidates, including BAT5, ABHD6 and FASN (Hadváry et al., 1988; Kridel et al., 2004; Hoover et al., 2008). Of the 9 serine hydrolases moderately inhibited by PalmB in this study (Figure   76 2.3E,F), only ABHD6 and BAT5 showed a robust decrease in FP-rho labeling by the cABPP assay in the presence of THL, consistent with previous findings (Hoover et al., 2008). We also saw a partial reduction in the FP-rho intensity for FASN, but observed little or no inhibition of APT1 or APT2 activity (1% inhibition towards APT2; 7% inhibition towards APT1) (Figure 2.5A,B, n = 3). Interestingly, when cells transfected with PSD-95-GFP were treated with THL, we found a partial yet significant inhibition of palmitate turnover on PSD-95-GFP (p < 0.008 at 4 hours of chase) (Figure 2.5C,D, n = 3), comparable to that observed with the general SH inhibitor methylarachidonyl fluorophosphate (MAFP) (p < 0.001 at 4 hours of chase) (Zhang et al., 2010) (Figure 2.5C,D, n = 3). These results suggest that THL targets a novel PSD-95 depalmitoylation activity. The partial inhibition observed with THL and MAFP suggests either (1) The target(s) of THL, such as BAT5, or others not assessed in this study, could play a modest part in PSD-95 depalmitoylation inhibition; or (2) The drug concentration used here was insufficient to sustain the continuous blockade of serine hydrolase activity due to it being metabolized. We found the latter case to be true for MAFP where chemical stability was the issue (data not shown); however, this still has to be examined for THL.          77  Figure 2.5 Tetrahydrolipstatin Inhibits BAT5 and ABHD6 and Causes Partial Inhibition of PSD-95 Depalmitoylation. (A,B) THL potently inhibits BAT5 and ABHD6 enzymatic activity. cABPP assay and in-gel fluorescence quantification were carried out as described in Figure 2.3(E,F) (n = 3). (C,D) THL partially but significantly inhibits PSD-95 depalmitoylation. Pulse-chase metabolic labeling in COS-7 cells expressing PSD-95-GFP, immunoprecipitation and click chemistry, in gel-fluorescence and image quantification were performed as described in Figure2.1 (n = 3, Two-tailed Student’s t-test). Note that (A) and (C) are composites of multiple lanes from in-gel fluorescence images of a single polyacrylamide gel scan. Error bars indicate SEM from three independent experiments. HDFPDMSOPalm BTHLFAM108A1FAM108B1FAM108C1APT1APT2BAT5ABHD6PNPLA6FASNFP-RhodamineA0. MAFP17-ODYAL-AHACHASE (2 Hrs)PSD95-GFPT=0 DMSO PalmB THL MAFPCHASE (4 Hrs)DMSOPalmBDMSO Palm B THL MAFP Palmitoylated PSD95(relative to Time 0)T=0 2 Hrs 4 HrsChase Time:% Activity Blockade by THL(Relative to DMSO)02080100FAM108C1FASNPNPLA6FAM108A1ABHD6FAM108B1BAT5APT2APT16040BCDp<0.001p<0.001p=0.008p<0.001p=0.005p=0.072  78 2.3.6 Inhibition of PSD-95 Depalmitoylation Alters its Localization We investigated the subcellular localization of PSD-95-GFP following inhibitor treatment to determine if the inhibition of palmitate turnover is reflected in a cellular phenotype. When overexpressed in COS-7 cells, PSD-95-GFP was diffusely localized in most cells, with a small proportion showing strong, perinuclear localization (Figure 2.6A,B, n = 3). The PAT inhibitor 2-bromopalmitate (2-BP) resulted in completely diffuse distribution of PSD-95-GFP, whereas co-overexpression with its PAT, DHHC2, caused a robust shift of PSD-95-GFP localization to a perinuclear compartment and to the PM (Figure 2.6A,B, n = 3). This suggests that a substantial pool of PSD-95 is not palmitoylated when overexpressed in COS-7 cells due to limited abundance or activity of its DHHC PAT.  Treatment with PalmB for 3 hours in situ caused a dramatic shift in PSD-95 to a perinuclear compartment, which could reflect the redistribution of the membrane-associated, palmitoylated form of the protein on vesicular trafficking pathways. The wide-spectrum SH inhibitors MAFP and HDFP caused a partial re-localization (Figure 2.6B, n = 3). A partial shift in PSD-95-GFP was also seen with THL, consistent with the moderate inhibition of depalmitoylation associated with this inhibitor in the click assay (Figure 2.5C,D). In contrast, inhibitors that showed little effect on PSD-95 depalmitoylation, including WWL70, BEL, and the APT1/APT2 inhibitors C83 and C115, did not induce PSD-95-GFP re-localization (Figures 2.2 and 2.4).  These observations demonstrate that PSD-95 localization parallels its palmitoylation status, indicating that changes in dynamic palmitate turnover result in a cellular phenotype.    79   Figure 2.6 Treatment with Inhibitors Causing PSD-95 Palmitoylation Changes Also Leads to Localization Shift of PSD-95-GFP. (A,B) Cos-7 cells expressing PSD-95-GFP were treated with the indicated small molecules in culture or, in the “+DHHC2” condition, was co-transfected with the PSD-95 PAT DHHC2. Cells were fixed 3 hours post-treatment for microscopy. (A) Representative microscopy images showing that in PSD-95-GFP expressing cells treated with DMSO, PSD-95-GFP is more diffuse, whereas PalmB and +DHHC2 conditions that enhance PSD-95 palmitoylation both leads to perinuclear PSD-95 accumulation, compared to a partial re-localization that is seen with THL. Cell nuclei were stained with Hoescht (dark blue). Scale bar, 10µm. (B) Bar graph representing percentage of COS-7 cells with perinuclear PSD-95-GFP as visualized by microscopy from (A). In each trial, 100 cells were counted (n = 3). Error bars indicate SEM from the three independent trials.    2.4 Discussion In the present work, we show that PSD-95 depalmitoylation occurs independently of the known thioesterases APT1 and APT2. APT1 and APT2 knockdown, or use of APT1/2-specific inhibitors, effectively blocked the depalmitoylation of N-HTT. However, these same treatments had no effect on the dynamic palmitate cycling of PSD-95. We find that the APT1/2 inhibitor PalmB targets other metabolic serine hydrolases, which explains its robust inhibition of palmitate turnover on PSD-95. By comparing the inhibition profile of three different small molecules, we identified a subset of uncharacterized serine hydrolases that we propose can function as new candidate PSD-95 depalmitoylation enzymes.  DMSO Palm B+DHHC2 THLA B% Cells withperinuclear PSD95-GFP100806040200DMSOMAFPHDFP THL2-BPPalmBWWL70BELC83&C115+DHHC2  80  APT1 was suggested to mediate the bulk depalmitoylation of many proteins at cellular membranes (Rocks et al., 2010; Vartak et al., 2014). However, palmitate removal from some substrates proceeds at very different rates. Using dual pulse-chase click chemistry to simultaneously monitor palmitate and protein turnover, we found the variation in turnover rate cannot be accounted for by cell type or methodological differences, but appears to be a characteristic feature of the substrate. The conformation of the substrate near the palmitoylation site could alter the efficiency of palmitate turnover and explain some of these differences. For example, the prolyl isomerase 12kDa FK-506 binding protein (FKBP12) causes a conformational change in Ras that enhances its depalmitoylation (Ahearn et al., 2011). APTs could also preferentially access their substrates at a particular cellular compartment. Differences in the rate of depalmitoylation could also arise if a single APT binds some substrates more efficiently than others. For some mSHs, substrate selectivity is conferred by residues proximal to the active site (Lukowski et al., 2014), while others, such as PPT2, restrict access to the substrate binding pocket (van Tilbeurgh et al., 1993; Calero et al., 2003). APT1 and PPT1 have deep substrate-binding pockets that can accommodate longer-chain modifications such as palmitate, in contrast with APT1L, which has a shallower binding groove and is unlikely to hydrolyse palmitate from proteins (Bellizzi et al., 2000; Devedjiev et al., 2000; Burger et al., 2012). However, structural features of APT1 likely to mediate differential substrate specificity have not yet been identified.  The existence of several different APTs, each with distinct substrate preferences, could also explain the different depalmitoylation rates. Using genetic knockdown and specific small   81 molecule inhibitors we found APT1 and APT2 act redundantly on N-HTT and GAD65. However, other reports have suggested these enzymes have different specificities. For example, the localization of GAP43 was affected by the overexpression of APT2 but not APT1 (Tomatis et al., 2010). In contrast, overexpression of APT1, but not APT2, reduced BK channel depalmitoylation (Tian et al., 2012), whereas both enzymes could target H-RasG12V when overexpressed (Tomatis et al., 2010). While the extent to which APT1 and APT2 display substrate selectivity is not yet clear, our finding that these APTs are dispensable for PSD-95 depalmitoylation shows that additional substrate-specific APT(s) must exist. These APTs may be exclusive for each substrate, or perhaps work together with APT1 and APT2 to regulate substrate depalmitoylation.  