MODULATION OF HIVGP120 ANTIGEN-SPECIFIC IMMUNE RESPONSES, IN VITRO AND IN VIVO, BY 9-TETRAHYDROCANNABINOL AND CANNABINOID RECEPTORS 1 AND 2 By Weimin Chen A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Microbiology – Environmental Toxicology – Doctor of Philosophy 2014 ABSTRACT MODULATION OF HIVGP120 ANTIGEN-SPECIFIC IMMUNE RESPONSES, IN VITRO AND IN VIVO, BY 9-TETRAHYDROCANNABINOL AND CANNABINOID RECEPTORS 1 AND 2 By Weimin Chen Approximately 25% of HIV patients use marijuana for its putative therapeutic benefit; however, it is unknown how cannabinoids affect the immune function of immunocompromised HIV patients at the early stage. The goal of these studies was to investigate the immunomodulatory effects of cannabinoids on the early stage of the anti-HIV immune responses. A surrogate in vitro mouse model was first established to determine the effects of cannabinoids on HIV antigen-specific T cell effector function. Specifically, CD8+ T cell proliferation and gp120-specific IFN production were induced, which were suppressed or enhanced by ∆9-tetrahydrocannabinol (THC), the predominant psychoactive component in marijuana, depending on the magnitude of the control response. To further elucidate the molecular and cellular mechanisms by which cannabinoids differentially modulate T cell responses, phorbol ester, phorbol 12-myristate 13-acetate (PMA), in combination with the ionophore, ionomycin (Io) or anti-CD3/CD28 antibodies were used as activation stimuli, and another marijuana-derived cannabinoid, cannabidiol (CBD), was also investigated. THC or CBD suppressed or enhanced IFN and IL-2 production by mouse splenocytes under optimal or suboptimal stimulation, respectively. Similar differential effects of cannabinoids on cytokine production were also observed on nuclear translocation of nuclear factor of activated T cells (NFAT) and with human peripheral blood mononuclear cells (PBMCs) in response to PMA/Io stimulation. However, THC and CBD elevated intracellular calcium, regardless of the stimulation level with PMA/Io, suggesting that the cannabinoid-induced calcium increase provides an appropriate signal for activation in suboptimally stimulated T cells but an anergiclike signal as a result of excessive calcium in optimally stimulated T cells. Together, these data demonstrate differential modulation by cannabinoids of a HIV antigen-specific response and identify a possible mechanism responsible for this effect. To investigate whether THC produced similar effects in vivo, a mouse model to induce HIVgp120-specific immune responses was then developed. The gp120-expressing plasmid, pVRCgp120, or the vector plasmid, pVRC2000, was injected intramuscularly into mice. Both the gp120-specific IFN and IL-2 responses were detected, when splenocytes were restimulated with gp120-derived peptide IIGDIRQAHCNISRA (#81), which was identified as being immunodominant among 211 tested peptides. In addition, T cell and non-T cell populations were all activated in response to pVRCgp120 stimulation followed by peptide restimulation, as evidenced by increased expression levels of surface markers (e.g., CD69, CD80, major histocompatibility complex II [MHC II]). The gp120-specific IFN response and the magnitude of cellular activation were enhanced by THC in C57Bl/6 wild-type (WT) mice but suppressed or not affected by THC in cannabinoid receptors 1 (CB1) and 2 (CB2) -/- -/- knockout (CB1 CB2 ) mice. -/- -/- Furthermore, CB1 CB2 mice exhibited an augmented immune response phenotype compared with WT mice on IFN production. Collectively, these in vivo findings suggest that THC modulates HIV antigen-specific immune responses in vivo in both a CB1/CB2-dependent and -independent manner. The significance of this dissertation work is that it provides novel insights into the understanding of cannabinoidmediated effects on antigen-specific immune responses and the associated mechanisms. ACKNOWLEDGEMENTS First of all, I would like to acknowledge my thesis advisor, Dr. Norbert Kaminski, for his excellent mentorship during my years in his laboratory. He inspired me with great projects and ideas, trained me to become a motivated and meticulous researcher, and provided tremendous support for my research and other activities at the Society of Toxicology, which made my Ph.D. experience in his laboratory productive as well as beneficial for my future career. I would also like to thank my guidance committee for providing helpful suggestions and insights during discussions of my dissertation research. Specifically, I thank Dr. Andrea Amalfitano for valuable discussions on HIV biology and vaccine research; Dr. Barbara Kaplan for being my go-to person in the lab and all her great inputs in many experiments and manuscript writings; Dr. James Pestka for sharing critical thinking on experimental designs; Dr. Kefei Yu for sharing his expertise in cell and molecular immunology; former committee member Dr. Ian York for nurturing my interest in the immunology field and providing valued suggestions on the model systems. Additionally, I would like to thank current and former Kaminski lab members: Dr. Schuyler Pike, Dr. Peer Karmaus, Dr. Thitirat Ngaotepprutaram, Dr. Ale Manzan, Dr. Haitian Lu, Dr. Ashwini Phadnis-Moghe, Dr. Priya Raman, Dr. Collin North, Natasha Kovalova, Jose Suarez Martinez, Jinpeng Li, Bob Crawford, Kimberly Hambleton, Tony Bach, Shawna D'Ingillo, Sara Fox, and Sonia Dantas, for providing me a wonderful working environment, helping me with many aspects of my projects, and being great friends. I also want to express my appreciation to the Department of Microbiology & Molecular Genetics and Center for iv Integrative Toxicology for giving me the opportunity to be part of the programs. They have been a great source of help and support all these years I have spent here. Last but not least, I would like to thank my parents for bringing me up and giving their encouragement in my oversea study and life. Their love is always a major source of strength to me. I would also like to thank my husband Dr. Kezhen Huang for his encouragement, enduring love and support all these years, which have made me through my Ph.D. and will leave an everlasting impression in my life, and my baby girl Leah (Zixuan) for being such an angel and bringing so much love, happiness & joy to my life. v TABLE OF CONTENTS LIST OF TABLES ................................................................................................................ ix LIST OF FIGURES .............................................................................................................. x KEY TO ABBREVIATIONS ............................................................................................... xiii INTRODUCTION ................................................................................................................ 1 I. Human Immunodeficiency Virus (HIV) ..................................................................... 1 A. HIV Classification and Genome .......................................................................... 1 B. HIV-1 Prevalence ................................................................................................. 2 C. HIV-1 Infection .................................................................................................... 3 II. Innate Immune Responses to HIV-1 .......................................................................... 4 A. Cytokines/Chemokines ........................................................................................ 4 B. DCs ....................................................................................................................... 5 C. NK Cells ............................................................................................................... 5 III. Antigen Processing and Presentation ........................................................................ 6 A. MHC I and MHC II Pathways ............................................................................. 6 B. DC Maturation ...................................................................................................... 8 IV. Adaptive Immune Responses to HIV-1 .................................................................... 8 A. CD8+ T Cell Responses ........................................................................................ 8 B. CD4+ T Cell Responses ........................................................................................ 10 C. B Cell Mediated Immune Responses ................................................................... 11 D. Implications for Vaccine Development................................................................ 12 V. T Cell Signaling Cascades ......................................................................................... 12 A. T Cell Activation Signaling Pathways ................................................................. 12 B. T Cell Stimulation In Vitro ................................................................................... 14 VI. Cannabinoids and Cannabinoid Receptors ............................................................... 14 A. Cannabis Use ........................................................................................................ 14 B. Cannabinoids ........................................................................................................ 15 C. Pharmacokinetics of THC .................................................................................... 18 D. Physiological Effects of THC .............................................................................. 19 E. Toxicity of THC ................................................................................................... 19 F. Cannabinoid Receptors ......................................................................................... 20 1) CB1 and CB2 ................................................................................................... 20 2) Signaling Events Associated with CB1/ CB2 .................................................. 21 3) Other Targets for Cannabinoid Ligands ......................................................... 25 VII. Cannabinoid and CB1/ CB2-Mediated Immune Modulation .................................. 26 A. T Cells .................................................................................................................. 27 B. B Cells .................................................................................................................. 29 C. NK Cells ............................................................................................................... 30 D. APCs .................................................................................................................... 31 E. In Vivo Host Resistance Models ........................................................................... 32 vi F. Involvement of CB1/ CB2 in Cannabinoid-Mediated Immune Modulation .......... 33 VIII. Rationale and Objectives........................................................................................ 36 MATERIALS AND METHODS .......................................................................................... 39 I. Chemicals and Reagents .............................................................................................. 39 II. Mice ............................................................................................................................ 39 III. Cell Culture Conditions ............................................................................................ 40 IV. Lentiviral Plasmid Construction ............................................................................... 40 V. DC2.4gp120 and EL4gp120 Cell Line Construction........................................................ 43 VI. In Vivo Immunizations Using Plasmids .................................................................... 43 VII. Real Time PCR ........................................................................................................ 44 VIII. Western Blotting Analysis ..................................................................................... 45 IX. Elicitation of Splenocytes with DC2.4gp120 Cells...................................................... 46 X. THC Treatment In Vivo .............................................................................................. 46 XI. Necropsy and Tissue Preparation.............................................................................. 47 XII. Enumeration of Cytokine Secreting Cells by ELISPOT Assays ............................. 50 XIII. Cell Staining for Flow Cytometry .......................................................................... 51 XIV. T Cell Purification.................................................................................................. 