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Palmitoyl-proteins : regulators of neuronal functions and potential targets for neuroprotection Yang, Guang 2011

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PALMITOYL-PROTEINS: REGULATORS OF NEURONAL FUNCTIONS AND POTENTIAL TARGETS FOR NEUROPROTECTION  by GUANG YANG B.Sc., Wuhan University, 2005  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY  in  The Faculty of Graduate Studies (Neuroscience)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  October 2011 © Guang Yang, 2011  Abstract Protein palmitoylation is an important post-translational lipid modification. While hundreds of palmitoyl proteins have been identified in neurons, little is known about how palmitoylation regulates these neuronal proteins and how it contributes to neuronal development and function. A special group of palmitoyl proteins, 23 mammalian zinc-finger DHHC-type containing (zD) proteins are potent palmitoyl acyltransferases (PATs) that catalyze protein palmitoylation. However, the physiopathological roles of these PATs in brain function are largely elusive.  AMPA receptor (AMPAR) subunit GluR1 and GluR2 are palmitoyl proteins. In this thesis, I have found that GluR1 and GluR2 show different palmitoylation properties in neurons. Palmitoylation regulate AMPAR stability in a subunit-selective manner in response to synaptic stimulations. In addition, c-jun N-terminal kinase 3 (JNK3), but not other JNK isoforms, has been identified in this thesis as a novel palmitoyl protein. Without palmitoylation, JNK3 is associated more strongly with the cytoskeleton and promotes axonal branching. This suggests a potential role of palmitoylation in modulating axonal development via isoform-specific regulation of JNK3.  I have further revealed that zD17 mediates neuronal responses in acute ischemic brain injury via a mechanism independent of its PAT activity. ZD17 directly interacts with JNK to form a signaling module for JNK activation. Pathological stressors induce the zD17-JNK interaction which promotes downstream neuronal cell death signals. I have developed novel peptides targeting the JNK-interacting motif on zD17 to selectively block the enhancement of the zD17-JNK interaction and the activation of JNK isoforms 2 and 3. Application of these peptides successfully blocks JNK activation and neuronal cell death pathways, protects cultured neurons from excitotoxicity, and dramatically reduces brain damage and behavioural deficits in a rat model of focal ischemic stroke. These findings indicate PAT zD17 as a key player in ischemic stroke, and suggest the potential therapeutic value of targeting palmitoyl proteins for neuroprotection.  ii  Preface For Chapter 2, a version of this chapter has been published (Guang Yang, Wei Xiong, Luba Kojic, Max S. Cynader. 2009. Subunit-selective palmitoylation regulates the intracellular trafficking of AMPA receptor. European Journal of Neuroscience, 30 (1): 35-46). I performed all biochemical experiments including the metabolic 3H-palmitate labelling, the ABE assay, and the deglycosylation assay etc. Dr. Wei Xiong and Dr. Luba Kojic were involved in establishment of chemical LTD protocol. AMPAR subunit constructs were provided by Dr. Huganir’s lab. I analyzed the data from all experiments, generated figures, and wrote the manuscript.  For Chapter 3, a version of this chapter has been published (Guang Yang, Yuan Liu, Kaiyun Yang, Rui Liu, Shanshan Zhu, Ainsley Coquinco, Wendy Wen, Luba Kojic, William Jia and Max S Cynader. Isoform-specific palmitoylation of JNK regulates axonal development. Cell Death and Differentiation. doi: 10.1038/cdd.2011.124). I generated all JNK3 palmitoylation mutant constructs. GFP-JNK3 WT was provided by Dr. Vsevolod V. Gurevich. I performed JNK palmitoylation analysis, cytoskeleton fractionation analysis, and axonal development experiments etc. Liu Yuan and I performed lipid rafts isolation. I analyzed the data from all experiments and generated figures.  For Chapter 4, a version of this chapter has been published online (Guang Yang and Max S. Cynader. 2011. Palmitoyl acyltransferase zD17 mediates neuronal responses in acute ischemic brain injury by regulating JNK activation in a signaling module. Journal of Neuroscience, 31 (33): 11980-91). I generated all zD17 deletion constructs, designed the peptides used in the study and performed all experiments including in vitro biochemical experiments and in vivo animal experiments. Dr. Wendy Wen provided assistance in animal experiments. I analyzed the data and generated figures.  iii  All animal experiments were approved by the Animal Care Committee of the University of British Columbia (Protocol A10-0118; Certificates RBH-634-09, RA-367-09, RSHX-280-09, 1097).  iv  Table of Contents Abstract .......................................................................................................................................... ii Preface ........................................................................................................................................... iii Table of Contents .......................................................................................................................... v List of Tables................................................................................................................................. ix List of Figures ................................................................................................................................ x List of Abbreviations................................................................................................................... xii Acknowledgments ...................................................................................................................... xiv Chapter 1. Introduction .............................................................................................................. 1 1.1 Protein Palmitoylation and Palmitoyl Acyltransferases .................................................... 2 1.1.1 Biochemical Basics of Protein Palmitoylation ................................................................. 2 1.1.2 Molecular Functions of Protein Palmitoylation ................................................................ 8 1.1.3 Strategies for Studying Protein Palmitoylation .............................................................. 12 1.1.3.1 Prediction of protein palmitoylation ........................................................................ 12 1.1.3.2 Metabolic labelling and acyl-biotin exchange ......................................................... 13 1.1.3.3 Large-scale identification of the palmitoyl-proteome ............................................. 16 1.1.3.4 Analysis of palmitoylation in the secretion pathway............................................... 17 1.1.3.5 Functional assessment of protein palmitoylation .................................................... 19 1.1.4 Palmitoyl Acyltransferases / PATs .................................................................................. 22 1.1.4.1 Substrate selectivity of PATs.................................................................................... 22 1.1.4.2 Multiple functions of PATs ...................................................................................... 23 1.1.5 Protein Thioesterases ...................................................................................................... 24 1.1.5.1 Turnover of palmitate .............................................................................................. 24 1.1.5.2 APT and PPT ........................................................................................................... 25 1.2 Physiological Roles of Palmitoylation in the Brain .......................................................... 26 1.2.1 Palmitoyl Proteins in the Brain ....................................................................................... 27 1.2.2 PATs in the Brain ............................................................................................................ 30  v  1.2.3 Regulation of Functional Synapses and Signal Transmission ........................................ 31 1.2.3.1 Synaptic plasticity and AMPAR .............................................................................. 31 1.2.3.2 Differential roles of subunits in AMPAR trafficking............................................... 32 1.2.3.3 Effects of protein palmitoylation on synaptic transmission .................................... 34 1.2.4 Regulation of Neuronal Development ............................................................................ 35 1.2.4.1 General background of neurite development ........................................................... 35 1.2.4.2 The role of the JNK pathway in regulating neurite development............................ 37 1.2.4.3 Palmitoyl proteins in regulation of neuronal development ..................................... 40 1.2.4.4 The c-jun N-terminal kinase family members as potential palmitoyl proteins ....... 41 1.2.5 Regulation of Protein Palmitoylation in Neurons ........................................................... 43 1.3 Protein Palmitoylation and PATs in Human Diseases ..................................................... 44 1.3.1 Protein Palmitoylation in Pathological Processes........................................................... 44 1.3.2 The Pathological Roles of PATs in Human Diseases ...................................................... 45 1.4 Stroke and Neuroprotection Therapy ............................................................................... 46 1.4.1 General Background of Stroke ....................................................................................... 46 1.4.2 Neuroprotection Therapy ................................................................................................ 48 1.4.2.1 Mechanisms underlying stroke-induced neuronal cell death .................................. 48 1.4.2.2 Role of JNK in neuronal cell death ......................................................................... 51 1.4.2.3 Therapeutic targets for neuroprotection therapy...................................................... 53 1.4.2.4 The challenge of targeting JNK and potential solutions .......................................... 54 1.5 Thesis Hypotheses and Objectives ..................................................................................... 55 1.5.1 AMAPR Palmitoylation may Have Subunit-Selectivity ................................................ 55 1.5.2 Specific JNK Isoforms may be Regulated by Palmitoylation ........................................ 56 1.5.3 How zD17 Is Involved in Regulating JNK Activation ................................................... 57 Chapter 2. Palmitoylation Contributes to Subunit-Selective Regulation of AMPA Receptors in Synaptic Plasticity ........................................................................................ 59 2.1 Introduction ......................................................................................................................... 60 2.2 Results .................................................................................................................................. 61 2.2.1 GluR1 Palmitoylation Requires Anterograde Transport ................................................. 61  vi  2.2.2 GluR2s Are Palmitoylated in the ER as Immature Receptors ........................................ 64 2.2.3 Blocking Palmitoylation Results in Reduction of GluR2 ............................................... 69 2.2.4 Palmitoylation at TMD2 of GluR2 Regulates GluR2 Stability ...................................... 71 2.2.5 Blocking Palmitoylation Leads to Lysosomal Degradation of GluR2 ........................... 72 2.2.6 Palmitoylation of AMPARs Is Regulated by Neuronal Activity .................................... 74 2.2.7 Acute Effects of NMDA and AMPA on AMPAR Palmitoylation .................................. 77 2.3 Discussion............................................................................................................................. 80 2.3.1 Different Palmitoylation Mechanisms for AMPAR Subunits......................................... 80 2.3.2 Role of GluR2 Palmitoylation in AMPAR Trafficking .................................................. 84 2.3.3 Implications for Synaptic Function ................................................................................ 86 Chapter 3. Isoform-Specific Palmitoylation of c-Jun N-Terminal Kinase (JNK) Regulates Axonal Development ........................................................................................................... 90 3.1 Introduction ......................................................................................................................... 91 3.2 Results .................................................................................................................................. 94 3.2.1 Palmitoylation Occurs Primarily on the JNK3 Isoform ................................................. 94 3.2.2 Axonal Branching Is Regulated by JNK3 Palmitoylation .............................................. 96 3.2.3 Palmitoylation Regulates JNK3 Trafficking to Cytoskeleton ...................................... 100 3.2.4 Wnt7a-Induced Axonal Branching Is Modulated by JNK3 Palmitoylation ................. 103 3.3 Discussion........................................................................................................................... 106 3.3.1 An Isoform-Specific Regulatory Mechanism for JNK ................................................. 106 3.3.2 Palmitoylation as a Trafficking Signal for JNK3 ......................................................... 107 3.3.3 A Novel Physiological Role of JNK3 in Axonal Branching ........................................ 109 Chapter 4. Interaction of zD17 with JNK Mediates Brain Injury in Ischemic Stroke, and Represents a Novel Therapeutic Target for Neuroprotection ........................................ 111 4.1 Introduction ........................................................................................................................ 112 4.2 Results ................................................................................................................................. 114 4.2.1 JNK Activation Is Regulated by the zD17-JNK Signaling Module ..............................114 4.2.2 Excitotoxicity Promotes Signaling Module Formation .................................................117  vii  4.2.3 Identification of JNK Binding Motifs on zD17 ............................................................ 122 4.2.4 Isoform- and Scenario-Selective Inhibition of JNK ..................................................... 124 4.2.5 Effective Neuroprotection against Excitotoxicity by NIMoE ...................................... 126 4.2.6 Targeting the zD17-JNK Module Protects Brains from Ischemic Stroke .................... 129 4.3 Discussion........................................................................................................................... 132 4.3.1 The zD17-JNK Interaction as a Therapeutic Target for Neuroprotection .................... 133 4.3.2 A Novel Function of zD17 Independent of its PAT Enzyme Activity .......................... 137 Chapter 5. Conclusion ............................................................................................................ 138 5.1 Summary of Findings ....................................................................................................... 139 5.1.1 Role of AMPAR Palmitoylation in Neuronal Plasticity ............................................... 139 5.1.2 Role of JNK Palmitoylation in Neuronal Development ............................................... 141 5.1.3 Role of PAT zD17 in Neuronal Cell Death ................................................................... 143 5.1.4 Potential Therapeutic Target for Neuroprotection ........................................................ 145 5.2 Future Directions .............................................................................................................. 146 5.2.1 Physiological and Pathological Roles of Palmitoyl-Proteins in the Brain ................... 146 5.2.2 Drug Discovery Based on Palmitoyl Proteins .............................................................. 148 5.3 Conclusions ........................................................................................................................ 149 Bibliography .............................................................................................................................. 151 Appendices ................................................................................................................................. 173 A1. Reagent and Samples ....................................................................................................... 173 A2. Experiment Procedures .................................................................................................... 176  viii  List of Tables Table 1. Human palmitoyl acyltransferases: isoforms and subcellular locations........................... 7 Table 2. Examples of neuronal palmitoyl proteins. ........................................................................ 8 Table 3. Regulation of protein stability by palmitoylation. ...........................................................11  ix  List of Figures Figure 1. Lipid modification of proteins. ....................................................................................... 2 Figure 2. Biochemistry of palmitoylation. ..................................................................................... 4 Figure 3. The cysteine rich domain domain of PATs identified in human and yeast. .................... 6 Figure 4. Biochemistry of acyl-biotin exchange assay................................................................. 15 Figure 5. Biochemistry of ω-alkynyl or ω-azido palmitate detection. ......................................... 16 Figure 6. Strategies for functional assessment of protein palmitoylation. ................................... 19 Figure 7. Gene expression patterns of PATs in the mouse brain. ................................................. 28 Figure 8. Development stages of cultured neurons. ..................................................................... 35 Figure 9. Wnt signaling pathway in regulating neuron development. ......................................... 36 Figure 10. Diagram of JNK activation pathway and regulatory mechanisms. ............................ 39 Figure 11. Anterograde transport in the early secretory pathway is important for GluR1 palmitoylation. .............................................................................................................................. 62 Figure 12. GluR2 subunits are primarily palmitoylated in the ER. ............................................. 64 Figure 13. GluR2 palmitoylation is enhanced by BFA treatment in exogenous cells.................. 65 Figure 14. GluR2 subunits are transported to the Golgi in the presence of palmitate attached in the ER. ................................................................................................................................................ 67 Figure 15. Preventing palmitoylation does not enhance GluR2 aggregation in COS7 cells and cultured neurons. ........................................................................................................................... 68 Figure 16. Blocking palmitoylation destabilizes the mature GluR2 pool. ................................... 69 Figure 17. Blocking palmitoylation enhances reduction of GluR2 WT in HEK293 cells. .......... 71 Figure 18. Blocking palmitoylation by 2-BrPA does not affect the mRNA level of GluR2. ....... 72 Figure 19. Reduced palmitoylation induces lysosome-dependent degradation of GluR2. .......... 73 Figure 20. Detection of total and surface GluR1 and GluR2 after blocking neuronal activity. ... 75 Figure 21. AMPAR palmitoylation is regulated by neuronal activity. ......................................... 76 Figure 22. Turnover rate of palmitate on GluR1 and GluR2. ...................................................... 77 Figure 23. AMPAR subunit palmitoylation is differentially affected by acute NMDA and AMPA treatment. ...................................................................................................................................... 78 Figure 24. JNK3 is palmitoylated at the COOH-terminus. .......................................................... 96 Figure 25. The palmitoylation-deficient JNK3 mutant promotes axonal branching and filopodia motility. ......................................................................................................................................... 98 Figure 26. Palmitoylation regulates the translocation of JNK3 to the Triton-insoluble actin cytoskeleton. ............................................................................................................................... 101  x  Figure 27. JNK3 palmitoylation modulates Wnt7a-regulated axonal branching. ...................... 104 Figure 28. Formation of the zD17-JNK3 signaling module. ......................................................115 Figure 29. JNK phosphorylation is regulated by PAT zD17. .....................................................117 Figure 30. zD17 physically associates with JNK3 and regulates JNK activation in an MKK7-dependent manner. ..........................................................................................................118 Figure 31. Regulation of the zD17-JNK3 signaling module in response to NMDA-induced excitotoxicity................................................................................................................................119 Figure 32. Identifying novel JNK-interacting motifs on zD17 and achieving scenario-selective inhibition of isoform JNK2/3 with peptides derived motif-E. .................................................... 121 Figure 33. Isoform- and scenario-selective inhibition of JNK by peptides derived from JNK binding motifs on zD17. ............................................................................................................. 122 Figure 34. Axonal development with peptide application. ........................................................ 125 Figure 35. Neuroprotection against NMDA excitotoxicity is achieved by targeting the zD17-JNK interaction. .................................................................................................................................. 126 Figure 36. Effects of peptide application on NMDA excitotoxicity-induced neuronal cell death. ..................................................................................................................................................... 128 Figure 37. JNK downstream pathways are affected by peptide application. ............................. 130 Figure 38. Blocking the JNK-zD17 interaction with NIMoEsh protects from ischemic brain injury in rats. .......................................................................................................................................... 131 Figure 39. Potential mechanisms of zD17-mediated JNK activation and the intervention for neuroprotection. .......................................................................................................................... 134  xi  List of Abbreviations 2-BrPA, 2-bromopalmitate Ala, alanine AMPA, α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid AMPAR, AMPA receptor ABP, AMPAR-binding protein APT, acyl protein thioesterase BFA, brefeldin A CHX, cycloheximide CoA, coenzyme A Cys, cysteine DMEM, Dulbecco’s modified eagle’s medium SDH, dishevelled ECL, electrochemiluminescence EPL, expressed protein ligation ER, endoplasmic reticulum FRAP, fluorescent recovery after photonbleaching Fz, frizzled GABA, γ-aminobutyric acid GAD, glutamic acid decarboxylase GAP43, growth associated protein 43 GFP, green fluorescent protein Gly, Glycine GODZ, Golgi apparatus-specific protein with a DHHC zinc finger domain GPCR, G-protein coupled receptors GPI, glycophosphatidyl inositol G protein, guanine nucleotide-binding protein GRAP1, glutamate receptor associated protein 1 GRIP, glutamate receptor interacting protein GRK6, G-protein coupled receptor kinase 6 HAM, hydroxylamine HIP14, huntingtin-interacting protein 14 HRP, horseradish peroxidase  xii  Htt, huntingtin IPTG, isopropyl β-D-1-thiogalactopyranoside JIP, JNK-interacting protein JNK, c-jun N-terminal kinase MAPK, motigen-activated protein kinase NEM, N-ethylmaleimide NMDA, N-methyl-D-aspartic acid NMDAR, NMDA receptor NOC, nocodazole NSF, NEM-sensitive factor PAT, palmitoyl acyltransferase PFA, paraformaldehyde PI4KIIα, phosphatidylinositol 4-kinase II α PICK1, protein interacting with C-kinase 1 PKC, protein kinase C PM, plasma membrane PPT, palmitoyl protein thioesterase PSD-95, postsynaptic density-95 Ser, serine Shh, sonic hedgehog SNAP25, synaptosomal-associated protein 25 SYT, synaptotagmin TMD, transmembrane domain TTX, tetrodotoxin WT, wildtype zD, zinc-finger DHHC containing protein  xiii  Acknowledgments I would like to express my deep and sincere gratitude to my supervisor and mentor, Dr Max S. Cynader, for his guidance during my study at the University of British Columbia. His immense knowledge, perpetual enthusiasm for science and research, and thoughtful insights have helped me on the right track for my research and kept inspiring me. As a excellent scientist with rich experience and the director of the Brain Research Centre, he not only taught me research skills, but also enlarged my vision on neurosciences. His understanding, eternal encouraging and endless support throughout my doctoral study have helped me to develop essential research abilities, and allowed me to pursue my research interests. He has been a scientist role model for me at all times.  I am deeply grateful to Dr. Ann Marie Craig, Dr. Weihong Song, and Dr. YuTian Wang, my committee members and advisors. This thesis would not have been possible without their guidance at all levels of my research and study. Learning from these outstanding scientists has provided me a priceless opportunity to develop my expertise at different areas, broaden my research perspectives, and define my career path. Their wide knowledge, patience, and encouragement have constantly supported me. A very special thanks goes out to Dr. Alaa El-Husseini, my previous committee member, who introduced me to the field of protein palmitoylation and gave me insightful comments on my research projects before we lost him in 2007. I would like to thank Dr. Lynn Raymond, for taking time out from her busy schedule to serve as my external examiner for my comprehensive exam, and for her helpful suggestions on my research and career development. I also wish to express my sincere thanks to Dr. William Jia for his continuous support on my study and important suggestions for my career. I am grateful to Dr. Luba Kojic, who was always accessible and willing to help me with my research projects. I learned a lot from frequent discussions with him and his vast knowledge in many areas. I warmly thank Swarni Sunner, whose kind help and great support at all times allowed me to focus on my research study and made my student life a lot easier. My deepest gratitude goes to my family for the support they provided me through my entire life and in particular, I must acknowledge my wife and best friend, Jia. Without their love, understanding, encouragement and support, I would not have finished my doctoral study and this thesis.  xiv  I would also like to thank Dr. Ruijun Kang and Dr. Kun Huang for teaching me essential experimental techniques and sharing research experiences when I started my projects, and Wendy Wen for her essential assistance in routine research material preparation which enabled me to finish my thesis projects smoothly. I am indebted to many colleagues during my study. I am especially grateful to Dr. Yuan Liu, Dr. Dong Li, and Huan Bao for their technical support.  I am also grateful to all members in the Cynader Lab, Ainsley, Aobo, Bryson, Carlo, Chengyong, Dong, Jade, Kaiyun, Melanie, Rui, Shanshan, Wei, for providing a stimulating environment in which to learn and grow, which helped enrich my experience. My thanks also goes to many other friends in the Brain Research Centre for their help, support and all the fun we have had, Alan, Bo, Chao, Denis, Dongchuan, Huili, Jing, Jingfei, Juliet, Jun, May, Mianwei, Ning, Sharmin, Shu, Simon, Ted, Xiaojun, Xuguang, Xuefeng, Yuan, Zhifang – there are too many to name you all.  xv  Chapter 1. Introduction  1  1.1 Protein Palmitoylation and Palmitoyl Acyltransferases  1.1.1 Biochemical Basics of Protein Palmitoylation In eukaryotic systems, protein palmitoylation represents one of four major types of lipid modifications, in addition to NH2-terminal myristoylation, COOH-terminal prenylation, and glycophosphatidyl inositol/GPI addition (El-Husseini and Bredt, 2002; Bijlmakers and Marsh, 2003). In contrast with irreversible myristoylation and prenylation, the post-translational palmitoylation process is reversible and is dynamically regulated (Figure 1).  Figure 1. Lipid modification of proteins.  This is achieved by covalently linking palmitate to active free thiol groups of cysteine (Cys) residues and forming labile thioeaster connections, known as S-palmitoylation (Bijlmakers and Marsh, 2003). The addition and removal of palmitate on proteins provides a way to dynamically regulate protein functions. Despite the common involvement of palmitate in this modification, 2  other saturated or unsaturated fatty acids including stearate, oleate and arachidonate have also been described as attaching to proteins via S-acylation in a similar way as palmitate does (Bijlmakers and Marsh, 2003). The divergence in chemical and physical properties of these lipids may have different impacts on protein trafficking and function. Occasionally, palmitate can also be stably linked to the amino group of a NH2-terminal Cys, such as secreted morphogenic factor sonic hedgehog (Shh) (Pepinsky et al., 1998) . This process is known as N-palmitoylation (Figure 2). Since modified proteins in most cases are found to be S-acylated with palmitate, I adopt the commonly used term palmitoylation in this thesis for this type of post-translational modification.  In the past two decades, significant advances have been made to reveal the mechanisms underlying protein palmitoylation. Before the identification of corresponding enzymes, it was thought that a non-enzymatic process accounted for protein palmitoylation in vivo. This idea was based on the observations that auto-acylation can occur spontaneously in vitro when introducing palmitoyl-coenzyme A (CoA) to synaptosomal-associated protein 25 (SNAP25) (Veit, 2000) or guanine nucleotide-binding protein (G protein) Gα (Duncan and Gilman, 1996). However, it remains unknown whether auto-acylation occurs in vivo and has physiological significance. Moreover, several issues have arisen to question this auto-acylation model. First, auto-acylation in vitro requires a high concentration of palmitoyl-CoA that usually does not exist in vivo (Leventis et al., 1997). Second, auto-acylation requires long reaction times, whereas the kinetics of palmitoylation of some proteins varies from minutes to hours (El-Husseini and Bredt, 2002). 3  Third, proteins such as β-actin which are not normally palmitoylated in vivo are able to be palmitoylated under auto-acylation conditions in vitro (Duncan and Gilman, 1996). These concerns about the auto-acylation model encouraged the screening for enzymes that might regulate protein palmitoylation.  Figure 2. Biochemistry of palmitoylation.  A significant breakthrough was achieved by identifying yeast proteins Akr1p and Erf2p-Erf4p as members of a protein family which catalyze protein palmitoylation, known as palmitoyl acyltransferases (PATs) (Lobo et al., 2002; Roth et al., 2002). Despite their disparity in protein structures, they contain similar Cys rich DHHC domains and affect palmitoylation of 4  yeast proteins in vivo, such as Ras2p and Yck2p (Lobo et al., 2002; Roth et al., 2002). The screening of proteins that share the similar DHHC domain identifies at least 23 potent PATs in mammalian genome, although their ability to palmitoylate proteins has not examined individually (Figure 3) (Mitchell et al., 2006).  These PATs share little sequence similarity outside of the DHHC domain and reside in different subcellular compartments including endoplasmic reticulum (ER), the Golgi apparatus, endosome and the plasma membrane (PM) (Table 1) (Ohno et al., 2006). This variety of subcellular location and protein structure suggests diverse loci where protein palmitoylation may occur and potential functional differences. Interestingly, a recent study has identified histone H3 variants as palmitoyl proteins (Wilson et al., 2010). Their Cys sites for palmitoylation are only accessible during active gene transcription, suggesting a role of dynamic palmitoylation in regulating chromatin modulations (Wilson et al., 2010). However, no known PATs are found to be present in the cell nucleus. Although histone H3 proteins may be palmitoylated outside the nucleus followed by shuttling into the nucleus, the question of how dynamic palmitoylation is achieved in the nucleus and the functional role of palmitoylation in histone regulation requires further investigation.  