Few other reports have demonstrated significant changes in depalmitoylation resulting from knockdown of APT1 and/or APT2. Instead, most studies have relied on use of PalmB or other inhibitors. A proteomics strategy using a PalmB-based activity probe suggested APT1, APT2, and PPT1 are the only serine hydrolases targeted by PalmB (Rusch et al., 2011). However, we have shown that PalmB has additional targets, including FASN, PNPLA6, ABHD6, BAT5, and FAM108A1/B1/C1 proteins. Recently, PalmB was found to also inhibit ABHD12 and MAGL (Monoacylglycerol lipase) by cABPP in the mouse brain membrane proteome (Savinainen et al., 2014). Thus PalmB is not as selective as previously believed, suggesting studies using this inhibitor to support a role for APT1/2 should be carefully evaluated. Inhibition of these other PalmB targets has the potential to indirectly affect palmitoylation dynamics, membrane lipid composition and signaling. For example, inhibition of FASN, which is responsible for palmitate synthesis in vivo, could alter palmitoylation   82 kinetics, although it is not expected to change depalmitoylation rates. PNPLA6, ABHD6, and ABHD12 are all highly expressed in neurons (Fiskerstrand et al., 2010; Marrs et al., 2010; Long and Cravatt, 2011), and both ABHD6 and ABHD12 physically associate with AMPAR at synapses where they have established functions in the metabolism of the endocannabinoid ligand 2-AG (Marrs et al., 2010; Schwenk et al., 2012; Blankman et al., 2013; Schwenk et al., 2014). Whether this signaling can indirectly affect PSD95 palmitate turnover remains to be examined.  The small number of enzymes found to be targets of the three inhibitors that block PSD-95 depalmitoylation - PalmB, HDFP and THL - represent strong candidates for new APTs. While BAT5 has lipase activity in vitro (Savinainen et al., 2014), its activity towards lipidated proteins has not been tested. Similarly, the cellular function of the three FAM108 family members is not known. We were unable to find an enzyme whose individual knockdown blocked PSD-95 depalmitoylation. Any residual activity may have been sufficient to maintain normal depalmitoylation rates, or the PSD-95 depalmitoylase may not have been previously identified as an HDFP target (Martin et al., 2011) due to cell type difference or low expression. Alternatively, APTs could exhibit widespread redundancy. The existence of several APTs with overlapping specificity could explain why small molecular inhibitors such as PalmB, HDFP and THL, which inhibit several different mSHs, have proven to be the most effective depalmitoylation inhibitors to date. The mSH superfamily is known to consist of >120 members, only half of which have been functionally annotated (Simon and Cravatt, 2010). Thus, it is likely that PalmB, HDFP, and THL inactivate   83 additional targets that were not analyzed in this study, and which may contribute directly or indirectly to protein palmitoylation.   The total number of cytosolic APTs is not known. APT1 and APT2 could act ubiquitously to maintain the correct subcellular distribution of palmitoylated proteins, while other substrate-specific APTs could act at key cellular locations to fulfill more specialized functions. Activity-dependent regulation of palmitoylation modulates the synaptic localization and interactions of PSD-95 and δ-catenin (Noritake et al., 2009; Fukata et al., 2013; Brigidi et al., 2014). While the stimulated translocation of DHHC enzymes that repalmitoylate proteins is central to this process, APTs are a critical element of the dynamic palmitoylation cycle and may be subject to regulatory inputs. For example, activity-stimulated depalmitoylation of Lck has been demonstrated, although it remains to be determined whether this results from changes in APT activity/localization or substrate conformation (Zhang et al., 2010). In neurons, changes in the local translation of APT1 were suggested to be a mechanism of activity-dependent depalmitoylation (Banerjee et al., 2009); however the substrates affected by such changes are not known. Given these observations, it will be imperative to identify all cellular APTs as a first step towards determining how substrate selectivity and activity-induced depalmitoylation is achieved.    The FAM108 proteins were recently identified as palmitoyl-proteins whose modification is important for their membrane localization (Martin and Cravatt, 2009). These proteins may be subject to an auto-regulatory feedback loop similar to that described for APT1 and APT2 (Kong et al., 2013; Vartak et al., 2014). Alternatively, the subcellular localization of these   84 mSHs may play a role in their substrate selectivity. Whether BAT5 and FAM108 proteins show activity-induced changes in localization or membrane association remains to be explored. Further characterization of these enzymes should provide more insights into their in vivo targets and functions.      85 Chapter 3: The Huntingtin-Binding Partner Optineurin is a Novel Interactor of the PAT HIP143   3.1 Introduction Palmitoylation is a post-translational modification where the 16-carbon saturated fatty acid, palmitate, is added to protein cysteine residues (Greaves and Chamberlain, 2011a). Palmitoylation is reversible and highly dynamic, as a result of palmitoyl-acyl transferases (PATs) that mediate palmitate attachment and acyl-protein thioesterases (APTs) that remove palmitate from proteins (Conibear and Davis, 2010). Dynamic protein palmitoylation is important for the trafficking and folding of various intracellular proteins. For example, the activity-dependent palmitoylation of post-synaptic density 95 (PSD-95) leads to a change in membrane association and conformation that allows it to scaffold α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptors (AMPARs) at the PSD (Fukata et al., 2013). Another well-characterized substrate is the Huntingtin protein (HTT), whose palmitoylation status is important for its proper folding (Yanai et al., 2006). The mutant form of HTT (mHTT), which contains an expanded polyglutamine tract, shows a reduced palmitoylation that may contribute to aggregate formation (Yanai et al., 2006). These aggregates are thought to be central to the pathogenesis of HTT disease by forming a toxic species (Yanai et al.,                                                 3 A portion of this chapter  (Figure 3.1) has been published. Butland , S.L.,Sanders, S.S., Schmidt, M.E., Riechers, S.P., Lin, D.T., Martin, D.D., Vaid, K., Graham, R.K., Singaraja, R.R., Wanker, E.E., Conibear, E., and Hayden, M.R. (2014) “The Palmitoyl Acyltransferase HIP14 Shares a High Proportion of Interactors with Huntingtin: Implications for a Role in the Pathogenesis of Huntington’s Disease.” Hum Mol Genet. 23:4142-4160. I performed the OPTN studies and prepared the related figures.   86 2006) or acting as an agent to upregulate cellular autophagic responses (Arrasate et al., 2004; Bjørkøy et al., 2005).   Recent studies suggest that dynamic palmitoylation is regulated through the localization and/or activity of the PATs. PATs are a family of highly conserved DHHC-motif-containing transmembrane enzymes that are responsible for substrate palmitoylation (reviewed in Mitchell et al., 2006). In neurons, dynamic palmitoylation of PSD-95 is achieved in part by local cycling of DHHC2 in dendritic spines in response to changing synaptic activity (Noritake et al., 2009). On the other hand, dendritic-localized DHHC5 was shown to be required for the activity-dependent palmitoylation of another substrate, δ-catenin (Brigidi et al., 2014). Phosphorylation of DHHC8 downstream of brain-derived neurotrophic factor (BDNF) signaling was also reported to increase PSD-95 palmitoylation, although whether this is through activity, conformation or localization changes was not examined (Yoshii et al., 2011).   Interaction of HTT with its PAT, HIP14, has been shown to enhance HIP14 activity towards other substrates (Huang et al., 2011), suggesting HTT is a regulatory co-factor of HIP14. Wild type, but not mutant HTT, is able to bind HIP14, leading to enhanced SNAP25 palmitoylation by HIP14 in vitro. In vivo, HIP14 autopalmitoylation and substrate palmitoylation are decreased in mice lacking one HTT allele (Hdh+/-) and in neurons with downregulated HTT expression (Huang et al., 2011). The mechanism by which HTT enhances HIP14 activity, however, remains unclear. Interestingly, Hip14-/- mice recapitulate   87 some cardinal features of Huntington’s disease (HD), including reduced striatal volume and motor deficits (Singaraja et al., 2011).   