53 XV. ELISA...................................................................................................................... 53 XVI. Calcium Determinations ........................................................................................ 54 XVII. Low Density Array Mouse Immune Panel ........................................................... 55 XVIII. Verification of Genotype ..................................................................................... 55 XIX. Statistical Analysis ................................................................................................. 56 EXPERIMENTAL RESULTS .............................................................................................. 58 I. In Vitro Cannabinoid-Mediated Effects on T Cell Response to HIVgp120 ................... 58 A. Expression of gp120 mRNA and protein in transduced DC2.4 and EL4 clones . 58 B. Elicitation of gp120-specific CD8+ T cell response ............................................. 59 C. Differential effects of cannabinoids on CD8+ T cell IFN production and proliferation ............................................................................................................... 67 D. Differential effects of cannabinoids on IL-2 response ......................................... 80 E. Differential effects of cannabinoids on cytokine production were correlated with effects on NFAT nuclear translocation ..................................................................... 86 F. Cannabinoid enhanced intracellular calcium regardless of the magnitude of cellular activation ................................................................................................................... 90 II. In Vivo Cannabinoid-Mediated Effects on Immune Responses to HIVgp120 .............. 93 A. pVRCgp120 induced gp120-specific cytokine production by ELISPOT assay .. 93 B. pVRCgp120-induced T cell activation ................................................................. 97 C. pVRCgp120-induced activation on non-T cells ................................................... 97 D. pVRCgp120-induced B cell responses................................................................. 98 E. THC treatment enhanced pVRCgp120-induced IFN production by lymphocyte populations in the spleens of WT but not CB1-/-CB2-/- mice ..................................... 103 F. THC treatment enhanced pVRCgp120-induced activation of T cells in WT but not CB1-/-CB2-/- mice........................................................................................................ 104 G. THC treatment enhanced pVRCgp120-induced B cell activation in WT but not CB1-/-CB2-/- mice........................................................................................................ 109 vii H. THC treatment enhanced pVRCgp120-induced activation of non-T/non-B cells in WT but not CB1-/-CB2-/- mice .................................................................................... 109 I. THC modulated mRNA expression of genes induced by pVRCgp120 ................ 110 DISCUSSION ....................................................................................................................... 114 I. In vitro cannabinoid-mediated effects on T cell responses to HIVgp120 ....................... 114 II. In vivo cannabinoid-mediated effects on immune responses to HIVgp120 .................. 119 III. Significance and concluding remarks ....................................................................... 126 APPENDIX ........................................................................................................................... 131 BIBLIOGRAPHY ................................................................................................................. 175 viii LIST OF TABLES Table 1. Sequences of PCR primer/probe sets ................................................................. 42 Table 2. Differential gene expression in response to pVRCgp120 injection and THC treatment in WT and CB1-/-CB2-/- mice ............................................................................. 113 Table 3. Grafting amount and size of SNPs ..................................................................... 150 Table 4. Grafting data from different size SNPs terminated with carboxylic acids ..... 151 ix LIST OF FIGURES Figure 1. Chemical structures of selected cannabinoids ................................................. 17 Figure 2. Simplified signaling pathways associated with CB1/CB2 ................................ 24 Figure 3. Experimental model of in vitro elicitation of gp120-specific T cells ............... 48 Figure 4. Protocol for the administration of THC or CO into the mice immunized with pVRCgp120/pVRC2000...................................................................................................... 49 Figure 5. Plasmid cassette constructs for expression of gp120 and puror in DC2.4 (top) or gp120 and neor in EL4 cells (bottom) ........................................................................... 60 Figure 6. Expression of gp120 transcript and protein in transduced DC2.4gp120 or EL4gp120 clones ..................................................................................................................... 61 Figure 7. Proliferation of gp120-specific CD8+ T cells was elicited in the in vitro model ............................................................................................................................................... 63 Figure 8. gp120-specific IFN production by CD8+ T cells was elicited in the in vitro model .................................................................................................................................... 64 Figure 9. Differentiation of gp120-specific CD8+ T cells was elicited in the in vitro model ............................................................................................................................................... 66 Figure 10. Differential effects of THC on IFN production ............................................ 68 Figure 11. Differential effects of THC on CD8+ T cell proliferation .............................. 71 Figure 12. THC-mediated effects on IFN response in a CB1/CB2-dependent manner 75 Figure 13. Effects of cannabinoids on IFN production in response to different stimuli ............................................................................................................................................... 78 Figure 14. Effects of cannabinoids on IL-2 production in response to different stimuli ............................................................................................................................................... 81 Figure 15. Enhancement of the IL-2 response by CBD in purified T cells .................... 83 Figure 16. Enhancement of cytokine response by CBD in different cellular populations ............................................................................................................................................... 84 Figure 17. Differential effects of CBD on IL-2 response in human PBMC ................... 85 x Figure 18. Cannabinoids differentially regulated IFN steady state mRNA expression ............................................................................................................................................... 87 Figure 19. Cannabinoids differentially regulated IL-2 steady state mRNA expression ............................................................................................................................................... 88 Figure 20. Cannabinoids differentially modulated nuclear translocation of NFAT2... 89 Figure 21. Cannabinoids increased intracellular calcium regardless of activation level ............................................................................................................................................... 91 Figure 22. gp120-specific cytokine production by splenocytes ....................................... 94 Figure 23. Identification of immunodominant gp120 peptide for restimulation .......... 96 Figure 24. Activation of T cells in response to pVRCgp120 stimulation ....................... 99 Figure 25. Activation of non-T cells in response to gp120 antigen stimulation ............. 100 Figure 26. B cell responses to pVRCgp120 stimulation ................................................... 102 Figure 27. THC-mediated enhancement on gp120-specific IFN response in WT but not CB1-/-CB2-/- mice ................................................................................................................... 105 Figure 28. THC-mediated enhancement on gp120-specific T cell activation in WT but not CB1-/-CB2-/- mice ............................................................................................................ 107 Figure 29. THC-mediated enhancement on gp120-specific B cell activation in WT but not CB1-/-CB2-/- mice ............................................................................................................ 111 Figure 30. THC-mediated enhancement on gp120-specific non-T/non-B cell activation in WT but not CB1-/-CB2-/- mice .............................................................................................. 112 Figure 31. Synthesis of different SNPs .............................................................................. 142 Figure 32. Surface chemistry of modified SNPs with different functional groups and at different sizes ....................................................................................................................... 149 Figure 33. Aqueous dispersion stability of LTM40PTrg4PEG SNPs in in vitro culture media .................................................................................................................................... 153 Figure 34. Aqueous dispersion stability of SNPs with -OH functional group in in vitro culture media ....................................................................................................................... 154 xi Figure 35. Aqueous dispersion stability of SNPs with –NH2 functional group in in vitro culture media ....................................................................................................................... 155 Figure 36. Aqueous dispersion stability of SNPs with -COOH functional group in in vitro culture media ....................................................................................................................... 156 Figure 37. SIINFEKL concentration-dependent activation of CD8+ T cell responses . 158 Figure 38. Enhancement by modified LTM40PTrg4PEG on CD8+ T cell responses induced by a suboptimal concentration of SIINFEKL .................................................... 163 Figure 39. Absence of enhancement by SNPs with different functional groups on CD4+ T cell responses under suboptimal OVA323-339 stimulation ................................................. 165 Figure 40. Effect of -COOH SNPs at different sizes on CD8+ T cell responses under suboptimal induction .......................................................................................................... 