5  Figure 3. The cysteine rich domain domain of PATs identified in human and yeast.  Although consensus motifs for palmitoylation have not been successfully identified, it has been noticed that protein palmitoylation displays high sensitivity and selectivity. Small variations in protein sequence or structure can result in alterations of palmitoylation status (Bijlmakers and Marsh, 2003). The presence of hydrophobic amino acids and the cluster of basic residues around potential Cys sites have been found to be important for palmitoylation (El-Husseini and Bredt, 2002). Mutations around palmitoylated Cys residues have been shown to affect palmitoylation efficiency (Liu et al., 1993). In addition to residue combination, other unclear factors also play a role. For example, one of two splicing variants of glutamate receptor interacting protein 1 (GRIP1) fails to be palmitoylated though both variants contain similar combinations of hydrophobic and basic residues around Cys sites (Yamazaki et al., 2001). Such differences have also been observed in the two spliced forms of glutamic acid decarboxylase (GAD65/67) (Solimena et al., 1994). Despite the uncertainty of the mechanisms underlying substrate selectivity, these lines of evidence suggest that protein palmitoylation is a delicately regulated process.  6  Table 1. Human palmitoyl acyltransferases: isoforms and subcellular locations.  7  1.1.2 Molecular Functions of Protein Palmitoylation A range of integral and peripheral membrane proteins, cytosolic proteins, and also secreted proteins have been reported to be palmitoylated (Bijlmakers and Marsh, 2003).  Table 2. Examples of neuronal palmitoyl proteins.  Protein  Motif  Palmitoylation function  Transmembrane Proteins GluR2  LVALIEFCYKSRAEAK (C-terminus) GAFMRQGCDISPRSL (loop)  release from Golgi and endocytosis  D1 receptor  LGCRGDK  receptor activity  Fas  VQKTCRKHR  actin cytoskeleton association Synaptic Vesicle Proteins  SYT I  VVTCCFCVCKKCL  presynaptic sorting  SYT VII  IVLCGLCHWCQRKL  lysosome targeting  SNAP25  LGJFCGLCVCPCNKLK  maintaining stable membrane association  Cell Adhesion Molecules NCAM  MDITCYFLNKCGLLMCIAVNLCGK  lipid raft targeting  Signaling Proteins H-Ras  GPGCMSCKCVLS  Golgi, PM and lipid raft targeting  Fyn  MGCVQCKDKE  PM and lipid raft targeting  GAP43  MLCCMRRT  neuronal compartment targeting  Paralemmin  KKHRCKCCSIM  axonal filopodia targeting Scaffolding Proteins  PSD-95  MDCLCIVT  dendrite and synapse targeting  ABP  LALCLQRL  spine targeting  Table modified from El-Husseini and Bredt, 2002.  8  One extensively studied impact of palmitoylation, like other lipid modifications, on targeted motifs or proteins is to increase hydrophobicity and thus promote their association with membrane structures. Fusing different domains derived from palmitoyl proteins to cytosol-distributed GFP is able to redirect it to the PM and other intracellular membrane compartments (Zacharias et al., 2002). This membrane targeting signal is used for dynamically regulating intracellular trafficking of several important neuronal proteins, including post-synaptic protein 95 (PSD-95) (El-Husseini et al., 2000; El-Husseini et al., 2002), AMPA receptor binding protein (ABP) (DeSouza et al., 2002), GAP43 (McLaughlin and Denny, 1999), GRIP (Yamazaki et al., 2001) and others (Table 2).  As neurons are polarized cells, palmitoylation of some neuronal proteins is found to be critical for their axonal or dendritic localization. For example, without normal palmitoylation, the dendritic protein, PSD-95, or axonal protein, GAD63, are no longer restricted to their individualcompartments (El-Husseini et al., 2000; Kanaani et al., 2004); instead, they are sorted to both dendrites and axons without preference. Within the PM, specific micro-domains enriched with lipids such as lipid rafts are regions where signalling proteins locate and initiate signal transduction (Simons and Toomre, 2000). Studies on CD8, Fas-associated protein with death domain (FADD), Fyn, H/N-Ras, and other lipid raft-associated proteins suggest that palmitoylation, in some cases, also acts as a specialized signal to aid proteins targeting to such micro-domains on the PM where they function (Rodgers et al., 1994; Arni et al., 1998; Perez and 9  Bredt, 1998; Prior et al., 2001; Zacharias et al., 2002; Chakrabandhu et al., 2007; Levental et al., 2010). This role of palmitoylation may not be solely due to the increase of hydrophobicity of proteins, as myristoylated or prenylated proteins are not targeted to lipid rafts (Zacharias et al., 2002).  It is not uncommon that palmitoylation occurs with other lipid modifications including prenylation and myristoylation to dynamically regulate protein association with lipid membrane structures (Table 2). One extensively studied example is H,N-Ras. Cytosolically localized H,N-Ras is first prenylated at the ER which allows transport to Golgi. Further palmitoylation at Golgi drives Ras trafficking to the PM where it can be depalmitoylated and delivered back to Golgi and ER. Thus, the dynamic process of palmitoylation regulates the cycling of Ras between the PM and intracellular membranes (Rocks et al., 2005). Paralemmin, an axon-targeting protein, is also dually modified by prenylation and palmitoylation (Gauthier-Campbell et al., 2004). Palmitoylation of kinase Fyn occurs after it is N-myristoylated at theN-terminus (van't Hof and Resh, 1997).  Besides serving as a PM/lipid-raft targeting signal, palmitoylation has been found to direct proteins to or from other subcellular compartments, including the lysosome (Flannery et al., 2010), actin cytoskeleton (Chakrabandhu et al., 2007), and the Golgi apparatus (Rocks et al., 2005) (Table 2). Recent data also suggest a role of palmitoylation in regulating protein 10  degradation in the proteosome by preventing ubiquitination (Gao et al., 1999; Percherancier et al., 2001; Wang et al., 2010). This strongly emphasizes the involvement of palmitoylation in regulating protein stability (Table 3).  Table 3. Regulation of protein stability by palmitoylation.  Protein  Degradation pathway  Mechanism  CCR5  proteosome  Prevent ubiquitination  A1 adenosine receptor  proteosome  Prevent ubiquitination  GPCR  proteosome  Prevent ubiquitination  Tlg1  vacuole  Prevent ubiquitination  Anthrax-toxin receptor  lysosome  Prevent targeting to lysosome  Recent studies have pinpointed a role of palmitoylation in regulating protein activity. The kinase activity of G-protein coupled receptor kinase 6 (GRK6) and phosphatidylinositol 4-kinase IIα (PI4KIIα) is found to be regulated by palmitoylation (Stoffel et al., 1998; Barylko et al., 2009). In some cases, palmitoylation has been shown to regulate aspects of protein-protein interactions. Palmitoylation of huntingtin (Htt) protein is found to be required for its interaction with Htt-interacting protein 14 (HIP14), which in turn affects the aggregation of Htt in the cytosol (Huang et al., 2009). Although palmitoylation should not be a necessary process for normal membrane targeting of integral membrane proteins, they are common substrates for palmitoylation (Bijlmakers and Marsh, 2003). It is possible that palmitoylation on these proteins may affect the structure of their cytosolic fragments as a hook on the PM for interacting with their appropriate partners . This is true in the case of AMPAR regulation. In this situation, 11  palmitoylation of GluR1 at the cytosolic COOH-terminus is found to affect GluR1’s interaction with protein N4.1, a protein associated with actin-cytoskeleton, which then drives GluR1 internalization from the PM (Lin et al., 2009).  1.1.3 Strategies for Studying Protein Palmitoylation Several approaches have been broadly used to assess biochemical and functional properties of protein palmitoylation in vitro and in vivo. Thanks to the development of bioinformatics and proteomics, we now are able to predict and identify a huge number of palmitoyl proteins. Using novel strategies to study protein palmitoylation have greatly enhanced our understanding of palmitoyl proteins and their physiopathological roles.  1.1.3.1 Prediction of protein palmitoylation  Although the identification of consensus motifs for palmitoylation has proven challenging, some patterns have been noticed based on known palmitoyl proteins. A potential palmitoylation site is always surrounded by several hydrophobic and/or charged amino acids, and the presence of hydrophobic residues (such as leucine/L, isoleucine/I, methionine/M) as observed in PSD-95 for example, are critical for efficient palmitoylation (El-Husseini et al., 2000; El-Husseini and Bredt, 2002). In many cases, proteins are dually palmitoylated at two adjacent Cys residues, or multiple Cys in nearby regions (Table 2). For many proteins that undergo myristoylation and  12  palmitoylation, Cys residues following the myristoylated glycine residue at N-termini are always palmitoylation sites (such as Fyn). Based on these and other observations, an open source software programme “CSS-Palm” has been published and used for predicting the possibility for palmitoylation of a particular protein and the potential palmitoylation sites using a mathematical algorithm (Ren et al., 2008). Many known palmitoyl proteins and their sites for palmitoylation are predicted with the CSS-Palm software and it provides useful guidance to palmitoyl protein candidates, though further experimental validation beyond the software’s prediction, should be always employed.  1.1.3.2 Metabolic labelling and acyl-biotin exchange  To assess the incorporation of palmitate to proteins, metabolic labelling with 3H- or 14Cpalmitate is the most commonly used method. The incorporation of palmitate is then examined by immunoprecipitation with antibodies against protein targets of interest, SDS-PAGE, and autoradiography (Yang et al., 2009). The sensitivity and selectivity of metabolic labelling allow the identification and confirmation of particular palmitoyl proteins. Using metabolic labelling, the kinetics of palmitoylation on proteins can be easily assessed using classical pulse-chase experiments (Yang et al., 2009). However, metabolic labelling does give several limitations. First, the palmitoylation status of endogenous proteins in vivo, such as occurs in the brain, is difficult to assess. Second, the labelling period is relatively long, usually several hours, and cannot be  13  used to examine palmitoylation that is rapidly changed during a period of minutes. Third, metabolic labelling is unsuitable for identification of a large quantify of unknown proteins and is unable to analyze protein palmitoylation in a large-scale. Fourth, short-time labelling only reflects the incorporation rate of labelled palmitate to proteins, but cannot provide information on the readiness status of the palmitoyl pool of proteins.  To overcome these drawbacks, Schmidt and colleagues have invented the acyl-biotin exchange assay (biotin-BMCC labelling) to assess protein palmitoylation (Schmidt et al., 1988; Drisdel and Green, 2004) (Figure 4). In this method, free thiols of Cys residues on extracted total proteins are blocked by N-ethylmaleimide (NEM), followed by removal of palmitate from proteins with hydroxylamine (HAM). Via the thiol reactive group of biotin (btn)-BMCC, the newly freed thiols of Cys are then labelled with biotin. Once the formerly-palmitoylated Cys is labelled with biotin, palmitoylation can be analyzed indirectly by pulling down biotin-labelled proteins (path 1) or by detecting biotin on the protein directly (path 2). The readiness status of the palmitoylated pool of proteins can be examined by btn-BMCC labelling (Schmidt et al., 1988; Drisdel and Green, 2004). By applying the palmitoylation inhibitor 2-bromopalmitate (2-BrPA), the turnover rate of palmitate on proteins can also be determined (Kang et al., 2008). Potential drawbacks of this method are the specificity and sensitivity, because the presence of free Cys residues may give false positive signals and the efficiency of relabeling of Cys residues may give false negative signals. Moreover, the fatty exchange assay cannot distinguish the different lipids 14  covalently attached to Cys residue. The reaction is also sensitive to pH conditions.  Figure 4. Biochemistry of acyl-biotin exchange assay. Two protocols were taken to determine protein palmitoylation. In protocol 1, after replacing palmitate with biotin, total palmitoylated proteins were isolated by streptavidin-labelled beads. In protocol 2, a particular protein was first isolated by immunoprecipitation, followed by Biotin-BMCC labelling. Total palmitoylation of the target protein was determined by blotting with avidin-HRP.  As a combination of above methods, labelling live cells with modified lipids provides an alternative way to study protein palmitoylation. Incubation with ω-azido palmitate or ω-akynyl palmitate has been shown to effectively substitute for endogenous palmitate and to label proteins at palmitoylation sites (Hang et al., 2007; Kostiuk et al., 2008; Hannoush and Sun, 2010) (Figure 5). Because the azide group is reactive to phosphine derivatives conjugated with tags or fluorophores, and the akynyl group is reactive to conjugated azide groups, labelled proteins can be then analyzed with corresponding strategies. By reducing potential false positives introduced by in vitro labelling, this method provides a higher specificity for the identification of palmitoyl proteins. 15  Figure 5. Biochemistry of ω-alkynyl or ω-azido palmitate detection.  1.1.3.3 Large-scale identification of the palmitoyl-proteome  The combination of the acyl-biotin exchange assay and mass-spectrometry highlights a novel strategy to identify massive quantity of palmitoyl proteins (Wan et al., 2007). Using newly developed methodologies, more than 30 new palmitoyl proteins have been identified from the yeast (Roth et al., 2006). The characterization of palmitoyl proteome has also been done recently in cultured mammalian neurons and about 200 neuronal proteins are identified as new palmitoyl protein candidates (Kang et al., 2008). Most of these new palmitoyl proteins are neurotransmitter receptors, transporters, adhesion molecules, and proteins involved in regulating transmitter 16  release and vesicle trafficking. Given the presence of 23 PATs in mammalian cells, however, more palmitoyl proteins remain likely undiscovered.  Recently, several large-scale screenings of fatty acyl modified proteins have been taken using modified methods including azido pamitate metabolic labelling and biotin labeling, leading to identification of new palmitoyl proteins in rat liver mitochondrial matrix (Kostiuk et al., 2008). Using a resin-based binding and purification method, palmitoyl proteins are retained on beads containing thiol-reactive thiopyridinyl groups, and then identified by mass spectrometry-based proteomics (Forrester et al., 2010).  In addition to screening new palmitoyl proteins, palmitoyl-proteome analysis provides a way to investigate substrate selectivity of a particular PAT. After knocking down one of the PATs, for example zD2 with siRNA, palmitoyl proteins are compared to control samples (Zhang et al., 2008). Proteins with a reduced palmitoylation level are then assigned to zD2 as its potential substrates. Although there are some concerns about the redundancy of PATs and complementary effects by knocking down a particular PAT, this strategy provides a valuable tool for examining PAT-substrate specificity.  1.1.3.4 Analysis of palmitoylation in the secretion pathway  PATs are resident at different subcellular compartments where they palmitoylate substrates 17  or have other functions. An intact secretion pathway has been shown to be important for palmitoylation of some proteins. Brefeldin A (BFA), a drug that destroys the cell’s Golgi apparatus, has been used to block secretion from the Golgi apparatus to the PM (Lippincott-Schwartz et al., 1989). Palmitoylation of several transmembrane and cytosolic proteins has been shown to be sensitive to BFA treatment, including proteins such as CD151 (Yang et al., 2002), SNAP25 (Gonzalo and Linder, 1998), GAP43 (Flannery et al., 2010) and others. The Golgi apparatus and post-Golgi intracellular compartments are then suggested to be the sites of their palmitoylation. BFA causes collapse of the Golgi apparatus and redistributes Golgi enzymes back to ER, resulting in palmitoylation of some ER resident proteins which are not normally palmitoylated (Lippincott-Schwartz et al., 1989). This process is mediated by microtubule systems which can be blocked by adding the microtubule depolymerizer nocodazole (NOC) (Lippincott-Schwartz et al., 1991). These pharmacological approaches together with the metabolic labelling assessment provide important information about the subcellular locations at which the palmitoylation of particular proteins occurs.  18  Figure 6. Strategies for functional assessment of protein palmitoylation.  1.1.3.5 Functional assessment of protein palmitoylation  In cultured cell systems, several strategies have been widely used to study molecular functions of protein palmitoylation (Figure 6). Brief incubation with the palmitoylation inhibitor 2-BrPA has been shown to reduce protein palmitoylation and change the corresponding properties of proteins (Figure 6A) (Webb et al., 2000; Draper and Smith, 2009). For example, 2-BrPA effectively blocks PSD-95 palmitoylation and stops its trafficking to postsynaptic sites (El-Husseini et al., 2002). The inhibition of 2-BrPA is reversible (El-Husseini et al., 2002). Although it is effective in blocking palmitoylation, 2-BrPA shuts down the palmitoylation system and does not have any specificity for one particular group of proteins. Moreover, because 2-BrPA affects the incorporation of palmitate, the elimination of palmitate on proteins by 2-BrPA 19  depends on the kinetics of protein palmitoylation. Proteins with a longer turnover rate of palmitate, such as synaptotagmin I (SYT-I), are expected to be less sensitive than those with rapid kinetics, such as PSD-95 (El-Husseini et al., 2002; Kang et al., 2004).  An alternative and commonly used way to manipulate protein palmitoylation is the creation of point mutation of Cys residues on which palmitoylation occurs. Based on the structural similarities of amino acids, alanine (Ala) or serine (Ser) are usually used as the replacements of Cys (Figure 6B). The point mutation eliminates palmitoylation and corresponding consequences. Using live-imaging fluorescent recovery after photonbleaching (FRAP), mutated Ras has been shown to accumulate at the Golgi apparatus, but not to be delivered to the PM (Rocks et al., 2005). Overexpressing mutated proteins as dominant negatives has also been shown to interfere with endogenous proteins. Mutation of Cys to Ser not only abolishes PSD-95 trafficking properties, but also reduces AMPAR accumulation at synapses, which is controlled by normal clustering of PSD-95 at postsynaptic sites (El-Husseini et al., 2000; El-Husseini et al., 2002). However, introducing exogenous proteins may cause potential artefacts and has drawbacks when used to examine endogenous protein palmitoylation.  Overexpression of certain PATs has facilitated the analysis of the function of protein palmitoylation in some studies by promoting protein palmitoylation (Figure 6C). Introducing the PAT zD15 promotes PSD-95 palmitoylation and enhances its targeting to postsynaptic membrane 20  and also enhances AMPAR accumulation (Fukata et al., 2004). Overexpression of the PAT zD17 promotes Htt palmitoylation and reduces aggregation of the poly-glutamine (poly-Q) repeats Htt mutant (Singaraja et al., 2002). However, given the ambiguous substrate selectivity of PATs, palmitoylation of multiple substrates by single PAT overexpression is to be expected, and this makes it challenging to study the physiological functions of palmitoylation of one particular protein.  As an alternative approach, generating a pseudo-palmitoylated protein has been used to study the molecular and physiological functions of palmitoyl proteins (Figure 6D). A COOH-terminal motif of paralemmin which is irreversibly prenylated and dually palmitoylated has been used to generate a fusion protein to mimic constant palmitoylation on PSD-95 (Kutzleb et al., 1998; El-Husseini et al., 2002). Expression of this pseudo-palmitoylated PSD-95 is able to restore intracellular trafficking of PSD-95 and the accumulation of AMPAR at postsynaptic sites even in the presence of the palmitoylation inhibitor 2-BrPA (El-Husseini et al., 2002).  In addition to the expression of mutated proteins by endogenous systems, microinjection of in vitro engineered and modified proteins has also been used to study the effects of palmitoylation on protein trafficking (Rocks et al., 2005). Synthetic peptides that are modified with lipids are fused to proteins expressed in vitro by expressed protein ligation (EPL) (Brunsveld et al., 2006). The fusion protein is then introduced to cells by microinjection. Taking 21  Ras proteins as an example, palmitoylated control protein and a hexadecylated protein with a noncleavable thioether bond that cannot undergo de/reacylation are transiently injected into cells and the cycling of Ras proteins between Golgi and the PM are monitored in living cells using microscopy (Rocks et al., 2005). This approach can provide valuable information on the function of palmitoylation in the regulation of protein trafficking.  1.1.4 Palmitoyl Acyltransferases / PATs 1.1.4.1 Substrate selectivity of PATs  In most, if not all, situations, one substrate can always be palmitoylated by multiple PATs and one PAT has multiple substrates (Fukata et al., 2004; Planey and Zacharias, 2009). Although the substrate selectivity of PATs is ambiguous, it has been observed that some PATs exhibit a preference to palmitoylate a particular group of proteins. In the case of PSD-95, several PATs have been identified as its catalytic enzymes in vitro, named as P-PATs (zD2, 3, 7, 15) (Fukata et al., 2004). Under different conditions, zD17 may also be a potential PAT for PSD-95 (Huang et al., 2004; Huang et al., 2009). Despite the lack of evidence to confirm that all these PATs are involved in palmitoylation PSD-95 in vivo, this highly suggests a redundancy of PATs in palmitoylating proteins. Interestingly, knocking down only one of P-PATs, such as zD15 or zD17, is sufficient to block PSD-95 palmitoylation in neurons (Fukata et al., 2004; Huang et al., 2004). It is still unknown how multiple PATs work co-ordinately to palmitoylate a selected group of 22  substrates in vivo. Among the 23 PATs, zD13 and zD17 are unique due to their special ankyrin domains located at their cytosolic NH2-termini (Singaraja et al., 2002). Ankyrin domains are known as regions for protein-protein interactions (Mosavi et al., 2004). In agreement with this function, zD17 has been found to interact with its substrates via ankyrin domains and this protein-protein interaction is necessary for palmitoylation of its substrates (Huang et al., 2009). As the structure of the NH2-terminus of zD13 is different from that of zD17, a different profile of protein interactions and substrate selectivity is expected. Though ankyrin domains are not observed in other PATs, certain profiles of the protein-protein interaction may exist to ensure substrate selectivity.  1.1.4.2 Multiple functions of PATs  PATs catalyze palmitoylation by receiving palmitate from a palmitate-CoA donor to the Cys residue within the DHHC domain and then transferring palmitate to their substrates (Mitchell et al., 2010). In addition to the function of catalyzing palmitoylation, some PATs have been found to act as ion channels in cells. PAT zD3 (Golgi-specific DHHC zinc finger protein/GODZ) also acts as a calcium channel while PAT zD17 and zD13 act as magnesium channels at the Golgi apparatus (Goytain et al., 2008; Hines et al., 2009). Their palmitoyl transferase activity is found to be necessary for their channel activity. Given the presence of 23 PATs in mammalian cells, their distinct subcellular localizations and broad range of protein sequences, these discoveries  23  highlight the idea that PATs may possess multiple functions. Identification of additional functions of PATs will facilitate our understanding of the physiological roles of PATs.  1.1.5 Protein Thioesterases 1.1.5.1 Turnover of palmitate  In contrast to myristoylation and prenylation, protein palmitoylation is reversible and is highly dynamic on some proteins, such as Ras and PSD-95 (Bijlmakers and Marsh, 2003). Different turnover rates of palmitate on distinct groups of proteins correlate with functions of palmitoylation on these proteins. In the case of PSD-95, the half-life of palmitate (2 hours) is significantly shorter than the half-life of PSD-95 protein (>12 hour) (El-Husseini et al., 2002). Such rapid depalmitoylation and repalmitoylation cycling allows dynamic regulation of PSD-95 trafficking to postsynaptic sites and of synaptic AMPAR clustering in response to stimuli (El-Husseini et al., 2000; El-Husseini et al., 2002). Indeed, synaptic stimulation with AMPA and/or NMDA that decluster postsynaptic structures enhance the dynamics of PSD-95 palmitoylation/depalmitoylation, whereas reducing synaptic activity by TTX, that leads to accumulation of AMPAR at postsynaptic sites, extends the half-life of palmitate on PSD-95 (El-Husseini et al., 2002).  In contrast, SYT-I is a transmembrane protein for docking transmitter vesicles at  24  presynaptic sites (Kang et al., 2004). The turnover rate of palmitate on SYT-I is relatively slow (>12 hours) (Kang et al., 2004). Rapid neuronal depolarization by AMPA or NMDA does not affect palmitate turnover on SYT-I. But long-term treatment with TTX modulates palmitoylation and trafficking of SYT-I (Kang et al., 2004). The palmitoylation/depalmitoylation of SYT-1 is thus implicated in regulating transmitter release. The turnover of palmitate on proteins is tightly regulated by depalmitoylation and palmitoylation, and is important for establishing the functions of palmitoyl proteins.  1.1.5.2 APT and PPT  While great advances have been made in understanding protein palmitoylation, little is known about the depalmitoylation process. In order to tightly control the rapid turnover of palmitate, presumably a group of depalmitoylation enzymes would exist, in collaborating with PATs to regulate palmitoylation. One such enzyme discovered for protein depalmitoylation is acyl protein thioesterase 1 (APT1) (Duncan and Gilman, 1998). APT1 has been found to catalyze depalmitoylation of Ras, Ga and eNOS in vitro. In addition, palmitoyl protein thioesterase-1 (PPT1) was also identified and found to show depalmitoylation activity for a range of palmitoyl peptides (Camp et al., 1994; Soyombo and Hofmann, 1997). And mutation of PPT1 leads to infantile neuronal ceroid lipofuscinosis (Vesa et al., 1995; Mitchison et al., 1998). In contrast to PATs, most of which are ER/Golgi resident proteins, PPT1 is primarily found in the lysosome, a  25  locus for protein degradation (Camp et al., 1994). It is still unknown whether PPT1 represents the major depalmitoylation enzyme in vivo. Given the subcellular distribution of the majority of palmitoyl proteins, a group of depalmitoylation enzymes that are located outside of the lysosome is expected and remains to be identified.  A recent study has identified an APT1 inhibitor called palmostatin B that is active in cellular assays (Dekker et al., 2010). Application of this APT1 inhibitor significantly blocks depalmitoylation of H,N-Ras, and also its activity, which may be used for further identification of anti-cancer therapies (Dekker et al., 2010). However, considering that only two acyl thioesterases are currently identified, the impact of using a general APT1 inhibitor on other palmitoyl proteins needs more investigation.  1.2 Physiological Roles of Palmitoylation in the Brain In yeast, palmitoylation has been shown to contribute to diverse physiological processes as several proteins are found to be regulated by palmitoylation (Roth et al., 2006). Recently, the physiological roles of palmitoylation in the brain has attracted a lot of attention due to the identification of several synaptic proteins as palmitoyl proteins. Abnormal palmitoylation of these proteins results in altered synaptic transmission and interference with synaptic plasticity (El-Husseini and Bredt, 2002). Discoveries of signalling molecules as palmitoyl proteins further 26  support a role for palmitoylation in regulating brain development and function.  1.2.1 Palmitoyl Proteins in the Brain Given that palmitoylation is important for protein trafficking, the contribution of palmitoylation to the regulation of synaptic proteins and proteins that are targeted to special neuronal compartments including dendrites and axons has been extensively studied. One group of palmitoyl proteins includes transmembrane receptors. G-protein coupled receptors (GPCR) are palmitoylated to directly target them to lipid rafts, where multiple signalling modules are located (Barnett-Norris et al., 2005; Resh, 2006). Malfunction of palmitoylation results in failure to drive GPCR to lipid rafts where they initiate appropriate signals (Barnett-Norris et al., 2005). The GABA-A receptor can be palmitoylated and normal palmitoylation of GABA-A receptor is necessary for its normal synaptic targeting and maturation of inhibitory synapses (Fang et al., 2006). Excitatory AMPARs can also be palmitoylated at multiple sites (Hayashi et al., 2005). Distinct effects of palmitoylation at different sites are observed (Hayashi et al., 2005; Yang et al., 2009). While palmitoylation at the transmembrane domain (TMD) 2 of AMPAR retains receptors at the Golgi apparatus, palmitoylation at the C-terminus controls AMPAR internalization from the PM. For these transmembrane proteins, palmitoylation always occurs at the TMD or the cytosolic domain. However, the role of palmitoylation on transmembrane proteins is still unclear. One potential role of palmitoylation is to change the protein structure and thus its availability for various protein-protein interactions or for other modifications, such as phosphorylation. 27  Figure 7. Gene expression patterns of PATs in the mouse brain.  Another group of palmitoyl proteins consists of cytosolic and membrane associated proteins. Palmitoylation has been shown to facilitate the targeting of these proteins to specific membrane domains, or to direct proteins to neuronal compartments such as dendrites and axons (El-Husseini and Bredt, 2002; Bijlmakers and Marsh, 2003). PSD-95 is a well documented palmitoyl protein which contributes to regulation of synaptic strength and plasticity. Dynamic palmitoylation of PSD-95 ensures its specific targeting to dendritic spine synapses and facilitates 28  clustering of AMPAR’s (El-Husseini et al., 2002). In contrast, the palmitoylation of GAD65, a presynaptic protein, specifies its transport to axons but not dendrites (Solimena et al., 1994). Blocking palmitoylation alters axon-specific targeting of GAD65 and redirects it to dendrites. Another example includes SYT-I, a presynaptic protein. Palmitoylation of SYT-I is necessary for axonal targeting of SYT-I and its association with presynaptic vesicles (Kang et al., 2004).  Secreted proteins represent another group of palmitoyl proteins. Examples are members of the family of Wnt proteins and Shh (Pepinsky et al., 1998; Kurayoshi et al., 2007). Wnt and Shh are morphogens involving in specifying differentiation patterns in many tissues including the brain (Logan and Nusse, 2004). Palmitoylation of these proteins has been found to affect their association with the PM and corresponding initiation of signals via their receptors (Pepinsky et al., 1998; Kurayoshi et al., 2007).  Recently, large-scale screening has been used to identify palmitoyl proteins in neurons (Kang et al., 2008). More than 200 proteins were identified as new palmitoyl proteins in addition to the 50 previously known palmitoyl proteins. Although several kinases have been identified as palmitoylated proteins, such as yck1 in yeast, Fyn and Lck in mammalian cells, the importance of palmitoylation in regulating kinases in the brain remains largely unknown. Cdc42, an upstream activator of the mitogen-activated protein kinase (MAPK) pathway, is found to be palmitoylated (Kang et al., 2008). Palmitoylation of cdc42 ensures its activation and promotes 29  filopodia formation and establishment. Since kinases are important for signalling transduction, palmitoylation provides a potential way to regulate trafficking and function of kinases.  