To elucidate the functional connection between HIP14 and HD and the potential mechanisms by which altered HIP14-HTT interaction may contribute to HD pathogenesis, a large-scale yeast-two-hybrid screen for HIP14 was recently performed (Butland et al., 2014). This screen identified a number of proteins that interact with both HIP14 and HTT, providing insight into the shared function of HIP14 and HTT. While several of these shared interactors are established HIP14 substrates, some were not previously documented to be modified by palmitate, including the known HTT interactor, optineurin (optic neuropathy inducing protein; OPTN) (Hattula and Peränen, 2000; del Toro et al., 2009).   OPTN is a ubiquitous, cytosolic multi-domain protein that is highly expressed in the brain (reviewed in (Ying and Yue, 2012)) and functions both as an autophagy receptor and as an adaptor that links cargo to myosin motors. OPTN contains a C-terminal UBAN (ubiquitin binding in ABINs (A20-binding inhibitor of nuclear factor-κB proteins) and NEMO (nuclear factor-κB essential modulator)) domain that binds K63-linked and linear ubiquitin chains (Wagner et al., 2008). The UBAN domain, together with an N-terminal LC3 (microtubule-associated protein light chain 3) binding domain, links ubiquitinated substrates to autophagosomal membranes (Wild et al., 2011). OPTN also interacts with intracellular protein aggregates, including HTT. However, this interaction is independent of the UBAN domain (Korac et al., 2013).     88 The interaction of OPTN with full-length HTT is required for its role as a cargo adaptor. HTT, OPTN and Rab8 form a complex that mediates post-Golgi trafficking of proteins by interacting with the motor protein Myosin VI (del Toro et al., 2009; Sahlender et al., 2005). mHTT destabilizes this interaction and leads to aberrant trafficking (del Toro et al., 2009). The HTT-OPTN interaction was mapped to the C-terminal region of OPTN encompassing the UBAN domain (Hattula and Peränen, 2000), but the role of ubiquitin in complex assembly was not determined.  Based on the documented roles of OPTN and HTT as cargo receptors, we hypothesized that OPTN and HTT bind HIP14 to direct its trafficking to different subcellular locales. This, in turn, could lead to changes in dynamic palmitoylation by bringing HIP14 in proximity to its substrates. Here, we show that the interaction between OPTN and HIP14 is enhanced in the presence of HTT, regardless of its CAG tract length. This interaction is mediated through the N-terminus of HTT and is highly dependent on the UBAN domain of OPTN. Thus far, we have identified mutations in the HIP14 ankyrin domain that decouple HIP14-HTT and HIP14-OPTN interactions. This will provide a useful tool for future studies to assess the role of OPTN on HIP14 trafficking in neurons.   3.2 Materials and Methods 3.2.1 Mammalian cDNAs HIP14-GFP, full length HTT15Q-HA, and full length HTT128Q-HA cloned in pCINeo were generous gifts from Dr. Michael Hayden. See Table 3.1 for nucleotide sequences used for amplification and preparation of the following plasmids used in this study: pFLAG-OPTN   89 was generated by polymerase chain reaction (PCR) amplification of the OPTN ORF from MGC Fully sequenced Human OPTN cDNA, clone ID 3831267 (Openbiosystems; http://www.ncbi.nlm.nih.gov/nuccore/ BC013876) and subcloning into EcoRI and XhoI sites of FLAG-NT (gift from Dr. Stefan Taubert, the University of British Columbia). FLAG-OPTND474N was generated from pFLAG-OPTN using the QuikChange Site-directed Mutagenesis Kit (Promega). HTTExon1-15Q/128Q-HA were generated by PCR amplification of amino acids 1-90 of FLHTT15Q and FLHTT128Q, respectively, with C-terminally attached HA epitopes, and subcloning into NheI and EcoRI sites of pCINeo vector (Promega). HTT1-548-15Q/128Q-HA were generated by PCR amplification of amino acids 1-548 of FLHTT15Q and FLHTT128Q, respectively, and cloned as described for HTTexon1-15Q/128Q-HA.  For the generation of HIP14 ANKRD mutants, Geneart fragments (Life Technologies) with sizes of 862 nucleotides encompassing part of the pCINeo backbone and 5’ sequence of HIP14-GFP were synthesized (refer to Appendix A). The fragments contain respective mutations and were flanked by XhoI and AfeI sites. Fragments were PCR amplified using HIP14MUT forward and reverse oligos (Table 3.1) and were subcloned into XhoI and AfeI-double digested HIP14-GFP plasmids.         90 Table 3.1 Cloning Oligos and Methods Used to Generate the Constructs for This Study.    3.2.2 Chemicals  Poly-L-lysine was obtained from Sigma-Aldrich. Lipofectamine 2000, sodium dedocyl sulfate (SDS) solution, and Biotin-azide were purchased from Life technologies. 17-ODYA was purchased from Cayman Chemical.   3.2.3 Cell Culture Conditions HEK293 cells were maintained and propagated in high glucose Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Life Technologies), 4mM L-Glutamine and 1mM sodium pyruvate, in a humidified incubator at 37°C, 5% CO2.    91 3.2.4 cDNA Transfections HEK293 cells were transfected with cDNAs as indicated in each experiment using Lipofectamine 2000 as per manufacturer’s instructions. Cells were grown in 100mm dishes (CoStar) pre-coated with poly-L-lysine (Sigma) and transfected at 90% confluence with a total of 12µg of cDNA (4µg per plasmid) per well. Cells were processed for co-immunoprecipitations or CuAAC/click chemistry studies as described below 48 hours post-transfection.  3.2.5 Co-Immunoprecipitations 48 hours post-transfection, HEK293 cells were lifted from plates by trypsinization. Cells were collected by spinning at 200 x g for 5 minutes in DMEM. The media were aspirated and cell pellets were frozen at -80°C. For co-immunoprecipitation, cells were lysed in mild TEEN buffer (50mM Tris-HCl, 50mM NaCl, 1% Triton-X 100, 0.5mM EDTA, 0.5mM EGTA, pH7.4). 1 – 1.5mg of lysates were incubated with indicated antibodies for 16 hours at 4oC. Protein G sepharose (GE Healthcare) was added to the antigen-antibody complexes for an additional 4-6 hours. Beads were washed and eluted samples were separated on 8% SDS-PAGE gels. Proteins were transferred onto nitrocellulose membranes and analysed by immunoblotting with primary antibodies [Mouse Anti-FLAG M2, 1:1,000 (Sigma); Rabbit Anti-GFP, 1:1,000 (Invitrogen); Mouse Anti-HA, 1:1,000 (Covance)] and secondary antibodies [Alexa Fluor® 680 Goat Anti-Rabbit, 1: 10,000 and Alexa Fluor® 800 Goat Anti-Mouse, 1:10,000]. Blots were scanned using the Li-Cor Odyssey® Infrared Imaging System.    92 3.2.6 Yeast-Two-Hybrid Yeast two-hybrid plasmids for OPTN and HTT were generated by PCR amplification of plasmid DNA and cloning into the BglII site of pGAD-C2 and pGBD-C2, for prey and bait plasmids, respectively (James et al., 1996). All constructs were sequence verified. Expression plasmids were transformed into the PJ69-4A (Prey) and P69J4<α> (bait) yeast strains and transformants were selected on SD minimal medium lacking leucine (for prey strains) or uracil (for bait strains). The clones were mated on YPD plates supplemented with adenine for three days to allow mating. The resulting diploids were replica plated onto SD minimal medium lacking leucine and uracil for growth for two more days, prior to replica plating onto SD minimal medium lacking leucine, uracil, and histidine supplemented with 3-amino-1,2,4-triazole (3-AT) to select for interactions. Plasmids were rescued from yeast colonies positive for interaction for plasmid sequence verification.   3.2.7 Metabolic Labeling and CuAAC/Click Chemistry 24 hours post-transfection, HEK-293 cells were briefly washed twice in phosphate-buffered saline (PBS) then lipid-starved for 1 hour in DMEM containing 5% charcoal-filtered FBS (Life Technologies). Cells were incubated with 50µM 17-ODYA for 3 hours in this media for metabolic labeling. Cells were briefly washed twice in PBS, then harvested in triethanolamine (TEA) lysis buffer [1% TX-100, 150 mM NaCl, 50 mM TEA pH 7.4, 2x EDTA-free HaltTM Protease Inhibitor (Thermo)], and lysates incubated for 16 hours at 4oC with Protein G sepharose (GE Healthcare) bound to IP antibody [Mouse Anti-FLAG M2, 4 ug per sample (Sigma)]. Following IP, beads were washed three times in modified RIPA buffer (150mM NaCl, 1% Sodium deoxycholate (w/v), 1% TX 100, 0.1% SDS, 50mM TEA   93 pH7.4), then resuspended in 50µL RIPA buffer containing 3.5µL of freshly premixed click chemistry reaction mixture. Final concentrations in reaction were: 1mM TCEP in dH2O, 1mM CuSO4·5H2O in dH2O, 100µM TBTA in DMSO, and 100µM biotin-azide in DMSO. Beads were washed three times with RIPA buffer and resuspended in 10µL SDS buffer (150mM NaCl, 4% SDS, 50mM TEA pH7.4), 4.35µL 4x SDS-loading buffer (8% SDS, 4% Bromophenol Blue, 200mM Tris-HCl pH 6.8, 40% Glycerol), and 0.65µL 2-mercaptoethanol. Samples were heated for 5 min at 95°C, and separated on 10% tris-glycine SDS-PAGE gels. Proteins were transferred to nitrocellulose and blotted with primary antibody [Rabbit Anti-FLAG, 1:1000 (Sigma-Aldrich)] and secondary antibody [Alexa Fluor® 800 Goat Anti-Rabbit, 1:10,000 (Invitrogen) and Alexa Fluor® 680 Streptavidin, 1:5,000 (Invitrogen)] to detect biotin and total protein. Blots were scanned using the Li-Cor Odyssey® Infrared Imaging System.  