167 xii KEY TO ABBREVIATIONS 2-AG 2-arachidonoyl glycerol AC adenylate cyclase AEA anandamide AFC antibody forming cell Akt activation of protein kinase B AHR airway hyperesponsiveness AP-1 activator protein-1 APC antigen presenting cell ART antiretroviral drug therapy BALF bronchoalveolar lavage fluid BCS bovine calf serum bmDCs bone marrow-derived DCs BSA bovine serum albumin cAMP cyclic adenosine 3’,5’-monophosphate CB1 cannabinoid receptor 1 CB2 cannabinoid receptor 2 CB1-/-CB2-/- CB1 and CB2 null CBD cannabidiol CBN cannabinol CCR chemokine co-receptor CD40L CD40 ligand CNS central nervous system xiii CO corn oil conA concanavalin A CRAC calcium release-activated channels CREB/ATF cAMP-response element binding protein/activation transcription factor CTL cytotoxic T lymphocytes DAG diacylglycerol DC dendritic cell DC2.4gp120 gp120-expressing DC2.4 cells DLS dynamic light scattering DYRK1A dual-specificity tyrosine-phosphorylation regulated kinase 1A EL4gp120 gp120-expressing EL4 cells ERK extracellular signal-regulated kinases FTIR fourier transform infrared spectroscopy Gi inhibitory heterotrimeric G proteins Gs stimulatory heterotrimeric G proteins GPCR G protein-coupled receptor GSK-3 glycogen synthase kinase-3 HIV Human Immunodeficiency Virus HLA human leukocyte antigen iCD3/CD28 immobilized anti-CD3 plus soluble anti-CD28 antibodies IFN interferon i.m. intramuscular Io ionomycin xiv IP3 inositol trisphosphate ITAMs the immunoreceptor tyrosine-based activation motifs JNK c-Jun N-terminal kinases KIR killer cell immunoglobulin-like receptor KO knockout KV the voltage-gated potassium channels LN lymph node LPS lipopolysaccaride MAPK mitogen-activated protein kinases MCP-1 monocyte chemotactic protein-1 NEAA non-essential amino acids neor neomycin resistance NFAT nuclear factor of activated T cells NF-B nuclear factor kappa-light-chain-enhancer of activated B cells NK natural killer NT no treatment OVA ovalbumin PBMC peripheral blood mononuclear cells PBS phosphate buffered saline pDC plasmacytoid DCs PEG polyethylene glycol P/I PMA/Io PIP2 phosphatidylinositol 4,5-bisphosphate PKA protein kinase A xv PMA phorbol 12-myristate 13-acetate PPAR peroxisome proliferators-activated receptors PTK protein tyrosine kinase puror puromycin resistance S SIINFEKL sCD3/CD28 soluble anti-CD3 plus soluble anti-CD28 antibodies SEM scanning electron microscopy SIV simian immunodeficiency virus SNP silica nanoparticle sRBC sheep red blood cell STIM stromal interaction molecule TAP transporters associated with antigen processing TCR T cell receptor TEM transmission electron microscopy TGA thermogravimetric analyses THC 9-Tetrahydrocannabinol TLR toll-like receptors TNF tumor necrosis factor TRPA1 transient receptor potential ankyrin 1 TRPC1 transient receptor potential canonical 1 TRPV1 transient receptor potential vanilloid 1 VH vehicle WT wild-type xvi INTRODUCTION I. Human Immunodeficiency Virus (HIV) A. HIV Classification and Genome HIV is a single-stranded (ss) RNA (positive-sense) virus, belonging to the retroviridae family. There are two known types of HIV, HIV-1 and HIV-2. HIV-1 is more infective and virulent, causing the majority of infections in the world, whereas HIV-2 is less infective upon exposure and mainly found in West Africa [1]. Therefore, only HIV-1 is discussed in this dissertation. HIV-1 viruses are divided into different groups, including the major group M, which is responsible for 90% of the infections worldwide. Group M viruses are further subdivided into various subtypes through A to K, also referred to as clades, based on their genome sequences [2]. Subtype B viruses are predominant in Western countries [2, 3]. HIV-1 genome contains nine identified genes, including gag, pol, env, tat, rev, nef, vif, vpr, and vpu, encoding a total of 19 proteins [4]. The structural proteins of the virus consist of Gag, protease, integrase, RNase H, reverse transcriptase, and envelope protein (Env), which have their particular functions. For example, the Gag protein, also called p55, consists of four smaller parts: the matrix (p17) responsible for particle stabilization [5] or nuclear transportation [6], the capsid (p24) to form the core of viral particles, the nucleocapsid (p9) involved in recognition of packaging signal [7], and p6 needed for interaction with Vpr protein [8]. In particular, the Env protein is cleaved to the transmembrane domain p41 and the surface protein gp120, which mediates the 1 entry of virus into target cells by interacting with the surface CD4 molecule as well as a chemokine co-receptor (CCR), CXCR4 or CCR5, depending on the viral tropism [9]. There are two regulatory proteins, including the transcriptional activator Tat involved in viral replication [10], and the RNA binding protein Rev associated with transportation of RNA from the nucleus to the cytoplasm [11]. In addition to the structural and regulatory proteins, HIV-1 genome also encodes the following accessory proteins. Nef protein is involved in decreasing surface CD4, major histocompatibility complex (MHC) I, and MHC II expression, therefore downregulating T cell activation and increasing HIV-1 infectivity [12-14]. Vif protein is required for HIV-1 replication in some cells [15]. Vpr protein causes cell accumulation at the G2/M division [16]. Vpu protein is known to increase viral release from infected cells [17]. B. HIV-1 Prevalence HIV-1 is associated with high mortality in the US. At the end of 2009, approximately 1.1 million people were infected with HIV-1 in the United States, and an estimation of half a million people have died due to acquired immune deficiency syndrome (AIDS) [18]. According to estimates by the Center for Disease Control, each year there are about 50,000 new HIV-1 infections in the United States, and an estimated 47,500 people were newly infected in 2010 (the most recent year that data are available) [19]. 2 C. HIV-1 Infection HIV-1 exposure mainly occurs through the mucosal route of transmission, including reproductive and gastrointestinal routes. Upon exposure in the mucosal tissues, the virus infects immune cells locally by interacting with CD4 receptor and a chemokine coreceptor, CCR5 or CXCR4 [20]. CCR5-tropic HIV-1 viruses are the predominant ones in chronically infected patients [21]. Since it is impossible to study the physiological events in the early stage of HIV-1 infection in humans in vivo, simian immunodeficiency virus (SIV) infection in rhesus macaque monkeys has been used as a model system to provide a fundamental understanding of HIV infection. + CD4 T cells have been demonstrated as the major initial targets for SIV replication in the lymphoid tissue in the mucosa [22]. Free virus and/or virus-infected cells then travel to the other lymphoid tissues + where they further infect more immune cells, including CD4 T cells, dendritic cells (DC), and macrophages. In the acute phase of HIV-1 infection, the viral load increases exponentially to reach a peak by 2-3 weeks, which accompanies a dramatic + + decrease of CD4 T cell counts, establishing a pool of latently infected CD4 T cells [23]. Meanwhile, HIV-1 specific immune responses are developed (see below), + followed by a decrease of viremia and a partial recovery of peripheral blood CD4 T cell counts [24]. The viral load falls over the next 3-5 months to reach a stable level, known as the viral set point [25], which determines the intensity, the magnitude, and the kinetics of disease progression in the chronic infection stage in the absence of + antiretroviral drug therapy (ART). Over the next 10 years, the CD4 T cell numbers 3 slowly decrease and the viremia slowly increases, along with any potentially opportunistic tumors and viral infections in the AIDS patients [26, 27]. Upon entry into host cells, HIV-1 viral RNA is reverse transcribed by the reverse transcriptase into double-stranded cDNA, which is then transported into the cell nucleus and integrated into the DNA genome of host cells by the viral integrase [28]. Upon integration, the virus may become latent to protect itself and the host cells from being detected by the immune system [28]. The integrated DNA provirus can also be transcribed in activated T cells, making new RNA and viral proteins that are packaged into new viruses [29, 30]. During the replication process, HIV-1 virus mutates rapidly, which makes the virus highly genetically-variable and thus increases the difficulty of developing HIV-1 vaccines [31]. II. Innate Immune Responses to HIV-1 A. Cytokines/Chemokines When there is HIV-1 infection, the innate immune response is the first line of defense. As the plasma viremia increases, toll-like receptors (TLRs), which recognize viral RNAs and proteins [32], are activated, leading to the production of acute-phase cytokines and chemokines. Type I interferons (IFNs), such as IFN, interleukin-15 (IL-15), and CXC-chemokine ligand 10 (CXCL10) are produced rapidly upon infection, followed by production of tumor necrosis factor (TNF) , IFN, and IL-18, and IL-10 [33]. Type I IFNs are known to have antiviral activities by inhibiting HIV replication [34]. Type I IFNs, IL-15, and IL-18 can also enhance innate and adaptive 4 immune responses. During the acute phase of HIV-1 infection, cytokines and chemokines can be produced by infected CD4 + T cells, DCs, monocytes, macrophages, natural killer (NK) cells, and NKT cells [35]. B. DCs DCs and macrophages were demonstrated to be infected either through the + + CD4 CCR5 route or through the DC-specific intracellular adhesion molecule-3- grabbing non-integrin (also known as CD209) receptor on the cell surface [36]. In particular, plasmacytoid DCs (pDCs) were shown to be activated in response to HIV-1 infection, which led to activation of TLR7 by viral ssRNA [37] as well as production + of IFN to inhibit HIV-1 replication and promote CD4 T cell expansion [38, 39]. + + Infected conventional DCs were found to further activate CD4 and CD8 T cells in the adaptive immune responses [40]. C. NK Cells NK cells also play an important role in controlling HIV viremia during acute infection. NK cells are activated by pro-inflammatory cytokines secreted by DCs and monocytes, including IL-15 and IFN [41]. Depending on the functionality of killer cell immunoglobulin-like receptors (KIRs), NK cells might have activating or inhibitory functions on HIV-1 replication. For example, NK cells that expressed the activating receptor KIR3DS1 binding to its human leukocyte antigen (HLA) class I ligands, HLA class I molecules of the HLA-Bw4-80I family, were found to inhibit 5 + HIV-1 replication in CD4 T cells [42] and therefore slowed down HIV-1 disease progression [43]. In contrast, expression of inhibitory receptor KIR3DL1 in combination with the same HLA family as mentioned above by NK cells might have an opposite effect [44]. In addition, NK cells can also secrete antiviral cytokines and chemokines as well as cytotoxic granules that mediate the antiviral activities to further induce apoptosis of infected cells. Furthermore, NK cells were demonstrated to eliminate immature DCs, therefore leaving the mature DCs to induce antiviral T cell responses [45]. III. Antigen Processing and Presentation A. MHC I and MHC II Pathways Antigen presentation links the innate and adaptive immune responses. As reviewed by Cresswell et al. [46], endogenous antigens are usually presented by MHC I molecules, whereas exogenous antigens are presented by MHC II molecules. However, exceptions exist under certain conditions (see below). Endogenous antigens, including viral proteins, intracellular bacteria, tumor antigens, and misfolded proteins, are escorted to the proteasome after ubiquitin tagging and are processed into peptide fragments in the cytosol. These antigenic fragments are then transported to the endoplasmic reticulum (ER) by transporters associated with antigen processing (TAP) to be further trimmed by ER aminopeptidases (ERAPs) to peptides of 8-10 amino acids in length and loaded onto MHC I molecules. This process is coordinated by the peptide-loading complex that contains several proteins. Finally, peptide-loaded 6 MHC I complexes are exported to the cell surface to allow for immune surveillance by + CD8 T cells. On the other hand, exogenous antigens, such as extracellular bacteria, parasitic worms, and free viruses, are endocytosed by antigen presenting cells (APCs). The endosome compartments fuse with lysosomes where the antigens are processed to fragments that are further degraded to peptides (~13-18 amino acids) by proteases in the lysosome MHC class II compartment (MIIC). Meanwhile, MHC II molecules assembled in ER traffic to MIIC where the invariant chain (li) is cleaved to class IIassociated li peptide (CLIP) in the peptide-binding groove prior to binding with high affinity-peptides. Then antigenic peptides bind to MHC II molecules, which are + transported to the cell surface to allow for immune surveillance by CD4 T cells (as reviewed in [47]). All nucleated cells express MHC I molecules, whereas only professional APCs, such as DC, macrophages, and B cells, express MHC II molecules. There are exceptions that extracellular antigens taken up via endosomes may be retrotranslocated into the cytosol and presented on MHC I molecules by some DC and macrophages, referred to as cross-presentation [46]. In addition, endogenously expressed proteins may traffic to the MIIC through autophagy for presenting on the MHC II molecules [48]. In the case of HIV-1 infection, viral proteins are processed to antigenic peptides in the APCs, which will then be presented in the context of MHC I + + and II molecules to be recognized by CD8 and CD4 T cells, respectively. 