1.2.2 PATs in the Brain Among the 23 mammalian PATs, 20 of them are abundant with various expression levels in the brain . Moreover, these PATs show distribution preferences among brain regions, implying the involvement of different PATs in divergent brain functions (Ohno et al., 2006) (Figure 7). In recent years, some PATs ha ve been implicated in brain functions via palmitoylation of important neuronal proteins. For example, PAT zD3 is able to palmitoylate GABA-A receptors and AMPARs in neurons and to regulate their synaptic trafficking and function (Fang et al., 2006). Although zD3 is not highly expressed in the brain compared to other PATs, loss of zD3 results in developmental deficiencies among inhibitory synapses in cultured neurons (Fang et al., 2006). Interestingly, excitatory synapses appear to develop normally though AMPAR is also a substrate for zD3 (Hayashi et al., 2005). Due to our lack of knowledge of the full substrate spectra of each PAT, defining functional associations between PATs and potential substrates remains a major challenge for studying the physiological roles of PATs in brain systems.  30  1.2.3 Regulation of Functional Synapses and Signal Transmission 1.2.3.1 Synaptic plasticity and AMPAR  Changes in synaptic strength are believed to be the mechanisms underlying the process of learning and memory, known as neural plasticity (Barry and Ziff, 2002). Postsynaptic AMPARs are critical for fast synaptic transmission in the mammalian brain (Malenka, 2003). In response to appropriate stimuli, AMPARs are able to insert into and be maintained at postsynaptic sites, resulting in the enhancement of transmission, known as long-term potentiation (LTP). In contrast, the persistent reduction of the number of AMPARs causes diminution of transmission, known as long-term depression (LTD). The underlying mechanisms responsible for regulating the trafficking of postsynaptic AMPARs are thus believed to contribute to the process of learning and memory (Barry and Ziff, 2002; Malenka, 2003; Collingridge et al., 2010).  In the past two decades, our understanding of these mechanisms has significantly expanded. The trafficking of AMPARs is now known to be regulated by various binding proteins, in response to external stimuli (Malenka, 2003; Shepherd and Huganir, 2007; Collingridge et al., 2010). The proteins include scaffolding proteins such as synapse-associated protein 97 (SAP97), PSD-95, ABP, stargazin, and others. Binding to these proteins helps AMPAR targeting to appropriate locations. Alterations of these proteins can, directly or indirectly, affect the clustering of AMPARs at synapses.  31  In addition to the regulation introduced by other binding proteins, AMPAR is regulated by post-translational modifications. Phosphorylation of AMPAR by various kinases has been shown to contribute to AMPAR channel properties and AMPAR synaptic insertion and retraction during synaptic plasticity. Phosphorylation of GluR1 at Ser845 by protein kinase A (PKA) enhances AMPAR currents by increasing channel open time probability (Roche et al., 1996) and direct synaptic incorporation (Esteban et al., 2003), while phosphorylation of GluR1 at Ser831 by CaMKII increases single-change conductance (Derkach et al., 1999). GluR1 phosphorylation at Ser818 by PKC also regulates AMPAR insertion during LTP (Boehm et al., 2006). For LTD, phosphorylation of GluR1 threonine (Thr) 840 by PKA/PKC and of GluR2 Ser880 by PKC (Chung et al., 2000) and tyrosine (Tyr) 869, 873 and 876 (Ahmadian et al., 2004) are critical for AMPAR internalization.  1.2.3.2 Differential roles of subunits in AMPAR trafficking  The AMPAR family consists of four types of subunits, GluR1-GluR4 (Ozawa et al., 1998). Various combinations of the four AMPAR subunits (GluR1~GluR4) account for different properties of heterotetrameric receptors, such as ion conductance (Kumar et al., 2002), channel opening properties (Ozawa et al., 1998) and synaptic targeting of AMPAR’s (Shi et al., 1999; Bredt and Nicoll, 2003; Malenka, 2003; Collingridge et al., 2010). The most common form of AMPAR is the heteromer of GluR1 and GluR2 subunits. These subunits have been shown to  32  independently contribute to different aspects of AMPAR synaptic functions. One intensively studied example is subunit-dependent AMPAR trafficking. It is believed that GluR1 drives AMPAR synaptic incorporation during LTP, whereas GluR2 plays a key role in AMPAR endocytosis during LTD (Shi et al., 1999; Ehlers, 2000; Lee et al., 2004; Isaac et al., 2007). This functional division of AMPAR trafficking is governed by subunit-specific regulations. For example, by means of a sequence at its C-terminus, GluR1 specifically interacts with SAP97, which facilitates surface delivery of AMPAR from intracellular compartments (Sans et al., 2001). In contrast, GluR2 is targeted to and stabilized at synapses by associating with stargazin (Vandenberghe et al., 2005), and ABP (Osten et al., 2000). At the synapses, AMPARs are also anchored by interacting with NEM-sensitive factor (NSF) via GluR2 (Luthi et al., 1999). However, in response to stimuli that induce LTD, proteins interacting with C-kinase (PICK1) drives PKC to synapses and phosphorylates GluR2 at Ser880, which dissociates ABP/GRIP from GluR2 and enhances the interaction between GluR2 and PICK1 (Collingridge et al., 2010). In addition, the NSF binding site on GluR2 is rapidly occupied by the AP2 adaptor complex which then recruits the clathrin-mediated receptor internalization machinery (Collingridge et al., 2010). Other subunit-specific phosphorylations catalyzed by distinct sets of kinases are also involved in regulating AMPAR trafficking. Differential processing of AMPAR subunits enables a high flexibility of AMPAR trafficking regulation and function.  33  1.2.3.3 Effects of protein palmitoylation on synaptic transmission  Several AMPAR binding proteins have been identified as palmitoyl proteins. These include PSD-95 (El-Husseini et al., 2000), ABP (DeSouza et al., 2002), and GRIP1 (Yamazaki et al., 2001). Palmitoylation of these proteins has been shown to affect their trafficking and compartment targeting to synapses (Greaves and Chamberlain, 2007). As these scaffolding proteins are critical for regulating AMPAR trafficking, alteration of their palmitoylation status can destroy the AMPAR clustering at the synapses, indicating an essential role of palmitoylation in neuronal plasticity.  Recently, AMPAR subunits have been found to be palmitoylated at two Cys sites: one at the TMD2 domain, the other at the COOH-terminus (Hayashi et al., 2005) (Table 2). Palmitoylation at these two sites leads to differential effects on AMPAR trafficking (Hayashi et al., 2005; Yang et al., 2009). While palmitoylation on GluR2 at the TMD2 enhances AMPAR retention in Golgi, palmitoylation on GluR1 at the COOH-terminus facilitates AMPAR internalization, possibly by promoting dissociation of surface AMPAR from 4.1N (Lin et al., 2009). This highlights new molecular mechanisms for subunit-dependent AMPAR regulation.  Palmitoylation also plays a role in regulating presynaptic functions, such as transmitter release. Presynaptic proteins SNAP25 and SYT-1 are vesicle-associated proteins involved in regulating transmitter release (Gonzalo and Linder, 1998; Kang et al., 2004). Blocking their 34  palmitoylation abolishes their normal targeting to presynaptic sites and interferes with transmitter vesicle transport.  Figure 8. Development stages of cultured neurons.  1.2.4 Regulation of Neuronal Development 1.2.4.1 General background of neurite development  Axonal and dendritic development is essential for the normal formation of neuronal connections and brain networks. In vitro, the development of neurons can be categorized into several stages (Figure 8): i) formation of lamina around cell bodies (d0.25); ii) growth of neurites (d1); iii) initiation and growth of axons (d1.5); iv) extension of dendrites (d5); v) maturation of synapses (>d10) (Dotti et al., 1988; Kaech and Banker, 2006). Several classes of growth factors and signalling molecules have been characterized as key regulators for neurite development, 35  such as neural growth factor (NGF) and brain-derived neurotrophic factor (BDNF) etc.  Figure 9. Wnt signaling pathway in regulating neuron development.  Members of the Wnt family of proteins represent one group of prominent regulators for neurite development (Salinas and Zou, 2008) (Figure 9). As ligands, Wnt family members bind to cell-surface receptors of the Frizzled (Fz) family and activate the binding partners such as the Dishevelled (DSH) family of proteins. Via the canonical pathway, upstream Wnt signals recruit β-catenin, a gene transcription factor, to control the expression of genes in the nucleus that are 36  involved in neurite development, such as pax2. This recruitment is achieved by inhibiting the axin/GSK-3/APC complex that promotes β-catenin degradation in the cytoplasm (Salinas and Zou, 2008). Further studies have uncovered a noncanonical pathway that exists downstream of Wnt signals to regulate cell polarity. In this novel pathway, Wnt stimulates either calcium signals or recruits the small GTPase Rho/Rac and c-jun N-terminal kinase (JNK) signalling molecules. These proteins are found to regulate actin filaments and the microtubule cytoskeleton which contributes to cell polarity and migration (Salinas and Zou, 2008). In the brain, the expression of two Wnt members, Wnt7a and b, are upregulated in the hippocampal formation and contribute to activity regulated axon and dendrite development in vitro and in vivo (Gogolla et al., 2009). It has been observed that Wnt7a regulates axonal branching and remodelling, while Wnt7b modulates the formation of complex dendritic arborization (Lucas and Salinas, 1997; Rosso et al., 2005). The Rho-Rac-JNK signalling pathway is found to mediate Wnt-induced neurite development, possibly through cytoskeleton modulation (Rosso et al., 2005; Davis et al., 2008). Moreover, Wnt signals have also been discovered to play a role in axon guidance in the corpus callosum (Keeble et al., 2006) and spinal cord (Lyuksyutova et al., 2003).  1.2.4.2 The role of the JNK pathway in regulating neurite development  JNK is one of three members of the MAPK family which also includes ERK and p38 (Weston and Davis, 2007; Haeusgen et al., 2009). JNK is activated by a kinase cascade  37  containing MAPK kinases (MAPKK/MKP2K), e.g. MKK4 and MKK-7, and MAPKK kinases (MAPKKK/MAP3K), e.g. ASK1, the mixed-lineage kinases (MLK1/2/3), MEKK1/4 and TAK1 (Weston and Davis, 2007) (Figure 10). JNK activation is regulated by phosphorylation on Thr and Tyr residues within the threonine-proline-tyrosine (TPY) motif by upstream kinases (Weston and Davis, 2007). In the brain, this phosphorylation responds to diverse internal and external stimuli, including ER stress, oxidative stress, inflammatory factors, excitatory transmitter toxicity (excitotoxicity), axonal transport blockage, protein aggregation and other stresses (Haeusgen et al., 2009). Therefore, JNK has been extensively studied in many neurological diseases, e.g. stroke, Alzheimer’s disease, Huntington’s disease (HD) (Haeusgen et al., 2009).  In addition to neuronal stress response, a role of JNK in neurite growth and development has been recently recognized (Coffey et al., 2000; Chang et al., 2003; Waetzig and Herdegen, 2003; Kimberly et al., 2005; Bogoyevitch, 2006; Tararuk et al., 2006; Ciani and Salinas, 2007). Inhibition of JNK kinase activity results in blockade of axon initiation and growth (Oliva et al., 2006). Moreover, deletion of JNK1 in mice causes abnormal development of microtubules and neurites (Chang et al., 2003). A pro-apoptosis JNK family member, JNK3, has also been shown to induce neurite formation (Waetzig and Herdegen, 2003). These observations strongly suggest the involvement of JNK in regulating neurite development.  38  Figure 10. Diagram of JNK activation pathway and regulatory mechanisms.  Another line of evidence comes from the study of the signalling pathways of growth factors and signalling molecules, including Wnt. In the brain, Wnt regulates the development of neurites via this non-canonical pathway involving JNK and cytoskeleton rearrangement (Rosso et al., 2005). Consistent with this notion, small GTPase Rho, Rac1, and cdc42 that are activators of JNK are major regulators of the cytoskeleton in neurons (Weston and Davis, 2007). On the other hand, several cytoskeleton modulators known to be involved in regulating neurite development are JNK substrates, such as the actin modulator paxillin, and microtubule regulators microtubule 39  associated protein (MAP)2 and MAP1b (Bogoyevitch and Kobe, 2006). Phosphorylation of MAP2/1b by JNK has been shown to mediate Wnt-induced axonal development (Ciani and Salinas, 2007).  1.2.4.3 Palmitoyl proteins in regulation of neuronal development  Secreted Wnt proteins are palmitoylated. Wnt palmitoylation does not affect the secretion of Wnt, but is important for the triggering of signalling at the cell surface, possibly by enhancing binding ability to its receptors (Komekado et al., 2007; Kurayoshi et al., 2007). A recent study has reported a role of palmitoylation in regulating small GTPase cdc42, a regulator of cytoskeleton networks and neurite development (Kang et al., 2008). In this study, Rac1 is also identified as a palmitoyl protein. But how palmitoylation may regulate Rac1 functions remains unclear.  SYT-I, the synaptic vesicle protein, has also been shown to participate in the regulation of neurite outgrowth of chick dorsal root ganglion (DRG) neurons (Flannery et al., 2010). Several palmitoylated proteins are found to be associated with the cytoskeleton and affect membrane dynamics and filopodia outgrowth. Two examples are GAP-43 and paralemmin (Strittmatter et al., 1994; Gauthier-Campbell et al., 2004). Further evidence suggests that palmitoylation may facilitate proteins targeting to specific active sites of the PM to regulate cell morphology and membrane dynamics that contributes to neurite development (Gauthier-Campbell et al., 2004). 40  1.2.4.4 The c-jun N-terminal kinase family members as potential palmitoyl proteins  Given a broad range of substrates at various subcellular locations and multiple pathophysiological roles it plays, JNK as an important kinase must be regulated tightly to ensure its diverse functions. How is its activation status regulated by particular stimuli? How does JNK achieve specific substrate selectivity? How is JNK targeted to the specific subcellular loci where it functions? To achieve these specificities, several mechanisms are proposed to regulate JNK activity, substrate recognition, and trafficking. The most well studied mechanisms is the regulation by JNK-interacting or scaffolding proteins (Weston and Davis, 2007) (Figure 10). These proteins interact with JNK as well as its upstream activators, and modulate JNK activation. Different interacting proteins are specific to some particular upstream signals by selecting and recruiting corresponding upstream activators. Moreover, some scaffolding proteins can guide JNK to particular subcellular compartments via protein-protein interaction. For example, the JNK-interacting protein (JIP) has been shown to recruit JNK to synaptic vesicles and to drive it to axons (Horiuchi et al., 2005; Whitmarsh, 2006). Specific protein interaction can serve as a mechanism to achieve JNK activation and trafficking specificity. Although more than a dozen JNK scaffolding proteins have been discovered, such as POSH and β-arrestin2 (Weston and Davis, 2007), it is expected that there would be more such proteins and/or other mechanisms exist in mammalian cells to regulate JNK, when considering the splicing isoform complexity in this large JNK family. 41  At least 10 JNK spliced variants from three genes are identified in mammals (JNK1α1, JNK1α2, JNK1β1, JNK1β2, JNK2α1, JNK2α2, JNK2β1, JNK2β2, JNK3β1, and JNK3β2) (Bogoyevitch, 2006). Although they share similar kinase domains and activation TYP motifs, different physiological functions have been identified for the various JNK isoforms. For example, JNK2 shows 10-fold greater kinase activity against the substrate c-jun than does JNK1. In line with this, JNK1 is found predominantly in the cytoplasm of neurons (Coffey et al., 2002), whereas JNK2 and JNK3 are suggested to play critical roles in activating transcription factors in the nucleus (Lindwall et al., 2004). Moreover, differential roles of JNK isoforms in mediating neurite initiation and elongation during axonal regeneration have been observed (Bogoyevitch, 2006). However, little is known about how JNK isoforms are specifically regulated to participate in different biological processes. Some clues to their functionally specific roles come from the differences among their protein sequences. The most distinct features are found at the NH2-termini and COOH-termini. JNK3 has an extended NH2-terminus and this domain is found to specifically interact with β-arrestin2, a scaffolding protein regulating JNK3 activation (Guo and Whitmarsh, 2008). Some isoforms have extended COOH-termini (Bogoyevitch, 2006). However, the function of this domain is unknown. In the preliminary study that leads to one part of this thesis, we found that the COOH-termini possessed potential sites for palmitoylation. This raised the possibility that this post-translational modification may serve as a novel mechanism for JNK regulation.  42  1.2.5 Regulation of Protein Palmitoylation in Neurons One advantageous property of palmitoylation as a post-translational modification is its reversibility and high dynamics. This may be particularly important for regulating proteins related to synaptic functions. In response to synaptic transmission, post-synaptic proteins are mobilized in and out of synapses rapidly to ensure precise regulation of synaptic functions. It has been shown that neuronal activity dynamically modulates the palmitoylation status of a broad range of neuronal proteins, including receptors, scaffolding proteins, kinases, signalling molecules and others (El-Husseini and Bredt, 2002). For example, the palmitoylated pool of PSD-95 is reduced in response to synaptic stimulations with AMPA or NMDA, which induce internalization of AMPARs (El-Husseini et al., 2002). Similar results are observed for AMPAR’s (Yang et al., 2009). Stimulation with AMPA or NMDA which causes synaptic depression, leads to rapid enlargement of a palmitoylated pool of GluR1 (Yang et al., 2009). In addition to the palmitoylated pool, the turnover rate of palmitate on these proteins is accelerated by these treatments (Yang et al., 2009). In contrast, another group of presynaptic proteins including SYT-I, SNAP-25 is insensitive to fast synaptic activation. Instead, their palmitoylation status is enhanced in response to long-term synaptic suppression, which leads to accumulation of synaptic vesicles as a compensation (Kang et al., 2004). Neuronal stress is another prominent factor which affects protein palmitoylation. Palmitoylation of neuronal nitric oxide synthase (nNOS) is found to be sensitive to excitotoxicity. These results suggest that a dynamic regulation of protein palmitoylation is important for proper function of proteins and normal neuronal functions. 43  Despite the obvious involvement of PATs in regulating protein palmitoylation in response to stimuli, little is known about the mechanisms underlying the regulation of PAT activity. One possible mechanism is by protein-protein interactions. Htt is a substrate of zD17, and it interacts with the ankyrin domains of zD17 (Huang et al., 2009). It has been shown that the Htt-zD17 interaction promotes PSD-95 palmitoylation while reducing their interaction inhibits zD17-mediated palmitoylation of its substrates, including PSD-95 and SNAP-25 etc (Huang et al., 2009). However, more regulatory mechanisms underlying PAT activity regulation require further investigations.  1.3 Protein Palmitoylation and PATs in Human Diseases  1.3.1 Protein Palmitoylation in Pathological Processes Given the importance of palmitoylation in ensuring proper protein trafficking and function, incorrect palmitoylation of certain proteins may contribute to pathological processes in human diseases. HD results from the expanded member of poly-Q-repeats of the Htt protein. It has been found that the poly-Q-repeats Htt mutant shows reduced palmitoylation compared to the wildtype control (Singaraja et al., 2002). Reduction of its palmitoylation promotes Htt aggregation in neurons which is cytotoxic and induces neuronal cell death. Furthermore, reduced palmitoylation of Htt leads to decreased interaction with zD17 and reduces palmitoylation of 44  other zD17 substrates, including PSD-95 (Huang et al., 2009). This would result in loss of synaptic clustering of AMPARs and cause transmission deficiency. However, whether abnormal palmitoylation of other proteins also exists in vivo and contributes to other human diseases requires further studies.  Incorrect palmitoylation of pro-death proteins, on the other hand, may impair neuronal death signals. For example, palmitoylation is necessary for Fas ligand (Fas-L) mediated cell death . Loss of palmitoylation results in cell survival (Guardiola-Serrano et al., 2011). Similarly a requirement of palmitoylation is observed for death receptor 4 in TRAIL (TNF-related apoptosis-inducing ligand)-induced death signal transmission (Rossin et al., 2009).  1.3.2 The Pathological Roles of PATs in Human Diseases Several PATs have been linked with human diseases. In brain diseases, for example, mutation of zD8 is found to associate with schizophrenia (Faul et al., 2005), while zD17 is related to HD (Singaraja et al., 2002). However, a major unsolved issue concerns the mechanisms underlying these PAT-associated diseases. Given that the substrate selectivity is ambiguous for most, if not all, PATs in mammals, it is hard to delineate how PATs are involved in one particular disease. One example is HD. Loss of palmitoylation of Htt and suppression of zD17 expression by siRNA are found to be critical for this disease in animal models, and zD17 is thus suggested to be the PAT for Htt and palmitoylation to be responsible for causing the disease 45  (Singaraja et al., 2002). However, zD17 has many other substrates including PSD-95, SNAP-25, AMPARs, GAP-43 and others (Huang et al., 2004). These known and possibly other unknown proteins are directly and indirectly related to synaptic transmission and neuronal cell death processes. Therefore, much still remains unknown regarding the underlying mechanisms of each PAT’s role in human diseases. Moreover, it is possible that a given PAT’s contribution to human diseases not simply restricted to its role in palmitoylation. For example, zD17 has been found to activate JNK pathways that play a critical role in cell death and neurological diseases (Chapter 4) (Harada et al., 2003).  1.4 Stroke and Neuroprotection Therapy  1.4.1 General Background of Stroke Stroke is a leading cause of death and serious long-term disability worldwide (WHO, 2007). It can be divided into hemorrhagic stroke and ischemic stroke. Among all stroke cases, hemorrhagic stroke accounts for 15% of total stroke cases (Donnan et al., 2008). This type of stroke is caused by rupture of blood vessels and bleeding in the brain. On the other hand, in ischemic stroke the supply of blood and oxygen to a part of the brain is reduced, due to occlusion of a vessel and this eventually causes the death of brain tissue in that area and forms the infarct core (Donnan et al., 2008). This irreversibly-damaged core is surrounded by a dysfunctional but 46  potentially viable hypoperfused region, known as the penumbra (Lee et al., 1999). Within a few hours after ischemia, neurons in the penumbra are challenged by excitotoxic and inflammatory processes resulting in delayed death (Lee et al., 1999). The massive loss of brain tissue caused by delayed neuronal death leads to the severe cognitive and physical impairments observed in stroke patients (Koroshetz and Moskowitz, 1996; Donnan et al., 2008).  Despite of the social and economic problems of stroke, effective treatments for stroke are largely lacking. The only available approved treatment for stroke involves thrombolytic therapy with Tissue Plasminogen Activator (tPA). It targets clots in vessels and reopens blood flow (Koroshetz and Moskowitz, 1996; Donnan et al., 2008). However, several issues arise from this treatment. It only works for ischemic stroke. In some cases, tPA application in ischemic stroke patients results in conversion to the hemorrhagic stroke with potentially negative consequences. Second, the time window for tPA application is very critical. The maximum effective time window is 3-4 hours. However, many patients cannot reach hospital within 4 hours. Third, tPA application will introduce secondary reperfusion damage to neurons in the penumbra. To overcome these drawbacks, neuroprotection may be an alternative strategy for stroke therapy (Koroshetz and Moskowitz, 1996; De Keyser et al., 2005; Tymianski, 2010). However, it has been noticed that in ischemic stroke, neuroprotective reagents cannot reach injured brain tissues if blood vessels in the vicinity are blocked by clots. It may well be that the combination of neuroprotection therapy with tPA thrombolysis will be the best approach to maximize neuronal 47  survival following stroke.  1.4.2 Neuroprotection Therapy 1.4.2.1 Mechanisms underlying stroke-induced neuronal cell death  It is now clear that the massive tissue damage that occurs in stroke is progressive and potentially reversible (Lo, 2009). Many studies have let to great advances in delineating important biochemical processes underlying stroke-induced neuronal cell death (Bossy-Wetzel et al., 2004; Bredesen et al., 2006). By targeting the pathways leading to neuronal cell death, several neuroprotective strategies have been proposed and some of them have been shown to be effective in treating stroke in animal models (Tymianski, 2010). It has been observed that during ischemic stroke, the release of excitatory transmitters such as glutamate and aspartate is dramatically increased, which causes pathological hyperactivity in the brain, known as excitotoxicity. Excessive release of excitatory amino acids (EEA) activates glutamate receptors such as NMDA receptors, which leads to a rapid influx of calcium into neurons. Abnormally high concentrations of intracellular calcium are lethal to neurons because they activates several downstream protein kinases, proteases, phospholipases and other cell death pathways (Bossy-Wetzel et al., 2004; Bredesen et al., 2006; Tymianski, 2010). For example, Calpain1 is a calcium-activated protease that contributes to the degradation of crucial cytoskeletal and regulatory proteins. Another example is phospholipase A2, which activates a positive feedback 48  loop to stimulate further release of glutamate during ischemic stroke. Calcium has also been found to activate some isoforms of nNOS, and to generate NO which causes oxidative stress in ischemic stroke. Other calcium channels also play a role in mediating calcium influx or release from intracellular stores. For example, EEA-caused depolarization activates voltage-dependent calcium channels which have been shown to contribute to excitotoxicity in ischemic stroke, and also ryanodine receptors enhance release of calcium from intracellular stores (Lee et al., 1999; De Keyser et al., 2005; Bredesen et al., 2006).  In additional to the calcium-induced biochemical processes, other NMDA-mediated signaling pathways involved in neuronal cell death have been also discovered. Direct recruitment of signaling modules to NMDA receptors via the protein-protein interaction initiates downstream signals for execution. PSD-95 is one scaffolding protein linking nNOS to NMDA receptors during ischemic stroke and this recruitment produces NO resulting in neuronal cell death (Aarts et al., 2002). The death kinase DAPK is also an NMDA receptor-interacting protein that has recently been identified. The direct interaction between DAPK and the NMDA receptor promotes DAPK activity and initiates downstream neuronal cell death signals (Tu et al., 2010).  The stroke insult also leads to mitochondrial dysfunction and activation of mitochondria-dependent death pathways, which are believed to contribute to neuronal cell death. A reduction of the activity of mitochondrial proteins has been noticed during ischemic stroke, 49  which may result in energy failure and eventually neuronal death (Lee et al., 1999; Bossy-Wetzel et al., 2004). As the membrane potential drops due to energy failure, cytochrome c is released from mitochondria and initiates the caspase-dependent apoptosis cascade. This results in activation of caspase-3, a protease involved in the execution phase of programmed cell death. When activated, caspase-3 cleaves several protein targets, including cytoskeletal proteins, DNA repair proteins, and others. Proteolysis of ICAD by activated caspase-3 results in activation of the endonuclease CAD and cleavage of DNA. Some signaling pathways leading to cytochrome c release and caspase-3 activation have been identified, including signaling by free radicals like NO, overload of intracellular calcium, activation of death receptors and others. In response to these pathological stimuli, the bcl-2 protein family members play a key role in controlling the release of cytochrome c from mitochondria. While bcl-2 and bcl-x-long function as anti-apoptotic factors to inhibit the release of cytochrome c, other family members such as bad, bax, and bcl-x-short promote cytochrome c release, and enhance apoptosis (Lee et al., 1999; De Keyser et al., 2005). There are more than 20 members in this bcl family. It is the balance of activity of anti-apoptotic and pro-apoptotic members that controls the release of cytochrome c from mitochondria and the subsequence activation of caspase-3 (Lee et al., 1999; Bossy-Wetzel et al., 2004; Bredesen et al., 2006).  Increasing evidence indicates that stroke-induced neuronal death develops slowly, especially in the penumbra (Lee et al., 1999; Lo, 2009). Production of new genes and proteins 50  has been found to contribute to the progression of stroke pathology. One example is the generation of inflammatory factors which causes inflammation and increases neuronal death (Barone and Feuerstein, 1999). An increase of the transcription and expression of TNF-α and IL-1β has been observed in cerebral ischemia. In line with this observation, TNF receptor production is also increased. Another example is the increase of the expression of the death receptor Fas and Fas ligand during ischemic stroke, which can also activate the caspase cascade and other neuronal death pathways (Barone and Feuerstein, 1999; Bossy-Wetzel et al., 2004).  1.4.2.2 Role of JNK in neuronal cell death  Acting as a mediator of extracellular and intracellular stress signals, JNK is broadly involved in excitotoxicity, inflammation, and neuronal cell death in ischemic stroke as well as other neurological diseases (Resnick and Fennell, 2004). It has been observed that JNK activity is strongly enhanced during ischemic stroke and JNK initiates both immediate and late-phase cell death events via phosphorylation and activation of substrates in both the nucleus and cytosol (Bredesen et al., 2006; Centeno et al., 2007; Weston and Davis, 2007). One of the JNK substrates, the transcriptional factor c-jun, when activated by JNK, promotes the transcription of several pro-death gene products, including TNF-α, Bim and cyclooxygenase 2 (COX2) (Bogoyevitch and Kobe, 2006). Other JNK substrates include the bcl family member, bad. Phosphorylation of bad by JNK triggers its translocation from cytosol to mitochondria, which promotes cytochrome  51  c release and activates the caspase cascade (Bogoyevitch and Kobe, 2006). Further evidence shows that JNK is able to be translocated to mitochondria and to phosphorylate bcl-xl to promote cytochrome c release, suggesting a key role of JNK in regulating neuronal cell death.  Compared to the ubiquitously expressed JNK1 and JNK2, JNK3 is predominantly expressed in the nervous system (Weston and Davis, 2007). In line with this tissue specificity, JNK3 has been indeed found to play a key role in mediating neuronal cell death not only in stroke, but also other neurodegenerative diseases including Alzheimer’s disease and Parkinson’s disease (Kuan and Burke, 2005; Bogoyevitch, 2006). JNK3-/- mice display increased resistance to ischemic insult, as well as MTPT toxicity. Although the JNK activators MKK4 and MKK7 are all involved in activating JNK in response to various stresses, TNF and excitotoxicity-induced JNK activation is particularly mediated by MKK7, suggesting that specialized pathways may exist for activating JNK and downstream signals in ischemic stroke (Centeno et al., 2007).  As JNK activation is critical for neuronal cell death, several inhibitors have been developed to target JNK or related cell death pathways for treating stroke (Kuan and Burke, 2005). CEP-1347 is an ATP-competitive inhibitor of the family of MKKK, MLKs. Blocking MLKs with CEP-1347 effectively inhibits JNK activation under pathological conditions and protects neurons against MTPT-induced cell death. Direct JNK inhibitors SP600125 and AS601245 have also been used in protecting neurons against neuronal stresses. Application of these drugs blocks the 52  JNK activation and downstream death signals in animal models of ischemic stroke (Kuan and Burke, 2005; Borsello and Forloni, 2007). Recently, a peptide inhibitor targeting the activation of the JNK substrate c-jun has been developed to block the interaction between JNK and c-jun (Borsello et al., 2003). This peptide JNKI is derived from the scaffolding protein JIP1. This peptide has been found to be effective in protecting neurons against ischemic stroke as well as in an animal model of Alzheimer’s disease (Borsello and Forloni, 2007).  