3.2.8 HIP14 Ankyrin Repeat Domain Structure Analysis The HIP14 ankyrin repeat domain (ANKRD) crystal structure with atomic coordinates was retrieved from the Protein Data Bank (http://www.rcsb.org/, PDB ID code: 3EU9) (Gao et al., 2009). Candidate protein-ligand and protein-protein binding sites were identified using the Active Site Detection software, Molecular Operating Environment (MOE™) Suite (Chemical Computing Group, Quebec, Canada). The program uses a fast α-shapes algorithm to score sites for the ligand binding propensity (Soga et al., 2007).    94 3.2.9 Statistical Analyses  Statistical analyses were carried out by performing Student’s two-tailed t-tests using Prism 6  (GraphPad Software, Inc., La Jolla, CA).  A p < 0.05 was considered to indicate a statistically significant difference.   3.3 Results 3.3.1 HIP14 Interacts with, but Does not Palmitoylate, OPTN We recently conducted a large-scale HIP14 interactome study to identify novel HIP14 interactors (Butland et al., 2014). The resulting data set showed a highly significant overlap between HIP14 interactors and 370 published interactors of HTT, where nearly half of the 36 shared interactors are implicated in HD, supporting a direct link between HIP14 and HD (Butland et al., 2014). We reasoned that these HTT-HIP14 shared interactors could include novel HIP14 substrates, and/or potential regulators of HIP14. One of these binding partners, OPTN, had previously been shown to interact with HTT to modulate post-Golgi trafficking of cargo proteins for plasma membrane secretion or lysosomal delivery (del Toro et al., 2009). Thus, the observation that OPTN and HTT also interact with HIP14 suggests that the three proteins act in a common pathway.  The interaction between HIP14 and OPTN was first detected in a systematic yeast two-hybrid screen of the ORFeome (Butland et al., 2014). The HIP14/OPTN interaction was later confirmed using a modified LUMIER (luminescence-based mammalian interactome mapping) assay, where the bait protein (HIP14) tagged with the Protein A (PA)-Renilla luciferase (RL) module and the prey protein (OPTN) tagged with the Firefly luciferase (FL)   95 module were co-produced in HEK293 cells and their interaction was detected by measuring the FL activity following IgG purification of PA (Barrios-Rodiles et al., 2005; Suter et al., 2013; Butland et al., 2014). To further validate this interaction, we performed co-immunoprecipitation from HEK293 cells overexpressing both FLAG-tagged OPTN and GFP-tagged HIP14. When we pulled down HIP14 using anti-GFP antibodies, we observed co-purification of FLAG-OPTN (Figure 3.1A, n = 3). Similarly, HIP14-GFP co-purified with FLAG-OPTN in FLAG pulldowns (Figure 3.1B, n = 3), confirming a physical interaction between the two proteins.  Because HIP14 is a PAT, we examined the palmitoylation status of FLAG-OPTN by click chemistry after labeling cells with 50µM of the palmitate analogue 17-octadecynoic acid (17-ODYA) for 3 hours. We found that OPTN is not palmitoylated when overexpressed in COS cells, even in the presence of co-overexpressed HIP14 (Figure 3.1C,D, n = 3). In contrast, the predicted HIP14 substrate SPRED2 (Sprouty-related, EVH1 domain-containing protein 2), which was also identified in the screen (Butland et al., 2014), was palmitoylated, and this palmitoylation was enhanced in the presence of HIP14-GFP (Figure 3.1C,D, n = 2). Together, these results demonstrate that HIP14 interacts with OPTN, but does not palmitoylate it, suggesting the interaction could serve another physiological function.       96   Figure 3.1 OPTN Interacts with HIP14 but is not a Palmitoylated Substrate. (A,B) Reciprocal co-immunoprecipitation between HIP14-GFP and FLAG-OPTN. Co-immunoprecipitation experiments were performed on HEK293 cells expressing HIP14-GFP and/or FLAG-OPTN, where in (A) HIP14-GFP was immunoprecipitated by anti-GFP antibody (n = 3); and in (B) FLAG-OPTN was immunoprecipitated by anti-FLAG antibody, and the resulting blots were probed for GFP and FLAG (n = 3). (C) OPTN is not palmitoylated, whereas SPRED2 is a HIP14 substrate. HEK293 cells expressing FLAG-OPTN or FLAG-SPRED2 were labeled with the palmitate analogue 17-ODYA for 23 hours, then FLAG-tagged proteins were immunoprecipitated using anti-FLAG antibodies. On-bead click chemistry with biotin-azide was carried out to detect OPTN and SPRED2 expression and palmitoylation by Western blot (n = 3 for OPTN; n = 2 for SPRED2). * indicates unspecific endogenous reactivity. (D) SPRED2 palmitoylation is potentiated by HIP14. Blots from (C) were quantified. Band intensities were normalized to the –HIP14 condition for each substrate.  Error bars indicate SEM from three (for OPTN) or two (for SPRED2) independent experiments.    3.3.2 HIP14 Interaction with OPTN is Enhanced by the Presence of HTT  As both OPTN and HIP14 have been shown to physically interact with HTT (del Toro et al., 2009; Huang et al., 2011), we wondered if HTT might compete for OPTN/HIP14 binding. To determine if the level of cellular HTT influences the OPTN-HIP14 interaction, we simultaneously overexpressed HTT-HA, HIP14-GFP and FLAG-OPTN in HEK293 cells. Immunoprecipitation of HIP14-GFP recovered both FLAG-OPTN and HTT-HA. Interestingly, in the presence of HTT-HA, we saw a significant increase in the amount of   97 FLAG-OPTN that co-purified with HIP14-GFP (Figure 3.2A,B, n = 3). The enhancement of the HIP14/OPTN interaction was independent of the HTT CAG tract length, as mHTT with a 128Q polyglutamine expansion was equally effective in increasing the recovery of FLAG-OPTN in HIP14-GFP pulldowns (Figure 3.2A,B, n = 3). The enhancement of the HIP14-OPTN interaction by exogenous HTT indicates that HTT is limiting for complex formation when HIP14 and OPTN are overexpressed, suggesting these proteins form a trimeric complex.  The smallest fragment of HTT that is sufficient for the full interaction with HIP14 has been mapped to residues 1-548, a region that includes the Cys214 palmitoylation site (Yanai et al., 2006; Sanders et al., 2014). In contrast, HTT1-90 (Exon 1) contains the site of CAG repeat expansion but does not bind HIP14 (Huang et al., 2011). When co-overexpressed with HIP14-GFP and FLAG-OPTN, expression of the N-terminal 548 residues of HTT (N-HTT) enhanced FLAG-OPTN pulldown by HIP14-GFP, similar to full-length HTT (Figure 3.2C,D, n = 2), whereas exon1 did not bind to HIP14, and did not increase OPTN pulldown (Figure 3.2C,D, n = 2). Co-expression of Exon1-128Q, which carries the polyglutamine expansion, similarly failed to enhance OPTN recovery (data not shown). These results suggest that the N-terminal 548 residues of HTT, but not the first 90 residues, are sufficient to promote formation of an OPTN-HIP14 complex.      98   Figure 3.2 Expression of Exogenous HTT Enhances the Interaction between HIP14 and OPTN.  (A,B) Expression of exogenous HTT enhances the HIP14/OPTN interaction regardless of the HTT polyglutamine tract lengths. Co-immunoprecipitation experiments were performed on HEK293 cells expressing HIP14-GFP and/or FLAG-OPTN, in the presence or absence of ectopic HA-HTT-15Q or HA-HTT-128Q. HIP14-GFP was immunoprecipitated by anti-GFP antibody, and the resulting blots were probed for GFP, FLAG, and HA. The amount of OPTN that co-IP’d with HIP14 was quantified from band intensities on the FLAG blot, normalized to the “No exogenous HTT” condition (n = 3, Two-tailed Student’s t-test). (C,D) Expression of HTT1-548, but not HTT exon1 (1-90), is sufficient for enhancement of the HTT/OPTN interaction. Co-immunoprecipitation experiments were carried out as described in (A,B) except with HA-HTT-Exon1 (“Ex1”) or HA-HTT1-548. The amount of OPTN that co-IP’d with HIP14 was quantified from band intensities on the FLAG blot, normalized to the “No exogenous HTT” condition (n = 2). Error bars indicate SEM from three (B) or two (D) independent experiments.    3.3.3 OPTN Interacts with HTT via its C-Terminal Domain OPTN interacts with full-length HTT via its C-terminus, which encompasses the UBAN domain (Hattula and Peränen, 2000). While OPTN recognizes HTT exon1 aggregates via its C100kDa70kDa55kDa35kDa25kDaWB:GFPWB:FLAGIP: GFPWB:HAHTT1-548HTT-Exon1HIP14-GFPIP (gα-GFP)Lysate (20μg)+HA-HTT+ +FLAG-OPTN1-548Ex1+++++++ ++++++1-548Ex1 1-548Ex1 1-548Ex1HTT-128QHTT-15QWB:GFPWB:FLAGIP: GFPWB:HA+HIP14-GFP +HA-FLHTT+ +FLAG-OPTN +++15Q 128Q+ ++ ++++15Q 128QIP (gα-GFP)Lysate (20μg)A01.02.03.0p=0.03p=0.02+HTTNoHIP14NoHTT 15Q+HTT128Q01.02.03.0+HTTNoHIP14NoHTT Exon1+HTT1-548Amount OPTN COIP’d with HIP14(Normalized to “No HTT”)Amount OPTN COIP’d with HIP14(Normalized to “No HTT”)BD  99 coil-coiled domain independent of ubiquitin binding (Korac et al., 2013), it is not known if this is also true for OPTN binding to full-length HTT. To determine if the HTT-OPTN interaction requires recognition of ubiquitin by the UBAN domain, we generated an OPTN C-terminal fragment (C-OPTN; 411-577aa) carrying the D474N mutation, which is predicted to disrupt ubiquitin binding (Ying and Yue, 2012). By the yeast-two-hybrid assay, N-HTT interacted with full-length OPTN, C-OPTN, as well as C-OPTN D474N under low stringency (3mM 3-AT) conditions (Figure 3.3A). At higher stringency (20mM 3-AT), however, the interaction with C-OPTN D474N was lost, and that with full-length OPTN was reduced (Figure 3.3A). Based on these observations, the binding of HTT and OPTN appears to be partially dependent on the ubiquitin-binding activity of the UBAN domain.   