7 B. DC Maturation DCs have been demonstrated to be more effective to present antigens to naïve T cells than other APCs [49]. Upon capturing antigens, DCs mature and migrate to lymphoid tissues to activate T cells. DC maturation involves up-regulation of MHC I and II molecules, co-stimulatory molecules, CD80 and CD86, and changes in cell morphology [49]. Proinflammatory cytokines TNF-, IL-1, IL-6, type I and type II interferons, and pathogen-associated molecules, such as ssRNA, dsRNA, and lipopolysaccaride (LPS) are known to induce DC maturation [49]. IV. Adaptive Immune Responses to HIV-1 A. CD8+ T Cell Responses + As the viremia increases to the peak level, HIV-specific CD8 T cells develope. T cells are activated upon the engagement of interactions between T cell receptor (TCR) and antigen-MHC complexes on APCs. The co-stimulatory signal from the binding of CD28 on T cells by CD80 and CD86 on APCs is also required for T cell activation (see below). Once T cells are activated, they clonally expand and differentiate to effector T cells. Virus-specific cytotoxic T lymphocytes (CTLs) can recognize HIVinfected cells presenting MHC I-restricted peptides on cell surface, and eliminate infected cells. Therefore, clearance of virus in the acute infection stage correlates with the appearance of HIV-specific CTLs [50]. CTLs produce and release granules containing cytolytic proteins, such as perforin and granzymes. Upon degranulation, they directly act on target cells presenting the same antigens and induce apoptosis of 8 target cells [51, 52]. An alternative route of triggering target cell apoptosis is mediated through the interaction between FasL on CTLs and Fas on infected target cells [53]. In addition, cytokines, such as IFN, TNF-, and IL-2, secreted by CTLs following recognition of target epitopes, can promote the cytolytic activity of CTLs and help maintain the growth, differentiation and survival of CTL [51, 52]. Moreover, CTLs can secret chemokines, such as MIP-1, MIP-1, and RANTES, which can bind to the CCR5 coreceptor on infected cells, therefore destroying the entry of viruses, and also facilitate the cytolytic activity of CTLs among other functions [54]. Recently, + presence of multifunctional CD8 T cells, which possess more than two functions simultaneously, including cytokine/chemokine production and degranulation, was detected in HIV elite controllers coincident with low viremia in the plasma [55]. Collectively, CTLs play a critical role in controlling HIV, resulting in a decrease of viral load in the plasma until reaching the set point in the acute phase of HIV infection. A more robust immune response after acute infection will lead to a lower viral set point, which might slow the disease progression to AIDS. Thus, the viral set point is an important indicator of the initial immune responses as well as the longerterm prognosis of the patient. Although CTL induction helps to set a low viral set point, the virus sequence still + mutates rapidly at multiple epitopes in order to escape from the recognition by CD8 T cells [56, 57]. Changes in amino acid sequence were often found to occur upstream of the sequence that could be recognized by T cells, and were suggested to be involved in antigen processing [58, 59]. T cell responses elicited against Env and Nef proteins in particular were indicated to control early viremia, while T cell responses against 9 other viral proteins, such as Gag and Pol, were more involved in maintaining the viral set point [56, 60]. Once the escape mutants were selected, the magnitude of the initial T cell responses decreased [56]. A lower viral set point was observed when T cell responses were induced against immunodominant viral epitopes [61]. In addition, it + has been suggested that CD8 T cell responses to conserved epitopes played a more important role in long-term control of HIV-1 infection [56, 62]. + B. CD4 T Cell Responses + Naïve CD4 T cells upon activation can proliferate and differentiate to T helper cells. Typically, CD4 + T helper cells can produce various soluble mediators, including cytokines and chemokines. Th1 cytokines, such as IL-2 and IFN, are secreted to maintain effective CTL functions (Th1 response). Th2 cytokines, such as IL-4, IL-5, IL-6, and IL-10, are secreted to assist in the humoral response (Th2 response). Particularly among patients with HIV infection, stronger CD4 responses were correlated with lower viral loads, whereas weaker CD4 responses were correlated with higher viral loads [63]. CD4 responses in HIV antiviral immunity + were determined by measuring CD4 T cell proliferation as well as production of IFN [63]. In addition to IFN, production of other cytokines, such as TNF- and IL+ 2, or chemokines, such as RANTES, MIP-1, and MIP-1, by CD4 T cells derived from peripheral blood mononuclear cells (PBMCs) of HIV patients in response to HIV antigens in vitro, was also detected [63-65]. Moreover, stronger CD4 helper responses 10 + were demonstrated to correlate with stronger CTL responses [66]. Since CD4 T cells + are targets of HIV infection, loss of CD4 T cells is rapid due to a combination of direct viral killing, increased rate of apoptosis, and CTL killing of infected cells. However, the CD4 + T cell arm is still partially functional according to above + investigations. Due to activation and proliferation of CD4 T cells, more cellular targets for HIV will be produced, which will further induce more HIV specific T cell responses [23, 67]. C. B Cell Mediated Immune Responses B cells also provide protection against HIV-1 by producing neutralizing antibodies, which have been demonstrated to block the interaction between the virus and the cellular receptors to prevent virus from entry into the cells [68]. B cell + activation requires CD4 T helper cells for providing the necessary signals, including one signal from cross-linking the antigens and the other signal from co-stimulation between CD40 ligand on T cells and CD40 on B cells, therefore the B cell response is usually delayed compared to other leukocytes [69]. Once B cells differentiate into plasma cells, they first produce IgM, and after affinity maturation through somatic hypermutation they undergo class switching to IgG and IgA [70]. Neutralizing antibodies against autologous virus have been shown to develop slowly, approximately 12 weeks or longer after HIV-1 transmission, and antibodies against heterologous virus were only effective in about 20% of the patients years after infection [71, 72]. In addition, neutralizing antibodies induced in clade B or clade C 11 HIV-1 infections were found to be specific only for the initial transmitted virus isolates [71, 73, 74]. Thus broadly reactive neutralizing antibodies are rarely generated. D. Implications for Vaccine Development In vaccine research, the focus of vaccine development against HIV-1 (the most common and pathogenic strain) is to induce both antigen-specific T cell responses and + neutralizing antibodies [75-77]. CD8 T cells play an important role on controlling the virus in the early stage of HIV-1 infection, when the immune system is still + functionally normal. An effective vaccine would be expected to induce CD8 T cell responses to multiple epitopes, especially to those that are conserved among different virus variants [78, 79]. In addition, broadly reactive antibodies need to be generated, which can target multiple neutralizing epitopes and induce a rapid secondary response. V. T Cell Signaling Cascades A. T Cell Activation Signaling Pathways T cells are activated via two signals, including the first signal mediated through TCR and the second signal mediated through the costimulatory molecule CD28, which then activate the cytosolic protein tyrosine kinases (PTKs), such as Lck and Fyn, to further phosphorylate the immunoreceptor tyrosine-based activation motifs (ITAMs) on the cytosolic side of the CD3 domain in the TCR complex (reviewed in [80]). Subsequently, zeta-chain associated protein kinase (Zap-70) is recruited to the 12 TCR/CD3 complex by ITAMs, and phosphorylated Zap-70 will further phosphorylate the downstream adaptor or scaffold proteins. Phospholipase C 1 (PLC1) is phosphorylated, leading to hydrolysis of phosphatidylinositol 4,5-bisphosphate (PIP2) to produce the second messengers, diacylglycerol (DAG) and inositol trisphosphate (IP3). DAG activates PKC and the mitogen-activated protein kinase (MAPK) pathways, which activate the transcription factor nuclear factor kappa-light-chainenhancer of activated B cells (NF-B) (reviewed in [81]), while IP3 induces intracellular Ca 2+ 2+ release from the ER, which causes extracellular Ca to be 2+ transported into cells via the calcium release-activated channels (CRAC). Ca -bound calmodulin activates the phosphatase calcineurin, which results in activation of transcription factors, including nuclear factor of activated T cells (NFAT), NF-B, and 2+ activator protein-1 (AP-1), with NFAT being the main target of the Ca (reviewed in [82]). signaling On the other hand, ligation of CD28 leads to activation of phosphoinositide 3-kinase (PI3K), which converts PIP2 to phosphatidylinositol 3,4,5trisphosphate (PIP3) resulting in activation of protein kinase B (Akt) (reviewed in [83]). Phosphorylated Akt inhibits glycogen synthase kinase-3 (GSK-3) from phosphorylating NFAT, and thereby promotes its nuclear translocation [84]. In addition, CD28 activates JNK pathway to phosphorylate transcription factors such as JUN and AP-1 [85]. Activation of the above transcription factors leads to increased expression of regulated genes, including cytokines and chemokines, such as IL-2, IFN and TNF [86, 87]. 13 B. T Cell Stimulation In Vitro Traditionally, T cells can be activated in vitro using pharmacological stimuli such as the phorbol ester, phorbol 12-myristate 13-acetate (PMA), in combination with the ionophore, ionomycin (Io). PMA can directly activate PKC bypassing the ligation of TCR [88, 89], while Io can directly increase the level of intracellular Ca 2+ [90]. In addition, a more physiologically relevant approach is used by stimulating T cells with anti-CD3 and anti-CD28 antibodies, which activates T cells through the TCR/CD28 signaling pathways. VI. Cannabinoids and Cannabinoid Receptors A. Cannabis Use Despite legal restriction, cannabis, also known as marijuana, is used for recreational purposes. More than 100 million Americans have tried cannabis at least once, and use of cannabis has increased particularly among young adults aged 18 to 25, about 21% of whom report using of marijuana every month [91]. Cannabis intoxication leads to relaxation, sleepiness, and mild euphoria (getting high), but might also increase the incidence of possible health issues, such as schizophrenia or depression [92]. As of 2013 in the US, 20 states and the District of Columbia, including Michigan in 2008, have legalized marijuana use for medical purposes. Medical marijuana is used in cancer patients to reduce nausea, vomiting, and pain that are associated with chemotherapy [93]. 14 In addition, marijuana is also used by approximately 25% of AIDS patients to stimulate appetite and ameliorate HIVassociated nausea, pain and wasting syndrome [94, 95]. However, effects of marijuana-derived compounds on immune function of these immunocompromised patients are still not well understood, which has led to our research interests. B. Cannabinoids Cannabinoids are a group of compounds originally extracted from the cannabis 9 plant.  -Tetrahydrocannabinol (THC) is the main psychoactive constituent among over 60 structurally-related congeners in the cannabis plant [96]. It is lipophilic and has a tricyclic ring structure, which contains a phenol ring with a 5-carbon alkyl chain, a central dihydrobenzopyran ring, and a mono-unsaturated cyclohexyl ring (Fig. 1) (see [97] for review). These terpenophenolic compounds are primarily produced by the female plants from their glandular trichomes [98]. In addition to THC, cannabidiol (CBD) and cannabinol (CBN) are also well studied compared with the other constituents. CBD is highly abundant in the fresh plant, making up about 40% of the whole extract, and CBN is the primary product during THC oxidation [99]. These phytocannabinoids exhibit varying binding affinities to the two identified cannabinoid receptors, cannabinoid receptor 1 (CB1) and 2 (CB2). For instance, THC exhibits similar affinity to CB1 and CB2 and is psychotropic, whereas CBD exhibits weak affinity to both receptors and is therefore, not psychotropic. CBN has weak affinity to CB1, but high affinity to CB2, which is still lower compared with THC, and therefore has little psychotropic activity [100-102]. 15 In addition to the plant-derived cannabinoids, there are endogenously produced cannabinoids by humans and animals, also known as endocannabinoids, which are ligands for CB1 and/or CB2 [103]. Two well-characterized endocannabinoids are Narachidonoylethanolamine or anandamide (AEA) and 2-arachidonoyl glycerol (2-AG) (Fig. 1). AEA is different in chemical structure from THC and is converted to the arachidonic acid, which is one of the most abundant fatty acid in the brain, by fatty acid amide hydrolase (FAAH). AEA exhibits affinity to CB1 and, to a lesser extent, CB2, whereas 2-AG binds to CB1 and CB2 with similar affinity [104]. Endocannabinoids are synthesized upon demand and function by maintaining homeostasis in the body related to neuronal system, immune system, metabolism, and energy [105]. The third category of cannabinoids is synthetic cannabinoids. These are the compounds produced in laboratories and structurally-related to the phytocannabinoids or having binding affinities to CB1 and/or CB2. For example, Dronabinol (Marinol) is synthetic THC, a drug approved by FDA to be used in cancer and AIDS patients for treating appetite loss, nausea, vomiting and pain [106]. Cyclohexylphenol (CP) 55,940, developed by Pfizer in the 1970s, and HU-210, developed by Raphael Mechoulam at the Hebrew University in the 1960s, are two other representative synthetic cannabinoids, which have structural similarity to THC, but possess more potency than THC [107] (Fig. 1). WIN 55,212-2 is another example of synthetic cannabinoid (Fig. 1), the non-classical prototypic aminoalkylindole cannabinoid, which is functionally similar but structurally different to THC [108]. 