1.4.2.3 Therapeutic targets for neuroprotection therapy  Based on the knowledge of cell death pathways acquired in past two decades, significant advances have been made in delineating cellular and molecular mechanisms of ischemic brain injury. By targeting crucial signalling cascades underlying excitotoxicity, inflammation and neuronal cell death, several strategies have shown promising neuroprotective effects in preclinical studies. These include the application of glutamate receptor antagonists, ion channel blockers, kinase inhibitors, free radical scavengers, and anti-inflammatory compounds (Koroshetz and Moskowitz, 1996; Fisher and Ratan, 2003; De Keyser et al., 2005). However, clinical success with these neuroprotective reagents has still not been achieved. In addition to the issues of the clinical trial design, our limited knowledge of signaling cascades underlying neuronal cell death hampers the development of drugs with high efficacy and low side-effects.  To overcome the side effects of general inhibition of targeted proteins, specially designed 53  peptides as an alternative novel strategy are now used to achieve the blockade of selective protein-protein interactions. In the case of NMDA receptor inhibition, a peptide has been designed to block the NMDA receptor-PSD-95 interaction, while another peptide has been used to block NMDA receptor-DAPK interaction (Aarts et al., 2002; Tu et al., 2010). Both peptides are effective in reducing brain damage in ischemic stroke.  1.4.2.4 The challenge of targeting JNK and potential solutions  Inhibitors targeting JNK have shown promising neuroprotection in neurological diseases, despite concerns of side-effects raised in clinical trials (Borsello et al., 2003; Kuan and Burke, 2005; Bogoyevitch, 2006; Borsello and Forloni, 2007). Studies on the three JNK isoforms have pinpointed different roles of JNK1, JNK2 and JNK3 in neuronal functions and neurological diseases. While JNK2-/-, JNK3-/- or JNK2-/- JNK3-/- mice show reduced vulnerability in models of Parkinson’s disease and ischemic stroke, JNK1-/- animals show little protective effect (Hunot et al., 2004). Moreover, the loss of JNK1 leads to disorganization of neuronal microtubules and neurodegeneration (Chang et al., 2003). These results strongly suggest that selective inhibition of stress-responsive JNK isoforms (primarily JNK2/3) would be beneficial for therapeutic targeting of JNK (Coffey et al., 2002; Bogoyevitch, 2006). Although the advantage of isoform-selective JNK inhibition has long been appreciated, the application of small molecule inhibitors with this feature in treating ischemic stroke has not been reported. The development of small molecules  54  with isoform selectivity has proven challenging since JNK isoforms are highly conserved at the ATP-binding sites. An alternative strategy is to take the advantage of JNK interacting proteins, some of which have been shown to have selective binding preferences to JNK isoforms. Using these proteins as targets, however, no JNK isoform-selective inhibition has been successfully tested as a therapeutic strategy in neurological diseases.  1.5 Thesis Hypotheses and Objectives  1.5.1 AMAPR Palmitoylation may Have Subunit-Selectivity Subunit-specific regulation of AMPAR trafficking and function has been observed. Several mechanisms underlying the subunit-specific AMPAR regulation have also been proposed and examined, including subunit-specific binding partners and subunit-unique phosphorylation and ubiquitination (Malenka, 2003). As AMPAR subunits show distinct regulatory and functional characteristics, it is possible that palmitoylation of AMPAR subunits may have different properties and functions.  Given the ambiguous substrate selectivity of PATs, AMPAR subunits may be catalyzed by other PATs in addition to the known zD3. The redundancy of PATs in palmitoylating a particular substrate has been observed in many other proteins, e.g PSD-95 (Fukata et al., 2004), δ opioid  55  receptor (Petaja-Repo et al., 2006), and GABA-A receptor (Fang et al., 2006). It is possible that some PATs might show preference for GluR1 and GluR2, two prominent subunits of the AMPAR. GluR1 has been shown to mature rapidly and transport from ER to Golgi soon after synthesis, whereas GluR2 stays in ER and is only slowly transported to the Golgi apparatus. As many mammalian PATs are resident proteins in ER and Golgi, it is possible that GluR1 and GluR2 are processed by different PATs and have distinct palmitoylation mechanisms. I thus examine the palmitoylation mechanisms of GluR1 and GluR2, and study the potential functional differences of palmitoylation in regulating GluR1 and GluR2.  1.5.2 Specific JNK Isoforms may be Regulated by Palmitoylation Given the strong evidence of JNK isoforms with different functions, understanding how JNK isoforms are specifically regulated is essential for elucidating JNK isoform-specific roles in multiple biological processes. Isoform-selective regulation has been observed with some JNK scaffolding proteins that regulate JNK activity and trafficking (Lee et al., 1999; Coffey et al., 2002). Another potential mechanism may be post-translational modification. In addition to phosphorylation, that we know can activate JNKs, little is known about whether JNKs are subject to other modifications.  Protein palmitoylation is one such candidate modification. Several kinases including PKC, casine kinase, Ras, and Fyn have been reported to be palmitoyl proteins. Protein palmitoylation 56  is involved in regulating different aspects including kinase activity, trafficking and protein interaction. Moreover, proteins with little sequence variance may differ in palmitoylation sensitivity and specificity, such as glutamate receptor interacting proteins 1 isoform a and b (GRIP1a/b) (Yamazaki et al., 2001), providing a potential way to achieve isoform-specific regulation.  Sequence alignment of JNK isoforms indicates that differences exist at NH2- and COOH-termini, potential regions for isoform-specific regulation. Indeed, the NH2- terminus of JNK3 has been shown to selectively interact with the scaffolding protein β-arrestin2 (Guo and Whitmarsh, 2008). The Cys residues that are likely targets for palmitoylation are located at COOH-termini of the p54 isoforms and are not found on the p46 isoforms. Using CSS2.0, I have identified JNK3 as potential palmitoylated proteins. I thus hypothesize that selective JNK isoforms are palmitoylated and palmitoylation contributes to isoform-selective regulation of JNK in neurons.  1.5.3 How zD17 Is Involved in Regulating JNK Activation Through its regulation of Htt palmitoylation and aggregation, zD17 has been implicated in chronic neurodegeneration in Huntington’s disease (HD) (Yanai et al., 2006). However, little is known about the role of zD17 in acute brain injury such as ischemic stroke, which involves neuronal cell death independent of Htt aggregation. In our pilot studies leading to this report, I 57  have found that zD17-induced JNK activation is independent of its PAT activity. Since JNK activation has been linked to cell death (Resnick and Fennell, 2004; Centeno et al., 2007; Weston and Davis, 2007), I hypothesize that zD17 may contribute to acute neuronal cell death in ischemic stroke via its palmitoylation independent activity involving JNK pathways.  58  Chapter 2. Palmitoylation Contributes to Subunit-Selective Regulation of AMPA Receptors in Synaptic Plasticity  A version of this chapter has been published Guang Yang, Wei Xiong, Luba Kojic, Max S. Cynader. 2009. Subunit-selective palmitoylation regulates the intracellular trafficking of AMPA receptor. European Journal of Neuroscience, 30 (1): 35-46  59  2.1 Introduction The AMPAR is critical for fast synaptic transmission in the mammalian brain. Various combinations of the four AMPAR subunits (GluR1–GluR4) account for the different properties of the heterotetrameric receptors (Ozawa et al., 1998). Although AMPAR heteromers are assembled soon after translation in the endoplasmic reticulum (ER), each subunit seems to be processed in parallel by different machinery, which independently contributes to AMPAR synaptic function (Greger et al., 2006). AMPAR subunits have been found to be palmitoylated at two cysteine sites: one at TMD2, the other at the C-terminus (Hayashi et al., 2005). This reversible lipid modification highlights new molecular possibilities for subunit-dependent AMPAR regulation. Indeed, palmitoylation of GluR2 has been shown to enhance AMPAR retention in the Golgi apparatus, whereas palmitoylation of GluR1 facilitates AMPAR internalization possibly by promoting dissociation of surface AMPAR from 4.1N, an actin-associated protein (Hayashi et al., 2005; Lin et al., 2009).  Similar to the situation with post-synaptic density 95 (PSD-95), AMPAR palmitoylation could be enhanced by GODZ (Golgi apparatus-specific protein with a DHHC zinc finger domain), a member of the PAT family (Fukata et al., 2004; Hayashi et al., 2005). Moreover, PSD-95 is also the substrate for many other PATs in neurons (Fukata et al., 2004). The redundancy of PATs in palmitoylating one substrate is also applicable to other proteins, such as  60  the δ opioid receptor, and GABAA receptor. It is thus possible that AMPAR could also be targeted to other PATs for palmitoylation and some PATs might show preference for particular subunits. Subunit-specific mechanisms of palmitoylation might thus be expected for AMPAR, in accordance with the different properties of AMPAR subunits.  In this chapter, we investigated potential differences in palmitoylation mechanisms of the AMPAR subunit GluR1 and GluR2, and explored the functional consequence of palmitoylation on these subunits.  2.2 Results  2.2.1 GluR1 Palmitoylation Requires Anterograde Transport Protein palmitoylation is a post-translational modification that can take place at multiple sites along the secretion pathway and on the cell surface (Bijlmakers and Marsh, 2003; Greaves and Chamberlain, 2007). To examine whether GluR1 palmitoylation requires de novo protein synthesis in the ER, neurons were pretreated with the protein synthesis inhibitor cycloheximide (CHX) for 1 h before incubation with 3H-palmitate for an additional 3 h. We found that 3  H-palmitate incorporation into GluR1 was attenuated to 50.1±7.9% of the untreated control  level upon CHX treatment (p<0.01, n=4; Figure 11A, B). Given that a large portion of newly  61  synthesized GluR1 exits the ER and is transported to the PM within several hours (Greger et al., 2002), CHX-induced reduction of GluR1 palmitoylation suggests that newly synthesized GluR1 might be available to be palmitoylated locally in the ER or other compartments along the secretion pathway.  Figure 11. Anterograde transport in the early secretory pathway is important for GluR1 palmitoylation. Rat neurons cultured for 14–17 days in vitro were pretreated with CHX (25 μg/mL), BFA (5 μg/mL) or NOC (20 μg/mL) as indicated for 1 h before labelling with 3H-palmitate for 3 h in the presence of the reagents. 3 H-palmitate intensity (3H-palm) was quantified and normalized to GluR1 input shown as immunoblotting (IB). GluR1 palmitoylation levels from treated groups were compared and normalized to untreated controls (A, B). (C,D) Incubating with NOC did not prevent the BFA-induced reduction of GluR1 palmitoylation. (E and F) 3 H-palmitate incorporation with GluR1 was sensitive to a combination of CHX and BFA. Mean ± SEM from three to four separate data sets are shown.  We thus further investigated the cellular compartments in which palmitoylation takes place, by testing the effect of brefeldin A (BFA) on GluR1 3H-palmitate incorporation. BFA blocks protein transport from the ER to the PM in mammalian cells by disrupting the structures and function of the Golgi apparatus and redirecting the traffic back to the ER (Lippincott-Schwartz et 62  al., 1989; Orci et al., 1993). As shown in Figure 11A and B, 1 h pre-incubation of BFA followed by 3 h 3H-palmitate labelling diminished GluR1 palmitoylation to 73.0±8.1% (p<0.01, n=4). The BFA induced redistribution of Golgi components to the ER could be prevented by the addition of nocodazole (NOC), a microtubule depolymerizer which has been used to block vesicle transport within the early secretion pathway (Lippincott-Schwartz et al., 1990). NOC, in combination with BFA, could thus distinguish the disruption of anterograde transport from the inhibition by retrograde transported contents from the Golgi. Whereas GluR1 palmitoylation was unaffected by NOC alone (84.1±8.8% of control, p=0.14, n=4; Figure 11A,B), BFA and NOC co-application attenuated GluR1 palmitoylation to a degree similar to BFA treatment alone (71.7±8.3% of control, p<0.05, n=3; Figure 11C,D). These results indicate that anterograde transport of GluR1 from the ER to the Golgi apparatus is important for GluR1 palmitoylation.  Since BFA caused less reduction than did CHX, we wondered whether BFA and CHX affected the same fraction of GluR1 subunits. To assess this, neurons were pretreated with a combination of CHX and BFA. CHX/BFA treatment reduced 3H-palmitate incorporation into GluR1 to 66.1±10% (p<0.05, n=3; Figure 11E, F). This result suggests that the BFA-sensitive fraction of GluR1 might be partially built up by immature, newly synthesized GluR1s resident in the ER, since the effect of CHX and BFA is not additive.  63  2.2.2 GluR2s Are Palmitoylated in the ER as Immature Receptors Figure 12. GluR2 subunits are primarily palmitoylated in the ER. (A and B) GluR2 palmitoylation was greatly reduced by incubating with CHX or NOC. In contrast to GluR1, BFA promoted GluR2 palmitoylation, which was insensitive to the microtubule destructor NOC (C, D). Mean ± SEM from three to four separate data sets are shown. *P < 0.05, **P < 0.01. (E) The mature and immature pools of GluR2s were separated by EndoH treatment. Mature GluR2s as indicated (M) were of a similar size to untreated controls. Immature GluR2s (IM) shifted more rapidly to a lower level as when treated with PGNF. (F) [3H]-palmitate-labelled GluR2s were immunoprecipitated and treated with EndoH and PNGF. Only the immature form of GluR2 can be detected with the 3H signal. Representative figures were from three independent experiments. (G) GluR1-associated GluR2s were depleted by co-immunoprecipitation with GluR1 antibody. Both GluR1 and GluR2 were detected in the precipitates (lane 1). The supernatant was further co-immunoprecipitated with GluR1 antibody to confirm that all the GluR1s and GluR1-associated GluR2s were completely collected in the first round (lane 2). The rest of the GluR2s (‘Free’ GluR2) were immunoprecipitated by GluR2 antibody from the supernatant with 0.1% SDS (lane 3). (H) Total and Free GluR2 were subject to the biotin-BMCC reaction to test for GluR2 palmitoylation. With equalized protein input, Free GluR2 showed a significant higher palmitoylation level (192 ± 20.5%, P < 0.01, n = 4) compared with total GluR2.  We next investigated the cellular mechanisms of GluR2 palmitoylation. In contrast to GluR1 subunits which are mainly confined to cell surface, a large proportion of GluR2 subunits stably reside in an intracellular pool of ER (Greger et al., 2002). Therefore, we first asked 64  whether the GluR2 pool available for palmitoylation is made up of newly synthesized receptors. As in the case of GluR1, inhibiting protein synthesis by CHX reduced the GluR2 incorporated 3  H-palmitate signal to 37.7±2.5% of untreated control (p<0.01, n=4; Figure 12A, B). In  comparison with GluR1, GluR2 palmitoylation is more sensitive to CHX, suggesting that newly synthesized GluR2 in the ER is the major target for palmitoylation. To understand whether GluR2 palmitoylation occurs locally in the ER or requires anterograde transport from the ER to the Golgi as does GluR1, we further examined the effects of BFA on GluR2 palmitoylation. To our surprise, destruction of the Golgi apparatus by BFA significantly increased 3H-palmitate incorporation on GluR2 (171.0±10.2% of control, p<0.01, n=4; Figure 12A, B). BFA also enhanced palmitoylation of heterologously expressed GluR2 in HEK293 cells (Figure 13).  Figure 13. GluR2 palmitoylation is enhanced by BFA treatment in exogenous cells. A) Representative figure showing a strong increase of GluR2 palmitoylation upon BFA treatment. HEK293 cells were transfected with GluR2 WT (16 hr) and incubated in the presence or absence of BFA (5 µg/ml) for 5 hr. Cell lysates were precleaned with protein-A sepharose 1 hr at 4 °C, and immunoprecipitated. Palmitoylation on GluR2 was detected using Btn-BMCC labelling via protocol 2. B) Quantitative data from experiments shown in A) after normalizing the palmitoylation signal (avidin-HRP) to GluR2 input. BFA significantly increased GluR2 palmitoylation level to 154.8±17.1% of untreated controls. Mean ± SEM% from five separate sets are shown. * P<0.05.  It has been reported that GODZ shows PAT activity for AMPARs (Hayashi et al., 2005). It is thus  65  possible that the increase of GluR2 palmitoylation is caused by increased accessibility of GluR2 to Golgi-located PATs which are not always available for GluR2 but redistributed to the ER upon BFA treatment (Lippincott-Schwartz et al., 1989; Lippincott-Schwartz et al., 1990). To address this issue, NOC was used to prevent BFA-induced retrograde transport of the GODZ and other Golgi enzymes. However, NOC failed to attenuate BFA-enhanced GluR2 palmitoylation (187.7±27.4% of control, p<0.05, n=3; Figure 12C, D), suggesting that other processes, rather than retrograde transport of Golgi-resident enzymes, are responsible for BFA-induced palmitoylation enhancement. Intriguingly, NOC treatment alone was found to reduce 3  H-palmitate incorporation on GluR2 to 43.4±7.4% of untreated control levels (p<0.01, n=4;  Figure 12A, B). These data not only suggest that an intact microtubule network is essential for palmitoylation of GluR2, but also suggest that blocking protein anterograde transport by BFA could compensate the loss of palmitoylation caused by inhibiting protein retrograde transport. Given that the ER residency time for newly synthesized GluR2 is relatively longer than that for GluR1 (Greger et al., 2002), the transport of GluR2 itself seems not to be crucial for palmitoylation after its synthesis. To test whether newly palmitoylated GluR2 resides in the ER as an immature subunit or is rapidly transported to late stages of the secretion pathway, we took the advantage of the processing of AMPAR by N-glycosylation. Membrane proteins undergo a high-mannose type N-linked glycosylation in the early secretion pathway, and some high-mannose oligosaccharides are then further modified to form complex oligosaccharides in the Golgi apparatus (Greger et al., 2002). Therefore, the level of high-mannose sugar can 66  indicate the maturation status of GluR2’s and their subcellular localization. Total AMPAR subunits were immunoprecipitated from lysates of neurons labelled with 3H-palmitate for 3 h and the precipitates were treated with endoglycosidase H (Endo H) or PNGaseF, enzymes catalyzing removal of only the high-mannose type or all types of N-linked glycosylation, respectively. Two bands were found for EndoH treated GluR2 (Figure 12E). Using autoradiography, only one band at the size that corresponded to immature GluR2 after the EndoH reaction was observed, indicating that the palmitoylation on GluR2 subunits primarily occurs in the ER (Figure 12F).  Figure 14. GluR2 subunits are transported to the Golgi in the presence of palmitate attached in the ER. GluR2 subunits were immunoprecipitated from samples of 14-d-old rat cortical neurons labelled with [3H]-palmitate for 24 h, and were then subject to the EndoH or PNGF reaction. The 3H signal was detected from both mature and immature forms of GluR2 subunits which were processed by EndoH. PNGF-deglycosylated GluR2 subunits were detected at the size as immature subunits.  Twenty-four hours of labelling showed a weak band at the mature GluR2 size, together with a strong signal at the immature size (Figure 14), suggesting that palmitoylated GluR2 is slowly transported to the Golgi from the ER, and possibly followed by a depalmitoylation process in the Golgi (Hayashi et al., 2005). In conclusion, the above data collectively demonstrate that neuronal mechanisms for GluR1 and GluR2 palmitoylation are quite distinct.  67  Figure 15. Preventing palmitoylation does not enhance GluR2 aggregation in COS7 cells and cultured neurons. A) COS7 cells were transfected with GluR2 (green) constructs as indicated. After 16 hr, cells were fixed, permeabilized and stained with ER marker calnexin (red). Similar subcellular distribution was observed among GluR2 WT and palmitoylation-deficient mutants. B) Effect of the sulfhydryl-specific cross-linker BMH on cell lysates from HEK293 cells expressing GluR2 WT and palmitoylation-deficient mutants. After cell lysis, lysates were incubated in the dark at room temperature for 45 min in the presence or absence of BMH (2.5 mM). Excess BMH was quenched by adding an equal volume of 2x reducing sample buffer (with 20mM DTT) and incubating for a further 10 min. Upon BMH crosslinking, bands representing GluR2 dimer (arrow), as well as GluR2 monomer (arrowhead), could be detected for all GluR2 constructs. Representative figure were chosen from two separate experiments. C) The percentage of crosslinked GluR2 dimmer is not significantly different between GluR2 WT and mutants (WT, 26.3±0.4%; C610S, 23.6±2.0%; C836S, 22.4±2.4%; C610,836S, 23.4±1.5%; n=2). D) and E) 2-BrPA does not promote GluR2 aggregation in neurons. 14-d-old cultured cortical neurons were 68  treated with 2-BrPA for 24 h. Cell lysates were then crosslinked and analyzed by SDS-PAGE gel. GluR2 dimmer could be detected at size ~200kDa upon BMH crosslinking. However, the ratio of crosslinked and uncrosslinked GluR2 indicated no significant change with 2-BrPA treatment (control, 54.8±2.3%; 2-BrPA, 53.8±2.1%; P=0.77; n=5). Mean ± SEM% are shown.  2.2.3 Blocking Palmitoylation Results in Reduction of GluR2  Figure 16. Blocking palmitoylation destabilizes the mature GluR2 pool. (A and B) Palmitoylation was broadly inhibited by 2-BrPA. 2-BrPA caused no significant change of GluR1and NR1 levels whereas it reduced GluR2 levels. (C) Effects of 2-BrPA on mature and immature GluR2. The ratio of mature/immature GluR2 decreased after 2-BrPA treatment, as shown in D. (E) Statistical analysis shows that 2-BrPA selectively reduced the mature GluR2 pool, after normalization to β-actin as shown in C. (F and G) Incubation with CHX (25 μg/mL, 24 h) after 1 h pretreatment was used to block de novo protein synthesis, which prevented the GluR2 decrease induced by 2-BrPA. (H and I). Mean ± SEM from three to four separate data sets are shown. *P < 0.05, **P < 0.01.  As the early secretion pathway is critical for GluR2 palmitoylation, we were interested to 69  determine whether palmitoylation contributes to ER-related GluR2 regulation. The ER is an important quality control site within the cell, where proteins are monitored for proper folding and oligomeric assembly (Hebert and Molinari, 2007). Consistent with the observations that palmitoylation deficient GluR2 mutants did not show aggregation in heterologous cell lines, blocking palmitoylation by 2-BrPA did not promote GluR2 aggregation in neurons (Figure 15). Instead, overall levels of GluR2 were reduced to 77.7±1.5% (p<0.01, n=4) compared with control following 2-BrPA treatment (Figure 16A,B). In contrast, blocking palmitoylation had no effect on overall GluR1 (101.2±5.9%, p=0.16, n=4) and NMDA receptor subunit 1(NR1) numbers (94.1±4.5%, p=0.46, n=4) (Figure 16A, B). Since the reduction is specific for GluR2, we further asked which pool of GluR2 is affected by blocking palmitoylation.  EndoH assays showed that only the mature, post-ER, GluR2 levels decreased after 2-BrPA treatment, leaving the immature, ER-resident, GluR2 population intact (mature, 72.3±3.6% of control, p<0.05; immature, 96.7±2.7% of control, p=0.35, n=3; Figure 16C,E). The percentage of mature versus immature GluR2 receptors was correspondingly decreased in response to the treatment (Control, 89.9±2.7%; 2-BrPA, 68.5±1.6%, p<0.01, n=3) (Figure 16D,E). Given that GluR2 is primarily palmitoylated in the ER soon after it synthesized (Figure 12A,B), the GluR2 stability may have already be determined in the ER by its palmitoylation status. To test this hypothesis, we examined the effect of protein synthesis inhibition on the reduction of GluR2 level induced by 2-BrPA. As expected, 2-BrPA could not induce further GluR2 decreases after 70  we pre-incubated neurons with CHX (CHX only, 59.1±3.5% of control; CHX and 2-BrPA, 58.8± 2.7% of control; p=0.75, n=3) (Figure 16F,G). However, we noticed that preventing protein synthesis caused more GluR2 reduction than did 2-BrPA treatment, suggesting the possibility of the existence of a pool of newly synthesized GluR2 which is not normally palmitoylated and therefore not affected by blocking palmitoylation.  2.2.4 Palmitoylation at TMD2 of GluR2 Regulates GluR2 Stability To further determine which palmitoylation site(s) is/are involved in controlling GluR2 stability, the inhibition of protein synthesis was used to determine the turnover rate of palmitoylation-deficient mutants overexpressed in HEK293 cells.  Figure 17. Blocking palmitoylation enhances reduction of GluR2 WT in HEK293 cells. Cells expressing GluR2 WT were treated with CHX in the presence or absence of 2-BrPA. After 24 h of incubation, cells were harvested and GluR2 level was tested by immunoblotting (A). In the presence of 2-BrPA, GluR2 WT level dropped from 84.02±12.8% to 46.89±5.0% of 0 h control (B). The level of GluR2 in the presence of 2-BrPA was similar to that of C610S (30.03±16.9%, P=0.32). Mean ± SEM% from three separate sets are shown. * P<0.05.  As shown in Figure 16H, the levels of GluR2 C610S (single mutant) and of C610,836S (double 71  mutant), but not of the C836S (single mutant), dramatically decreased when protein synthesis was arrested by CHX for 8 h (WT, 83.8±5.7% of the 0 h control; C610S, 41.5±4.1%, p<0.01; C836S, 81.4±7.7%, p=0.8; C610,836S, 51.5±6.6%, p<0.01, n=5) or 24 h (WT, 66.1±7.2% of the 0 h control; C610S, 28.9±5.5%, p<0.01; C836S, 61.2±6.3%, p=0.63; C610,836S, 44.7±2.6%, p<0.05, n=5) compared with the control GluR2 WT. Furthermore, incubating the cells with 2-BrPA promoted the reduction of WT GluR2 to the same level as is observed with C610S mutant (Figure 17). Taken together, the rapid turnover rate of the C610S and of the C610,836S mutants suggests that palmitoylation on cysteine 610 is critical for the stability of GluR2.  2.2.5 Blocking Palmitoylation Leads to Lysosomal Degradation of GluR2 2-BrPA incubation did not cause a decrease of GluR2 mRNA level (Figure 18). We wonder whether degradation mechanisms may be responsible for 2-BrPA-induced decrease of GluR2.  Figure 18. Blocking palmitoylation by 2-BrPA does not affect the mRNA level of GluR2. Preventing palmitoylation by 2-BrPA did not change GluR2 mRNA level (untreated GluR2, 99.8±2.2%; 2-BrPA, 101.8±2.8%; P=0.6). Mean ± SEM% from three separate sets are shown.  We thus examined the effect of proteasome and lysosomal inhibitors on 2-BrPA treated neurons. Incubation with 2-BrPA and the proteasome inhibitors clasto-lactacystin-β-lactone (lactone) (78.5±3.2% of control, p<0.01, n=6) or MG-132 (73.4±3.1% of control, p<0.01, n=6) (Figure 72  19A,B) did not block the degradation of GluR2. However, co-applying the lysosomal inhibitors, leupeptin (88.8±7.1% of control, p=1.6, n=6) or chloroquin (99.3±4% of control, p=0.86, n=6) prevented the reduction of GluR2. The proteosome inhibitor N-acetyl-leucyl-leucyl-norleucinal (ALLN) also partially rescued the decrease of GluR2 (90.0±6.4%, p=0.16, n=6) possibly because of its nonspecific inhibition of lysosomal function (Fuertes et al., 2003).  Figure 19. Reduced palmitoylation induces lysosome-dependent degradation of GluR2. (A and B) Lysosome inhibitors but not proteasome inhibitors rescued GluR2 from degradation induced by 2-BrPA. Lactone (1 μm), MG132 (5 μm) and ALLN (25 μm) were used as proteasome inhibitors while chloroquine (100 μm) and leupeptin (50 μm) were employed to inhibit lysosome functions. Incubation (24 h) with chloroquine and leupeptin rescued GluR2 degradation induced by 2-BrPA. ALLN partially blocked the reduction possibly due to its side-effect of inhibiting enzymes in the lysosome. (C and D) The effect of PMA on total GluR2 level. PMA (1 μm) was used to enhance cargo budding from the Golgi and total protein levels of GluR2 were measured at the indicated times. No significant reduction of GluR2 was observed within 24 h. (E and F) PMA applied with 2-BrPA to neurons for 24 h could not prevent the 2-BrPA-induced GluR2 degradation. Mean ± SEM from three to six separate data sets are shown. *P < 0.05, **P < 0.01.  73  It has been suggested in earlier studies that GluR2 export from Golgi apparatus requires depalmitoylation of Cys610 on TMD2. Therefore, the fast degradation of unpalmitoylated GluR2 via the lysosome may be due to either enhanced AMPAR cycling rate by promoting receptor budding from the Golgi or by directly targeting those AMPARs to the lysosome. To address this question, we enhanced AMPAR budding from the Golgi apparatus by treating neurons with phorbol 12-myristate 13-acetate (PMA), which that has been used to stimulate protein transport from the Golgi to the PM (Fernandez-Ulibarri et al., 2007). In the presence of PMA (3 h, 98.1±4.0% of the 0 h control, p=0.78; 8 h, 92.8±4.0%, p=0.08; 24 h, 95.2±4.0%, p=0.58; n=3; Figure 19C,D), GluR2 expression level was not reduced significantly. However, incubating neurons with 2-BrPA in the presence of PMA for 24 h did not prevent the degradation of GluR2 (82.0±6.0% of untreated control, p<0.05, n=5), suggesting that PMA itself does not affect lysosomal degradation of GluR2 (Figure 19E,F).  2.2.6 Palmitoylation of AMPARs Is Regulated by Neuronal Activity Neuronal activity is a major regulator of AMPAR subcellular distribution, concentration and properties. We thus asked whether palmitoylation of AMPARs is regulated by neuronal activity. TTX was employed to block action potential generation. CNQX and APV, which block AMPAR and NMDAR, respectively, were used together to prevent postsynaptic miniature EPSC’s. It is known that long-term (48 h) global activity blockade by TTX or APV/CNQX gradually promotes a homeostatic increase in glutamate receptor currents (O'Brien et al., 1998). Our 74  observed a marked increase of total and surface GluR1, as well as a slight increase of GluR2 (Figure 20). To compliment the metabolic labelling, we used a Btn-BMCC labelling method to examine the total palmitoylated pool of AMPAR subunits.  Figure 20. Detection of total and surface GluR1 and GluR2 after blocking neuronal activity. A)-D) Total protein levels of GluR1 and GluR2 subunits from 14-d-old rat cortical neuron cultures were detected after treatments with TTX or APV/CNQX for 48 h. Protein were extracted by directly adding 1x protein loading buffer and boiled at 100oC for 8 min, and then analyzed with 8% SDS-PAGE gel. A) and B) Chronic TTX and APV/CNQX application dramatically increased total GluR1 level to 162.8±21.6% and 187.8±31.2% of untreated controls, respectively. C) and D) Total GluR2 level also significantly increased after these treatments (TTX, 123.8±3.5% of control; APV/CNQX, 118.4±4.2% of control). E)-H) Surface GluR1 and GluR2 subunits were detected by biotinylation assays as previously reported. Surface proteins were biotinylated, isolated with streptavidin-beads, and analyzed with SDS-PAGE gels. E) and F), The number of surface GluR1’s increased markedly after blocking neuronal activity by TTX and APV/CNQX. After normalization to the surface protein marker LRP, surface GluR1’s were enhanced to 135.3±1.1% of the untreated controls by TTX. Similarly, APV/CNQX increased the number by 143.0±6.7%. The effects of blocking neuronal activity on surface GluR2 are shown in panel G) and H). In contrast to GluR1, no enhancement of surface GluR2 was observed (TTX, 94.3±7.9%, P=0.54). Although APV/CNQX slightly decreased the surface GluR2 level to 89.9±10.8% of untreated control, no significant difference was detected (P=0.45). * P<0.05, ** P<0.01. 75  Figure 21. AMPAR palmitoylation is regulated by neuronal activity. (A and B) GluR2 palmitoylation was increased upon blocking global neuronal activity by TTX. GluR1 palmitoylation was slightly increased due to TTX treatment, but was not statistically different from the untreated control. GluR1, GluR2 and SYTI total palmitoylation were unaffected by APV/CNQX manipulation. (C and D) Autoradiographic data showing the half-life of palmitate on GluR2 after blocking neuronal activity. TTX increased the calculated half-life from ∼6 h to ∼14 h, while APV/CNQX increased it to more than 30 h. Mean ± SEM from three separate data sets are shown. *P < 0.05, **P < 0.01. After normalization to total AMPARs Input, we noted that the palmitoylated pool of GluR2 grew to 131.0±6.1% (p<0.05,n=3) after TTX, but not APV/CNQX application (97.9±3.7% of untreated control, p=0.92, n=3), and that the total palmitoylated pools of GluR1 (TTX, 111.7±7.9%, p=0.18; APV/CNQX, 105.0±5.7%, p=0.63, n=3) and SYTI (TTX, 97.3±8.8%, p=0.95; APV/CNQX, 102.5±9.7%, p=0.99, n=3) remained unchanged (Figure 21A, B). We further examined the half-life of newly added palmitate on GluR2 using pulse-chase methods. In control neurons, GluR2 labelling indicated a half-life of approximately 6 h (Figure 21C, D; 76  Figure 22). Chronic TTX manipulation markedly prolonged the half-life of palmitate on GluR2 to approximately 17 h. Moreover, chronic APV/CNQX incubation extended the half-life of palmitate even more, for longer than 30 h (Figure 21C, D). In combination, the pool size of palmitoylated AMPARs and the retention time of palmitate on AMPARs appear to work interactively as a dynamic system and contribute to activity-regulated AMPAR trafficking and stability.  