However, the D474N mutation also abolished the interaction of OPTN with itself (Figure 3.3B). Because OPTN forms homo-oligomers via its UBAN domain in an ubiquitin-independent manner (Schwamborn et al., 2000; Ying et al., 2010; Gao et al., 2014), this suggests that the D474N mutation may impair other protein-protein interactions. In addition, C-OPTN shows reduced interaction with full-length OPTN (Figure 3.3B). Note that the reduced interactions are not due to a significant reduction in protein levels, as the expression of full-length OPTN, C-OPTN, and C-OPTN D474N appear similar in our yeast strains (Figure 3.3C).      100  Figure 3.3 An Intact OPTN C-Terminus is Important for its Interaction with HTT and HIP14.  (A) The N-terminus of HTT interacts strongly with C-terminus of OPTN. A yeast-two-hybrid assay was performed with indicated strains on SD plates lacking uracil and leucine containing 3mM or 20mM 3-amino-1,2,4-triazole (3-AT) to select for interactions. (B) OPTN self-association is abolished with the D474N mutation. Yeast-two-hybrid assay was performed as described in (A). Selection was performed with 3mM 3-AT. (C) The OPTN411-577 D474N is not severely destabilized. Western blot of lysates from yeast-two-hybrid prey strains containing each OPTN clone used in (A) and (B). (D,E) The interaction between FLAG-OPTN D474N and HIP14-GFP is significantly destabilized, even in the presence of ectopically expressed HA-HTT. Co-immunoprecipitation experiments were carried out as described in Figure 3.2(A) except with WT full-length OPTN or OPTN D474N, and HA-HTT-128Q was excluded. The amount of OPTN or OPTN D474N that co-IP’d with HIP14 was quantified from band intensities on the FLAG blot, normalized to the “WT” condition (n = 2). Error bars indicate SEM from two independent experiments.    We next tested if OPTN D474N is deficient in HIP14 binding, by co-overexpressing HIP14-GFP with full length, FLAG-tagged OPTN, or the D474N mutant form, in the presence or absence of exogenous HTT-HA. We found that the interaction between HIP14 and OPTN was almost ablated by the D474N mutation. However, in the presence of HTT-HA, this interaction was weak yet increased by two fold (Figure 3.3D,E, n = 2). Taken together, these WT01.  +HTTD474NNoHIP14BAIT BAITPREYEmptyEmptyN-HTT16QN-HTT16QEmptyOPTN-FLOPTN411-577OPTN411-577(D474N)[3-AT]: 3mM 20mMA+HIP14-GFPIP (gα-GFP)Lysate (20μg)+HA-FLHTT + +WB:GFPWB:FLAGIP: GFPWB:HAD474NWTFLAG-OPTN+ ++ +D474NWT+ ++ ++ ++ +D EAmount OPTN COIP’d with HIP14(Normalized to WT)70kDa55kDa35kDa25kDa100kDaEmptyOPTN-FLOPTN411-577OPTN411-577 D474NOPTN-FLOPTN411-577BM.Wt (kDa)BAITPREYEmptyEmptyOPTN-FLOPTN411-577OPTN411-577(D474N)OPTN-FLEmptyOPTN-FLOPTN411-577OPTN411-577(D474N)C  101 results suggest that the interaction between OPTN, HTT and HIP14 strongly depends on OPTN having an intact and functional C-terminus.  3.3.4 Mutation of the HIP14 Ankyrin Repeat Domain (ANKRD) Causes Selective Loss of the OPTN Interaction HIP14 binding to HTT is mediated by its ankyrin repeat domain (ANKRD) (Huang et al., 2011). Our large-scale HIP14 interaction screen suggested that this domain was also responsible for OPTN binding (Butland et al., 2014). Structural analyses have predicted two distinct surfaces on the HIP14 ANKRD domain with the potential to bind proteins or small molecules between ankyrin repeats 4 and 7. One of these is an aromatic cage that is predicted to bind trimethylated lysines, whereas the second site is a hydrophobic groove that lies opposite the aromatic cage (Gao et al., 2009; Gupta et al., 2011). Analysis of the ANKRD structure using a fast α-shapes algorithm (MOE™ Suite 2010) (Soga et al., 2007) revealed no additional candidate protein-ligand or protein-protein binding sites (Figure 3.4A). Subsequently, we used a structure-guided approach to mutate critical residues in these two sites. Four mutations were made in HIP14-GFP that were predicted to disrupt the aromatic cage structure: Met191Lys, Trp196His, Tyr199Ala, and Trp231His (“HIP14-GFPmut1”). The second site, described as a hydrophobic groove, was altered by making Leu211Asp and Val217Asp mutations (“HIP14-GFPmut2”). In addition, we constructed another variant of HIP14-GFP containing all six mutations (“HIP14-GFPmut3”) (Figure 3.4B).  We performed co-immunopreciptations to determine if OPTN and/or HTT bind to either site on the HIP14 ANKRD. Mutation of the aromatic cage alone did not affect binding of OPTN   102 or HTT to HIP14 (data not shown). However, mutation of the hydrophobic groove selectively destabilized binding of OPTN to HIP14 (Figure 3.4C,D, n = 1). Combining the two sets of mutations further decreased OPTN binding, but also destabilized HIP14 (Figure 3.4C,D, n = 1). Therefore, the hydrophobic groove is important for the interaction of OPTN with HIP14.      Figure 3.4 Mutations in HIP14 Ankyrin Repeats Region Selectively Destabilizes Interaction with OPTN but Maintains Interaction with HTT. (A) HIP14 ANKRD crystal structure showing the two candidate protein-protein / protein-ligand interaction sites: An aromatic cage (M191, W196, Y199, W231) and a binding pocket (L211, V217, A247). (B) A list of the three HIP14 mutants used in this study and their mutations. (C,D) The HIP14 mutants Mut2 and Mut3 show severely destabilized binding of OPTN, but maintained binding of HTT. Co-immunoprecipitation experiments were performed on HEK293 cells expressing FLAG-OPTN, HA-HTT, and HIP14-GFP wildtype or mutants (Mut2 or Mut3). HIP14-GFP was immunoprecipitated by anti-GFP antibody, and the resulting blots were probed for GFP, FLAG, and HA. The amount of OPTN or HTT that co-IP’d with HIP14 was quantified from band intensities on the FLAG and HA blots, respectively, normalized to the “WT HIP14” condition (n =1).  Mut3WB:GFPWB:FLAGIP: GFP WB:HAHIP14-GFPFLAG-OPTN + +HA-FLHTT++ + +++IP (gα-GFP)Lysate (20μg)+ ++ + + + + ++ +Mut2 Mut3 Mut2C DAmount of OPTN or HTT(normalized to “ WT HIP14”)CO-IP’d with HIP141. NO HIP14 WT HIP14 HIP14Mut2 Mut3 HIP14MUT1 – Aromatic Cage Aromatic Cage Binding Pocket - Binding to trimethylated lysines.  - Chemical & protein docking site.  HIP14 MUTANTS: MUT2 – Binding Pocket MUT3 – Combined Mutation M191K, W196H, Y199A, W231H L211N, V217N M191K, W196H, Y199A, V211N, V217N, W231H AMUT1 – Aromatic Cage  HIP14 MUTANTS: MUT2 – Binding Pocket MUT3 – Combined Mutation M191K, W196H, Y199A, W231H L211N, V217N M191K, W196H, Y199A,  V211N, V217N, W231H B  103 OPTN association with HTT was shown to mediate vesicular trafficking from the Golgi (del Toro et al., 2009). If OPTN serves as an adaptor to promote the transport of HIP14-containing vesicles, loss of the HIP14-OPTN interaction might result in HIP14 trafficking defects. Thus far, we have observed HIP14 localization at Golgi outposts in some neurons (Figure 3.5). The identification of a HIP14 mutant selectively defective in OPTN binding will be important for future studies to determine the role of OPTN in HIP14 trafficking.   Figure 3.5 Wild-Type HIP14 can be Found on Golgi Outposts in Cultured Primary Rat Neurons. HIP14-GFP co-localizes with the Golgi marker GM-130 staining for Golgi outposts (white arrows). Immunofluorescence of primary rat neurons (13 days in vitro) transfected with HIP14-GFP. Fourty-eight hours post transfection, the neurons were fixed and immunocytochemistry was carried out with anti-GM-130 antibody to mark Golgi structures and anti-GFP antibody to stain HIP14-GFP. Scale bar, 10µm.     3.4 Discussion We previously identified an interaction between HIP14 and OPTN (Butland et al., 2014). Here, we show this HIP14-OPTN interaction is strengthened by the presence of HTT, suggesting that HTT and OPTN bind HIP14 as a complex. Because HTT and OPTN are both cargo adaptors, the binding of HTT/OPTN to HIP14 might direct the transport of HIP14-containing Golgi-derived vesicles to other cellular compartments. HIP14-GFP MERGEGM-130  104  Only a small fraction of HTT/OPTN/HIP14 appeared to form a trimeric complex, suggesting the assembly of these proteins is transient, or relies on additional cellular factors. Pairwise interactions between HTT/OPTN, HTT/HIP14 and HIP14/OPTN have been observed in the yeast-two-hybrid system (Faber et al., 1998; Singaraja et al., 2002; Butland et al., 2014), suggesting each of these interactions is direct. In further characterizing these interactions, we found domains on each protein component that are important for the formation of this complex.  