16 Figure 1. Chemical structures of selected cannabinoids. 17 C. Pharmacokinetics of THC Cannabis products are usually administered by two main routes, inhalation or oral consumption, and therefore the pharmacokinetic profile of THC is different. With inhalation by smoking cannabis cigarettes, THC is absorbed through the lungs and rapidly reaches the bloodstream and the brain within minutes [109]. The bioavailability of THC in the plasma ranged between 10 to 35%, depending on the number of puffs and the extent of breath holding during smoking [109, 110]. On the other hand, the absorption of THC after oral use is very slow, and peak plasma concentration reached up to 6 hours post administration, so the bioavailability of THC is less than when using the same amount for inhalation [111]. Within the low pH environment of the stomach and the gut, THC was found to degrade to CBD [112]. The plasma concentration of THC decreases as THC rapidly distributes to various tissues including liver, heart, lung, kidney, spleen, thyroid, fat, and pituitary glands, but only about 1% distributes to the brain [113]. Since THC is highly lipophilic, the tissue clearance half-life of THC was found to take about 7 days, and complete clearance after a single dose might take up to 30 days [114]. Further metabolism of THC mainly happens in the liver where it is hydroxylated and oxidized by the enzymes of the cytochrome P450 (CYP) complex, [115, 116]. The major metabolite after THC hydroxylation in humans is 11-OH-THC, and subsequent oxidation produces THC-COOH, which may be glucuronidated to 11-nor-9-carboxy-THC glucuronide [117]. In addition, 11-OH-THC may also be conjugated with fatty acids to be stored in tissues [118]. 18 D. Physiological Effects of THC THC has modulatory effects on various physiological systems, as reviewed in [119, 120]. For example, THC can lead to euphoria, anxiety, dysphoria, and increased sensory perception. The effect of THC on the cognition and motor functions may include fragmented thinking, disturbed memory, ataxia, and slurred speech. THC may also cause muscle relaxation, appetite stimulation, analgesia, and neuroprotection in hypoxia in the nervous system. In addition, THC may trigger hypertension, vasodilation, and enhanced heart rate in the cardiovascular system. Moreover, impaired cell-mediated and humoral immune responses have been demonstrated to be mediated by THC (see below for details). Furthermore, growth retardation, and impairment of fetal development might be associated with THC use as well. E. Toxicity of THC Studies have been conducted to compare acute oral toxicity of THC in different species. In rats, oral THC resulted in a median lethal dose (LD50) of 800-1900 mg/kg. In dogs and monkeys, oral THC at doses up to 3000 mg/kg and 9000 mg/kg, respectively, did not induce any fatality, but animals showed symptoms of drowsiness and ataxia [121]. In humans, no fatal cases have been demonstrated due to toxicity following acute THC treatment; however, death might be induced due to THCtriggered myocardial infarction [122, 123]. The harmful effects of medical marijuana use are suggested to include impaired cognition, decreased motor function, and cardiovascular diseases, such as hypertension and vasodilation, but these adverse effects are considered minimal or comparable to the adverse effects caused by other 19 medications [124, 125]. Whether long-term use of cannabis could cause any physical or cognitive dysfunction is still controversial, which might depend on the existing health condition of the individuals. F. Cannabinoid Receptors 1) CB1 and CB2 Both CB1 and CB2 are Gi/o protein-coupled receptors (GPCR) [97]. CB1 was originally cloned from rat cerebral cortex [102], and its human homolog was identified soon after [126]. The amino acid sequence of CB1 is very conserved among different mammalian species, with 81% homology between human and rat, and 93% homology between rat and mouse [101, 127-129]. CB1 is expressed throughout the body, but mainly expressed in the central nervous system (CNS), including cerebellum, hippocampus, and basal ganglia, to suppress the release of various neurotransmitters that are associated with multiple functions, such as cognition, memory, motor and metabolism [97, 130]. The psychotropic effect of THC is mediated through CB1 [102]. CB2 was originally cloned from HL60 after CB1 was identified [101]. The amino acid sequence of human CB2 shares 44% homology with CB1 [101]. CB2 is predominantly expressed in the periphery, including the immune system [101], and was recently found in neurons and glial cells of the CNS [131, 132]. In the immune system, CB2 expression is higher than CB1, with human B cells having the highest mRNA levels of CB2 than the other immune cells, including NK cells, monocytes, 20 + + neutrophils, CD4 T cells, and CD8 T cells [133, 134]. In addition, the expression levels of CB1 and CB2 can be altered under certain conditions. For example, CB1 mRNA expression was up-regulated in activated human T cell line [135], whereas CB1 mRNA and protein levels were down-regulated during B cell differentiation [136]. 2) Signaling Events Associated with CB1/CB2 As members of the GPCR family, cannabinoid receptors have the transmembrane structure with seven -helices, an extracellular glycosylated N-terminus, and an intracellular C-terminus [101, 102]. Both CB1 and CB2 are known to couple to Gi/o proteins, but they might have differential binding affinities. For example, human CB1 expressed in Spodoptera frugiperda cells showed similar binding affinity to Gi and Go proteins (the pertussis toxin sensitive G proteins), whereas CB2 was only found to bind to Gi protein [137]. Gi is known as an inhibitory G protein to activate downstream signaling events (see below), and Go is a similar protein in the Gi class, but its function is yet to be determined. In addition, the nature of the ligands for CB1 and CB2 determine the magnitude of Gi/o protein activation. For example, THC partially activated Gi protein through CB1; however WIN 55,212-2, HU-210, and AEA induced CB1-mediated activation of Gi to a much higher extent [137]. On the 21 other hand, HU-210 was the only agonist that could elicit CB2-mediated Gi activation to the maximal level. It has been demonstrated that stimulation of Gi/o protein by cannabinoids inhibited the activity of adenylate cyclase (AC), which decreased the levels of intracellular cyclic adenosine 3’,5’-monophosphate (cAMP) [134, 138]. The schematic description of signaling events mediated by CB1 and CB2 is illustrated in Fig. 2. The level of cAMP is critical for activation of protein kinase A (PKA) to further phosphorylate the cAMP-response element binding protein/activation transcription factor (CREB/ATF) family of transcription regulators, including CREB, ATF, and cAMP response element modulator (CREM) [138]. CREB is the best studied among the three transcription factors, and it could associate with other transcription factors for binding to cAMP regulatory element (CRE) DNA sequences in the promoter region of various genes [139, 140]. In fact, cannabinoids were demonstrated to inhibit PKA activity and the binding of CREB/ATF to CRE [140]. In addition, CREB might form heterodimers with transcription factors in the Fos and Jun family to bind to the promoter region of inducible regulatory factors, such as AP-1 [141, 142]. CBN was shown to suppress PMA/Io-induced CREB binding at the AP-1 proximal site in the IL-2 promoter region, which inhibited IL-2 expression [140]. 2+ Furthermore, cannabinoids, including THC, were found to increase intracellular Ca levels in resting splenic T cells, which was further enhanced in activated splenocytes potentially mediated through the calcium channel TRPC1 [143, 144]. CBN was shown to inhibit the activation of ERK MAPK, which functions downstream of the 22 calcium signaling pathways [145]. Moreover, it has also been reported that PKA can phosphorylate IB, resulting in dissociation of IB from NFB, to facilitate NFB activation and its bind to the NFB site in the promoter region of various genes, such as IL-2 [146]. By extension, suppression of PKA activity by cannabinoids might represent a mechanism that could explain impairment of NFB. In addition to binding to Gi/o proteins, studies have shown that under certain conditions cannabinoid receptors could also interact with the stimulatory G protein, Gs, which in turn increased AC activity upon CB agonist binding. It was shown that pertussis toxin treatment of CB1 attenuated HU-210-mediated inhibition of AC activity to increase cAMP accumulation and unmasked an agonist-mediated stimulation of the enzyme in the presence of a costimulant such as forskolin [147, 148]. WIN 55,212-2 at high concentrations was found to increase cAMP accumulation in the absence of pertussis toxin or costimulant, which was blocked by CB1 antagonist SR141716A [149]. It is still unclear whether CB1 simultaneously binds to Gs and Gi proteins or different populations of CBs are involved. In addition, different AC isoforms have also been suggested to contribute to the differential effects mediated by cannabinoids. Activation of the cannabinoid receptors CB1 or CB2 by the agonists, HU-210 and WIN 55,212-2, inhibited the activity of AC isoforms I, III, V, VI, and VIII, whereas activation of CB1 and CB2 by cannabinoids stimulated the activity of AC isoforms II, IV, and VII [150]. 23 Figure 2. Simplified signaling pathways associated with CB1/CB2. 24 3) Other Targets for Cannabinoid Ligands It has been demonstrated that some effects of cannabinoids are not mediated through CB1 or CB2, suggesting the existence of other targets for cannabinoid ligands, which are classified as nonclassical cannabinoid receptors. The established targets include GPCRs, ion channels and nuclear receptors. For example, orphan GPCRs, such as GPR18, GPR55, and, GPR119, have recently been found to interact with phytocannabinoids, endocannabinoids, or synthetic cannabinoids, although the functions of these GPCRs remain unclear [107]. GPR55 shares low amino acid sequence similarity to CB1 (13.5%) and CB2 (14.4%), and it is most closely related to lysophosphatidic acid (LPA) receptors (30% similarity) [151, 152]. GPR55 has been shown to couple with G13, Gq, G12, or G13, which activate 2+ GTPase RhoA and PLC for signal transduction, and therefore affect intracellular Ca , actin cytoskeleton, and growth of neural pathways (reviewed in [97]). THC and Narachidonoyl glycine have been shown as ligands of GPR18 to affect intracellular Ca 2+ mobilization [153]. Oleylethanolamide (OEA) has high affinity for GPR119, which is involved in controlling glucose-dependent insulin release in the pancreatic tissues [154]. In addition, the transient receptor potential vanilloid 1 (TRPV1) has been reported to interact with AEA, N-arachidonoyl dopamine, and a lower affinity with CBD, but not 2-AG. These agonists affect intracellular Ca 2+ levels, cation currents in the neurons, and release of vasodilatory peptides from sensory neurons [155, 156]. Apart 25 from TRPV1, other TRPV channels and other TRP channels, such as TRPV2, -3, -4, 5, -6, TRP ankyrin 1 (TRPA1), TRP canonical 1 (TRPC1) have also showed evidence of being activated by cannabinoids (reviewed in [97]). Moreover, endogenous cannabinoids, such as AEA, can activate calcium, potassium, and sodium channels (reviewed in [97]). Other than the putative receptors described above, peroxisome proliferatorsactivated receptors (PPARs) have been found to interact with endocannabinoids, phytocannabinoids, and synthetic cannabinoids, such as AEA, 2-AG, THC, and WIN55212 (review in [97]). For examples, 2-AG was found to suppress IL-2 production through impairment of NFAT activity, due to binding of a putative cyclooxygenase (COX)-2 metabolite of 2-AG, 15-deoxy-12,14-prostaglandin J2glcyerol ester, to PPAR receptor [157, 158]. So far, there is not enough evidence indicating these other receptors or channels as identified cannabinoid receptors, so they are considered as putative targets for cannabinoid ligands, and therefore, further investigations are needed. VII. Cannabinoid and CB1/CB2-Mediated Immune Modulation Over forty years of studies have demonstrated that THC possesses broad modulatory effects on the immune system in a CB1/CB2-dependent and -independent manner. In this section, the effects of THC and the role of CB1/CB2 on multiple immune cell populations will be discussed. 