Figure 22. Turnover rate of palmitate on GluR1 and GluR2. A) Representative data of pulse-chase experiments showing the half-life of palmitate on GluR1, GluR2, SYT I and PSD-95. B) The calculated half-life of palmitate is ~6.1 h for GluR2, ~ 9.8 h for GluR1, ~3.0 h for PSD-95 and ~35.2 h for synaptotagmin I. Data used to calculate the half-life of palmitate on GluR1 and GluR2 were from four and five independent experiments, respectively.  2.2.7 Acute Effects of NMDA and AMPA on AMPAR Palmitoylation Besides chronic activity manipulation, we wondered whether palmitoylation on AMPARs is also involved in rapid receptor regulation. It has been shown that 1 h of glutamate stimulation, which induces AMPAR internalization (Mangiavacchi and Wolf, 2004), accelerates 77  depalmitoylation of AMPARs (Hayashi et al., 2005). It is thus possible that acute postsynaptic AMPAR stimulation could regulate AMPAR palmitoylation.  Figure 23. AMPAR subunit palmitoylation is differentially affected by acute NMDA and AMPA treatment. Neurons were acutely treated with NMDA (50 μm for 3 min) or AMPA (100 μm for 10 min) and were returned to Neurobasal medium for the indicated times. The palmitoylation level was tested using the biotin-BMCC assay for GluR1 (A) and GluR2 (C). The normalized palmitoylation level was then plotted according to the indicated time points for GluR1 (B) and GluR2 (D). Mean ± SEM from three to five separate data sets are shown. *P < 0.05, **P < 0.01.  To test this hypothesis, we examined the time course of AMPAR palmitoylation level after acute NMDA (50 µM for 3 min) or AMPA (100 µM for 10min) application to cultured cortical neurons. AMPA or NMDA treatment led to different palmitoylation profiles. Following NMDA treatment, GluR1 palmitoylation was rapidly enhanced within 3 min to 126.5±9.9% compared to 0 h control  78  (p<0.05), and was then significantly reduced with a minimum of 77.6±7.6% (p<0.05) at 30 min. After that, although the palmitoylation level of GluR1 slowly increased, it was still significantly lower than the 0 h control at the end of 2 h after treatment (80.8±6.1% of baseline; p<0.05; n=5; Figure 23A,B). In contrast, AMPA treatment caused a dramatic reduction of GluR1 palmitoylation after 3 min (72.3±6.2% of 0 h control; p<0.05), followed by a recovery within 30 min (91.3±0.1%; p=0.28). After a recovery, a gradual decrease of palmitoylation level was observed till the end of the experiment. However, the difference was not statistically different from the 0 h control (2h, 86.5±7.5%; p=0.14; n=3; Figure 23A,B). NMDA treatment also induced a slight increase of GluR2 palmitoylation at 3 min (110.4±15.2%; p=0.51), and then a reduction to 83.7±1.0% (p=0.07) at 30 min. Both changes were not statistically significant in contrast to the effects on GluR1. Instead of a slow recovery observed with GluR1, a steady decrease in GluR2 palmitoylation level to 71.9±3.6% of 0 h control was noticed at 2 h after the treatment (p<0.01, n=4; Figure 23C,D). AMPA application promoted an increase to 107.4±11.5% (p=0.54, n=4) of GluR2 palmitoylation after3 min, followed by a slow decrease to 91.±7.7% (p=0.0.29) of control after 2 h, which was not statistically significant either compared to control or to NADA treatment (Figure 23C, D). It is interesting to note that whereas NMDA and AMPA caused opposite changes of GluR1 palmitoylation, they showed no significant effect on GluR2 palmitoylation within the early 30 min after treatments. In this period, both NMDA and AMPA effectively changed GluR1 palmitoylation only, suggesting a differential role of palmitoylation in fast regulation of AMPAR subunits. Moreover, at the late stage of treatments NMDA, but not 79  AMPA, caused a significant reduction of GluR2 palmitoylation, suggesting a delayed regulatory role of GluR2 palmitoylation.  2.3 Discussion Increasing evidence indicates that AMPAR trafficking is regulated by distinct mechanisms separately mobilizing GluR1 and GluR2 (Chen et al., 2000; Bredt and Nicoll, 2003; Ahmadian et al., 2004; Lee et al., 2004). This specificity is achieved by recruiting subunit-unique domains or modifications. Here, we report that the mechanisms and functional consequences of palmitoylation on AMPAR subunit GluR1 and GluR2 are also subunit-specific.  2.3.1 Different Palmitoylation Mechanisms for AMPAR Subunits An important finding of the present study is that altering the early secretory pathway by CHX, BFA and NOC shows distinct effects on GluR1 and GluR2 palmitoylation. From our GluR1 results, we found that BFA attenuates GluR1 palmitoylation, a reduction that could not be prevented by blocking retrograde transport from the Golgi to the ER by the microtubule depolymerizing reagent NOC. This discovery supports the idea that anterograde transport from the ER to Golgi complex is important for the palmitoylation of a distinct subset of GluR1. Given that the ER residency time of GluR1 is short (Greger et al., 2002), this fraction of GluR1 probably comprises newly synthesized subunits, which is consistent with our finding that the 80  effects of BFA and CHX on GluR1 palmitoylation were not additive (Figure 11E). An interesting question is where this fraction of GluR1 is palmitoylated. It has been reported that NOC alone induces Golgi fragmentation and reassembles Golgi stacks at peripheral ER-exit sites (Lippincott-Schwartz et al., 1990). Until Golgi stacks are regenerated, Golgi enzyme processing of proteins exported from the ER is inhibited for several hours (Lippincott-Schwartz et al., 1990). In our experiments, GluR1 palmitoylation was unaffected by NOC application, suggesting that the BFA-sensitive fraction of GluR1 has been already palmitoylated before GluR1 enters the Golgi apparatus, probably at ER-exit sites. The palmitoylation of GluR1 is very similar to that of another membrane protein, the δ opioid receptor (Petaja-Repo et al., 2006), which is also palmitoylated at ER-exit and also shows dependency on anterograde transport. It is noteworthy that all treatments used in our study to alter normal functions of the early secretory pathway could not completely abolish GluR1 palmitoylation. It is thus possible that GluR1 could be palmitoylated at other cellular locations besides the ER and the region between the ER and the Golgi complex.  In contrast to GluR1, GluR2 palmitoylation does not require anterograde transport (Figure 12). Rather, GluR2 subunits are palmitoylated locally in the ER and newly synthesized GluR2 receptors are the major substrates, given the high sensitivity of GluR2 palmitoylation to CHX and the stable status of newly synthesized GluR2 in the ER (Greger et al., 2002). These conclusions are supported by EndoH sensitivity data, which show that newly palmitoylated 81  GluR2 receptors are mainly confined to the ER. AMPAR tetramer assembly occurs in the ER where GluR2s associate primarily with GluR1 or GluR3. Consistent with ER-palmitoylation of GluR2, we found that most of the palmitoylated GluR2s are not associated with GluR1 (Figure 12H). This is not surprising as it has been shown that GluR1/2 receptors are found predominantly on the cell surface, whereas GluR2/3s mostly reside in the ER (Greger et al., 2002). It thus seems that the rapid trafficking of GluR1/2 from the ER to the PM may require depalmitoylation of GluR2 (Hayashi et al., 2005). A surprising finding is the strikingly opposite effect of Golgi apparatus destruction on GluR1 and GluR2 palmitoylation. In fact, the majority of proteins show reduced palmitoylation upon BFA treatment as with GluR1, rather than an increase, such as p63/CKAP4 (Mundy and Warren, 1992). Similar to p63, the enhancement of GluR2 palmitoylation by BFA seems not to be due to the redistribution of Golgi-located PATs. Other possibilities could not be excluded, such as accumulation of substrates in the ER, or accumulation of PATs in the ER, or even that BFA itself may behave as an activator of PATs (Mundy and Warren, 1992). As ER-residency time for GluR2 is long, the accumulation of GluR2 in the ER is not likely to be the key factor for the increased palmitoylation. In fact, p63 has been reported to be a substrate for zD2, a PAT localized to both the ER and Golgi complex (Ohno et al., 2006; Zhang et al., 2008). It has also been shown that many proteins resident in the early secretory pathway, like zD2, are constantly transported via small cargos between cisternal ER, pre-Golgi intermediate and Golgi stacks (Appenzeller-Herzog and Hauri, 2006) and the retrograde trafficking of these cargos relies on molecular motors and the cytoskeleton 82  (Lippincott-Schwartz et al., 1990). BFA might cause accumulation of cycling ER-resident PATs such as zD2, which thus enhance palmitoylation of proteins in the ER, such as p63 and GluR2. On the other hand, the microtubule depolymerizer NOC, which blocks retrograde transport of PATs from the Golgi back to the ER, would reduce the amount of PATs in the ER and correspondingly attenuate protein palmitoylation. We observed a reduction of GluR2 palmitoylation upon NOC treatment, suggesting that an intact retrograde transport microtubule network is necessary for GluR2 palmitoylation. Although the recycling PAT hypothesis is plausible, there are some other possibilities. Perhaps BFA or NOC itself are activators or inhibitors of PATs. Thus, identification of ER/Golgi-resident PATs for GluR2 in the future would help to elucidate these possibilities. It is noteworthy that depalmitoylation processes are not considered in this scenario. Two well-documented deplamitoylation enzymes of the palmitoyl protein thioesterase (PPT) family, PPT1 and PPT2, have been shown to localize at lysosomes whose components are not redistributed to the ER when treated with BFA (Lippincott-Schwartz et al., 1991; Verkruyse and Hofmann, 1996), implying a limited role for PPTs in the early regulation of AMPAR palmitoylation. However, future studies on PPTs might provide new insights into this issue. In summary, our data collectively show that palmitoylation of GluR1 and GluR2 subunits takes place at different cellular subcompartments and follows distinct rules.  83  2.3.2 Role of GluR2 Palmitoylation in AMPAR Trafficking One advantage of subunit-specific palmitoylation rules is that they ensure diverse and multiple functions of palmitoylation on AMPAR. Here, this idea is further supported by our evidence that ER-palmitoylation specifically on GluR2 prevents AMPAR sorting to lysosome for degradation. In neurons, we found that blocking palmitoylation by 2-BrPA specifically reduced GluR2 but not GluR1 subunit number. Moreover, this reduction could be eliminated by applying lysosome but not proteosome inhibitors. Indeed, many presynaptic and postsynaptic proteins which are critical for AMPAR trafficking, such as SYTI, GRIP and ABP, have been reported to be palmitoylated (Greaves and Chamberlain, 2007). Broad inhibition of palmitoylation on these proteins by 2-BrPA may in part contribute to GluR2 redistribution in neurons. However, a decreased stability of palmitoylation-deficient GluR2 mutants in heterologous cell lines supports the notion that palmitoylation on GluR2 plays a direct role in regulating AMPAR sorting and stability (Figure 19).  One purpose of active protein degradation is elimination of malfunctioning proteins in cells (Hebert and Molinari, 2007). Palmitoylation-deficient proteins are sometimes recognized as misfolded proteins and are degraded via an ER-associated degradation (ERAD) pathway (Hebert and Molinari, 2007). However, we did not observe GluR2 aggregation in the ER after blocking palmitoylation (Figure 17), and deficient mutants show normal electrophysiological properties (Hayashi et al., 2005). GluR2 palmitoylation in the ER also seems not to be involved in receptor 84  maturation given that blocking palmitoylation: (i) did not affect the pool of immature GluR2 pool, suggesting a normal ER exit rate for AMPARs; but (ii) reduced the pool of mature GluR2 pool, indicating that non-palmitoylated receptors are transported to the Golgi before being targeted for degradation (Figure 16). Considering that GluR2 palmitoylation mostly occurs in the ER, we propose that the palmitoylation of newly synthesized GluR2 defines the degradation track of AMPARs. This is supported by our data, which show that inhibition of protein synthesis can completely prevent GluR2 degradation caused by 2-BrPA-induced block of palmitoylation. Given the half-life of palmitate on GluR2 subunits, some GluR2 subunits might be targeted for degradation after blocking protein synthesis and palmitoylation. However, the loss of this putative fraction of GluR2 is too subtle to be detected by our techniques.  Two sites on AMPAR have been found to be palmitoylated, one at TMD2 and the other at the C-terminus (Hayashi et al., 2005). Blocking only palmitoylation on TMD2 promotes GluR2 degradation (Figure 18). As depalmitoylation of the TMD2 site within the Golgi apparatus was suggested to promote receptor budding (Hayashi et al., 2005), it is possible that blocking palmitoylation enhances GluR2 budding from the Golgi and correspondingly increases GluR2 cycling to the lysosome for degradation. However, this is less likely as our experiments have demonstrated that PMA-promoted budding of AMPARs from the Golgi apparatus does not reduce total receptor levels. Although PMA may have some side effects, it is unlikely to inhibit GluR2 recycling (Iwakura et al., 2001; Barry and Ziff, 2002), lysosome targeting (Sapin et al., 85  1997; Trivedi et al., 2006) or lysosome processing (Figure 19). Thus, non-palmitoylated GluR2 is more likely to be directly sorted to lysosome for degradation. We suggest that palmitoylation on the GluR2 TMD2 site (C610) might play two distinct roles: first, early palmitoylation in the ER stabilizes AMPAR in the early secretory pathway and ensures its forward transport to other parts of the Golgi apparatus for surface delivery; second, once the receptor reaches its destination at the Golgi apparatus, its requires a depalmitoylation step to be delivered to the surface.  2.3.3 Implications for Synaptic Function Similar to many other synaptic proteins, such as PSD-95 (Perez and Bredt, 1998; El-Husseini and Bredt, 2002), the processing of AMPAR subunits is tightly controlled by neuronal activity (O'Brien et al., 1998). We report here that chronic inhibition of action potentials and neurotransmitter release by TTX can increase the pool of palmitoylated GluR2 (Figure 21B). According to our findings on the palmitoylation-dependent AMPAR sorting, this may be accompanied by an increase of total GluR2 number in the cell, as the lysosome-targeted pool of GluR2 is reduced due to stabilization of GluR2 subunits with palmitoylation. Indeed, we detected an increase of GluR2 subunits upon chronic TTX treatment, although it was not as dramatic as previously reported (Wierenga et al., 2005). Blocking spontaneous miniature excitatory postsynaptic current activity by APV/CNQX also increased total GluR2. However, APV/CNQX had no effect on the proportion of palmitoylated GluR2 (Figure 21B), suggesting a different mechanism for the increase of total GluR2. In a previous study, it has been shown that 86  blocking action potentials diminishes protein synthesis whereas blocking minis enhances protein translation (Sutton et al., 2004). Therefore, the APV/CNQX-induced increase of GluR2 protein is likely to be a result of net increase of protein synthesis instead of reducing GluR2 lysosome sorting. On the other hand, both TTX and APV/CNQX differentially prolonged the half-life of palmitate on GluR2 (Figure 21D). The increased stability of palmitate on GluR2 after blocking neuronal activity may reflect a reduced surface trafficking rate of Golgi-resident AMPARs, as depalmitoylation of GluR2 is required for exit from the Golgi apparatus (Hayashi et al., 2005). This notion is not surprising as chronic activity blockade has been shown to ‘slow down’ the system by reducing overall protein synthesis (Sutton et al., 2004), AMPAR trafficking rates (Ehlers, 2000) and protein turnover (O'Brien et al., 1998; Yi and Ehlers, 2005; Ibata et al., 2008). It is interesting to note that APV/CNQX treatment did not change the pool of palmitoylated GluR2. In contrast, it prolonged the half-life of palmitate on GluR2 more effectively than did TTX (Figure 21D). We speculate that the balance of palmitoylation and depalmitoylation processes may contribute to the differences that we observed. It is possible that blocking action potential by TTX causes a greatly reduced depalmitoylation process, which leads to both enlargement of the palmitoylated pool of GluR2 and a prolonged half-life of palmitate, whereas APV/CNQX reduces both palmitoylation and depalmitoylation processes, which only affects the palmitate half-life. Other mechanisms cannot be excluded and further study is needed to address these potential underlying mechanisms. However, the differential effects of TTX and APV/CNQX indeed suggest that action potentials and miniature synaptic events may play 87  different roles in GluR2 stability and trafficking by controlling palmitate half-life and the palmitoylated pool of GluR2. It also suggests that the increase of the palmitoylated pool and the stabilization of palmitate on GluR2 may have distinct functional consequences. It should be noted that we always detected an extra band with a slower migrating rate than GluR1 and GluR2 in the BMCC fatty acyl exchange assay (protocol 2). We are currently uncertain about the identity of this band (Figure 15). It is possible that such bands represent glycosylated forms of GluR subunits or are due to antibody-unspecific binding effects.  Palmitoylation of AMPAR subunits responded quickly to synaptic activity. Acute application of NMDA or AMPA, which mobilizes postsynaptic AMPARs, differentially regulates palmitoylation on AMPAR GluR1 and GluR2 subunits (Figure 25). Although the specific function of these rapid and diverse responses of GluR1 palmitoylation to NMDA and AMPA is currently unknown, they may partially contribute to activity-induced fast receptor trafficking (Hayashi et al., 2005). In contrast, different effects of NMDA and AMPA on GluR2 palmitoylation were observed after 30 min. While AMPA showed a small impact on GluR2 palmitoylation, NMDA caused a significant reduction of palmitoylation. The different palmitoylation outcomes from AMPA and NMDA treatments may be explained by their distinct effects on AMPAR stability. It has been shown that acute activation of NMDAR and AMPAR can distinctively regulate AMPAR trafficking (Shi et al., 1999; Ehlers, 2000; Lee et al., 2004). Whereas AMPAR stimulation promotes recycling of surface AMPARs, NMDAR activation 88  enhances degradation of internalized AMPARs and reduces their total level (Lee et al., 2004). Our findings that the NMDA-induced decrease of GluR2 palmitoylation leads to AMPAR degradation supports the idea that palmitoylation of GluR2 in the ER is needed to control the transport of AMPAR from the ER to the PM, while reduced palmitoylation targets GluR2 for degradation via the lysosomal pathway. Considering the presence of ER and Golgi in dendritic shafts and even at the base of spines (Horton and Ehlers, 2003), AMPAR palmitoylation, together with the mechanisms controlling the expression of surface AMPAR, may be involved in the establishment of a functional balance of synaptic AMPARs, therefore regulating synaptic plasticity.  89  Chapter 3. Isoform-Specific Palmitoylation of c-Jun N-Terminal Kinase (JNK) Regulates Axonal Development  A version of this chapter has been published online  Guang Yang, Yuan Liu, Kaiyun Yang, Rui Liu, Shanshan Zhu, Ainsley Coquinco, Wendy Wen, Luba Kojic, William Jia and Max S Cynader. Isoform-specific palmitoylation of JNK regulates axonal development. Cell Death and Differentiation. doi: 10.1038/cdd.2011.124  90  3.1 Introduction JNK family consists of JNK1, JNK2 and JNK3 subgroups, which participate in diverse biological and pathological processes (Davis, 2000; Weston and Davis, 2002; Haeusgen et al., 2009). At least 10 JNK isoforms varying in sizes, with most about 46 kDa and 54 kDa are produced from Jnk1-3 genes, giving rise to p46 and p54 types of JNK isoforms (Bogoyevitch, 2006). In the nervous system, JNKs (particularly isoform JNK3) have been extensively studied as key players in apoptosis and neurodegeneration (Yang et al., 1997; Saporito et al., 2000; Morishima et al., 2001; Savage et al., 2002; Kuan et al., 2003; Hunot et al., 2004). However, accumulating evidence supports a physiological role of JNK in regulating neurite formation and morphogenesis (Coffey et al., 2000; Chang et al., 2003; Waetzig and Herdegen, 2003; Bjorkblom et al., 2005; Kimberly et al., 2005; Tararuk et al., 2006; Ciani and Salinas, 2007). Pharmacological inhibition of JNK activity blocks axogenesis in hippocampal neurons, arguing for an essential role of JNK in neurite development (Oliva et al., 2006). Growth factors and signalling molecules, including secreted proteins of the Wnt family, have also been found to activate JNK for remodeling dendrites and axons, a process that relies on cytoskeletal rearrangement (Lucas and Salinas, 1997; Moriguchi et al., 1999; Rosso et al., 2005; Ciani and Salinas, 2007; Salinas and Zou, 2008). This leads to the identification of several actin- or microtubule-associated proteins as JNK substrates which modulate different aspects of cytoskeletal activity (Bogoyevitch and Kobe, 2006). However, all 10 JNK isoforms share the 91  same kinase domain and activation mechanism (Davis, 2000; Bogoyevitch and Kobe, 2006). The differential roles of JNK isoforms in neurite development and the mechanisms underlying isoform-specific regulation are poorly understood.  Analysis of animals null for particular JNK isoforms provides the first evidence to support isoform-specific roles of JNKs in the brain. Mice with Jnk1-/- show abnormalities of neurite development (Chang et al., 2003), while mice null for Jnk1 and Jnk2 show embryonic lethality due to severe neurological defects (Kuan et al., 1999; Sabapathy et al., 1999). Jnk2-/- mice, but not Jnk1-/- mice, also show enhanced protection against brain damage in Parkinson’s disease models (Hunot et al., 2004). JNK1 and JNK2 are thus implicated in regulating brain development, as well as neuronal death. In contrast, Jnk3-/- mice apparently develop normally but show enhanced resistance to stress-induced neurodegeneration (Yang et al., 1997; Morishima et al., 2001; Kuan et al., 2003; Hunot et al., 2004). JNK3 is thus considered as a key regulator of neuronal cell death, rather than of neuronal development. However, recent studies highlight a physiological role of one particular JNK3 isoform, p54, in neuronal differentiation and neurite growth (Waetzig and Herdegen, 2003; Kimberly et al., 2005), raising the possibility that each isoform may be differentially regulated to achieve selective functions. Understanding how JNK isoforms are specifically regulated is essential for elucidating JNK isoform-specific roles in multiple biological processes.  92  Isoform-selective regulation has been observed with some JNK scaffolding proteins that regulate JNK activity and trafficking (Morrison and Davis, 2003; Bogoyevitch, 2006). Another potential mechanism may be the post-translational modification. In addition to phosphorylation, that activates JNKs, little is known about whether JNKs are subject to other modifications (Davis, 2000; Weston and Davis, 2002). Protein palmitoylation is one such candidate modification. It dynamically regulates protein trafficking and function by reversibly attaching the lipid palmitate to cysteine (Cys) residues (El-Husseini and Bredt, 2002; Bijlmakers and Marsh, 2003; Linder and Deschenes, 2007). This modification is catalyzed by the family of palmitoyl acyl transferases (PATs) containing 24 members, and is sensitive to protein sequence and structure (El-Husseini and Bredt, 2002). Proteins with little sequence variance may differ in palmitoylation sensitivity and specificity (Yamazaki et al., 2001; Bijlmakers and Marsh, 2003; Yang et al., 2009), such as glutamate receptor interacting proteins 1 isoform a and b (GRIP1a/b) (Yamazaki et al., 2001), providing a potential way to achieve isoform-specific regulation. Sequence alignment of JNK isoforms indicates that differences exist at NH2- and COOH-termini, potential regions for isoform-specific regulation. Indeed, the NH2- terminus of JNK3 has been shown to selectively interact with the scaffolding protein β-arrestin2 (McDonald et al., 2000; Guo and Whitmarsh, 2008). The Cys residues that are likely targets for palmitoylation are located at COOH-termini of the p54 isoforms and are not found on the p46 isoforms.  In this chapter, we demonstrate that JNK3 p54 is the predominant isoform that is 93  palmitoylated in neurons. With this isoform-specific regulation, JNK3 is found to play a role in axon branching and filopodia motility, possibly via modulating cytoskeletal components, as palmitoylation affects JNK3 trafficking to the triton-insoluble actin cytoskeletal fraction. This palmitoylation-regulated function of JNK3 responds to Wnt signals and is necessary for Wnt7a-induced axonal branching. Our results demonstrate that palmitoylation is a JNK isoform-specific regulation and suggest that the rapid and dynamic palmitoylation on JNK3 plays an essential role in axonal growth and branching during development.  3.2 Results  3.2.1 Palmitoylation Occurs Primarily on the JNK3 Isoform After metabolic labelling of neuronal cultures, we detected 3H-palmitate incorporation into JNK3, which was eliminated by co-incubation with the palmitoylation inhibitor 2-bromopalmitate (2-BrPA) (Figure 24A). In vitro application of hydroxylamine (HAM), which breaks covalent bonds and releases 3H-palmitate from the protein, also abolished the signal, confirming that JNK3 is a palmitoylated protein in neurons. A pulse-chase strategy was used to examine kinetics of palmitoylation on JNK3.We found a highly dynamic cycling of palmitate on JNK3 with a calculated half-life of about 4 h (Figure 24B). The selectivity of PATs on JNK3 palmitoylation was then tested by expressing several PATs individually with GFP or GFP tagged 94  JNK3 in heterologous cells. PATs zD15 and zD20 significantly promoted JNK3 palmitoylation (Figure 24C). However, introducing zD23 or a zD15 mutant without PAT activity (zD15∆) failed to enhance JNK3 palmitoylation (Figure 24C,D), suggesting that JNK3 may be the substrate of a selective group of PATs in neurons.  Among the JNK p54 isoforms with an extended COOH-terminus, JNK1 and JNK3, but not JNK2, have two Cys residues located at the end of the terminus (Figure 24D). We next investigated whether these Cys residues are potential sites for JNK3 palmitoylation by replacing Cys with Ser. In the presence of PAT zD15, loss of one Cys was enough to abolish JNK3 palmitoylation, indicating that both Cys residues of JNK3 are required for normal palmitoylation. This implied that JNK2 with only one Cys residue may not undergo palmitoylation. Indeed, while JNK3 palmitoylation was clearly detected, we were unable to detect JNK2 p54 palmitoylation above the background control, assessed in heterologous cells (Figure 24E). To our surprise, we also found that JNK1 p54 was not a palmitoylated protein though it has two Cys residues at the COOH-terminus (Figure 24E). This point is considered further in the discussion section. Thus, the JNK3 p54 is the major isoform that is palmitoylated in neurons.  95  Figure 24. JNK3 is palmitoylated at the COOH-terminus. A, Metabolic labelling with 3H-palmitic acid (3H-palm) shows that JNK3 is palmitoylated in neurons. Normal rabbit IgG served as the immunoprecipitation control. The arrowhead indicates JNK3. B, Palmitate on JNK3 has a short half-life. Pulse-chase experiments show a short half-life (calculated ~4.29 h, n=5) of palmitate on JNK3. C, PATs show selectivity in palmitoylating JNK3. D, Two Cys residues at the COOH-terminus of JNK3 are critical for palmitoylation. The sequence alignment of the last 15 amino acids at the COOH-termini of JNK p54 isoforms reveals potential Cys residues (bold letters) for palmitoylation. E, The palmitoylation status of JNK p54 isoforms JNK1α2, JNK2α2, and JNK3α2 are shown and compared to background (HAM minus). The schematic diagram at the bottom shows that the JNK3 p54 is the major isoform that is palmitoylated.  3.2.2 Axonal Branching Is Regulated by JNK3 Palmitoylation We first suspected that palmitoylation may affect JNK3 phosphorylation under stress conditions (Stoffel et al., 1998; Weston and Davis, 2002). However, similar phosphorylation 96  levels of JNK3 wildtype (JNK3 WT) and palmitoylation-deficient JNK3 mutant (JNK3 CS) were observed in heterologous cells challenged with or without osmotic-stress, implying that palmitoylation may not be involved in the JNK3-regulated stress response.  Recent studies have suggested a physiological role of JNK3 in neuronal differentiation and neurite growth (Waetzig and Herdegen, 2003; Kimberly et al., 2005). To study the potential role played by JNK3 palmitoylation in axonal development, we transfected cultured hippocampal neurons with indicated constructs together with DsRed2 at 5 DIV for 48 h. DsRed2 expression filled the neuron and was used to track morphological changes of axons. At 7 DIV, neurons generated long and branched axons which were clearly labelled with both DsRed2 and the axonal marker Tau-1, and several short dendrites around soma where only DsRed2 signals were detected (Figure 25A). Axons from neurons overexpressing JNK3 WT showed similar structures to the DsRed2 control. In contrast, overexpressing JNK3 CS dramatically promoted axonal complexity with an increase of branch numbers (Figure 25B). We quantified axonal branching by measuring the axonal branch number of different branching orders (Bodmer et al., 2009). Neurons overexpressing JNK3 CS showed more branches at all branching orders (secondary, 6.53 ± 0.26, P<0.01; tertiary, 2.66 ± 0.23, P<0.01; higher, 0.44 ± 0.11, P<0.01) and doubled the total branch number, compared to neurons with DsRed2 alone (secondary, 3.27 ± 0.15; tertiary, 0.69 ± 0.11; higher, 0.06 ± 0.03) or JNK3 WT (secondary, 3.76 ± 0.19; tertiary, 0.83 ± 0.17; higher, 0.03 ± 0.02) (Figure 25C). Moreover, compared to controls, JNK3 CS induced a 97  significant increase in total branch length (Figure 25C), although the length of the primary axon decreased.  Figure 25. The palmitoylation-deficient JNK3 mutant promotes axonal branching and filopodia motility. A, Long and branched axons are observed in cultured hippocampal neurons at 7 DIV. Axons are 98  labelled with the axonal marker Tau-1 (green) and DsRed2 (red). Scale bar: 10 µm. B, Overexpression of the palmitoylation-deficient JNK3 mutant (JNK3 CS) increases axon length and branch complexity. Scale bar: 10 µm. C, Axonal branch numbers and total length of axons are enhanced by the JNK3 CS mutant. The tip number of secondary, tertiary and higher order of branches, the total branch number of axons (DsRed2 alone/blue, 5.02 ± 0.22, n=131; WT/red, 5.58 ± 0.28, n=66; CS/green, 10.61 ± 0.43, n=81) and total axonal length (fold change to DsRed2 alone, WT, 1.07 ± 0.04; CS, 1.42 ± 0.04) are shown. t-test. ** indicates P<0.01. D, The density of axonal filopodia is not affected by JNK3 CS. No significant changes in the CS group are observed (fold change to DsRed2 alone, WT, 0.96 ± 0.05; CS, 1.02 ± 0.04, P=0.85 to DsRed2 alone, P=0.40 to WT). Scale bar: 5 µm. E, Axonal filopodia motility is promoted by JNK3 CS. The motility of axonal filopodia is further transformed into a single image and shown below. Scale bar: 5 µm. Regions of marked dotted-squares are magnified and time-lapse images are shown with indicated time points within 135 sec. Four labelled filopodia, as selected examples in the CS group, are more mobile than those in the WT group. Scale bar: 5 µm. The motility of axonal filopodia (10 filopodia per neuron from 20 neurons) is quantified with the motility index (MI) (DsRed alone, 0.35 ± 0.02, n=200; WT, 0.40 ± 0.02, n=200, P=0.12; CS, 0.70 ± 0.03, n=200 P<0.01).  To test whether enhanced axonal branching was due to increased formation of filopodia, we assessed the density of axonal filopodia. As shown in Figure 25D, filopodial density measured in neurons overexpressing JNK3 CS did not differ from controls. However, time-lapse recording showed that axonal filopodia were more dynamic on extension and retraction in JNK3 CS neurons (Figure 25E). To directly visualize the dynamics of filopodia, we adopted a protocol to sum up intensity differences of each pixel between successive frames during the period of recording (Chang and De Camilli, 2001). Compared to controls, axonal filopodia in neurons overexpressing JNK3 CS were shown to display higher motility (Figure 25E), which was further quantified with a motility index (MI) (Chang and De Camilli, 2001).  