Past mapping efforts suggested the interaction between OPTN and HTT occurs via the C-terminal domain of OPTN and N-terminus of HTT (Hattula and Peränen, 2000), consistent with the results presented here. The UBAN and zinc finger domains within the OPTN C-terminus are reported to form a NEMO-like domain that binds specifically to K63-linked and linear ubiquitin chains (Zhu et al., 2007; Laplantine et al., 2009). We found the interaction between OPTN and HTT was weakened, though not abolished, by mutation of D474 within the UBAN site, which is predicted to disrupt ubiquitin binding (Zhu et al., 2007). HTT is ubiquitinated at the N-terminal lysine residues K6, K9, and K15 (Steffan et al., 2004), and mHTT was shown to preferentially contain K63-linked ubiquitin chains (Bhat et al., 2014). Thus it is possible that OPTN binds the ubiquitinated N-terminus of HTT. However, these ubiquitinated residues are also present in the HTT exon1 fragment, which does not enhance interaction or bind HIP14 (Huang and El-Husseini, 2005; Korac et al., 2013). Moreover, we found the D474N abolished the interaction with OPTN with itself, even though oligomerization of OPTN via the UBAN domain does not require ubiquitination (Gao et al.,   105 2014). Thus the D474N mutation may indirectly alter the interaction of OPTN with HIP14 and/or HTT. For example, the D474N mutation causes OPTN to relocalize from the Golgi to the cytoplasm (Mankouri et al., 2010). Because HIP14 is a transmembrane enzyme predominantly observed in the Golgi (Huang et al., 2004) and HTT is membrane-associated via palmitoylation (Young et al., 2012), the observed loss of HIP14-OPTN association in cultured cells may be in part due to mislocalization of OPTN.  HIP14 is the most conserved of 23 mammalian PATs, and one of the only two PATs that contain an ANKRD, the other being HIP14L (Young et al., 2012). We mapped the OPTN binding site to a hydrophobic groove in the HIP14 ANKRD. We found that point mutations within this groove disrupted the interaction with OPTN, but did not affect the binding of HTT. However, others have shown that the N-terminal fragment of HTT also interacts with the HIP14 ANKRD (Huang et al., 2011). Thus, if OPTN and HTT co-assemble with HIP14 they are likely to bind to adjacent, non-overlapping regions of the HIP14 ANKRD. Ankyrin repeats assemble into a highly structured platform that supports multiple protein-protein interactions (Mosavi et al., 2004). Aside from HTT and OPTN, both HIP14 and HIP14L have been reported to activate the JNK (c-Jun N-terminal kinase) pathway (Harada et al., 2003). In particular, HIP14 binds JNK3 via its ANKRD, resulting in JNK pathway activation independent of HIP14 PAT activity (Yang and Cynader, 2011). Interestingly, JNK3 phosphorylation of the microtubule motor kinesin-1 has been shown to reduce kinesin-1 binding to microtubules (Morfini et al., 2009). An in vitro peptide scan mapped JNK3 binding to residues that lie between HIP14 ankyrin repeats 3 and 5 (Yang and Cynader, 2011), adjacent to the OPTN binding site reported in this study. Whether this close proximity   106 could allow OPTN to contribute to the regulation of JNK3 as a mechanism to regulate trafficking remains to be examined.  The HIP14 ANKRD was recently shown to be necessary for the interaction and palmitoylation of the presynaptic substrates SNAP25 and CSP (Lemonidis et al., 2014). Thus, it is likely that the ANKRD domain engages in a variety of interactions. Importantly, replacing the HIP14 ANKRD with that of HIP14L prevents acylation of both SNAP25 and CSP despite strong binding, suggesting that these ANKRDs are not interchangeable for these substrates (Lemonidis et al., 2014). On the other hand, HIP14L and HIP14 bind and palmitoylate HTT comparably (Huang et al., 2011; Sutton et al., 2013). It will be interesting to determine if the aromatic cage, hydrophobic groove, or other sites in the HIP14 ANKRD are responsible for binding substrates such as SNAP25 and CSP. The relationship between substrate binding and HTT/OPTN binding will also require more detailed characterization.  The HIP14L-HTT interaction prompts the question whether HIP14L may also form a trimeric complex with HTT and OPTN. Hip14l-/- mice also develop adult-onset progressive pathology and some motor deficits resembling Hip14-/- mice (Sutton et al., 2013). While Hip14l-/- mice do not exhibit reduced PSD-95 palmitoylation, they do show reduced SNAP25 palmitoylation (Sutton et al., 2013). There is currently no evidence thus far that HTT regulates HIP14L activity. It will be interesting to examine whether this mechanism is conserved to modulate dynamic palmitoylation of other HIP14L substrates.     107 What is the role of the OPTN-HTT-HIP14 interaction? HIP14 is found at the Golgi in non-neuronal cells but at the synaptic membrane in neurons, suggesting its trafficking is regulated in a cell type-specific manner (Ohyama et al., 2007; Stowers and Isacoff, 2007). One hypothesis is that OPTN and HTT coordinately regulate the trafficking of HIP14-containing vesicles to the synapse. HTT is an established cargo adaptor that scaffolds both minus-end-directed and plus-end-directed motors on microtubules, by binding directly to the intermediate chain of dynein (DIC) (Caviston et al., 2007), and indirectly to kinesin through huntingtin-associated protein 1 (HAP1) (McGuire et al., 2006). These interactions allow HTT to facilitate bidirectional organelle and vesicle transport. Interestingly, HTT also binds the early endosome marker Rab5 through huntingtin-associated protein 40 (HAP40) to mediate the actin-based transport of early endosomes (Pal et al., 2006). Thus, HTT has been proposed to act as a scaffolding protein that regulates the transition between microtubule and actin motors by interacting with either a HAP40/optineurin/myosin complex and or a HAP1/kinesin/dynein complex (Fu and Holzbaur, 2014).   An alternate hypothesis is that OPTN and HTT regulate trafficking of HIP14-containg vesicles to autophagosomes. The role of OPTN as an autophagy receptor is well established (Wild et al., 2011), and OPTN has been shown to mediate the autophagic clearance of mHTTexon1-derived aggregates (Tumbarello et al., 2012; Korac et al., 2013). However, we found that mHTTexon1 was not sufficient to enhance the HIP14-OPTN interaction, suggesting that aggregate clearance does not explain the formation of the HTT-HIP14-OPTN complex observed here.     108 If HTT regulates HIP14 localization to enhance its activity at the synapse, this could explain why HTT mutation reduces the palmitoyation of some HIP14 synaptic substrates (Singaraja et al., 2011). HTT regulation of HIP14 activity has also been described in vitro using purified proteins (Huang et al., 2011). It is possible HTT regulates both HIP14 localization and activity. Our study strongly suggests a common functional link between OPTN, HIP14 and HTT. However, the role of this interaction in regulating HIP14 trafficking or function remains to be elucidated.      109 Chapter 4: Discussion & Conclusions   4.1 Summary of Major Findings The overall objective of this thesis was to identify additional enzymes or cofactors that play a role in mediating the dynamic palmitoylation cycles of different substrates. It was not previously known if the two established depalmitoylases, APT1 and APT2, are the only palmitoylthioesterases present in the cytosol. Meanwhile, there was limited evidence to suggest that the activity of the 23 highly conserved DHHC-CRD containing PATs is regulated. The overall hypothesis of this thesis is that additional APTs play a role in the dynamic depalmitoylation of proteins, and there are also additional modulators of PAT activity. For the first time, we show that cytosolic mSHs other than APT1 and APT2 contribute to the depalmitoylation of PSD-95. Meanwhile, we identified and validated the cargo adapter protein, OPTN, as a novel interactor of the PSD-95 PAT, HIP14. Our results suggest that OPTN and HTT form a trimeric complex with HIP14 and may regulate its localization to modulate its access to substrates. These findings add a new layer of complexity to current models of cellular palmitoylation/depalmitoylation, where ubiquitous depalmitoylation by APT1/2 and Golgi-centred repalmitoylation by PATs are thought to work in sync to achieve a constant flux of proteins between the Golgi and the plasma membrane.    