26 A. T Cells T cells have been demonstrated as one of the most sensitive targets of cannabinoids. THC has been demonstrated to modulate various T cell functions, including T cell activation, proliferation, and production of cytokines. For example, THC and other cannabinoids were found to suppress mitogen-induced T cell activation, proliferation, and production of IL-2, IFN, and other cytokines [140, 159162]. In addition, inhibition of DNA binding activity of transcription factors regulating IL-2, e.g. NFAT and AP-1, as well as temporal dysregulation of 2+ intracellular Ca signaling in resting T cells by THC or CBN have been suggested, at least in part, as mechanisms for cytokine suppression [143, 144, 159, 161, 163]. Effects of THC on CTL functions as well as the associated mechanisms have also been extensively studied. For example, cytolytic function of CTLs was reduced by THC and 11-OH-THC in response to allogeneic or 2,4,6-trinitrophenyl (TNP)modified stimulators in vivo and in vitro, which was due to suppression of + differentiation and maturation of precursor CD8 T cells or cytolytic activity of CTL after binding to the target cells [164]. Moreover, CTL activity to herpes simplex virus type 1 (HSV1)-infected cells as well as CTL cytoplasmic polarization toward the infected target cells were suppressed by THC [165]. Furthermore, kinetic and time of addition studies indicated that the effect of THC on T cell functions occurred in the early stage during activation [166]. This was consistent with a recent finding using an in vitro P815 allogeneic model to elicit CTL responses based on MHC mismatch, which revealed that THC suppressed CTL activation, cytolytic activity, and IFN production, when present during the elicitation phase but not the effector phase [167]. 27 + Likewise, CD4 T cell-derived cytokine production was modulated by THC and other cannabinoids. In vivo, both acute and chronic THC treatment in mice was found to cause a decrease in serum IL-2 and IFN levels as well as the number of T cells [168]. In addition, with allogeneic DC stimulation in vitro, T cell proliferation and Th1 cytokine (IFN, IL-2) production were suppressed by THC, but Th2 cytokine (IL-4, IL-5) expression was increased by THC, which were attenuated by CB2 antagonist, SR144528, suggesting a modulation of the balance between Th1/Th2 cytokines in the T cells by THC in a CB2-dependent manner [169]. Interestingly, depending on experimental conditions, such as cannabinoid concentration, cell types, and specific immune stimuli, enhancement of T cell responses by cannabinoids has also been reported in a number of studies. For example, it was first demonstrated that THC treatment of anti-CD3 stimulated spleen cells resulted in an enhancement in cellular proliferation, whereas using cells from lymph nodes or concanavalin A (conA) or phytohemagglutinin for stimulation led to THC-mediated suppression [170]. In addition to differences between various stimuli and organ sources, the age of animals was also suggested to play a role in cannabinoid-mediated differential effects [171]. Moreover, CP55,940 at nanomolar concentrations was found to increase the expression of IL-8 and beta-chemokine monocyte chemotactic protein-1 (MCP-1) in CB2-transfected HL60 cells, which was sensitive to pertussis toxin [172]. Although HL60 cells are not T cells, these studies are mentioned here to support the differential effects of cannabinoids in T cells. Low concentrations of cannabinoids were also observed to produce enhancement in B cells 28 [173]. Furthermore, positive regulation of IL-2 expression by cannabinol has been observed when EL4 T cells were suboptimally activated with low concentrations of PMA, due to binding of NFAT to a distal site in the IL-2 promoter region [174, 175]. In addition, CBN-mediated enhancement was also observed with THC, CP55,940 and CBD, which was not abrogated by CB2 antagonist SR144528, suggesting a CBindependent mechanism. B. B Cells Since B cells express higher levels of CB2 compared with other immune cells as discussed earlier, THC and other cannabinoids have shown modulatory effects on B cell functions, such as proliferation and antibody production responses, mediated through this receptor. For example, many studies have found that cannabinoids suppress plasma cell formation in response to the T cell-dependent antigen, sheep red blood cells (sRBCs), both in vivo and in vitro [162, 176-178]. To identify the cellular targets for THC, further studies have shown that the antibody forming cell (AFC) response to T cell-independent antigen, DNP-Ficoll, or polyclonal B cell activator, LPS, was not suppressed by THC, and T cell proliferation in response to sRBCs was indeed suppressed by THC, collectively suggesting that THC might target T cells rather than directly affecting B cells [179]. However, THC and other cannabinoids, such as 2-AG, were also found to suppress proliferation of B cells in response to LPS [180, 181]. In addition, recent studies have shown that the CD40 ligand (CD40L)induced IgM AFC response was suppressed by THC [162]. Moreover, studies from this laboratory have demonstrated that THC decreased IgM production by human 29 peripheral blood B cells activated by CD40L plus recombinant cytokines (IL-2, -6, and -10). THC also suppressed expression of activation marker CD80 on B cells as well as STAT3 phosphorylation [182], which is involved in B cell survival, proliferation, and differentiation [183]. Furthermore, in human studies, serum immunoglobulin levels in chronic marijuana smokers were altered; however, which immunoglobulin isotype, including IgG, IgM, IgA, IgD, and IgE, was changed in particular is still controversial among different studies [184-186]. Interestingly, enhancement of B cell responses by cannabinoids has also been observed. For example, ligation of CD40-induced proliferation of B cells isolated from human tonsillar tissue, which was in fact enhanced rather than suppressed by THC at nanomolar concentrations [173]. C. NK Cells NK cell function is also susceptible to modulation by THC. The lytic activity by splenic NK cells on target cells was found to be suppressed by THC and its primary metabolite 11-OH-THC in a dose-dependent manner [187]. Similarly, the cytolytic activity of IL-2 stimulated cloned NK cells against YAC-1 and EL4 target cells and proliferation of NK cells were suppressed by THC and 11-OH-THC, but the binding of cloned NK cells to target cells was not inhibited by THC [188]. Suppression of IL2 receptor expression by THC was suggested as a mechanism by which THC mediated inhibition on the killing activity of NK cells [189]. In addition, migration of NK cells induced by 2-AG was abolished by CB2 antagonist, SR144528, which suggested that the 2-AG-mediated effect occurred in a CB2-dependent manner [190]. Also, THC at 30 micromolar concentrations attenuated the protein and mRNA production of TNF from human NK cells in response to C. albicans [191]. ConA-induced IFN production by NK cells was reduced by THC, and both CB1 and CB2 antagonists reversed the effect [192]. D. APCs APCs are critical cellular populations to link the innate and adaptive immune responses. Antigen processing and presentation as well as cytokine production by APCs were reported to be modulated by THC and other cannabinoids [193]. For instance, DCs and macrophages are known to express CB1 and the voltage-gated potassium (KV) channels [194, 195]. The CB1 agonist arachidonylcyclopropylamide as well as AEA attenuated KV channel-mediated outward currents in DCs, which resulted in a decrease of surface MHC II expression on DCs and therefore a reduced T cell response, suggesting the modulation of DC function by cannabinoids via CB1 signaling [196]. Most recently, THC was found to suppress LPS-induced maturation of bone marrow-derived DCs (bmDCs), as evidenced by reduced MHC II, CD80, and + CD86 expression, which in turn decreased IFN production by CD8 T cells after the co-culture of bmDCs and splenocytes [197]. In addition, THC was shown to suppress gene expression as well secretion of cytokines, such as IL-1, IL-1, TNF, and IL-6, by activated macrophages [198-200]. Moreover, THC was demonstrated to inhibit NO production in macrophages in the CNS, by affecting NOS gene expression or synthesis via acting on the transcription factor NF-B/RelA [146, 201]. Furthermore, the level of CB2 expression in the macrophages was suggested to be altered depending 31 on the activating stage of the cells. THC and other CB2 agonists were reported to inhibit chemotaxis as well as antigen processing and presentation in macrophages that were at the responsive or primed stage [202, 203]. E. In Vivo Host Resistance Models Since THC and other cannabinoids have been shown to modulate immune function in humans and animals, the possible adverse effects on human health and disease have been a concern. Animal models have been used to study the effects of cannabinoids on host resistance to infection with viral and microbial pathogens. Due to the immunosuppressive properties of THC, increased host susceptibility to infections has been reported in a number of studies. For example, in a mouse model with Legionella pneumophila infection, serum levels of Th1-promoting cytokines, IL-12 and IFN, were increased in Balb/c mice, which were suppressed by THC treatment administered prior to bacterial challenge. In contrast, Legionella-induced production of Th2-promoting cytokine, IL-4, was further increased by THC treatment, which was also observed in IL-4 knockout mice, suggesting that THC-mediated suppression of Th1 immunity to Legionella infection was due to a decrease in IL-12 and IFN production but not an increase in IL-4 production [204]. Moreover, it was indicated that both CB1 and CB2 played a role in THC-mediated effects on cytokine production by using CB antagonists [205]. Recently, this laboratory has established a mouse model of influenza virus challenge to study the effects of THC and the role of CB1 and CB2 on host resistance 32 to influenza. THC increased viral load in the lungs, which was, in part, through + + decreasing the infiltration of CD4 , CD8 , and macrophages to the lung [206]. In -/- -/- addition, CB1 and CB2 double knockout mice (CB1 CB2 ) showed a lower viral burden in the lung than wild-type mice. THC increased the viral load in both genotypes when compared to the respective vehicle controls. Also, inflammatory cell -/- -/- infiltration, T cell activation and cytokine production were increased in CB1 CB2 mice, when compared to wild-type mice [207, 208]. These findings suggest that THCmediated effects on airway immune responses to influenza may be through CB1/CB2dependent and -independent mechanisms. Further studies have implied that the -/- -/- exacerbated immune response phenotype in CB1 CB2 mice might be due to a more mature function of APCs [208], and THC treatment suppressed APC-stimulated + cytokine production by CD8 T cells in a CB1/CB2-dependent manner [197]. F. Involvement of CB1/CB2 in Cannabinoid-Mediated Immune Modulation To study mechanisms for immune modulation by cannabinoids, one crucial factor that needs to be considered is the role of cannabinoid receptors involved in the biological activities of cannabinoids. Receptor antagonists, receptor agonists, or receptor knockout mice are the important and widely used approaches in identifying receptor-specific effects of cannabinoids. Both CB1/CB2-dependent and -independent mechanisms have been reported to be involved in cannabinoid-mediated immunomodulatory effects. For example, suppression of PMA/Io-induced IL-2 33 production in mouse splenocytes by cannabinoids, including CBN, CBD, and WIN55,212, was not attenuated by a CB1 antagonist SR141716A and a CB2 antagonist SR144528, suggesting a CB1/CB2-independent mechanism [144]. In addition, elevated intracellular calcium concentration in the presence of cannabinoids in resting splenocytes was attenuated by both CB1 and CB2 antagonists, suggesting a CB-dependent mechanism [144]. However, the role of CB1 and CB2 in cannabinoidmediated actions based on receptor agonists and antagonists studies has been controversial. For example, SR141716A and SR144528 have been found to have inverse agonist activity by inhibiting GPCR signaling [209-211]. Other studies have shown that SR141716A and SR144528 exhibited partial agonist properties mimicking the actions of CB1/CB2 agonist, HU-210, by decreasing left ventricular developed pressure and maximum rates of contraction and relaxation [212]. Additional studies have also suggested off-target effects of CB antagonists. CB1 or CB2 antagonists 2+ attenuated THC-, CBN-, and HU-210-mediated elevation of intracellular Ca levels - in T cells, which was in fact independent of CB1 and CB2 as demonstrated using CB1 /- -/- CB2 mice, suggesting the existence of putative receptors that are also sensitive to CB