99  3.2.3 Palmitoylation Regulates JNK3 Trafficking to Cytoskeleton Palmitoylation is an essential signal for facilitating protein association with lipid rafts which act as platforms for signal transduction. JNK has been shown to be recruited to lipid rafts where it can initiate downstream pathways (Gajate and Mollinedo, 2005; Nieto-Miguel et al., 2008). We thus examined the role of palmitoylation in lipid raft targeting of JNK3. Lipid rafts are the PM micro domains rich in cholesterol, which resist extraction with detergents (Simons and Toomre, 2000).  To isolate lipid rafts, cell lysates treated with Triton X-100 were subject to Optiprep gradient density centrifugation (Vetrivel et al., 2004). The lipid raft marker caveolin-1 showed an intensity peak at fractions 8 and 9 (Figure 26A, left panel). Treating cells with methyl-β-cyclodextrin (MβCD), a drug known for cholesterol depletion and lipid raft destruction (Vetrivel et al., 2004), reduced the presence of caveolin-1 at fractions 8 and 9, suggesting the enrichment of lipid rafts in these fractions. The distribution of JNK3 WT and JNK3 CS in fractions 8 and 9 showed no significant difference (Figure 26A, right panel). Similar results were found with JNK3 WT co-expressed with PAT zD15 to promote JNK3 palmitoylation. Moreover, unpalmitoylated JNK2 p54 was also present in lipid rafts. Hence, palmitoylation does not regulate lipid raft association of JNK3. Nevertheless, compared to JNK3 WT, an increase of JNK3 CS and a small decrease of JNK3 WT co-expressed with zD15 were detected at fraction 13, the high-density triton-insoluble fraction enriched with actin-associated cytoskeleton (Bodin 100  et al., 2005; Chakrabandhu et al., 2007). This implies that palmitoylation may affect JNK3’s association with the actin cytoskeleton. Signals for actin cytoskeleton modulation are essential for axonal development (Luo, 2002).  Figure 26. Palmitoylation regulates the translocation of JNK3 to the Triton-insoluble actin cytoskeleton. A, JNK3 CS does not affect JNK3 trafficking to lipid rafts. Quantifications of the immunoblot intensity of the lipid raft marker caveolin-1 or JNKs from each fraction are shown at the bottom. Fractions 8 and 9 (f 8, 9) are marked as lipid rafts. B, JNK3 is associated with actin-rich regions in Cos7 cells and neurons. Arrows indicate the colocalization of endogenous JNK3 (green) or GFP-JNK3 with the actin cytoskeleton (red) detected with TR-phalloidin. C, JNK3 is present in 101  the Triton X-100 insoluble-cytoskeleton fraction. The presence of JNK3 in the insoluble fraction is calculated by the insoluble/soluble ratio (fold change to control, Cyt.D, 0.45 ± 0.06; Lat.A, 0.38 ± 0.11). t-test. * indicates P<0.05. Representative figures are from three independent experiments. D, The JNK3 CS mutant localizes to membrane ruffles containing the actin cytoskeleton. Arrows indicate colocalization of JNK3 (green) with the actin cytoskeleton (red) at submembrane structures. Scale bar: 5 µm. E, Palmitoylation regulates JNK3 translocation to the Triton X-100 insoluble fraction. The insoluble/soluble ratio are shown (WT, 0.67 ± 0.08; CS, 1.46 ± 0.24; WT+zD15, 0.25 ± 0.07). t-test. ** indicates P<0.01.  Although JNK is known to modulate the actin cytoskeleton and several actin-binding proteins have been identified as JNK substrates, the association of JNK3 with the actin cytoskeleton has not been well investigated. We first examined the colocalization of GFP-JNK3 and the actin cytoskeleton, as detected by fluorescence-labelled phalloidin, in heterologous Cos7 cells. JNK3 is a soluble protein which distributes homogenously within the cell. We noticed that a portion of JNK3 accumulated at actin-rich PM ruffles and on stress-fibers (Figure 26B). This colocalization of JNK3 with actin structures was abolished by treating cells with the actin depolymerizer cytochalasin D (Cyt.D). To exclude the signal from soluble JNK3, 0.5% Triton X-100 was used to extract cells, which preserved proteins bound to the actin cytoskeleton in cells and released unbound ones. After extraction, JNK3 signals were mostly colocalized with the actin cytoskeleton, as detected by Texas Red (TR)-phalloidin, suggesting that a portion of JNK3 is associated with the actin cytoskeleton (Figure 26B). To analyze this quantitatively, we isolated the Triton-insoluble actin cytoskeleton fraction from neurons and heterologous cells, followed by detection of JNK3 (Bodin et al., 2005). Consistently, JNK3 was detected in the Triton-insoluble fraction (Figure 26C), which was sensitive to the actin depolymerizer Cyt.D or latrunculin A 102  (Lat.A). We next investigated whether palmitoylation affected JNK3 association with the actin cytoskeleton. Compared with WT, JNK3 CS showed stronger signals at membrane ruffles enriched with the actin cytoskeleton (Figure 26D). This was further supported by detecting a twofold increase of JNK3 CS in the Triton-insoluble fraction, compared to WT (Figure 26D). Consitently, enhancing JNK3 palmitoylation by PAT zD15 caused a 50% reduction of Triton-insoluble JNK3. These results suggest that JNK3 palmitoylation may affect axonal branching partially via regulating the actin cytoskeletal association of JNK3.  3.2.4 Wnt7a-Induced Axonal Branching Is Modulated by JNK3 Palmitoylation The Wnt family member Wnt7a is expressed in the hippocampal formation (Gogolla et al., 2009). It has been shown to modulate axonal development (Lucas and Salinas, 1997), and to activate the non-canonical JNK pathway (Salinas and Zou, 2008), raising the possibility that JNK3 palmitoylation may play a role in Wnt7a-induced axonal branching. We incubated 5 DIV hippocampal neurons with recombinant Wnt7a for 6 h and assessed JNK3 palmitoylation using Btn-BMCC labelling (Drisdel and Green, 2004). Wnt7a treatment dramatically reduced JNK3 palmitoylation to 50% of the untreated control, while the palmitoylation status of total proteins showed no obvious change (Figure 27A). The Wnt7a treatment also caused JNK3 translocation to the Triton-insoluble cytoskeleton fraction (Figure 27B). Incubation with Wnt7a for 48 h promoted axonal branching in neurons overexpressing JNK3 WT (secondary, 5.40 ± 0.26; 103  tertiary, 2.47 ± 0.27; higher orders, 0.24 ± 0.09; total, 9.50 ± 0.69) or DsRed2 alone (secondary, 5.40 ± 0.26; tertiary, 2.47 ± 0.27; higher orders, 0.24 ± 0.09; total, 9.13 ± 0.48), compared with the untreated control (secondary, 3.50 ± 0.29, P<0.01; tertiary, 0.52 ± 0.13, P<0.01; higher, 0.00 ± 0.00, P<0.05; total, 5.02 ± 0.37, P<0.01) (Figure 27C,D).  Figure 27. JNK3 palmitoylation modulates Wnt7a-regulated axonal branching. A, JNK3 palmitoylation is regulated by Wnt7a. Brief application of Wnt7a (200 ng/ml, 6 h) to neuronal cultures reduces JNK3 palmitoylation (fold change to control, Wnt7a, 0.56 ± 0.11). t-test. ** indicates P<0.01. The global palmitoylation status is not changed. B, Wnt7a increases the presence of JNK3 in the insoluble fraction (fold change to control, Wnt7a, 1.52 ± 0.11). C, 104  Wnt7a-induced axon branching is saturated by JNK3 CS and blocked by pseudo-palmitoylated JNK3 (Parlm). Scale bar: 10 µm. D, Differential effects of JNK3 CS and JNK3 Parlm on axonal branching induced by Wnt7a. The total axonal branches (DsRed2 only+Wnt7a/red, 9.13 ± 0.48, n=62; WT+Wnt7a/green, 9.50 ± 0.69, n=41; CS+Wnt7a/purple, 10.33 ± 0.60, n=46; F=1.28, P=0.28) and total axonal length (fold change to DsRed2 only, DsRed2 only+Wnt7a, 1.53 ± 0.43; WT, 1.55 ± 0.76; CS, 1.63 ± 0.63; F=0.34, P=0.71) are shown. The JNK3 Parlm group shows fewer axonal branches (4.25 ± 0.28, n=56; F=39.78, P<0.01) and reduced axonal length (fold change, 0.94 ± 0.35; F=19.14, P<0.01). One-way ANOVA-test. * and ** indicate P<0.05 and P<0.01, respectively. E, Pseudo-palmitoylated JNK3 inhibits normal axonal development. Neurons transfected with JNK3 Parlm develop fewer axonal branches (DsRed2 alone/black, 5.02 ± 0.22; Parlm/grey, 3.70 ± 0.25, n=33) and shorter axons (fold change to DsRed2 alone, Parlm, 0.78 ± 0.05). t-test. Scale bar: 10 µm.  However, Wnt7a stimulation did not further enhance axonal branching in JNK3 CS-transfected neurons (secondary, 6.08 ± 0.42; tertiary, 2.83 ± 0.30; higher, 0.45 ± 0.15; total, 10.33 ± 0.60), compared with JNK3 CS-transfected neurons without Wnt7a treatment (Figure 25C, Figure 27C,D). This suggests that overexpressing JNK3 CS simulated the effect of Wnt7a on axonal branching and thus prevented any further enhancement when Wnt7A was administered. Next, we fused the prenylated and dual palmitoylated motif of paralemmin to the COOH-terminus of JNK3 CS to mimic a constant pseudo-palmitoylation on JNK3 (JNK3 Parlm) (Kutzleb et al., 1998; El-Husseini et al., 2000; El-Husseini et al., 2002).The effects of Wnt7a on axonal branching were completely blocked in neurons overexpressing JNK3 Parlm (secondary, 2.75 ± 0.21, P<0.01; tertiary, 0.5 ± 0.12, P<0.01; higher, 0.00 ± 0.00, P<0.01; total, 4.25 ± 0.28, P<0.01), compared to WT treated with Wn7a (Figure 27C,D), suggesting that JNK3 palmitoylation modulates Wnt7a-induced axonal branching. Furthermore, overexpressing JNK3 Parlm was found to reduce axonal branching and total axon length in untreated neurons, 105  compared with controls (Figure 27E). Together, these results indicate that rapid and dynamic palmitoylation of JNK3 may be an essential regulator of axonal development.  3.3 Discussion Although the identification of JNK isoform-specific substrates has greatly improved our understanding of mechanisms underlying the differential roles played by JNK isoforms (Bogoyevitch, 2006; Bogoyevitch and Kobe, 2006), much remains unknown about how JNK isoforms themselves are differentially regulated (Davis, 2000; Weston and Davis, 2002). Our results demonstrate protein palmitoylation as a novel isoform-specific mechanism for JNK3 regulation, and reveal a role of JNK3 palmitoylation in axonal branching in response to Wnt signals.  3.3.1 An Isoform-Specific Regulatory Mechanism for JNK Based on previous studies (McDonald et al., 2000; Bogoyevitch, 2006; Guo and Whitmarsh, 2008), we assumed that JNK isoform-specific regulation may occur at regions unique to certain isoforms. We compared protein sequences of JNK isoforms. In the brain, Jnk1 and Jnk2 predominantly produce p46 and p54 isoforms, respectively, while Jnk3 produces isoforms of both sizes (Bogoyevitch, 2006). The COOH-termini of p54 isoforms of JNK2 and JNK3 are different from other isoforms, raising the possibility that this region may be manipulated by 106  certain mechanisms for isoform-specific regulation. One and two Cys residues are found on JNK2 p54 and JNK3 p54, respectively. We thus suspect that these isoforms may be modified by palmitoylation. By using metabolic labelling and point mutation analysis, we confirmed that JNK3 p54 is a palmitoylated protein and both Cys residues are required for palmitoylation. In contrast, JNK2 p54 which contains only one Cys residue, is not palmitoylated. Interestingly, although JNK1 p54 has similar COOH-terminal sequence with JNK3 p54, JNK1 failed to be palmitoylated in our experiments. It has been noticed that palmitoylation is sensitive to variation in protein sequence and structure (Yamazaki et al., 2001; Bijlmakers and Marsh, 2003). Small differences of amino acids at COOH-termini between JNK1 and JNK3, or different protein structures may account for their distinct palmitoylation status. However, we do not exclude the possibility that the JNK1 p54 isoform may be palmitoylated under certain circumstances. Since products of the Jnk1 gene in neurons are primarily p46 isoforms that lack extended COOH-termini for palmitoylation (Amura et al., 2005), JNK3 p54 is therefore the major isoform that is palmitoylated under our experimental conditions. This novel JNK3 palmitoylation provides evidence for the idea that certain JNK isoforms may undergo other post-translational modifications, in addition to the extensively studied phosphorylation.  3.3.2 Palmitoylation as a Trafficking Signal for JNK3 JNK proteins are ubiquitously distributed within the cell. In response to internal and external stimuli initiated from diverse origins, JNK is activated and transduces signals by 107  phosphorylating its substrates located at various loci within the cell, such as c-jun in the nucleus, MAP2 on microtubules, paxillin on the actin cytoskeleton, Bcl2 in mitochondria, and others (Weston and Davis, 2002; Bogoyevitch and Kobe, 2006). Given the simultaneous presence of JNK isoforms involved in different signalling pathways, mechanisms with isoform selectivity would be advantageous to precisely regulate JNK activity and to mobilize JNK isoforms to the appropriate cellular components. Protein palmitoylation may be one candidate mechanism. In neurons, palmitoylation has multiple effects on protein trafficking, lipid raft association and other protein functions (El-Husseini and Bredt, 2002). Despite the prominent role played by JNK3 in the stress response, loss of palmitoylation neither affects JNK3 phosphorylation in response to stress, nor changes its distribution in the nucleus and mitochondria where JNK3 activates apoptosis pathways (Weston and Davis, 2002). Thus, JNK3 palmitoylation may not directly participate in regulating the JNK3-mediated stress response. Our data further suggest that palmitoylation is not involved in controlling lipid raft association of JNK3 (Figure 26A). In fact, lipid raft association of unpalmitoylated JNK2 p54 and JNK1 p46 are normal. To our surprise, palmitoylation-deficient JNK3 CS shows an increase of translocation to the actin cytoskeleton, whereas promoting JNK3 palmitoylation by PAT zD15 reduces the translocation, assessed by confocal microscopy and cell fractionation (Figure 26E). This suggests a role of palmitoylation in regulating JNK3 association with the actin cytoskeleton. Several proteins that bind to and modulate the actin cytoskeleton have been identified as JNK substrates, such as paxillin (Huang et al., 2003). Our results provide a potential link between JNK3 palmitoylation 108  and cytoskeleton modulation, but future research is needed to elucidate underlying mechanisms.  3.3.3 A Novel Physiological Role of JNK3 in Axonal Branching The regulation of neuronal cytoskeleton is essential for axonal development (Chang and De Camilli, 2001; Luo, 2002). In agreement with the potential role of JNK3 palmitoylation in actin cytoskeleton modulation, overexpression of palmitoylation-deficient JNK3 CS in hippocampal neurons results in more elaborate axonal branches (Figure 25). JNK3 CS is also able to promote axonal filopodia motility, an actin-based process. The rapid kinetics of JNK3 palmitoylation provides the possibility that JNK3 regulates these highly dynamic processes (Figure 24B). Members of the Wnt family of proteins represent one group of prominent regulators for neurite development, which directly regulate cytoskeleton networks via recruiting JNK in the noncanonical Wnt pathway (Lucas and Salinas, 1997; Rosso et al., 2005; Ciani and Salinas, 2007; Salinas and Zou, 2008). The expressions of two Wnt members Wnt7a and b are substantial in the hippocampal formation (Gogolla et al., 2009). Previous studies have found that Wnt7a regulates axonal branching and remodelling (Lucas and Salinas, 1997), while Wnt7b modulates the formation of complex dendritic arborization (Rosso et al., 2005). In response to Wnt7a signals, a decrease of JNK3 palmitoylation and a increase of JNK3 translocation to the actin-cytoskeleton fraction were observed (Figure 27A,B), suggesting that depalmitoylation of JNK3 may participate in the Wnt7a signalling pathway for axonal development regulation. This is supported by our findings showing that JNK3 CS mimics and simulates the effects of Wnt7a on axonal 109  branching (Figure 27). In this condition, overexpressing JNK3 CS may have hijacked endogenous pathways that are normally regulated by Wnt7a-induced JNK3 depalmitoylation. To test this hypothesis, it would be ideal to have a JNK3 that is irreversibly palmitoylated, as it may resist Wnt7a-induced depalmitoylation and thus stop downstream signalling pathways. A COOH-terminal motif of paralemmin which is irreversibly prenylated and dual palmitoylated has been shown to mimic constant palmitoylation on proteins (Kutzleb et al., 1998). This pseudo-palmitoylation is able to restore intracellular trafficking of cytosol protein PSD-95 even in the presence of the palmitoylation inhibitor 2-BrPA (El-Husseini et al., 2002). Overexpression of pseudo-palmitoylated JNK3 Parlm is able to completely abolish the effects of Wnt7a, indicating the involvement of JNK3 palmitoylation in Wnt7a signalling pathways for axonal development. Interestingly, Jnk3-/- mice and neurons overexpressing JNK3 WT (Figure 25) show normal axonal development (Kuan et al., 2003; Hunot et al., 2004). Only manipulations of JNK3 palmitoylation produce abnormal branching phenotypes (Figure 25B,C; 29E). This implies that JNK3 is not a constitutive mediator of axonal development, but may act as an important modulator of axonal development in response to upstream signals via palmitoylation. In sum, our findings reveal a novel isoform-selective mechanism underlying JNK regulation, and provide evidence for a physiological role of JNK3 in axonal branching.  110  Chapter 4. Interaction of zD17 with JNK Mediates Brain Injury in Ischemic Stroke, and Represents a Novel Therapeutic Target for Neuroprotection  A version of this chapter has been published. Guang Yang and Max S. Cynader. 2011. Palmitoyl acyltransferase zD17 mediates neuronal responses in acute ischemic brain injury by regulating JNK activation in a signaling module. Journal of Neuroscience, 31 (33): 11980-91.  111  4.1 Introduction Stroke is a leading cause of death and disability worldwide (WHO, 2007). In acute ischemic stroke, the most common form of stroke, the supply of blood and oxygen to a part of the brain is reduced, which eventually causes the death of brain tissue and forms the infarct core (Donnan et al., 2008). This irreversibly-damaged core is surrounded by a dysfunctional but potentially viable hypoperfused region, known as the penumbra. Within a few hours after ischemia, neurons in the penumbra are challenged by excitotoxic and inflammatory processes resulting in delayed death (Fisher and Ratan, 2003; Bossy-Wetzel et al., 2004).  Zinc-finger DHHC containing 17 (zD17, or known as HIP14) belongs to a family of palmitoyl acyltransferases (PATs) which catalyze protein palmitoylation, a post-translational lipid modification affecting protein trafficking and function (El-Husseini and Bredt, 2002; Singaraja et al., 2002; Huang et al., 2004). Through its regulation of the poly-Q-repeat mutated huntingtin (Htt) protein’s palmitoylation and aggregation, zD17 has been implicated in neurodegeneration in Huntington’s disease (Yanai et al., 2006). However, whether zD17 is also involved in neuronal cell death in the absence of mutated Htt protein is not known. In addition to acting as a PAT, zD17 seems to have other roles (Goytain et al., 2008), including the activation of c-jun N-terminus kinase (JNK) pathways (Harada et al., 2003). JNK activation is broadly involved in excitotoxicity, inflammation, and neuronal cell death in ischemic stroke as well as  112  other neurological diseases (Resnick and Fennell, 2004; Weston and Davis, 2007). It is thus possible that zD17 may contribute to acute neuronal cell death, for example in ischemic stroke, via JNK pathways, rather than via controlling Htt aggregation. In our pilot studies leading to this report, we found that zD17-induced JNK activation is independent of its PAT activity. We thus investigate this PAT-independent role of zD17 in regulating ischemic stroke-induced acute neuronal cell death.  JNK initiates both immediate and late-phase cell death events via phosphorylation and activation of substrates in both the nucleus and cytosol (Bredesen et al., 2006; Centeno et al., 2007; Weston and Davis, 2007). Inhibitors targeting JNK have been shown promising neuroprotection in neurological diseases, despite concerns of side-effects raised in clinical trials (Kuan and Burke, 2005; Bogoyevitch, 2006; Borsello and Forloni, 2007). Studies on the three JNK isoforms have pinpointed different roles of JNK1, JNK2 and JNK3 in neuronal functions and neurological diseases. While JNK2-/-, JNK3-/- or JNK2-/- JNK3-/- mice show reduced vulnerability in models of ischemic stroke, JNK1-/- animals show little protective effect (Hunot et al., 2004). Moreover, the loss of JNK1 leads to disorganization of neuronal microtubules and neurodegeneration (Chang et al., 2003). These results strongly suggest that selective inhibition of stress-responsive JNK isoforms, (primarily JNK2/3) would be beneficial for therapeutic targeting of JNK (Coffey et al., 2002; Bogoyevitch, 2006). However, the design of JNK inhibitors with clear isoform selectivity and the therapeutic application of these inhibitors in ischemic stroke 113  have not been achieved. Here, we report a role of zD17 in mediating ischemic neuronal death by recruiting JNK pathways, and develop a neuroprotective JNK inhibitor with the isoform and scenario-selectivity.  4.2 Results  4.2.1 JNK Activation Is Regulated by the zD17-JNK Signaling Module To examine the impact of PATs on JNK activation, we expressed several neuronal PATs individually with JNK3 in HEK293 cells. Under both resting and osmotic stress conditions, zD17 greatly facilitated JNK3 phosphorylation while other tested PATs showed no effects (Figure 28A, 31). We further examined the involvement of zD17’s PAT activity by employing a PAT activity deficient mutant of zD17 (zD17∆) (Huang et al., 2004). Similar to wildtype zD17, zD17∆ strongly enhanced JNK3 phosphorylation (Figure 28B). These results indicate that zD17 is selectively involved in activating JNK in a PAT activity-independent manner. Because zD17 does not have a canonical kinase domain, we asked whether zD17 might interact directly with JNK and modulate its activity. Using co-immunoprecipitation, we found that JNK3 associated with zD17, but not with other similar PATs (zD15 and zD20) (Figure 28C). This association of JNK3 was independent of zD17’s PAT activity, as both zD17 wildtype and zD17∆ immunoprecipitated JNK3 (Figure 28D). In vitro, purified JNK3 was capable of binding purified 114  myc-zD17 (Figure 28E) and vice versa (Figure 30A). Thus, zD17 is a binding partner of JNK.  Figure 28. Formation of the zD17-JNK3 signaling module. A, ZD17 promotes JNK3 phosphorylation. Under both resting and osmotic stress (400 mM sorbitol, 30 min) conditions, the phosphorylation level of GFP-JNK3 in heterogeneous HEK293 cells is enhanced by co-expression of myc-zD17, but not PAT myc-zD15, zD20, or zD23. B, ZD17 promotes JNK3 phosphorylation in a PAT activity-independent manner. JNK3 phosphorylation is enhanced by both wildtype zD17 (fold change to JNK3 only; 9.98 ± 1.00; P<0.01) and the PAT-activity deficient mutant zD17∆ (10.61 ± 1.09; P<0.01). t-test. Quantifications show means ± s.e.m. C, JNK3 associates with zD17 in HEK293 cells. GFP-JNK3 is present in the zD17 immunopreciates enriched by anti-myc antibody, but is not detected in zD15 or zD20 immunoprecipitates. D, ZD17 associates with JNK3 independent of its PAT activity. Both zD17 wildtype and zD17∆ are capable of interacting with JNK3, as assessed by co-immunoprecipitation. E, Purified GST-JNK3 physically binds to myc-zD17 in vitro. F, ZD17 selectively interacts with MKK7. Only FLAG-MKK7 is present in immunoprecipitates. G, 115  MKK7 contributes to the zD17-mediated phosphorylation of JNK3. Co-expression of FLAG-MKK7ki (kinase inactive mutant) in HEK293 cells reduces the phosphorylation level of GFP-JNK3 promoted by myc-zD17 (fold change to JNK3 only; JNK3+zD17, 2.28 ± 0.23; JNK3+zD17+MKK7wt, 3.10 ± 0.43; JNK3+zD17+MKK7ki, 1.48 ± 0.22, P <0.01 compared to JNK3+zD17). H, ZD17 recruits MKK7 and JNK3 in a signaling module. GFP-JNK3 and FLAG-MKK7 are expressed in the HEK293 cells, with or without myc-zD17. The presence of MKK7 in the JNK3 immunoprecipitates is enhanced by co-expression of zD17 (fold change to JNK3+MKK7; JNK3+zD17+MKK7, 2.41 ± 0.17, P <0.01).  Because zD17 promoted JNK phosphorylation and activity (Figure 30B), we examined the role of several mitogen-activated protein kinase kinases (MAPKKs), upstream activators of JNK, in a potential signaling module (Weston and Davis, 2007). When MAPKKs were expressed in HEK293 cells, zD17 was able to co-immunoprecipitate the JNK activator MKK7. In contrast, the JNK activator MKK4, and the p38 activators MKK3 and MKK6, did not associate with zD17 (Figure 28F). To further confirm the functional involvement of MKK7, we employed kinase-inactive MKK7 (MKK7ki) (Merritt et al., 1999). Since MKK7ki interacted with zD17 similar to MKK7 wildtype (Figure 30C, D), we expressed MKK7ki as a dominant negative, resulting in attenuated JNK3 phosphorylation (Figure 28G). Moreover, the association of JNK3 and MKK7, assessed by co-immunoprecipitation, was strongly enhanced in the presence of zD17 (2.41 ± 0.17 fold, P <0.01) (Figure 28H, 31E). These results indicate that zD17 recruits MKK7 and JNK to form a signaling module for JNK activation.  116  Figure 29. JNK phosphorylation is regulated by PAT zD17. A, zD13 interacts with JNK3, but zD13 can only slightly promote JNK phosphorylation compared with zD17. GFP-JNK3 was transfected with myc-zD13 or myc-zD17. t-test. Quantifications show mean ± s.e.m. * and ** indicate P<0.05 and P<0.01, respectively. B, The effects of zD13 on JNK1 and JNK2 are similar to those on JNK3.  4.2.2 Excitotoxicity Promotes Signaling Module Formation As an important transducer of stress signals in the neuron, JNK is activated in response to various stresses such as excitotoxicity and inflammation (Borsello and Forloni, 2007). We thus asked whether the zD17-JNK signaling module is recruited in these scenarios. Inflammation induced by the cytokines, tumor necrosis factor-α (TNF-α) or interleukin-1β (IL-1β), as well as excitotoxicity induced by glutamate or N-methyl-D-aspartic acid (NMDA) stimulation robustly promoted the zD17-JNK3 interaction in cortical neuronal cultures (Figure 31A, B). Moreover, in comparison with stress-responsive isoforms JNK2 and JNK3, the JNK1-zD17 interaction was less sensitive to either NMDA or glutamate treatment (Figure 31C). Since excitotoxicity-induced JNK activation is predominantly mediated by MKK7 (Centeno et al., 2007), we focused on excitotoxicity as our cellular model to study mechanisms underlying activation of the zD17-JNK  117  signaling module and its contribution to neuronal cell death.  Figure 30. zD17 physically associates with JNK3 and regulates JNK activation in an MKK7-dependent manner. A, zD17 physically binds to JNK3 in vitro. GST-zD17 and GST purified from E.coli. were incubated with purified JNK3 (20 μg/ml), followed by probing with anti-GST (left) or anti-JNK3 (right). B, Co-expressing myc-zD17 promotes GFP-JNK3 phosphorylation and activation in HEK293 cells. Phosphorylation (fold change compared with JNK3 alone; JNK3+zD17, 10.1 ± 0.5, P <0.01; sorbitol, 27.3 ± 1.3; JNK3+zD17+sorbitol, 54.1 ± 2.3, P <0.01 compared with sorbitol alone) and activation (fold change compared with JNK3 alone; JNK3+zD17, 4.8 ± 0.2, P <0.01; sorbitol, 12.4 ± 0.2; JNK3+zD17+sorbitol, 15.3 ± 0.6, P <0.01 compared with sorbitol alone) of JNK3 were tested with anti-p-JNK and kinase assays (KA), respectively. C and D, zD17 interacts with MKK7 wildtype (wt) and the kinase inactive mutant (ki). FLAG-MKK7wt or FLAG-MKK7ki was co-expressed with myc-zD17 in HEK293 cells. Their interaction was examined by co-immunoprecipitation with anti-myc (C) or anti- FLAG (D), followed by detection with anti-FLAG (C) or anti-myc (D), respectively. E, zD17 promotes the formation of the JNK3-zD17-MKK7 module. FLAG-MKK7wt and GFP-JNK3 were expressed in HEK293 cells. The presence of JNK3 in MKK7-containing complex was detected by co-immunoprecipitation as indicated. Co-expression of myczD17 significantly enhances the presence of JNK3 in the complex. F, zD17 promotes JNK3 activation. Compared to GFP-JNK3 only, co-expression myc-zD17 enhances JNK3 phosphorylation in the zD17-containing complex (1st IP with anti-myc). The unbound JNK3 from the supernatant of 1st IP was collected by immunoprecipitated (2nd IP with anti-GFP). Representative figures are from three independent 118  experiments. We examined the activation pathways by which the zD17-JNK signaling module is regulated by excitotoxicity. ZD17 has been shown to interact with many different substrates, and it may be that changes in interaction of zD17 with one or more of its substrates affects its availability for interaction with JNK3 (Huang et al., 2009).  Figure 31. Regulation of the zD17-JNK3 signaling module in response to NMDA-induced excitotoxicity. A, NMDA-induced excitotoxicity promotes zD17-JNK3 interaction in neuronal cultures. NMDA stimulation enhances the zD17-JNK interaction assessed by co-immunoprecipitation (fold change to control. NMDA, 1.49 ± 0.09, P<0.01; NMDA+MK801, 0.96 ± 0.14; P=0.78). B, The cytokines TNF-α and IL-1β enhance the zD17-JNK3 interaction in neuronal cultures. TNF-α (50ng/ml, 6 h) and IL-1β (10ng/ml, 16 h) treatments induced a 1.53 ± 0.11 and 1.88 ± 0.23 fold 119  increase of zD17-JNK3 interaction, respectively. C, Glutamate and NMDA enhance the zD17 interaction with JNK isoform 2 and 3. The enhancement on JNK2 (Fold increase compared with control; Glutamate, 1.7 ± 0.2; NMDA, 2.1 ± 0.4) and on JNK3 (Glutamate, 2.6 ± 0.2; NMDA, 2.5 ± 0.3) are higher than that on JNK1 (Glutamate, 1.4 ± 0.1; NMDA, 1.4 ± 0.1). D, Binding to PSD-95 interferes with the zD17-JNK3 interaction (fold change; PSD-95, 0.82 ± 0.03; P<0.01; PSD-95C3,5S, 1.25 ± 0.07; P<0.05), and reduces JNK3 phosphorylation (fold change; PSD-95, 0.64 ± 0.11; P<0.05; PSD-95C3,5S, 1.19 ± 0.21; P=0.42) in HEK293 cells. E, In neuronal cultures, blocking palmitoylation with 2-bromopalmitate (2-BrPA, 100 μM, 6 h) promotes the zD17-JNK3 interaction (fold change; 2-BP, 1.59 ± 0.05). Disrupting the NR2B-PSD-95 association with NR2B9c (1 μM, 30min before NMDA treatment) reduces the NMDA excitotoxicity-induced zD17 interaction with JNK3, examined 15 min after NMDA challenge (fold change; NMDA, 1.64 ± 0.04; NR2B9c, 1.12 ± 0.1; NR2Baa, 1.48 ± 0.1, P=0.23 to NMDA). t-test. * and ** indicates P<0.05 and P<0.01, respectively. Error bars show means ± s.e.m.  One substrate of zD17, postsynaptic density-95 (PSD-95) is especially relevant, because excitotoxicity has been shown to alter its palmitoylation and protein-interaction profile (Kornau et al., 1995; El-Husseini et al., 2002; Kang et al., 2008). In HEK293 cells, overexpression of PSD-95 impeded the zD17-JNK3 interaction and JNK3 phosphorylation (Figure 31D). This impact of PSD-95 was completely abolished by mutating its palmitoylation sites, which eliminated the interaction with zD17 (Huang et al., 2009). We also used a broad spectrum palmitoylation inhibitor 2-BrPA to inhibit the zD17 interaction with its substrates in cultured neurons (Huang et al., 2009; Yang et al., 2009). This treatment robustly enhanced the zD17-JNK3 interaction (Figure 31E). Excitotoxicity has been shown to mobilize PSD-95 to NMDA receptors (NMDAR), which initiates neurotoxic signaling (Kornau et al., 1995; Aarts et al., 2002). A previously described peptide, NR2B9c, was applied to block the PSD-95-NMDAR association (Aarts et al., 2002). This peptide, but not its mutated control NR2Baa, also impeded  120  the enhancement of the zD17-JNK3 interaction (Figure 31E). Taken together, these results suggest that the PSD-95 may contribute, at least partially, to the regulation of the zD17-JNK interaction in response to excitotoxicity by competing with JNK in binding zD17.  Figure 32. Identifying novel JNK-interacting motifs on zD17 and achieving scenario-selective inhibition of isoform JNK2/3 with peptides derived motif-E. A, JNK-interacting motifs are embedded at the N-terminus of zD17. Purified GST-tagged zD17 fragments are its intracellular domains (CD1, zD171-310; CD2, zD17405-479; CD3, zD17550-632). B, Motif-D, motif-E and the third ankyrin repeat in between (as shown in diagram) were fused with GST and purified. After incubation with JNK3, only motif-D and motif-E bound to JNK3. C, The effect of the peptide NIMoE (derived from motif-E) on the zD17-JNK interaction. Application of NIMoE (1 μM, 30 min before NMDA treatment) blocked NMDA-induced enhancement of the zD17-JNK2/3 interaction. D, The effect of peptides on the zD17-JNK3 interaction. NIMoE and NIMoEsh, but not control peptides NIMoEscr and NIMoEmut, blocks the NMDA-induced enhancement of the zD17-JNK3 interaction. t-test. Quantifications show mean ± s.e.m.  121  4.2.3 Identification of JNK Binding Motifs on zD17 To develop blockers of the zD17-JNK interaction, we first identified two novel JNK binding motifs on zD17. We purified three cytosolic domains (CD1, 2 and 3) of zD17 with a GST tag, and found that, in vitro, JNK3 binding predominantly occurred at CD1 (Figure 32A).  