110 4.2 Functional Redundancy in APTs PSD-95 has a considerable basal palmitate turnover rate and shows synaptic activity-dependent palmitoylation changes in neurons (El-Husseini et al., 2002); however, the identity of the enzyme that mediates its depalmitoylation was unexplored. Here, we showed that PSD-95 undergoes constitutive palmitate turnover in heterologous COS-7 cells. We also observed that PSD-95 depalmitoylation was blocked by PalmB treatment, but not by simultaneous downregulation of APT1/APT2 activity. While we have confirmed endogenous APT1 and APT2 expression in COS-7 cells (data not shown), we have yet to examine the expression of other serine hydrolases that represent candidate depalmitoylases (BAT5, FAM108A1, FAM108B1, FAM108C1), by either Western blotting or qPCR.   Thus far, three independent studies, including this one, reported overlapping yet distinct PalmB targets (Rusch et al., 2011; Savinainen et al., 2014). Factors that could influence the detection of a target include whether it is expressed in the cell type used, its expression level, and technical variation. The fact that Savinainen et al. did not identify FAM108A1/B1/C1 as PalmB targets by activity profiling of the mouse brain proteome could be due to their low endogenous activity. However, they identified MAGL as a PalmB target (Savinainen et al., 2014). MAGL is not expressed in the T cell hybridomas used for the HDFP SILAC-ABPP study (Martin et al., 2011) and hence was not included in our analysis. To comprehensively define all PalmB mSH targets, cABPP assays could be carried out in a variety of cell types. Alternatively, selective screening of an established mSH construct library (Bachovchin et al., 2010) could be used to circumvent the issue of low expression levels and/or tissue-specific   111 expression. Taken together, these findings highlight potential limitations of using PalmB to study APT1/APT2 depalmitoylation efficacy.  The PalmB targets that represent candidate PSD-95 depalmitoylases (BAT5, FAM108A1, FAM108B1, FAM108C1) are previously unannotated mSHs. A FAM108B1 homologue, AHO-3, was recently characterized in Caenorhabditis elegans (Nishio et al., 2012). AHO-3, which is highly conserved in animal species, shows 61%–80% amino acid sequence similarity with FAM108 proteins. AHO-3-related proteins in animal species have highly conserved N-terminal clusters of cysteine residues, which are palmitoylated in worm and mammalian cells, and are required for proper localization (Martin and Cravatt, 2009; Nishio et al., 2012). In worms, palmitoylation of AHO-3 mediates its localization to sensory endings and is required for thermotactic plasticity via ODR-3, a G-coupled receptor alpha subunit that is predicted to be palmitoylated. Expression of FAM108B1 in a worm aho-3 mutant can rescue this function (Nishio et al., 2012); therefore, it will be interesting to assess whether AHO-3 has depalmitoylase activity in worms.  Other studies have addressed the role of protein depalmitoylation in lower organisms. In Saccharomyces cerevisiae, no significant phenotypic alterations were observed when the APT1 homologue YLR118c was deleted. Because deletion of YLR118c completely abolished depalmitoylation of G(alpha), but not H-Ras, additional thioesterases are likely to exist in yeast (Duncan and Gilman, 2002). Interestingly, the Toxoplasma TgASH2-4 gene family, proposed to have thioesterase activity by Kemp et al. (Kemp et al., 2013), is highly related to the uncharacterized Saccharomyces cerevisiae protein YNL320w (unpublished   112 observations). Our analyses indicate that the most closely related mammalian homologues of YNL320w include BAT5, FAM108s, ABHD12, and ABHD13 (unpublished observations). A dendrogram of mammalian mSHs (see Figure 1.5) indicates these last three belong to the same clade, which is distant from that containing APT1 and APT2 (Bachovchin et al., 2010). These genes may represent a novel subfamily of mSHs that emerged by duplication and divergence of an ancestral YNL320w-related protein. Because the biological role of YNL320w remains untested, a closer examination of the localization and function YNL320w in yeast, and its homologue in Toxoplasma, is warranted.   A sequence comparison of the human and mouse mSH proteome suggests that FAM108B1 and FAM108C1, and, to some extent, FAM108A1 (which is less conserved between mouse and humans), are highly conserved and thus may have redundant functions (Simon and Cravatt, 2010). Hence, a triple knockdown of the FAM108 family may be necessary to determine if they contribute to PSD-95 depalmitoylation. Supporting this hypothesis, my preliminary data showed that individual knockdown of each FAM108 protein, or BAT5, in COS-7 cells caused no change in PSD-95 palmitate turnover rate (data not shown). Functional redundancy could make it challenging to determine the role of FAM108 and BAT5 proteins in higher organisms.  Even if FAM108 proteins are highly redundant, they could mediate the depalmitoylation of distinct cellular substrates. Redundant APT proteins may be important for several reasons. First, APTs may be expressed in a cell-type dependent manner. This has been previously observed in CHO cells, which lack APT1 expression (Tomatis et al., 2010). Second, APTs   113 may have different affinities for distinct substrates. Third, some APTs may be differentially regulated. Changes in the local protein synthesis of APT1 (Banerjee et al., 2009) were suggested to in part explain the rapid stimulus-dependent decrease in the palmitoylation levels of some neuronal substrates in mouse brains (Kang et al., 2008b). Fourth, differential subcellular distribution of the enzymes may serve to mediate substrate depalmitoylation at distinct locations. Interestingly, in heterologous cells, APT1/2 are cytosolic and Golgi-localized (Vartak et al., 2014), while FAM108 proteins are localized predominantly to the plasma membrane (Martin and Cravatt, 2009), suggesting that if FAM108 proteins do have depalmitoylation activity, they may display different substrate specificities. As another approach to determine if FAM108 proteins have depalmitoylase activity towards PSD-95, we examined PSD-95 palmitate turnover under overexpression of each candidate mSH. Our preliminary data suggest that APT1 overexpression moderately inhibited PSD-95 depalmitoylation as observed by pulse-chase/click chemistry, an effect that was reproduced by overexpression of each of the FAM108 proteins (data not shown). These findings suggest that APT1 may also have PSD-95 depalmitoylating activity that is compensated for by the FAM108s (and/or other unidentified PSD-95 thioesterases) in knockdown studies. This is currently under investigation.  One major caveat of this study is that PSD-95 is expressed only in the nervous system. Hence, it would be highly relevant to examine PSD-95 palmitate turnover in neuronal cell types. Additional tools and techniques can be used to define the mechanism of PSD-95 depalmitoylation in neurons in both constitutive and neuronal activity-dependent contexts. Because selective inhibitors to FAM108 proteins and BAT5 are not available, achieving   114 complete APT knockdown in neurons will require multiple siRNAs targeting these proteins in the presence of APT1 and APT2 small molecule inhibitors. In neurons, PSD-95 palmitoylation results in changes in its localization and conformation, the latter of which can now be studied using the recently developed conformation-specific intrabody, PF-11 (Fukata et al., 2013). In neurons, an increased PSD-95-GFP co-localization with post-synaptic density markers upon PalmB treatment has been observed (Okunola Jayfious and William Green, Pers Comm.). Therefore, in future studies PF-11 could be used in live neurons depleted of multiple APTs in different combinations to determine the amount of PSD-95 clustering in the synapse. More importantly, as synaptic glutamate receptor activation has been demonstrated to enhance PSD-95 depalmitoylation, knockdown of FAM108s/BAT5 and/or APT1/2 could be used in combination with potassium chloride treatment to assess the contribution of these enzymes to activity-dependent depalmitoylation, using PSD-95 clustering as a readout (El-Husseini et al., 2002). Ultimately, once the depalmitoylase(s) responsible for PSD-95 is identified, future work will focus on finding additional substrates.  It will also be important to determine the in vivo roles of these enzymes by developing animal models where physiological and electrophysiological studies can be used to study neurological phenotypic alterations.  