antagonists [163]. -/- -/- CB1 CB2 mice (on a C57Bl/6 background) have been used as an experimental tool to study the role of both CBs in cannabinoid-mediated immune modulation, which would avoid the issues with antagonists as described above. Although the 34 specific CB played a role in mediating the effects in these studies can not be discerned -/- -/- by using CB1 CB2 mice, the involvement of one or both of these receptors can be -/- -/- determined. As described by Springs et al. [162], CB1 CB2 mice do not show any phenotypic defects, except for a higher tendency to develop conjunctivitis in aged animals. The proportions of various splenic cellular populations, including T cells, B -/- -/- cells, and macrophages, were similar between wild-type and CB1 CB2 mice. Also, mitogen-induced lymphocyte proliferation and cytokine production (IL-2 and IFN) were comparable between the two genotypes. In response to sRBC in vivo, the IgM -/- -/- AFC response was suppressed by THC in wild-type mice but not in CB1 CB2 mice. In vitro, the LPS-induced IgM response was not modulated by THC in both genotypes, but the IgM response induced by CD40L was suppressed by THC in wild-/- -/- type but not in CB1 CB2 mice. Together, these findings suggest that the THC- mediated effects on immune responses may involve both a CB1/CB2-dependent and independent mechanism. In addition, the lack of CB1 and CB2 potentially leads to an enhanced immune -/- - response. For example, in a model for cutaneous contact hypersensitivity, CB1 CB2 /- mice developed chronic ear ulceration with contact to nickel-containing ear clips, and exhibited exacerbated ear swelling compared with wild-type mice in response to the contact sensitizer, 2,4-dinitrofluorobenzene [213]. Additionally, CB antagonists SR141716A and SR144528 further enhanced allergic inflammation, whereas CB 35 agonist, THC, attenuated the responses, suggesting the involvement of endocannabinoids in the protection against allergic responses [213]. Similarly, in a mouse model challenged with influenza, there was a more robust immune response in -/- -/- CB1 CB2 mice in the absence of THC, as evidenced by a lower viral load in the lung when compared with wild-type mice. -/- -/- inflammation and lung injury in CB1 CB2 THC also increased pulmonary mice when compared with wild-type mice [206, 207]. VIII. Rationale and Objectives As described above, approximately 25% of HIV/AIDS patients use marijuana to ameliorate nausea, pain, and wasting syndrome that accompany with HIV infection [94]. However, the effects of cannabinoids on immune function of these immunocompromised patients are still not well understood. To date, only a few studies have demonstrated that short-term or chronic THC treatment neither increased + + HIV viral load nor decreased CD4 and CD8 lymphocyte counts using patients or nonhuman primate models, therefore suggesting that cannabinoid treatment does not have adverse effects on disease progression in the early stage of infection [214-216]. In-depth studies are needed to investigate the involvement of cannabinoids and cannabinoid receptors on immune responses to HIV antigens, especially on antigenspecific responses. The objective for the studies presented in this dissertation was to test the hypothesis that THC impairs the anti-HIV immune responses in a CB1/CB2dependent mechanism. 36 The specific aim for the first part of this dissertation research was to characterize the effect of THC and the role of CB1/CB2 on the elicitation and effector functions of antigen-specific CD8 mechanisms. + T cells, and to investigate the associated molecular A novel mouse model of HIVgp120 antigen presentation was established, which was designed to elicit a broad antiviral immune response against multiple gp120-derived antigenic epitopes, to closely resemble aspects of host antiviral immunity in the initial phase of HIV infection. Since both cell-mediated and humoral immune responses against gp120 have been reported previously, gp120 was used as a representative HIV protein against which to induce an immune response in our models [217-219]. A C57Bl/6 mouse-derived DC line, DC2.4, and the T lymphoma cell line, EL4, were transduced with gp120 (DC2.4gp120 and EL4gp120, respectively), and used as APCs and target cells to restimulate CTL, respectively. DC2.4gp120 cells were co-cultured with mouse spleen cells to activate T cells. The effect of THC on CTL elicitation and IFN production was investigated. The cellular and molecular mechanisms responsible for THC-mediated effects were then determined using more titratable T cell activators: specifically, PMA/Io or antiCD3/CD28. These studies provide a possible mechanism to explain the widely reported discrepancy regarding cannabinoid effects on immune responses in the literature. The specific aim for the second part of this dissertation research was to determine the effect of THC and the role of CB1/CB2 on antigen-specific immune responses in 37 vivo. In vaccine research, the focus of vaccine development against HIV-1 is to induce both antigen-specific T cell responses and neutralizing antibodies [75-77]. Plasmid DNA encoding viral proteins is often used as a more potent vaccine than using viral proteins directly in animal models to induce CTL, Th cell, and antibody responses [220]. Here, plasmid pVRCgp120, encoding HIV envelope protein gp120, is used as the immunogen, which is injected intramuscularly into mice in this model system. To assess gp120-specific responses, splenocytes from treated mice were restimulated with immunodominant gp120-derived peptides. The effect of THC was -/- -/- studied in C57Bl/6 wild-type mice as well as CB1 CB2 mice. Our findings provide an in-depth understanding of cannabinoid effects on HIV antigen-specific immune responses in vivo. 38 MATERIALS AND METHODS I. Chemicals and Reagents THC and CBD were obtained from the National Institute on Drug Abuse (Bethesda, MD). RPMI 1640 media, penicillin/streptomycin, MEM non-essential amino acids (NEAA), sodium pyruvate, and L-glutamine were obtained from Gibco Invitrogen (Carlsbad, CA). Restriction endonucleases and ligase were purchased from New England Biolabs (Ipswich, MA). Unless otherwise specified, all other reagents were purchased from Sigma (St. Louis, MO). II. Mice Female C57Bl/6 or B6C3F1 mice were purchased from Charles River (Portage, MI). -/- -/- Female CB1 CB2 mice on C57Bl/6 background were bred at the University Laboratory -/- -/- Animal Resources (Michigan State University). CB1 CB2 mice were a kind gift from Dr. Andreas Zimmer at the University of Bonn, Germany. Mice were used between the ages of 8-12 weeks and randomly assigned to experimental groups. They were provided with food (Purina Certified Laboratory Chow) and spring water ad libitum. Rooms were kept on a 12-hr light/dark cycle at 21-24°C and 40-60% humidity. All experiments followed the guidelines set forth by Michigan State University Institutional Animal Care and Use Committee. 39 III. Cell Culture Conditions DC2.4 (a kind gift from Dr. Kenneth Rock at University of Massachusetts), EL4, DC2.4gp120, and EL4gp120 cells were maintained in complete RPMI media (10% bovine calf serum [BCS; HyClone, Logan, UT]; 100 units/ml penicillin and 100 µg/ml streptomycin [Pen/Strep]). HEK293T cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM) with 10% BCS and Pen/Strep. Splenocytes were isolated aseptically from female C57Bl/6 mouse spleens unless otherwise specified and made into single-cell suspensions. In the elicitation phase of the gp120 response (see below), “CTL media” (RPMI plus 5% BCS, Pen/Strep, 2 mM MEM NEAA, 2 mM sodium pyruvate, 2 mM Lglutamine, and 50 µM 2-mercaptoethanol) supplemented with 5 ng/ml recombinant mouse IL-12 was utilized (R&D Systems, Inc., Minneapolis, MN). In the effector phase, 2% BCS and 1 ng/ml IL-12 were substituted in the CTL media. In the other experiments, splenocytes were cultured in complete RPMI media with 2% BCS, Pen/Strep and 50 µM 2-mercaptoethanol. Human leukocyte packs were obtained commercially from anonymous donors (Gulf Coast Regional Blood Center, Houston, TX). PBMC were enriched from each pack by density gradient centrifugation using Ficoll-Paque Plus (GE Healthcare, Piscataway, NJ). Human PBMC were cultured in complete RPMI media with 2% BCS and Pen/Strep. IV. Lentiviral Plasmid Construction HIV gp120 cDNA, derived from the HIV-1 IIIB (HXBc2) strain, was amplified from plasmid, pVRC2000-gp120 [221] (a kind gift from Dr. Norman Letvin, Beth Israel Deaconess Medical Center, Boston, MA) using primers gp-FP and gp-RP (Table 1). To 40 generate pLexPuro-gp120, the gp120 amplicon was digested with SpeI and NotI (underlined in gp-FP and gp-RP, respectively) and ligated into the pLexMCS plasmid r containing the IRES-puromycin resistance (puro ) gene cassette (Trans-Lentiviral™ pLex Packaging System, Open Biosystems, Huntsville, AL). A similar plasmid, pLexNeor gp120, was made by replacing the IRES-puro cassette of pLexMCS with an IRESr r neomycin resistance (neo ) cassette and inserting gp120 as above. The IRES-neo cassette was amplified from plasmid pIRESneo3 (Clontech Laboratories, Inc. Mountain View, CA) using primers IRES5’ and Neo3’ (Table 1) with respective XhoI and HpaI sites. Each ligate was transformed into competent E. coli cells, Stbl3™ (Invitrogen) or 10-beta (C3019, New England Biolabs). Transformants were selected on LB plus ampicillin plates. Individual colonies were expanded in LB broth plus ampicillin and 0.5% glycerol. Plasmids were isolated using a HiSpeed Plasmid Maxi kit (Qiagen, Valencia, CA). 41 Table 1. Sequences of PCR primer/probe sets. Primer/probe Sequence (5’ – 3’) sets gp-FP TACGTACTAGTTGGCAAGAGAAGGCCAGAAAC gp-RP TGACTGCGGCCGCTCCTCAGCTCTTCTCGCTCT IRES5’ AAAAAACTCGAGTATGCATCTAGGGCGGCCAATTC neo3’ AAAAAAGTTAACTTCAGAAGAACTCGTCAAGAAGGCGA gp120 (3’ end) CCTTGCCCACCTTCTG gp120 (5’ end) CACCAGCACCACCTCC puror CCGGGAACCGCTCAAC neor TCCGGTGCCCTGAATG 42 V. DC2.4gp120 and EL4gp120 Cell Line Construction Packaging plasmids (Trans-Lentiviral™ pLex Packaging System, OpenBiosystems) were co-transfected with either pLexPuro-gp120 or pLexNeo-gp120 into HEK293T cells according to the manufacturer’s protocol. Lentiviral particles were collected and r incubated with 6 mg/ml polybrene at RT for 5 min. The polybrene/particle (gp120-puro r or gp120-neo ) mixtures were added to DC2.4 or EL4 cells, respectively, in 24-well plates (750 l/well), and centrifuged at 800 x g for 2h. Media was replaced with complete RPMI media and incubated for 48h. Transduced cells were cultured in 1.5 g/ml puromycin for DC2.4gp120 cells and 400 g/ml of G418 for EL4gp120 cells. Cloning by limiting dilution was performed by seeding heterogeneous mixtures of transduced cells at approximately one cell per three wells in 96-well U-Bottom plates (200 l/well) to select individual clones, which were cultured for screening by Real Time PCR (primer/probe sets in Table 1) and Western blotting. VI. In Vivo Immunizations Using Plasmids The control vector plasmid pVRC2000 was constructed by digestion of the pVRCgp120 plasmid with the restriction enzyme, ScaI (New England Biolabs, Ipswich, MA), to remove the gp120 cassette and ligated with T4 DNA ligase (New England Biolabs). Each plasmid was transformed into competent E. coli 10-beta cells. Transformants were selected on LB plus kanamycin plates. expanded in LB broth plus kanamycin. Individual clones were Plasmid DNA was isolated using Endofree plasmid Mega Kit (Qiagen, Valencia, CA), according to the manufacturer’s instructions. 