Figure 33. Isoform- and scenario-selective inhibition of JNK by peptides derived from JNK binding motifs on zD17. A-C, Identification of novel JNK-interacting motifs on zD17. A, The in vitro peptide scan of potential JNK-interacting motifs on zD17 N-terminal cytosolic domain. Interaction-positive regions are plotted on the diagram showing the N-terminal part of zD17. Bold black lines with marks (A-F) indicate the potential JNK-interacting motifs from the peptide scan and their locations on zD17 are labelled beside the lines. The ankyrin domains for protein interaction were 122  plotted as rounded rectangles with locations indicated. B, Two motifs located in different ankyrin domains are critical for the zD17-JNK interaction. Myc-tagged full-length zD17 (FL) and fragments of zD17 with assorted deletions are indicated at bottom, and embedded potential interaction motifs (A-F) are labelled. The interaction ability (plus or minus) of these constructs is plotted. C, The locations on zD17 from which peptides NIMoD, NIMoE and NIMoEsh are derived, shown in a simulated structure of zD17 ankyrin repeats. 3D structure data was obtained from National Center for Biotechnology Information (PDB: 3EU9) and processed with PyMOL software (v1.3). D, NIMoE blocks the NMDA-induced enhancement of the zD17-JNK interaction without affecting baseline interaction in neuronal cultures (normalized to control. NMDA, 173.6 ± 16.9%, P<0.05, NMDA+NIMoE, 106.5 ± 1.2%, P =0.16; NIMoE: 103.5.7 ± 6.2%, P=0.64, NIMo-D, 54.1± 16%, P<0.01, t-test). E, The effect of peptides on JNK3 basal activity. JNK3 is immunoprecipitated from cell lysates, followed by the detection of kinase activity with a kinase assay (KA). Pretreating cells with a known JNK inhibitor SP600125 (10 μM) or NIMoD (1 μM) for 2 h caused a significant (fold change to control, 0.56 ± 0.06) or partial (0.72 ± 0.07) reduction of JNK basal activity, respectively. NIMoE (1 μM) has no significant impact on basal activity of JNK3 (0.89 ± 0.10, P=0.33). F, NIMoE abolishes NMDA-induced activation of JNK2 and JNK3 but not JNK1. JNK is immunoprecipitated from the cell lysate, followed by the determination of JNK activity by kinase assays (KA) (normalized to 0 h. JNK1: 0.5 h, 61.0 ± 5.4%, 1h, 137.8 ± 20.8%; JNK1+ NIMoE: 0.5 h, 104.3 ± 2.6, 1 h, 168.5 ± 3.3%; JNK2: 0.5 h, 164.3 ± 11.5%; 1 h, 179.2± 11.2%; JNK2+NIMoE: 0.5 h, 103.7 ± 5.0%; 1 h, 90.9 ± 10.9%; JNK3: 0.5 h, 141.7 ± 7.8%; 1 h, 172.1± 18.9%; JNK3+NIMoE: 0.5 h, 90.6 ± 5.3%; 1 h, 47.0 ± 17.6%). t-test. ** indicates P<0.01. Line graphs show means ± s.e.m.  To obtain further information, we used a peptide array containing 15mers with a 5aa shift to cover CD1 and identified regions on the membrane to which JNK3 bound (Figure 33A). Six potential binding regions in three categories were detected: N-terminus (motifs-A, -B, -C), ankyrin repeats (motifs-D,-E), and submembrane (motif-F). These zones are merely candidate regions and may not reflect actual binding domains. Thus, we generated a series of zD17 deletion mutants and examined their capability of interacting with JNK3 by co-immunoprecipitation (Figure 33B). Deleting CD2 and CD3 did not affect the zD17-JNK3 interaction. Deletion of motif-F did not reduce, but instead increased, the co-immunoprecipitation of JNK3. Removing 123  motif-E diminished the interaction. Strikingly, further deleting motif-D, or selectively removing motifs-D and -E from full-length zD17 completely eliminated the interaction (Figure 33B), suggesting the involvement of these two motifs in binding JNK3. Motifs-D and -E are located within the second and the fourth ankyrin repeats, respectively (Figure 33C). Although zD17 contains five ankyrin repeats, our data indicate that JNK3 binds specifically to motifs-D and -E of zD17 with a preference for motif-D (Figure 32B).  4.2.4 Isoform- and Scenario-Selective Inhibition of JNK We next determined whether targeting these two motifs could inhibit the excitotoxicity-induced zD17-JNK interaction and JNK activation. We synthesized peptides comprising motif-D (TPLHWATRGGHLSMV; Novel Interaction Motif D/NIMoD) and motif-E (MTPLMWAAYRTHSVDPTRLL; NIMoE) and fused them to the cell-membrane transduction domain of the HIV-1 Tat protein to allow the peptides to penetrate the cell-membrane (Schwarze et al., 1999). Based on results from the peptide array, we optimized NIMoE into a shorter 10mer peptide (WAAYRTHSVD, NIMoEsh) which included primarily the loop region of motif-E (zD17196-205) (Figure 33C). BLAST searches showed that the sequence of NIMoE/NIMoEsh is found only within zD17 (Figure 32E), suggesting the specificity of these peptides. Bath application of NIMoD to neuronal cultures significantly diminished the co-immunoprecipitation of zD17 with JNK3, resulting in a reduction of JNK3 basal activity (Figure 33D,E). NIMoE showed no effect on the baseline of the zD17-JNK3 interaction and JNK3 activity, however, 124  NIMoE, but not scramble and mutated controls, selectively blocked the enhancement of the interaction and of JNK3 activity induced by NMDA (Figure 32D, 35D,E). It also eliminated the enhancement of the JNK2-zD17 interaction and JNK2 activity (Figure 32C, 35F). In contrast, NIMoE did not inhibit the normal activation of JNK1 (Figure 33F).  Figure 34. Axonal development with peptide application. A, Incubation with SP600125 affects axon initiation and extension. Axons are labelled with the axon marker Tau-1. Axon initiation rate (normalized to control, SP600125, 18.2 ± 4.1%, n=3) and axon length (control, 161.2 ± 3.9; SP600125, 84.9 ± 1.6%, n=4) are shown. B, NIMoE (4 μM) does not block axon initiation (normalized to control, NIMoE, 98.6 ± 0.2%, n=3, P=0.69) or extension (control, 160.0 ± 8.9; NIMoE, 161.1 ±13.6, n=3; P=0.96). t-test. ** indicates P<0.01. Error bars show means ± s.e.m. Scale bar: 100 μm.  Because one concern with the use of pan-JNK inhibitors as therapeutics is their non-selective inhibition of JNK1, which is essential for normal neuronal development (Oliva et al., 2006), we tested the effect of NIMoE on axonal development, a process that is inhibited by 125  SP600125. In contrast to results with this pan-JNK inhibitor (Figure 34A), axon initiation and extension were not affected by chronic incubation with NIMoE in cultured neurons (Figure 34B).  4.2.5 Effective Neuroprotection against Excitotoxicity by NIMoE  Figure 35. Neuroprotection against NMDA excitotoxicity is achieved by targeting the zD17-JNK interaction. A, NMDA excitotoxicity-induced neuronal cell death is prevented by NIMoE (1 μM). The phase 126  contrast image, PI staining and nuclear Hoechst staining are shown as indicated. The arrow heads indicate the integrity of neurites in the upper panel (phase) and the shape of nucleus in the lower panel (Hoechst). Scale bar, 20 μm. B, Detection of cell death with LDH assays. C, NMDA-induced c-Jun phosphorylation is blocked by NIMoE treatment. C-jun phosphorylation was quantified in (D). E, NMDA-induced caspase-3 cleavage is blocked by NIMoE. Caspase-3 was blotted with an antibody either recognizing both p32 and p17, or only the p17 cleaved form of caspase-3. The p17 form is quantified in (F). One-way ANOVA-test. * and ** indicate P<0.05 and P<0.01, respectively. Error bars show means ± s.e.m. G, Translocation of cytochrome c. (Cyt.c) and Bax are blocked by NIMoE. Cytosolic (Cyto.) and mitochondria (Mito.) fractions were isolated as mentioned in Methods, followed by detection of Cyt.c and Bax with SDS-PAGE.  JNK activation is critical for excitotoxicity-induced neuronal death (Centeno et al., 2007). Application of excitotoxic NMDA causes degeneration of neurites, nuclear condensation, membrane permeabilization, and lactate dehydrogenase (LDH) release in neuronal cultures (Figure 35A,B). Pretreatment with NIMoE or NIMoEsh effectively preserved neuron morphology and significantly prevented excitotoxicity-induced cell death as measured by LDH release and propidium iodide (PI) staining, whereas NIMoEscr, NIMoEmut and a control peptide derived from an unrelated region of zD17 (255NVKGESALDLAKQ267; DIPep1) showed no protective effects in neuronal cultures (Figure 35A,B, 38A,B). NIMoE also protected neurons against a broad range of NMDA concentrations (10 μM to 100 μM) (Figure 36C, D).  JNK2/3 activation mediates neuronal death via two major output pathways: by phosphorylating c-jun to facilitate transcription of pro-death genes, and by activating the mitochondrial caspase-3 pathway to induce apoptosis (Weston and Davis, 2007). NIMoE and NIMoEsh were able to prevent NMDA-induced c-jun phosphorylation (Figure 35C,D, 39A). We 127  found, furthermore, that caspase-3 cleavage was also effectively blocked by NIMoE and NIMoEsh, while control peptides had no effect (Figure 35E,F, 39B). JNK-mediated caspase-3 cleavage depends on translocation of the pro-apoptotic protein Bax to mitochondria and subsequent release of cytochrome c (Cyt.c) (Tsuruta et al., 2004). We found that NMDA-induced translocation of both Bax and Cyt.c was inhibited by NIMoE and NIMoEsh (Figure 35G, 39C).  Figure 36. Effects of peptide application on NMDA excitotoxicity-induced neuronal cell death. A, NMDA-induced neuronal death was prevented with NIMoE/NIMoEsh. 14 DIV rat cortical neuronal cultures were subject to propidium iodide (PI) staining 24 h after NMDA treatment (20 μM for 1 h). In comparison with controls, NMDA toxicity caused a significant destruction of the neurites, a dramatic increase of PI stained cells and shrinkage of the cell nucleus as shown with Hoechst staining. Both the increase of PI-positive cells and nuclear shrinkage were prevented by pre-application of MK801 (10 μM) , JNKI, NIMoE and NIMoEsh. In contrast, control peptides NIMoEscr, NIMoEmut, DIpep1 showed no protective effects. Peptides were applied at concentrations of 1 μM, and cultures were incubated for 30 min before the NMDA insult, 128  followed by PI and Hoechst staining after 24 h. Scale: 20 μm. B, Quantification of LDH release in NIMoEsh protected cultures. NIMoEsh significantly reduced NMDA-induced LDH release (LDH reading, control, 0.23 ± 0.01, NMDA, 0.56 ± 0.02, NMDA+NIMoEsh, 0.21 ± 0.01). C, Dose response of NMDA-challenged neuronal cultures to NIMoE. NIMoE with indicated dosages was applied to neuronal cultures and LDH release was then quantified (NMDA with 0 μM, 0.56 ± 0.03; 0.1 μM, 0.58 ± 0.04, P=0.66 in comparison with 0 μM; 0.25 μM, 0.43 ± 0.29; 0.5 μM, 0.30 ± 0.21; 1 μM, 0.21 ± 0.15; 2 μM, 0.18 ± 0.13). D, NIMoE protects neuronal cultures against a range of NMDA concentrations. Neuronal cultures were pretreated with NIMoE, and then challenged with NMDA at the indicated concentrations. LDH release was examined (NMDA at 0 μM, 0.22 ± 0.01 (-NIMoE), and 0.12 ± 0.01 (+NIMoE); at 10 μM, 0.43 ± 0.02 and 0.20 ± 0.02; at 20 μM, 0.56 ± 0.03 and 0.20 ± 0.02; at 50 μM, 0.60 ± 0.03 and 0.30 ± 0.02; at 100 μM, 0.60 ± 0.02 and 0.40 ± 0.03). t-test. Quantifications show mean ± s.e.m.  4.2.6 Targeting the zD17-JNK Module Protects Brains from Ischemic Stroke We next explored the potential of applying this strategy in vivo in a model of transient ischemic stroke (Longa et al., 1989). Adult male Sprague-Dawley rats were subjected to left middle cerebral artery occlusion (MCAo) for 2 h, followed by 22 h reperfusion (Figure 38A). The zD17-JNK3 interaction in the injured hemisphere (L) remained enhanced up to at least 6 h after the ischemic insult, implying a broad time window for potential intervention. Because of the consideration of its potential for therapeutic applications, we used the shorter form peptide NIMoEsh in our in vivo experiments. A single intravenous injection of NIMoEsh 30 min before MCAo effectively attenuated the enhancement of the zD17-JNK3 interaction, and dramatically reduced the total infarct size (by about 80%; Figure 38B-D).  129  Figure 37. JNK downstream pathways are affected by peptide application. A, NIMoEsh but not NIMoEscr blocks NMDA-induced c-jun phosphorylation in neuronal cultures. 1 μM NIMoEsh or NIMoEscr was added 30 min before the NMDA insult. Cells were collected at the indicated times and phosphorylated c-jun was examined with anti-p-c-jun. B, Activation of caspases-3 is blocked by NIMoEsh but not NIMoEscr. C, The effects of the peptides on translocation of bax and Cyt.c. 21 DIV cultured hippocampal neurons were subject to NMDA insult for 30 min, in the presence of the indicated peptides. Mitochondria were labelled with MitoTracker-TR (red). After fixation, bax and Cyt.c were detected with specific antibodies (green). Scale: 10 μm. We further assessed the effects of post-surgical interventions at 2 h or 4 h. Compared with the infarct volume of non-treated, saline-treated, and scramble peptide-treated groups, NIMoEsh still showed effective protection for the ischemic brain (Figure 38D). The behavioural deficits, as evaluated with neurological scores (Bederson et al., 1986; Watanabe et al., 2004) were substantially reduced by NIMoEsh administration (Figure 38E). The quantitative adhesive  130  removal test (ART) further confirmed a large preservation of somatosensory functions in NIMoEsh-treated rats (Figure 38F) (Bouet et al., 2009). The improvement of behavioural performance persisted over 14 days in ischemic rats injected with NIMoEsh 2 h post-surgery, assessed in a double-blinded manner (Figure 38G,H). Together, these results suggest that targeting the zD17-JNK module to prevent JNK activation is effective in protecting the brain from ischemic injury and in improving behavioural outcomes.  Figure 38. Blocking the JNK-zD17 interaction with NIMoEsh protects from ischemic brain injury in rats. 131  A, The diagram shows the procedures for MCAo experiments. SD rats received sham (control), saline, NIMoEscr, or NIMoEsh by a single i.v. injection at indicated time before or after the onset of ischemia. B, Ischemia induces an enhancement of the zD17-JNK3 interaction, which is blocked by NIMoEsh. Peptide administration 30 min before the ischemic insult significantly blocked the enhancement of interaction induced by MCAo in the injured ipsilateral hemisphere (fold change normalized to the contralateral side; MCAo, 1.64 ± 0.13, P<0.05; MCAo+NIMoEsh, 0.85 ± 0.15, P=0.33). t-test. C, Representative TTC stained brain sections from saline and NIMoEsh treated (30 min before MCAo onset) groups. D, Quantification of total infarct size and volume (inset bar-chart). NIMoEsh treatments (-0.5 h, 43.43 ± 9.82, n=7; +2h, 41.95 ± 8.22, n=10; +4 h, 78.74 ± 28.50, n=6; F=20.71, P<0.01 compared with control groups) dramatically reduced total infarct volume, while saline or NIMoEscr had little effect (control, 240.70 ± 35.67, n=9; saline, 268.89 ± 40.47, n=4; NIMoEscr, 212.60 ± 37.6, n=5; F=0.38, P=0.69). One-way ANOVA-test. E, Neurological scores indicate an improved behavioural outcome in ischemic rats treated with NIMoEsh. Neurological functions were examined 22 h after the onset of MCAo. Saline or NIMoEscr treated groups showed similar deficits (control, 16.43 ± 0.95, n=7; saline, 18.00 ± 1.35, n=4; NIMoEscr, 18.60 ± 1.24, n=5; F=1.65, P=0.23). Administration of NIMoEsh significantly reduced neurological scores (-0.5 h, 8.00 ± 0.72, n=7; +2 h, 8.50 ± 0.75, n=10; +4 h, 11.20 ± 1.49, n=6; F=21.98, P<0.01 compared with control groups). One-way ANOVA-test. F, NIMoE treatment improves somatosensory functions as assessed with the adhesive removal test (ART). MCAo eliminates responses on the contralateral side in control, saline-treated and NIMoEscr-treated animals (control, 13.57 ± 4.30 sec, n=7; saline, 7.42 ± 3.21 sec, n=4; NIMoEscr, 10.80 ± 2.06 sec, n=5; F=0.64, P=0.54). Pre-treatment or post-treatment infusion of NIMoE led to an improvement in somatosensory functions (-0.5 h, 48.62 ± 8.24 sec, n=7; +2h, 62.1 ± 8.60 sec, n=10; +4 h, 60.83 ± 8.13 sec, n=6; F=11.28, P<0.01). One-way ANOVA-test. G, H, Functional tests show improved behavioural outcomes in NIMoEsh-treated animals after MCAo injury. NIMoEscr or NIMoEsh were given 4 h after the MCAo onset. Neurological scores (NIMoEscr, n=8; day-1, 19.25 ± 0.80; day-3, 13.63 ± 1.03; day-7, 7.13 ± 0.58; day-14, 4.38 ± 0.50; NIMoEsh, n=7; day-1, 11.43 ± 1.96; day-3, 5.86 ± 1.14; day-7, 1.71 ± 0.47; day-14, 0.85 ± 0.34) and ART (NIMoEscr, n=8; day-1, 12.63 ± 3.14; day-3, 34.13 ± 6.88; day-7, 59.13 ± 5.67; day-14, 81.88 ± 4.44; NIMoEsh, n=7; day-1, 52.57 ± 6.82; day-3, 73.86 ± 9.63; day-7, 94.86 ± 7.33; day-14, 101.43 ± 3.35). t-test. Bonferoni correction. * and ** indicates P< 0.05 and P< 0.01, respectively. Error bars show means ± s.e.m.  4.3 Discussion ZD17 is a PAT which catalyzes a post-translational lipid modification known as 132  palmitoylation. Although much is known about the broad effects of palmitoylation on protein trafficking, stability, and function, our understanding of the pathophysiological roles of PATs in the nervous system remains poor (Bijlmakers and Marsh, 2003). In addition to its well-studied function in protein palmitoylation, we report in this chapter that PAT zD17 possesses a novel function which appears unique in the PAT family. Via its ankyrin domain, a domain used for protein-protein interaction and signaling, zD17 interacts with JNK and regulates neuronal cell death in acute ischemic stroke. By manipulating this novel zD17-JNK interaction, we provide a therapeutic strategy for acute ischemic brain injury with high efficacy and the potential for reduced side-effects (Figure 39).  4.3.1 The zD17-JNK Interaction as a Therapeutic Target for Neuroprotection Our in vivo and in vitro assays indicate that zD17 is a binding partner of JNK. One of JNK’s activators MKK7, a stress mediator, is also selectively and functionally associated with zD17 (Figure 28D). This suggests that MKK7-zD17-JNK forms a signalling module for JNK activation in response to stressors, including the ischemic stress that we report in this chapter. Similar to zD17, several JNK-interacting proteins, or JNK scaffold proteins, have been suggested to play a role in neurological diseases (Morrison and Davis, 2003; Weston and Davis, 2007). Using these proteins as therapeutic targets has attractive advantages as it may inhibit JNK activation in a scenario-selective manner and thus reduce side-effects caused by global JNK inhibition. Despite the identification of JNK-binding motifs from several JNK scaffold proteins 133  (Morrison and Davis, 2003), successful manipulation of their interactions with JNK, and blockade of pathological activation of JNK has not been achieved. In the case of the newly developed peptide inhibitor JNKI, the JNK-binding motif on JIP, JNK-interacting protein, was designed to block the JIP-JNK interaction (Borsello et al., 2003). However, it was found to be more effective in blocking the interaction of JNK with its substrate c-jun and showed no effect on JNK activation (Borsello et al., 2003). This may be because the JNK-binding motif on the scaffold protein JIP shares the same sequence with the JNK substrate c-jun (Borsello et al., 2003). In contrast, the JNK-binding motifs that we have identified here appear to be unique to zD17. As we demonstrate with the peptide NIMoE, this provides an opportunity to manipulate JNK activation selectively under stress conditions.  Figure 39. Potential mechanisms of zD17-mediated JNK activation and the intervention for neuroprotection. A, In response to excitotoxic and other stresses, zD17 interacts with JNK and recruits MKK7 to activate JNK (primarily JNK isoforms 2 and 3) by phosphorylation. Activated JNK turns on downstream cell death pathways in the nucleus and mitochondria. Excitotoxicity promotes the 134  zD17-JNK interaction partially through the dissociation of zD17 from PSD-95 that competes with JNK in binding zD17. Other pathways that may contribute to the regulation of the JNK-zD17-MKK7 module remain to be elucidated. B, The tat-fused peptides (NIMoE/NIMoEsh) derived from the JNK-binding motif on zD17 blocks the excitotoxic-induced interaction of zD17 and JNK, and thus prevents JNK activation and neuronal cell death.  Another challenge for JNK inhibition as a neuroprotective therapy for ischemic stroke or other neurological diseases is JNK isoform selectivity (Bogoyevitch, 2006). Although the advantage of isoform-selective JNK inhibition has long been appreciated, the application of small molecule inhibitors with this feature in treating ischemic stroke has not been reported (Resnick and Fennell, 2004; Bogoyevitch, 2006). The development of small molecules with isoform selectivity has proven challenging since JNK isoforms are highly conserved at the ATP-binding sites. An alternative strategy is to take the advantage of JNK interacting proteins, some of which have been shown to have selective binding preferences to JNK isoforms (Kelkar et al., 2000; McDonald et al., 2000). Using these proteins as targets, however, no JNK isoform-selective inhibition has been successfully developed as a therapeutic strategy in neurological diseases.  ZD17 shows several attractive attributes to make it an appealing target. First, the zD17-JNK interaction responds to diverse neuronal stresses including excitotoxicity and inflammation (Figure 31), implying that the zD17-JNK signaling module may contribute to JNK activation at different stages in the ischemic process. In fact, an increase of the zD17-JNK interaction is detected up to at least 6 h after ischemia onset (Figure 38), when the stage of excitotoxicity is 135  already believed to be waning (Lo, 2009). This suggests a broad time window for therapeutic intervention. Second, the activation mechanism of the zD17-JNK signaling module suggests a central position of zD17 in sensing upstream stress signals and generation of outputs to JNK pathways. As we shown with PSD-95, substrate binding represents one candidate mechanism that links excitotoxicity to JNK activation via zD17 (Figure 31). Other mechanisms may also contribute to the regulation of zD17-mediated JNK activation. It is interesting that another PAT, zD13, is also able to interact with JNK though it is not efficient as zD17 in promoting JNK phosphorylation (Figure 29). Further studies on the role of zD13 and other potential regulators on JNK activation would bring more insights on the regulatory mechanism of the zD17-JNK signaling module. Third, the stress-induced enhancement of the zD17-JNK module shows a marked preference for JNK isoforms 2 and 3. Fourth, activation of JNK2/3 and downstream cell death pathways are largely mediated by the zD17-JNK module, and finally, a specific JNK-binding motif on zD17 (motif-E) is selectively recruited to enhance the zD17-JNK interaction under stress conditions.  By targeting zD17 with peptides derived from motif-E (NIMoE/NIMoEsh), we show that i) JNK can be inhibited with selectivity to isoforms JNK2/3; ii) inhibition of JNK is only effective when the zD17-JNK interaction is enhanced under stress conditions; iii) normal functions of JNK pathways are preserved, as we assessed with studies of axonal growth; iv) excitotoxicity-induced neuronal cell death is markedly prevented in cultured neurons; v) brain 136  damage and behavioural deficits in rats subjected to ischemia are substantially reduced over a broad time window. These studies establish the zD17-JNK3 interaction as a candidate target with reduced side-effects for ischemic stroke. However, several further steps, such as recruitment of additional stroke models and different animal species, are still required to more fully validate the therapeutic potential of this new target (STAIR, 1999).  4.3.2 A Novel Function of zD17 Independent of its PAT Enzyme Activity How are the multiple functions of zD17 connected to contribute to neuronal cell death? By studying activation mechanisms of the zD17-JNK module in excitotoxicity, we provide some clues on this issue. Although zD17-promoted activation of JNK is independent of PAT activity, substrate binding to zD17, which is dependent on the palmitoylation status of the substrates, regulates the zD17-JNK module and activation of JNK. This suggests that the different functions of zD17 are systematically connected, and may directly or indirectly contribute to the pathogenesis of neurological diseases. Altered palmitoylation and interaction profiles of zD17 substrates, as well as activation of JNK pathways, have been noted in several neurological diseases. Further investigations on how zD17 relates to its substrates to regulate JNK activation in different pathological conditions may lead to new approaches to treat neurological diseases.  137  Chapter 5. Conclusion  138  5.1 Summary of Findings  5.1.1 Role of AMPAR Palmitoylation in Neuronal Plasticity Neuronal plasticity requires a rapid regulation of synaptic transmission and structural organization of synapses (Barry and Ziff, 2002). Trafficking of AMPARs to and from postsynaptic membranes is one of the key regulatory steps for establishing neuronal plasticity (Bredt and Nicoll, 2003; Malenka, 2003; Collingridge et al., 2010). My research has identified that palmitoylation is a potential mechanism for regulating AMPAR subunits and may contribute to neuronal plasticity. By assessing the sensitivity of palmitoylation of GluR1 and GluR2 subunits to reagents that differentially affect protein synthesis and secretion, I have found that AMPAR palmitoylation shows subunit-selective characteristics. While GluR1 subunits are palmitoylated along the anterograde secretion pathway rapidly after being synthesized, GluR2 subunits are predominantly palmitoylated in ER as immature receptors. Given the continuity of ER and Golgi membrane systems, a constant cycling of enzymes between ER and Golgi may be important for the balance of GluR2 palmitoylation, since blocking such cycling by destructing microtubules, which prevents enzymes flowing back to ER and accumulates these enzymes in Golgi, dramatically reduces GluR2 palmitoylation. I have also shown that GluR1’s are able to be palmitoylated on the PM. In contrast, palmitoylation of GluR2 subunits on the PM is not detectable in same experiment conditions. Consistent with these findings, majority of GluR2 subunits have been shown to be retained in ER after being synthesized and become heteromers 139  with GluR3 subunits. This fraction of GluR2 seems to be the major portion for palmitoylation as depletion of GluR1-associated GluR2 increases the average level of palmitoylation of GluR2 subunits.  What is the functional role of AMPAR palmitoylation? It has been suggested that palmitoylation controls AMPAR trafficking from Golgi to the PM (Hayashi et al., 2005). Depalmitoylation at the TMD2 of AMPAR subunits enables their release from Golgi whereas palmitoylation of GluR1 C-terminus triggers endocytosis machinery (Hayashi et al., 2005). This is supported by my discovery that GluR1 subunits are palmitoylated at multiple sites along the secretion pathway. Given GluR2’s predominantly reside and are palmitoylated in ER, I have shown that palmitoylation contributes to the early regulation of GluR2 trafficking and stability. The TMD2 of GluR2, but not GluR1, is critical for this regulation. AMPARs are sorted for lysosomal degradation if GluR2’s are not palmitoylated after being synthesized. Palmitoylation seems to serve as a quality control mechanism that tags on GluR2 subunits and decides the destiny of AMPARs. Consistent with differential properties and functions of palmitoylation of AMPAR subunits, I have found that GluR1 and GluR2 subunits respond differently to synaptic activity. While GluR1 palmitoylation is rapidly altered by synaptic stimuli, GluR2 palmitoylation shows a gradual decrease in response to NMDA stimulus that induces loss of synaptic AMPARs. Together these findings indicate that AMPAR GluR1 and GluR2 subunits have distinct palmitoylation properties, that palmitoylation may regulate AMPAR trafficking from different 140  aspects on AMPAR subunits, and that palmitoylation provides a potential mechanism for neuronal plasticity regulation.  5.1.2 Role of JNK Palmitoylation in Neuronal Development JNK family members have been broadly associated with various physiological and pathological contexts in the brain, including neurite development, synaptogenesis, stress response, and neuronal cell death etc (Davis, 2000; Weston and Davis, 2007). Increasing evidence has highlighted that JNK1, JNK2 and JNK3 isoforms are preferentially involved in these biological processes (Bogoyevitch, 2006). However, regulatory mechanisms ensuring such isoform-specific functions are poorly understood. My results have suggested that palmitoylation is a potential isoform-selective mechanism for such regulation. In contrast to most JNK isoforms, only JNK3 p54 isoform with an extended C-terminus is palmitoylated in our experimental conditions. While JNK1 and JNK2 are ubiquitously expressed, JNK3 is predominantly detected in the heart, testis and the brain. A number of studies have confirmed a role of JNK3 in mediating neuronal stress response and cell death processes. However, JNK3 palmitoylation does not directly affect stress-induced JNK3 activity and functions, as the palmitoylation-deficient mutant shows similar stress response as the wildtype JNK3. Instead, palmitoylation regulates the intracellular distribution of JNK3. Non-palmitoylated JNK3 shows higher level in association with the actin cytoskeleton, while over-palmitoylated JNK3 becomes more soluble. A variety of proteins that are involved in different aspects of neuronal development have been identified as 141  JNK subunits (Bogoyevitch and Kobe, 2006). JNK has also long been implicated in regulating cytoskeleton organization (Chang et al., 2003; Oliva et al., 2006; Ciani and Salinas, 2007). As critical kinases involved in diverse processes, JNK isoforms must be tightly regulated with temporal and spatial specificity. My findings suggest that palmitoylation may provide a potential way for such regulation in an isoform-selective manner.  In accord with its functions in directing JNK3 intracellular distribution, I have found that JNK3 palmitoylation is important for axonal branching, a cytoskeleton-regulated process. Introducing the palmitoylation-deficient JNK3 mutant into hippocampal neurons at an early development stage dramatically enhances axonal branching. This increase is not the result of net increase of the density of axonal filopodia. However, the motility of these filopodia is enhanced with this JNK3 mutant. This may reflect that the growth machinery is accelerated, or the stabilization of filopodia may be affected which results in abnormal growth of unstable protrusions. In contrast, expression of a JNK3 mutant mimicking constant palmitoylation (JNK3-Palm) significantly reduces axonal branching compared to wildtype JNK3, supporting the role of JNK3 palmitoylation in regulating axonal development.  To extend the understanding of the role of JNK3 palmitoylation in axonal development, I have further examined the impact of Wnt signaling pathways. Wnt has been shown to regulate dendritic development via JNK-dependent pathways (Rosso et al., 2005). Wnt7a reduces JNK3 142  palmitoylation in hippocampal neurons and correspondingly alters JNK3 distribution on the actin cytoskeleton. While Wnt7a is able to promote axonal branching, expression of the palmitoylation-deficient JNK3 mutant mimics this promotion and prevents further enhancement induced by Wnt7a, and expression of JNK3-Palm completely blocked axonal branching even in the presence of Wnt7a. These results support a role of JNK3 palmitoylation in modulating Wnt7a-mediated axonal branching. Together, these findings suggest palmitoylation as a novel mechanism for regulating JNK3 in an isoform-selective manner and suggest a role of JNK3 palmitoylation in regulating axonal development.  5.1.3 Role of PAT zD17 in Neuronal Cell Death PATs are enzymes catalyzing protein palmitoylation. Recent studies have highlighted that certain PATs also obtained other functions in addition to serving as enzymes for protein palmitoylation (Goytain et al., 2008). PAT zD17 has been shown to activate the JNK pathway via an unknown mechanism (Harada et al., 2003). Because I have found that JNK3 is a palmitoyl protein and palmitoylation does not affect its activity, I wonder if zD17 may regulation JNK activity via a PAT activity-independent mechanism. By assessing the activity of JNK3 wildtype or palmitoylation-deficient mutant in the presence of zD17 wildtype or the enzyme activity-deficient zD17 mutant, I have confirmed that zD17-mediated activation of JNK3 is independent of zD17’s PAT activity. Rather than acting as a PAT, zD17 directly interacts with JNK and its upstream activator MKK to form a signaling module for activating JNK. ZD17 143  selectively recruits MKK7, one of two JNK activators, and this recruitment is important for zD17-mediated JNK activation. MKK7 has been associated with excitotoxicity and inflammatory stress-induced JNK activation (Kuan et al., 2003). Indeed, the formation of the zD17-JNK signaling module is significantly promoted by excitotoxicity and inflammatory factors. This suggests that zD17 may mediate JNK activation in response to various stress signals. Moreover, under stress conditions, zD17 shows preferences in binding stress-responsive JNK isoforms JNK2 and JNK3. This suggests that zD17 may play a special role in mediating pathological activation of JNK2/3 in stress circumstances.  