4.3 The Role of OPTN in Regulating HIP14 Trafficking and Function Our results suggest that dynamic palmitate cycling of PSD-95 is in part due to the depalmitoylating activity of redundant APTs. However, it is not known if APT activity or localization is modulated in response to synaptic activity. Instead, activity-dependent changes in PSD-95 palmitoylation in dendritic spines have been attributed to local trafficking of   115 DHHC2 (Noritake et al., 2009). PSD-95 is also subject to palmitoylation by a handful of other PATs including DHHC3, DHHC7, DHHC8, DHHC15, and HIP14 (DHHC17) (Fukata et al., 2004; Mukai et al., 2008; Singaraja et al., 2011). Interestingly, even though DHHC2 and DHHC3 have been reported as the major PSD-95 PATs in the hippocampus (Noritake et al., 2009), PSD-95 palmitoylation is significantly reduced in Hip14-/- mice in vivo, suggesting that HIP14 also plays a substantial role in PSD-95 palmitoylation (Singaraja et al., 2011). In addition, hippocampal long-term potentiation (LTP), which involves rapid activity-dependent changes in the localization of several synaptic proteins, including PSD-95, is impaired in Hip14-/- animals (Milnerwood et al., 2013). These findings raise the possibility that HIP14 may palmitoylate PSD-95 in an activity-dependent fashion.  How could HIP14 activity be regulated? Previously, Huang et al. showed that HTT modulates HIP14 PAT activity both in vitro and in vivo, and this is highly dependent upon the interaction between the two proteins (Huang et al., 2011). Several models could explain this observation. First, HTT could function as an allosteric cofactor that directly stimulates HIP14 catalytic activity by causing a conformational change that brings substrates closer to the DHHC active site (Huang et al., 2011). Second, HTT, which is largely made up of HEAT (Huntingtin, Elongation factor 3, protein phosphatase 2A, TOR1) repeats, may act as a scaffolding protein to link HIP14 to its substrates (Takano and Gusella, 2002; Li et al., 2006; Palidwor et al., 2009). Third, since HTT interacts with the molecular motor proteins myosin, dynein, and kinesin directly or indirectly to support cargo trafficking in and out of synapses (Fu and Holzbaur, 2014), HTT may promote the transport of HIP14-containing vesicles to various subcellular locations. While the observation that HTT enhances the palmitoylation of   116 purified SNAP25 by HIP14 in a cell-free system supports the first model (Huang et al., 2011), these models are not mutually exclusive as other HTT-HIP14 interactors may serve to further modulate HIP14 enzymatic function in vivo.  In this dissertation, we showed that HIP14 physically interacts with OPTN, a cargo adaptor that associates with HTT and directs post-Golgi trafficking in complex with Rab8 (del Toro et al., 2009). The HTT-OPTN-Rab8 complex has been implicated in clathrin-dependent trafficking to both PM and lysosomes in striatal-derived cells, most likely through binding to myosin VI, a motor protein that is vital for exocytosis (del Toro et al., 2009; Sahlender et al., 2005; Bond et al., 2011) Hence, we propose that in the striatum, HIP14 is a cargo for this trafficking complex, and that the HIP14, together with HTT and OPTN, form mobile trimeric complexes that travel on myosin motors to different subcellular locations as a way to mediate dynamic palmitoylation. One limitation of the current study is that the trimeric complex was observed in heterologous cells when all three components were overexpressed. Nevertheless, in COS-7 cells, HIP14 has been observed in cytoplasmic vesicles, although it is predominantly localized to the Golgi (Huang et al., 2004; Hines et al., 2010).  In addition, rapid transport of HIP14 in SNAP25-containing vesicles has been observed in COS-7 cells (Huang et al., 2004), suggesting this mechanism may occur even in non-neuronal cells.  Transport of the OPTN-HTT-HIP14 complex may be important for activity-dependent palmitoylation of PSD-95 and other synaptic proteins in a neuronal context. Consistent with this hypothesis, HIP14 is highly enriched in pre- and post-synaptic termini in Drosophila melanogaster neurons where it mediates the palmitoylation of presynaptic CSP and SNAP25   117 (Stowers and Isacoff, 2007; Ohyama et al., 2007). Also, transient relocalization of HIP14 to the presynaptic membrane upon synaptic vesicle depletion in stimulated shibirets1 (mammalian homologue of dynamin; van der Bliek and Meyerowitz, 1991) mutants at the restrictive temperature has been observed (Ohyama et al., 2007). Whether this is also true in mammalian systems remains to be examined. Previous analyses revealed HIP14 immunogold labeling in diverse vesicular structures present in the soma, axon, and dendrites of mouse neurons (Huang et al., 2004). Furthermore, partial co-localization of HIP14 with clathrin, the AP-1 component γ-adaptin, the synaptic vesicle marker SV2, and the late endosome markers Rab7 and, notably, Rab8, has been observed in neuronal NT2 and rat hippocampal cells (Singaraja et al., 2002; Huang et al., 2004; Stowers and Isacoff, 2007). Further work will be needed to determine whether HIP14 copurifies with the HTT-OPTN-Rab8 complexes in neuronal cells.  Multiple complementary approaches will be valuable to determine whether HIP14, like DHHC2, shows increased dendritic spine localization and plasma membrane insertion upon activity changes in neuronal cultures (Noritake et al., 2009; Fukata et al., 2013). Activity-dependent synaptic accumulation of HIP14 can be examined using total internal reflection fluorescence microscopy (TIRFM) after ionotropic glutamate receptor activity blockade by kynurenic acid (Noritake et al., 2009). The requirement for OPTN or HTT in HIP14 trafficking in neurons could be tested in parallel by siRNA-mediated knockdown. It will also be important to determine if HIP14 regulates local substrate palmitoylation at synapses. This is technically challenging, but with the recently developed PSD-95 palmitoylation-dependent intrabody, this can be determined at least for PSD-95, a known HIP14 substrate (Singaraja et   118 al., 2011; Fukata et al., 2013). In addition, as OPTN has been shown to form oligomers (Gao et al., 2014) and HIP14 is known to self-interact (Butland et al., 2014), it will be important to determine the stoichiometry of OPTN, HTT, and HIP14 within a complex. This can be determined by blue-native (BN)-PAGE or sedimentation velocity. Lastly, because OPTN is differentially expressed in distinct subpopulations of neurons (Okita et al., 2012), it would be interesting to determine if OPTN expression levels are correlated with HIP14 localization in various cell types. Given that HIP14 palmitoylates numerous substrates, it will be vital to dissect the mechanism by which OPTN regulates HIP14 activity and localization to achieve dynamic palmitoylation.    In HD, mHTT has been reported to exert its pathogenic effect by disrupting multiple cellular processes including transcription, protein degradation, synaptic signaling, mitochondrial metabolism, and protein/organelle trafficking (Labbadia and Morimoto, 2013). In particular, full-length mHTT-111Q was shown to displace OPTN from the Golgi apparatus and interfere with post-Golgi trafficking in striatal cell lines (del Toro et al., 2009). We did not observe a reduction in HIP14-HTT-OPTN complex formation in the presence of full-length mHTT-128Q in HEK293 cells. Cell type differences may explain the discrepant HTT/mHTT-OPTN interactions observed, especially as subsets of neuronal cells are uniquely vulnerable to mHTT (Her and Goldstein, 2008). Therefore, this will also have to be addressed in the future by examining the effect of mHTT in striatal cells on HIP14-HTT-OPTN complex formation.    119 4.4 Conclusions Dynamic palmitoylation depends on the activity and localization of both PATs and APTs. In this dissertation, we defined a novel OPTN/HIP14 interaction and speculate that trafficking of a HTT/OPTN/HIP14 trimeric complex may contribute to dynamic substrate palmitoylation at the synapse. In light of our findings that APTs other than APT1 and APT2 mediate PSD-95 depalmitoylation, further studies will be required to decipher the mechanisms that underlie the activity-dependent changes in substrate palmitoylation that accompany synaptic plasticity.  One of the best-studied substrates whose dysregulated palmitoylation leads to pathological phenotypes is mHTT. HIP14 and HIP14L are the two major PATs for HTT (Huang et al., 2011); however, they show reduced interaction with mHTT which subsequently leads to decreased mHTT palmitoylation, increased mHTT misfolding, and formation of cellular inclusions (Singaraja et al., 2002; Yanai et al., 2006). With this defined PAT-substrate specificity, a therapeutic approach would be to upregulate HIP14 or HIP14L enzymatic activity to promote mHTT palmitoylation. However, enhancing PAT activity could prove challenging; alternatively, blocking depalmitoylation enzyme activity using specific inhibitors could provide a more feasible approach for enhancing protein palmitoylation. In this dissertation, we showed that PalmB is able to robustly inhibit HTT depalmitoylation through inhibition of APT1 and APT2 activity. 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