To understand how this signaling module is regulated, I have focused on excitotoxicity because one of zD17 substrates, PSD-95, has been shown to play a role in excitotoxicity-induced neuronal cell death (Aarts et al., 2002). Interestingly, I have found that PSD-95 compete with JNK3 in binding zD17. Excitotoxicity induces the PSD-95 interaction with NMDAR subunit NR2B, which may increase the availability of zD17 for JNK3 interaction. This idea is supported by the result showing that blocking NMDA receptor-PSD-95 interaction impedes the excitotoxicity-induced JNK3-zD17 interaction. In other circumstances, the formation of the zD17-JNK3 signaling module may be regulated by other mechanisms.  To test whether zD17-mediated JNK3 activation is important for neuronal cell death under pathological conditions, I have mapped JNK3 binding sites on zD17, and identified two novel 144  JNK-interacting motifs, which are different from previous known JNK-binding motifs. By using peptides derived from these motifs to compete with endogenous JNK in binding zD17, I have found that one of the motifs (motif-D) is important for constant JNK binding, whereas the other one (motif-E) is only recruited in stress circumstances. This suggests that motif-E may be designed for maximizing pathological JNK activation. Application of peptides derived from motif-E (NIMoE/NIMoEsh) show no effect on basal JNK activity and related functions, such as neurite development. However, blocking the zD17-JNK interaction with these peptides robustly prevents excitotoxicity-induced activation of JNK2/3, activation of downstream pathways including c-jun phosphorylation and caspase-3 cleavage, and correspondingly neuronal cell death. These results indicate that zD17 is a critical player in neuronal cell death via regulating JNK pathways through the formation of the zD17-JNK signaling module.  5.1.4 Potential Therapeutic Target for Neuroprotection Given the importance of the zD17-JNK signaling module in mediating stress-induced neuronal cell death in cultured neurons, I have further examined the involvement of the zD17-JNK interaction in a rat model of ischemic stroke. MCA occlusion causes the massive release of excitotoxic glutamate, followed by gradual increase of inflammatory factors. Correspondingly, the zD17-JNK3 interaction is enhanced for at least 6 hours after the MCAo onset. Blocking the pathological enhancement of the zD17-JNK3 interaction by NIMoEsh prevents MCAo-induced neuronal cell death, tissue damage and behavioral deficits. These results 145  suggest that the zD17-JNK signaling module also play an important role in mediating cell death under pathological conditions, at least in ischemic stroke. Application of NIMoEsh does not affect the basal interaction between zD17 and JNK3. This special property provides a potential way to specifically modify JNK activity in pathological conditions. Together with the discovery of the interaction preference with JNK2 and JNK3, two stress-responsive JNK isoforms, my findings suggest that the zD17-JNK signaling module is a potential therapeutic target for neuroprotection. JNK has long been considered as a drug target for stroke and neurodegenerative diseases. However, inhibitors without isoform selectivity show side-effects that hamper their clinical applications. Targeting the zD17-JNK interaction on the motif-E, as I have demonstrated using the peptide NIMoEsh, could offer a way to inhibit JNK activity in an isoform- and activation scenario-selective manner.  5.2 Future Directions  5.2.1 Physiological and Pathological Roles of Palmitoyl-Proteins in the Brain Recent studies using large-scale screening technologies have identified hundreds of palmitoyl proteins in the brain and other cells (Kang et al., 2008), suggesting a broad impact of palmitoylation on cellular and physiological functions in the brain. Indeed, studies focusing the particular neuronal proteins have highlighted critical roles of protein palmitoylation in regulating 146  synaptic transmission, axonal transport, neuronal development, and cell death etc (El-Husseini and Bredt, 2002). In this thesis, I have studied how palmitoylation play a role in regulating AMPAR subunits and JNK3, and their potential physiological implications. Although the functions of palmitoylation have been studied with many proteins, more newly identified neuronal palmitoyl proteins are still waiting for investigation. How many proteins are palmitoylated in neuron and other brain cells? What are the molecular functions of palmitoylation on these proteins? How does palmitoylation of these proteins contribute to physiological and pathological processes in the brain? In addition to focusing on a single palmitoyl protein, the combination of proteomics (such as the ABE-MS method) and molecular and cellular biology techniques would help to answer these questions.  Compared to palmitoylation substrates, the physiological and pathological roles of PATs in the brain are poorly understood. The most studied mammalian PATs are zD3, zD13 and zD17. The difficulty of identifying their physiopathological roles in the brain is partially due to ambiguous substrate selectivity. The improvement on substrate identification techniques would help solve this problem. On the other hand, together with recent studies on zD17 and some other PATs, my research has highlighted that certain PATs may also possess additional functions rather than serving as a PAT for palmitoylation. Understanding whether other PATs also have PAT activity-independent functions would help depict their roles in the brain.  147  Interesting questions raised with additional functions of PATs are that how these different functions of one PAT are inter-connected with each other and how these functions are integrate for certain biological processes in the brain. For example, zD17 is able to palmitoylate JNK while it also mediates JNK activation which is independent of palmitoylation status. How zD17-mediated JNK palmitoylation and activation is integrated and how these functions contribute to neuronal development and cell death are still unknown. Further investigation on this would help better understanding the roles of PATs in brain functions.  5.2.2 Drug Discovery Based on Palmitoyl Proteins Increasing evidence supports a critical role of palmitoylation in regulating brain functions (El-Husseini and Bredt, 2002). Altered palmitoylation profiles would be expected in brain diseases as abnormal function of certain PATs may contribute such alteration. For example, loss of zD17-mediated palmitoylation of Htt is thought to partially contribute to the progression of HD (Singaraja et al., 2002). Thus, PAT-selective modulators (stimulators or inhibitors) may be useful as potential therapies. However, little is known about how PATs are regulated. Phosphorylation, as well as other modifications may be involved in regulating PATs. In the case of zD17, I have shown that it directly interact with JNK and regulates its activity. It is possible that JNK is able to modulate zD17 PAT activity by phosphorylating zD17. Indeed, several PATs contain potential sites for phosphorylation as predicted by bioinformatics methods. Depicting the mechanisms underlying PAT regulation would help identifying potential drug targets for brain 148  diseases.  Moreover, as I have shown that the zD17-JNK interaction is critical for JNK activation under pathological conditions, it is possible to screen libraries of small molecules which may be used as therapeutics for stroke and other neurodegenerative diseases with high efficacy and the potential for reduced side-effects. This raises the idea that additional functions of PATs may provide new drug targets for brain diseases.  5.3 Conclusions Protein palmitoylation is an important post-translational modification for regulating protein functions. Studies on neuronal palmitoyl proteins have greatly expanded our understanding of the molecular mechanisms of brain physiology and pathology. Focusing on AMPAR subunits and JNK isoforms, my research further emphasized a broad role of palmitoylation in neuronal development and function, and suggests protein palmitoylation as a sensitive and selective way for protein regulation. As a special group of palmitoyl proteins, PATs have long been suggested to play critical roles in brain functions, though few studies elucidate their pathophysiological functions in the brain. My research on PAT zD17 has demonstrated the contribution of zD17 in mediating neuronal cell death via JNK signaling pathways in ischemic stroke, which further supports the pathophysiological importance of PATs in the brain. Moreover, it highlights that 149  certain PATs may possess additional functions independent of their palmitoyl acyltransferase activity. 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Neuron 47:629-632. Zacharias DA, Violin JD, Newton AC, Tsien RY (2002) Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells. Science 296:913-916. Zhang J, Planey SL, Ceballos C, Stevens SM, Jr., Keay SK, Zacharias DA (2008) Identification of CKAP4/p63 as a major substrate of the palmitoyl acyltransferase DHHC2, a putative tumor suppressor, using a novel proteomics method. Mol Cell Proteomics 7:1378-1388.  172  Appendices A1. Reagent and Samples  Cell Cultures. All animal experiments were approved by the Animal Care Committee of the University of British Columbia. Primary cortical or hippocampal neuronal cultures were prepared from Wistar rat (UBC Animal Care Centre, Vancouver, BC, Canada) brains at embryonic days 18–19. Cultures were maintained in Neurobasal medium supplemented with B27 and 0.5 mM glutamax (Invitrogen) as described previously . For biochemical studies, cortical neurons were seeded in six-well culture plates with a density of 7.5 x 105 cells per well. For LDH assays and cell death detection, 24-well culture plates were used with a neuron density of 2.5 x 105 cells per well. Neurons of 14–17 days in vitro (DIV) were used for experiments. For immunostaining, hippocampal neurons were seeded on glass coverslips in twelve-well plates with a density of 1.25 x 105 cells per well. To assess axonal development and synapse formation, hippocampal neurons at 0 DIV and 21 DIV, respectively, were treated with 2 μM NIMoE for 3 d (media were changed daily to maintain peptide concentration relatively constant). To assess axonal development, hippocampal neurons at 5 DIV were transfected with pDsRed2 plus other indicated constructs at a 3:1 ratio with Lippofectamin2000 (Invitrogen). Recombinant Wnt7a proteins (R & D Systems, MN, USA) was added at 5 DIV for 48 h. At 7 DIV, neurons were fixed, stained and analyzed. HEK 293 cells were maintained in DMEM (Invitrogen) supplemented with 10% fetal bovine serum and 1%  173  penicillin/streptomycin. For protein expression in cell lines, cells were transfected with Calcium Phosphate Transfection reagent (Promega, San Luis Obispo, CA, USA) and used 24 h later as described previously.  Materials. [9,10-3H] Palmitic acid (57 Ci/mmol) and biotin-BMCC were purchased from PerkinElmer Life Sciences. L-JNKI, brefeldin A (BFA), nocodazole (NOC) and chloroquine were obtained from Alexis Biochemicals (San Diego, CA, USA). MG-132, ALLN, leupeptin, and cycloheximide (CHX), and the actin depolymerizers cytochalasin D (Cyt.D, 1 µM) and latrunculin A (Lat.A, 1 µM) were obtained from Calbiochem (San Diego, USA). Endoglycosidase H (EndoH) and N-Glycosidase F (PNGF) were from NEB (Ipswich, MA, USA). Thrombin was obtained from BIOPUR (Bubendorf, Switzerland). Bismaleimidohexane (BMH) was from Pierce (Rockford, IL, USA). CNQX and APV were from Ascent Scientific (Princeton, NJ, USA). Optiprep was purchased from Axis Shield (Dundee, Scotland). Other assay reagents without indication were obtained from Sigma.  Plasmids. The GST-tagged full-length and fragments of zD17 constructs were created by cloning zD17 and fragment cDNA sequences into a pGEX-4T-1 vector with a myc epitope sequence at the N-terminus. The GST-JNK3 construct was cloned into the same vector. Myc-tagged fragments of zD17 were also cloned into pcDNA3 to generate expression constructs. GFP-JNK1 and GFP-JNK2 expression constructs were created by cloning cDNA sequences into pEGFP-C1 174  plasmids. The vectors encoding myc-zD13, myc-zD15, myc-zD17, myc-zD17∆, myc-zD20, myc-zD23, GFP-PSD95, GFP-PSD95C3,5S, GFP-SNAP25 and HA-SYTI were generously provided by Dr. El-Husseini’s lab. The GFP-JNK3 construct was a gift from Dr. Vsevolod V. Gurevich. The AMPAR palmitoylation deficient mutant constructs (Hayashi et al., 2005) were a gift from Dr. R. Huganir. The constructs expressing FLAG-JNK3; FLAG-MKK3, FLAG-MKK4, FLAG-MKK6, FLAG-MKK7wt, and FLAG-kiMKK7 were purchased from Addgene.  Antibodies. The primary antibodies used to detect JNK1, activated-caspase3, β-actin and FLAG (DDDDK) epitopes were obtained from Abcam (Cambridge, MA). Anti-zD17, anti-synaptophysin and anti-GFP were purchased from Sigma. Anti-GST and anti-Tau1 were purchased from Abm (Richmond, BC, Canada) and Millipore, respectively. The antibody against PSD-95 was obtained from Synaptic System (Goettingen, Germany) or ABR (Golden, CO, USA). Rabbit polyclonal antibody against synaptotagmin I was purchased from Synaptic Systems. Purified IgG was supplied by Jackson ImmunoResearch. We used rabbit polyclonal antibodies against rat GluR1 or GluR2 as described previously (Ahmadian et al., 2004). All other primary antibodies were purchased from Cell Signalling Technology (Danvers, MA). Secondary antibodies were HRP-conjugated anti-mouse (Cell Signalling Technology), anti-rabbit (Perkin Elmer), anti-GFP (Santa Cruz Biotechnology), and anti-myc (Invitrogen). Fluorescent Alexa 488- and Alexa 568-conjugated anti-mouse IgG1, IgG2a and anti-rabbit IgG were obtained from Invitrogen.  175  A2. Experiment Procedures Protein Preparation and Western Blotting. Cultured neurons were collected by adding 1 x loading buffer containing 10 % glycerol, 50 mM Tris-HCl (pH 6.8), 2 % SDS and 0.01 mg/ml bromophenol blue directly with plastic scrapers. Samples were then boiled for 8 min and analyzed with SDS-PAGE gel. For immunoprecipitation, neurons were washed with ice-cold PBS and collected by centrifugation at 5,000 x g. The pellets were then resuspended in 1 ml of lysis buffer containing 50 mM Tris-HCl (pH 7.4), 1 mM EDTA, 1 mM EGTA, 150 mM NaCl, 1% Triton X-100 and 0.1% SDS supplemented with Complete Protease Inhibitor Tablets (Roche Applied Science, Indianapolis, IN, USA) and 1mM PMSF. For co-immunoprecipitation, cultured cells were washed with ice-cold PBS followed by incubation with 0.5 ml Gentle Lysis Buffer (GLB) containing 25 mM Tris-HCl (pH 7.4), 2 mM EDTA, 1 mM EGTA, 10 mM NaCl, 0.5% Triton X-100 and 10% Glycerol supplemented with the Complete Protease Inhibitor Tablets and 1mM PMSF. Total protein concentration was determined with the BioRad Protein Assay kit. Cells were homogenized by passing through a 28G1/2” needle 10 times on ice. Insoluble material was removed by centrifugation at 10,000 x g for 15 min at 4 °C. Lysates were first precleaned by incubating with Protein A-Sepharose beads (Roche Applied Science) for 1 h at 4°C, followed by incubation with indicated antibodies (>16 h at 4 °C) and protein A Sepharose beads (2 h at 4°C). Lysates of rat brain tissues were precleaned with IgG and protein A-beads.The supernatant was then sequentially incubated with primary antibodies (>16 h at 4 °C). After the addition of 50 µl of protein A (or G)-Sepharose bead, samples were incubated 176  for >16 h at 4°C. Immunoprecipitates were washed three times with lysis buffer; boiled in 2 x loading buffer with 1 mM dithiothreitol (DTT) for 2 min; and analyzed with SDS-PAGE as described previously (Kang et al., 2004). For autoradiography, protein samples were separated by 8% SDS-PAGE and incubated with Amplify reagent (Amersham Biosciences, Sydney, Australia). Gels were dried under vacuum and exposed to Hyperfilm (Amersham Biosciences) with intensifying screens at -80 °C for 30~50 days. The relative intensities of the bands on autoradiograms and Western blots were analyzed and quantified using Image J (NIH).  3  H-Palmitate Metabolic Labelling. To label palmitoylated proteins, cultured neurons were  incubated in Neurobasal medium containing 0.8 mCi/ml 3H-palmitate for 3 h or 24 h. 2-bromopalmitate (2-BrPA, 100 µM) was added in the medium 2 h before labelling to inhibit palmitoylation (Yang et al., 2009). For pulse-chase experiments, neurons were first labelled with 3  H-palmitate for 3 h, and then incubated for indicated periods in conditioned Neurobasal medium  supplemented with unlabelled palmitate (100 µM). JNK3 was then immunoprecipitated and incubated with or without 1M hydroxylamine (HAM) for 1 h at RT before analysis with SDS-PAGE. To block neuronal activity, cells were pretreated with CNQX (100 µM) and APV (100 µM), or TTX (100 nM) for 2 days before labelling (Kang et al., 2004). For experiments employing BFA (5 µg/ml), NOC (20 µg/ml) and CHX (25 µg/ml) (Lippincott-Schwartz and Cole, 1995; Petaja-Repo et al., 2006), cells were pretreated with these drugs 1 h before labelling and the drugs were present during the labelling. 177  Biotin-BMCC Labelling. As described previously (Drisdel and Green, 2004), a direct method has been used to detect protein palmitoylation. We employed a modified version of this method in this study. Briefly, cultures were lysed as shown above. For path 1 (Figure 39), N-ethylmaleimide (NEM) was added to cell lysates to a final concentration of 50 mM for 2 h at 50°C to block free sulfhydryl groups on proteins. Samples were then cleaned with Micro Bio-Spin6 Columns (Bio-Rad, Hercules, CA, USA) following the manufacturer’s instructions. Purified samples were treated with 1M hydroxylamine (HAM) and 1 µM Btn-BMCC (Pierce) in 50 mM Tris, pH7.0 for 2 h at 25°C to label palmitoylation sites. To purify biotin labelled proteins, the lysate was incubated with acetone at -20°C for 30 min and centrifugated at 10,000 x g (protocol from Pierce). The pellets were then resuspended in 0.1 ml lysis buffer containing 2% SDS. After the elution of protein precipitates, lysis buffer was added to a final volume of 1 ml. A 30 µl sample solution was used as “Input”, and boiled with 4 x loading buffer for SDS-PAGE analysis. Biotin-conjugated proteins in the rest of sample solution were purified by incubating with 50 µl Streptavidin-Sepharose (Pierce) overnight at 4°C, then eluted in 2 x loading buffer, and analyzed by SDS-PAGE together with Input. For path 2, primary antibodies were added to cell lysate containing 50 mM NEM (>16 h at 4°C), followed by 50 µl Protein A (or G)-Sepharose bead incubation (>16 h at 4°C). Immunoprecipitates were then washed, treated with 1 M HAM and 1 µM Btn-BMCC in lysis buffer for 2 h at 25°C, and boiled with 2 x loading buffer. Samples were divided into two parts, separated by SDS-PAGE gel, and blotting with either target protein antibodies or Avidin-HRP (Jackson Immunoresearch, West Grove, PA, USA). 178  Kinase Assays. JNK activity was measured with an in vitro kinase assay as described elsewhere (Khatlani et al., 2007). Briefly, cell lysates were prepared and JNK was immunoprecipitated with specific antibodies. The immnoprecipitates were then resuspended in 1 x kinase buffer (Cell Signalling Technology) supplemented with 200 μM ATP and 250 μg/ml GST-c-jun (1-89), and incubated at 30°C for 30 min. Reactions were terminated by the addition of SDS-PAGE sample buffer. Phosphorylated c-jun was resolved by SDS-PAGE and detected with anti-p-c-jun.  Deglycosylation of AMPA Receptors. Instruction for the use of EndoH and PNGF (NEB) were followed to digest proteins, with small modifications in procedures. Briefly, cultured neurons were lysed in lysis buffer in the presence of 0.5% SDS. The Sample lysate was boiled for 10 min and then incubated with EndoH or PNGF at 37°C for 10 h. For experiments involving detection of palmitoylation on AMPARs, samples were immunoprecipitated before EndoH and PNGF digestion. 2 x loading buffer was finally added to samples, which were analyzed with 6% SDS-PAGE.  BMH Crosslinking of AMPA Receptors. Protein crosslinking was conducted as previously described (Atlason et al., 2007) with some modifications. Briefly, 50 mM BMH stocking solution in DMSO was prepared freshly before experiments. Cells were collected and resuspended in 500 µl lysis buffer without SDS. The lysate was then separated into two parts: 100 µl was used as uncrosslinked control and 400 µl lysate was added BMH to a final concentration of 2.5 mM, and 179  incubated in the dark at room temperature for 45 min. Excess BMH was quenched by adding 4x loading buffer and incubating for 10 min. Samples were then boiled at 100°C and analyzed by 8% SDS-PAGE.  Quantitative Real-Time PCR (RT-PCR).  RT-PCR was employed to detect GluR2 mRNA  levels in rat cortical neuron cultures treated with 100 µM 2-BrPA for 24 h. Total RNA was isolated using TRIzol reagent (Invitrogen) and was incubated with DNase I (Invitrogen) according to the manufacturer’s instructions. The primers for GluR2 were 5’-TCAGGGTAAAGAACCACCACA-3’ and 5’-TTTCCTTGGGTGCCTTTATGCG-3’. The primers for β-actin were 5’-ACGAGGCCCAGAGCAAGAG-3’ and 5’-TCTCCATGTCGTCCCAGTTG-3’. RNA samples (~0.2 µg) were mixed with MultiScribe™ reverse transcriptase (Applied Biosystems) and SYBR Green PCR Supermix (Bio-Rad) containing 500 nM primers in each 25 µL reaction. The reaction was performed in a 7300 Real Time PCR System (Applied Biosystems). The RT-PCR conditions included an initial reverse transcription step of 30 min at 50°C, a denaturation step of 10 min at 95°C, followed by 40 cycles of PCR consisting consisting of 15 s at 95°C, 1 min at 60°C, and finally an extra dissociation step of 15 s at 95°C, 30 s at 60°C and 15 s at 95°C. Average threshold cycle (Ct) values from the PCR reactions for GluR2 were normalized against the average Ct values for β-actin from the same RNA sample. Fold inductions were calculated using the 2−(∆Ct) formula, where ∆∆Ct = ∆Ct(2-BrPA)- ∆Ct(control), ∆Ct = Ct(GluR2)-Ct(actin).  180  Cell Fractionation. For isolation of the Triton-insoluble fraction, cells were washed with cold PBS and lysed with lysis buffer with 1% Triton X-100 for 5 min at RT. Cells were then scraped into a tube and homogenized by pipeting with a 200ul tip. After centrifugation for 10 min at 13,000 x g at 4°C, the pellet was resolved with 1 x SDS loading buffer and was defined as the cytoskeleton fraction. The same volume of samples from the supernatant and pellet fractions was then subjected to SDS-PAGE analysis.  Mitochondrial Fractionation. Cultured neurons were lysed on ice for 10 min in TEEN-SKM buffer containing 20 mM Tris-HCl (pH7.5), 10 mM KCl, 1.5 mM MgCl2, 1 mM EGTA, 1 mM EDTA, 1 mM DTT supplemented with the Complete Protease Inhibitor tablets. The lysates were homogenized with a 281/2 gauge syringe for a total of 8 times on ice, followed by centrifugation twice at 700 g for 10 min at 4°C. The supernatant was then centrifuged at 10,000 g for 30 min at 4°C to enrich mitochondria in the pellet. The cytosolic fraction was collected from the supernatant after further centrifugation at 100,000 g for 1 h at 4°C. 1 x and 4 x LB were added to the mitochondrial and cytosolic fractions, respectively. Equivalent amounts of samples were resolved by SDS-PAGE and blotted with antibodies against Cytochrome c. and Bax.  LDH Assays and Cell Death Detection. Neuronal cultures were challenged with indicated concentrations of NMDA for 1 h, followed by 24 h survival. The release of lactate dehydrogenase (LDH) was measured with an in vitro toxicology assay kit (Sigma) according to the manufacturer’s 181  instructions. Spectrophotometric measurement was performed on a Multilabel Plate Reader (Envision® 2103, Perkin Elmer). The LDH reading represents the primary absorbance at a wavelength of 490 nm after subtraction of background absorbance at 690 nm. LDH readings were then converted to a percentage of neuronal death by dividing by the readings from cultures incubated with 1% triton for 15min representing maximal LDH. Following the LDH assay, propidium iodide (PI) (Sigma) was added to the medium at a final concentration of 1 μg/ml and neurons were stained for 30 min at 30°C. After fixation with 4% paraformaldehyde, neurons were washed with 1xPBS and stained with Hoechst 33342 (Sigma). The fluorescence of PI and Hoechst were examined with laser microscopy and analyzed with ImageJ.  Affinity Binding Assay and Peptide Array. An affinity binding assay was used to assess direct interaction between JNK3 and zD17. GST-fused full-length zD17 (GST-zD171-632), zD17 fragments (GST-zD171-310, GST-zD17405-479, GST-D17550-632, GST-zD17125-140, GST-zD17140-190, GST-zD17190-210) and JNK3 (GST-JNK3) were purified from E.Coli BL21 with Glutathione Sepharose 4B (GE Healthcare). Purified GST fusion proteins were resolved on SDS-PAGE and transferred onto nitrocellulose membranes. To prepare bait proteins, purified GST-myc-zD17 and GST-JNK3 were digested with thrombin overnight at room temperature, followed by clearance with β-Aminobenzamidine-agarose (Sigma) for 1 h at 4°C. The bait proteins were then prepared in affinity binding buffer (TBST with 5% skim-milk and 4% sucrose) at a concentration of 10 μg/ml. After blocking with affinity binding buffer at room temperature for 4 h, the membrane was 182  incubated with bait proteins overnight at 4°C, and washed three times with TBST. Bound bait proteins were detected with primary antibody against myc-epitope or JNK3, and HRP-conjugated secondary antibody. For mapping detailed interaction motifs, a peptide spot array was synthesized by PepMetric Technologies (Vancouver, Canada). The array contained overlapping peptides (15-mer peptides with five amino acids shift) to cover the N-terminal cytosolic domain of zD17 (zD171-325). The array membrane was initialized by washing twice with methanol for 10 min at room temperature, followed by three washes with TBST. The conditions for the preparation of array membrane, incubation with purified JNK3, and detection of bound bait protein, were the same as those of the affinity binding assays.  Immunostaining and Laser Microscopy. Cultured hippocampal neurons on the glass coverslips were used for immunocytochemistry and time-lapse studies. Immunostaining was conducted as described previously (Yang et al., 2009). For staining of the actin cytoskeleton, coverslips were incubated with TR-phalloidin in PBS for 1 h at RT. To assess axonal development, cultured hippocampal neurons were stained with anti-Tau-1 antibody. For mitochondrial detection, neuronal cultures were incubated at 37°C for 30 min with 2 μM MitoTracker-TR (Invitrogen). Coverslips were mounted with ProLong Gold Antifade Reagent containing DAPI (Invitrogen). Fluorescence was captured with a 60x objective affixed to an Olympus Fluoview 2000 confocal microscope or using a Zeiss Axiovert 200 microscope. For time-lapse imaging, 6 DIV neurons transfected with indicated constructs were mounted on a home-made chamber filled with 30°C 183  Neurobasal (minus phenol red) with supplements. Images were captured every 5 sec over 150 sec using a 60x oil immersion lens on a Zeiss Axio Observer Z1 microscope. All images were analyzed with ImageJ and processed using Adobe Photoshop.  Morphometric Analysis. The tip number and the length of axonal branches were quantified using ImageJ with NeuronJ add-on, after tracing axonal branches. The primary axon is defined as the longest continuous path from the soma, and the secondary axons are defined as branches initiated from the primary axon and etc. The terminal points of all axonal branches are defined as axon tips. To illustrate motility of axonal filopodia, the difference of pairs of successive frames from 30 time-lapse images were extracted by ImageJ arithmetically (Chang and De Camilli, 2001). The resulting 29 images were added up to generate a final image showing the total differences. A greater number of pixels represent higher motility. To quantify filopodia motility, the motility index (MI), which measures the areas that a filopodium occupies over time, was used as described previously (Chang and De Camilli, 2001). Briefly, images were adjusted using a threshold throughout the entire time-lapse stack and then binarized. 10 filopodia from each neuron were randomly chosen for further analysis. The areas that a filopodium occupied throughout the stack were measured and calculated using the formula: MI= [area (max)-area (min)]*2/[area (max)+area (min)].  Transient Ischemia in Rats. Adult male Sprague-Dawley (SD) rats weighing 250–290 g were 184  employed for transient middle cerebral artery occlusion. Rats were anaesthetized with 5% isofluorane and maintained with 2% isofluorane in 70% N2O and 30% O2 using a face mask. 5 mg/kg ketoprofen was given pre-surgery to block pain. Rectal temperature was maintained at 37°C during surgery with a homeothermic blanket system (Harvard apparatus, Holliston, MA). The scalp was incised at the midline and the skull was exposed. The skull was thinned with a dental drill at a region ipsilateral to the expected ischemia zone (2 mm posterior and 5 mm lateral from the bregma). A laser probe of the Doppler flowmeter (Perimed, Järfälla, Sweden) was placed at this location during surgery to detect cerebral blood flow (CBF) (Nagel et al.). We induced transient MCAo using a method of intraluminal vascular occlusion described elsewhere (Longa et al., 1989; Belayev et al., 1996; Aarts et al., 2002). Briefly, a poly-L-lysine coated 3-0 monofilament nylon suture (Harvard apparatus, Holliston, MA) with a rounded tip was advanced from the left common carotid artery into the internal carotid artery until a sudden drop of CBF was noted, indicating the blockade of the origin of the MCA. The success of induction of ischemia was confirmed by a fall of CBF to less than 25% of baseline level and by neurological assessment scores during MCAo (1 h after the onset). Two hours after MCAo, rats were re-anaesthetized and the suture was removed to allow reperfusion, confirmed by the increase of CBF at the same area.  Peptide Administration. Peptides were synthesized by PepMetric Technologies or GL Biochem (Shanghai, China). Peptides were prepared freshly in saline on the day of experiment at a stock concentration of 1 mg/ml. 1 mg/kg peptides were administrated at indicated times by a single 185  intravenous injection into the rat tail. NIMoEscr or NIMoEsh were injected at 4 h after MCAo onset to randomly chosen rats. For long-term studies, peptides were given 2 h after MCAo onset. We injected NIMoEscr and NIMoEsh, assessed brain damage and performed functional evaluations in a double-blind manner. To examine the delivery of the peptide into the brain, either 5 mg/kg FITC-labelled NIMoE, or saline, was intravenously injected 1 h before perfusion. The rats were then perfused with PBS followed by fixation solution (4% paraformaldehyde in PBS). Brains were removed, postfixed in fixation solution for 2 h, and then soaked in 20% sucrose–PBS buffer at 4oC for 24 h. After rapid freezing in Tissue-Tek embedding medium (Sakura, Torrance, CA) on dry ice, 20 μm sections were cut on a cryostat (Leica CM3050, Ontario, Canada) and examined for FITC fluorescence by a laser microscopy (Zeiss, Axiovert 200).  Histological Assessment. Rats were allowed to survive for 24 h. After decapitation, the brains were immediately removed and serially sectioned in the coronal plane at a thickness of 2 mm with a slicer matrix (Zivic, Pittsburgh, PA). A total of 8 sections were collected and then incubated in PBS with 1% 2,3,7-triphenyltetrazolium chloride at 37oC for 10 min. The stained slices were then fixed in 4% paraformaldehyde and digitised with a color flatbed scanner (MFC-8860DN, Brothers). The infarct area within each section was traced and quantified using ImageJ. The infarct volume was calculated using the formula: V = d(A1+A2+...+A8) where V is the infarct volume (mm3), d is the distance between sections, and A is the infarct area within each section.  186  Functional Tests. All animals were tested for neurological function during (1h after onset) and 24h after MCAo. We evaluated motor, sensory and coordination capacities using modified Neurological Severity Scores (Bederson et al., 1986; Watanabe et al., 2004). The motor system tests comprised 7 components, including motor initiation test, free activity, posture, walking, tail suspension, hindlimb flexion and pushing test. Somatosensory tests included a tactile test and forelimb placing test. Coordination functions were assessed with 3 tests including a foot fault test, edge test, and balance beam. Performance in the tests was evaluated on a cumulative scale from 0 to 2 (10 tests), except for two tests (tail suspension and balance beam) in which scores ranged from 0 to 3and 0 to 4, respectively. The scores from each test were summed and represented as a single overall neurological score (0 to 27). The adhesive-removal test was used for further evaluation of somatosensory deficits (Bouet et al., 2009). A piece of adhesive tape was wrapped around the right forepaw. The time spent in removing the tape from the paw within 2min was recorded.  Statistics. All values in text and figures are presented as mean ± standard error of the mean. Student t-test or one-way ANOVA was performed using Excel software (Microsoft). The limit of statistical significance was set at a P value < 0.05.  187  

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