43 -/- -/- Both C57Bl/6 and CB1 CB2 mice (n=5/group) were randomly placed into different treatment groups one week prior to the start of the experiment. The prime-boost immunization scheme, the route of administration and the amount of DNA per injection were based on studies in the literature and then optimized for the present investigation in order to stimulate robust antigen-specific immune responses [222-225]. Mice were injected intramuscularly (i.m.) in each quadricep muscle with 50 g pVRCgp120 or pVRC2000 plasmid DNA in 50 l final volume of Endofree water. Injections were performed twice per mouse over a one-week interval (day 1 and day 8). VII. Real Time PCR For gp120-transduced cells, total RNA was extracted using the SV Total RNA Isolation System (Promega, Madison, WI) following manufacturer’s protocols. For 6 cytokine production, splenocytes (5 x 10 cells) were treated with vehicle (VH, 0.1% ethanol) or CBD for 30 min at 37°C, and activated with 10x P/I (4 nM/0.05 µM) or 1x P/I (40 nM/0.5 µM) for 2, 6 or 24 hr at 37°C. Total RNA was isolated using the TRI Reagent method as previously reported [159]. In brief, cells were harvested, placed in TRI Reagent and frozen at -80°C. Upon thaw, bromochloropropane was added (0.1 ml per 1 ml TRI reagent) and phases were separated by centrifugation at 12,000 x g for 15 min. After isolation of the aqueous phase, RNA was precipitated with isopropranol, washed twice with 75% ethanol, dried at room temperature and suspended in nuclease-free water. In all experiments, equal amounts of RNA were reverse transcribed with random primers using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA). Real time PCR amplification and detection was performed with Taqman 44 primer/probe sets for mouse IL-2 (Mm00434256_m1, Applied Biosystems), or IFN (Mm00801778_m1, Applied Biosystems) on a 7900 HT Fast Real Time PCR System (Applied Biosystems). Assessment of genomic DNA contamination was performed using controls without reverse transcriptase (RT) during the cDNA reaction and demonstrated more than 10 Ct value difference between samples with and without RT for 18S and no detectable signal or above 35 Ct value for target genes in samples without RT. Foldchange values were calculated using the Ct method [226]. VIII. Western Blotting Analysis Cytosolic protein was isolated from DC2.4gp120 and EL4gp120 clones. Briefly, cells were washed with phosphate buffered saline, lysed with lysis buffer (1% Igepal CA-630, 0.5% sodium deoxycholate, 50 mM Tris, 150 mM NaCl, pH 8.0, supplemented with EDTA containing protease inhibitors [Complete, Mini Protease Inhibitor Cocktail Tablets, Roche Applied Science, Indianapolis, IN]) at ≤ 4 x 107 cells/ml on ice for 5 min, and centrifuged at 25,000 x g at 4°C for 5 min. Supernatants were collected, and 20 g protein for each sample was loaded per lane into a 7% denaturing polyacrylamide gel. Proteins from parental DC2.4 or EL4 were negative controls and 500 ng purified gp120 protein (HIV-196ZM651 gp120, NIH AIDS Research & Reference Reagent Program) was 7 the positive control. Nuclear protein was isolated from splenocytes (5 x 10 cells) that were treated with VH or CBD for 30 min at 37°C, then activated with 10x or 1x P/I for 30 min at 37°C as previously described [227]. Protein extract was then loaded into an 8% denaturing polyacrylamide gel to ensure resolution of several bands that correspond to 45 NFAT2 [228]. All protein samples were transferred to nitrocellulose membranes using a semi-dry electroblotter in transfer buffer (0.025 M Tris, 0.2 M glycine, 10% methanol for NFAT or 20% methanol for gp120). Membranes were blocked in 5% milk or 1% bovine serum albumin (BSA) in tris-buffered saline in Tween (0.01 M tris, 0.15 M NaCl, 0.1% Tween-20) overnight at 4°C. Gp120 was detected using a polyclonal HIV-1 gp120 antibody (Prosci, Inc., Poway, CA), and NFATc1 (NFAT2) was detected using an antimouse NFAT2 antibody (Affinity Bioreagents/Thermo Scientific, Rockford, IL), followed by horseradish peroxidase-conjugated sheep anti-mouse IgG (GE Healthcare, Buckinghamshire, United Kingdom) and west pico chemiluminescent substrate development (Thermo Scientific). IX. Elicitation of Splenocytes with DC2.4gp120 Cells The experimental model for in vitro elicitation of gp120-specific T cells is illustrated in Fig. 3. DC2.4gp120 cells were irradiated with 35 gray of X-ray to prevent proliferation, 7 washed 3 times, and adjusted to 1 x 106 cells/ml. Splenocytes (1 x 10 cells/ml) were treated with THC (1, 5, 10, and 15 M) or VH (0.1% ethanol) for 30 min, and then cocultured with DC2.4gp120 cells (100 l of each cell type in 96 U-bottom plates) for 5 days of elicitation in 5% CTL media. X. THC Treatment In Vivo THC was obtained from the National Institute on Drug Abuse (NIDA, Bethesda, MD). Two days post the injection of pVRCgp120 or pVRC2000 plasmid (day 1 and day 8), 46 mice received corn oil (CO) vehicle or THC (75 mg/kg/day per mouse) by oral gavage for three consecutive days (day 3-5 and day 10-12) (Fig. 4). The dose and dosing regimen were chosen based on historical data [206, 207, 229, 230], and the resulted THC concentrations in mouse serum are expected to be comparable with the finding from a previous study, in which 75 mg/kg of THC for 5 consecutive days resulted in 66.2 ng/ml of THC in mouse serum [206]. This is physiologically relevant to human plasma levels of THC, which typically ranged from 57 to 268 ng/ml in marijuana smokers [229]. There were total of four experimental groups for each genotype: CO-pVRC2000, COpVRCgp120, THC-pVRCgp120 using n=5/group and naïve (NA) using n=3/group. XI. Necropsy and Tissue Preparation Three days following the last THC dose, mice were sacrificed by decapitation for immunological assays. Trunk blood was collected into plasma separator tubes on ice (BD Medical, Franklin Lakes, NJ), and centrifuged at 3500 rpm (2536 x g) for 15 min at 4C. Splenocytes were isolated from the spleen and made into single-cell suspensions in icecold RPMI 1640 medium (Gibco Invitrogen, Carlsbad, CA). Splenocytes were centrifuged at 300 x g for 4 min and resuspended in RPMI supplemented with 2% bovine calf serum (BCS), 100 U/ml penicillin, 100 g/ml streptomycin, 2 mM non-essential amino acids, 2 mM sodium pyruvate, 2 mM L-glutamine, and 50 M 2-ME (Gibco Invitrogen). 47 Figure 3. Experimental model of in vitro elicitation of gp120-specific T cells. 48 Figure 4. Protocol for the administration of THC or CO into the mice immunized with pVRCgp120/pVRC2000. 49 XII. Enumeration of Cytokine Secreting Cells by ELISPOT Assays 7 For in vitro studies, after five days of co-culture, elicited effector splenocytes (2 x 10 6 cells/ml) were restimulated with EL4gp120 or parental EL4 target cells (1 x 10 cells/ml) in 2% CTL media. 50 l of each for effectors and targets were plated in ELISPOT wells overnight. A CTL only group (no target restimulation) was included as a control to assess background nonspecific CTL activity. 6 For in vivo studies, 1 x 10 /well splenocytes in 100 l of 2% BCS RPMI medium were restimulated with gp120-derived peptides ex vivo. A 211-peptide library, comprising the HIV-1 IIIB envelope protein region, was obtained from the NIH AIDS Reagent Program (https://www.aidsreagent.org/). These peptides are 15 amino acids in length, with 11-amino acid overlaps between sequential peptides. Splenocytes were restimulated ex vivo overnight with 100 l media containing one of the following treatments: 1). 53 pools of 4 peptides (except for 3 peptides in pool 53) from the library (0.2 g/well of each peptide) individually; 2). 0.2 g/well of the most immunogenic peptide from the library; 3). VH control for the peptides (0.1% dimethyl sulfoxide [DMSO]); 4). no treatment (NT). ELISPOT assays were performed as described previously [231]. Ninety six-well Multiscreen-HA filter plates (Millipore, Billerica, MA) were pre-coated with purified mouse anti-IFN or anti-IL-2 antibody (10 g/ml, BD PharMingen, San Diego, CA) overnight, washed and blocked with phosphate buffered saline (PBS) containing 5% bovine serum albumin (BSA), prior to the addition of cells (described above). ELISPOT plates were then washed and incubated with biotin-conjugated mouse anti-IFN or IL-2 antibody (1 g/ml, BD PharMingen), followed by the incubation with a 1/1000 dilution of 50 streptavidin peroxidase (Sigma, St. Louis, MO). Spots were developed with AEC Chromogen Kit (Sigma) and enumerated with the Cellular Technology ImmunoSpot system (Cellular Technology, Shaker Heights, OH). XIII. Cell Staining for Flow Cytometry 6 For in vitro studies, splenocytes (5 x 10 /ml) were labeled with 5 M CellTrace Violet dye (Cell Proliferation Kit, Invitrogen) following the manufacturer’s protocol. Proliferation was assessed on days 1-5 post co-culture of labeled splenocytes with DC2.4gp120 cells, and IFN production was assessed after restimulation as described above. In some studies, splenocytes were treated with VH (0.1% ethanol) or CBD at various concentrations for 30 min, then activated with 1x P/I (4 nM/0.05 µM) for 18-24 hr at 37°C. Cells were incubated with LIVE/DEAD Fixable Near-IR or Aqua Dead Cell Stain (Invitrogen) to assess cell viability according to the manufacturer’s instructions. Fc receptors were blocked with purified rat anti-mouse CD16/CD32 (BD Pharmingen) in FACS buffer (1 x HBSS containing 1% BSA and 0.1% sodium azide), and stained for extracellular and intracellular proteins using the following antibodies from Biolegend (San Diego, CA): PE/Cy7- or PE/Cy5-CD8 (clone 53-6.7), FITC-CD62L (clone MEL-14), APC/Cy7-CD3 (clone 145-2c11), PE/Cy7-CD4 (clone GK1.5), FITC-CD49b (clone DX5), APC-IL-2 (clone JES6-5H4), and PE-IFN (clone XMG1.2) following previously described protocols [208]. Brefeldin A (Biolegend) was added during the last 6 h of restimulation incubation to block cytokine secretion allowing the identification of cytokine producing T cells. Violet staining from each day was compared to the day 1 control, and a decrease of fluorescence intensity is indicative of cell proliferation. Single 51 stain controls were included in all experiments to compensate for fluorescence interference between detectors. Cells were assessed on a FACS Canto II flow cytometer (BD Biosciences, San Jose, CA), and analyzed with FlowJo v8.8.6 (Tree Star, Ashland, OR) or Kaluza 1.1 (Beckman Coulter, Miami, FL) software. Linear regression was performed using GraphPad Prism version 4.0a (Graphpad Software, San Diego, CA). 6 For in vivo studies, splenocytes (1 x 10 /well) isolated from different treatment groups were restimulated ex vivo with the most immunogenic peptide from the gp120 library (0.2 g/well) overnight in 2% BCS RPMI medium in a 96-well U-bottom plate. VH for the peptide (0.1% DMSO) or NT were included as controls. A mouse IFN secretion assay (Miltenyi Biotec Inc., Auburn, CA) was used to quantify IFN secretion by splenocytes 6 after peptide restimulation, according to the manufacturer’s instructions. Briefly, 1 x 10 splenocytes were centrifuged at 300 x g for 10 min at 4C and incubated with mouse IFN catch reagent in cold MACS buffer (PBS containing 0.5% BSA and 2 mM EDTA, pH 7.2) for 5 min on ice. Splenocytes were then diluted with warm (37C) medium and incubated for 45 min at 37C under slow continuous rotation. Subsequently, cells were washed with cold buffer and incubated with mouse antibodies (Biolegend, San Diego, CA) specific for IFN (PE), CD8 (PE/Cy7), CD4 (PE/Cy5), and NK1.1 (APC) for 10 min on ice. Next, cells were stained with LIVE/DEAD Fixable Near-IR cell stain to assess cell viability, fixed with Cytofix, and resuspended in FACS buffer. For cell activation studies, peptide restimulated splenocytes were first stained with LIVE/DEAD Fixable Aqua cell stain, followed by FcR blocking and cell staining for extracellular proteins using the following 52 antibodies from Biolegend: PE-CD69, PE/Cy5-CD80, FITC-MHC II, PE/Cy7-CD8, APC-CD4, and Pacific Blue-CD19. Cells were assessed and analyzed as described above. XIV. T Cell Purification T cells were purified from splenocytes by negative selection using the Pan T Cell Isolation Kit according to the manufacturer’s instructions (Miltenyi Biotec, Auburn, CA). Purity of T cells was determined using immunofluorescence analysis with anti-CD3 antibody (clone 145-2c11, BD Pharmingen), and generally exceeded 98%. Purified T cells were subsequently used for ELISA analysis. XV. ELISA 5 For cytokine ELISA, in 48-well plates, splenocytes (8 x 10 cells/well) or human 6 PBMC (2 x 10 cells/well) were treated with VH, THC or CBD for 30 min at 37°C, then activated with 10x or 1x P/I for 18-24 hr at 37°C. In some studies, cells were activated with anti-CD3 plus anti-CD28 antibodies (BD Pharmingen). For optimal stimulation, plates were coated with 0.1 g/well purified NA/LE hamster anti-mouse CD3e antibody (clone 145-2C11) overnight at 4C, washed twice with RPMI, and overlaid with splenocytes in the presence of 0.8 g/well purified NA/LE hamster anti-mouse CD28 antibody (clone 37.51) for 2 days at 37C. For suboptimal stimulation, splenocytes were activated with 0.8 g/well of each of anti-CD3 and anti-CD28. Quantification of IL-2 and IFN was determined by ELISA as previously described [159]. Briefly, recombinant purified mouse IL-2 or IFN (BD Pharmingen) or human IL-2 (BD Pharmingen) served as 53 standards from which the amount of each cytokine in the samples could be determined. Ninety-six well ELISA plates (Immulon 4 HBX strips, Thermo Scientific, Milford, MA) were coated with capture antibodies, purified anti-mouse IL-2 or IFN or anti-human IL-2 antibody. Detection antibodies were biotinylated anti-mouse IL-2 or IFN or anti-human IL-2 (BD Pharmingen). Color development was performed using streptavidin peroxidase followed by tetramethylbenzidine. Reactions were stopped with 6N H2SO4, after which samples were read at 450 nm using the BioTek Synergy HT (BioTek, Winooski, VT). For antibody ELISA, serum anti-gp120 antibody response from immunized mice was quantified by ELISA. ELISA plates were coated with 50 l/well of 2 g/ml purified recombinant IIIB gp120 (NIH AIDS Reagent Program) in 0.1 M NaHCO3 buffer (pH 9.6) at 4C overnight. Plates were washed with PBS containing 0.05% tween-20 and blocked with 3% BSA-PBS for 1.5-2 h at 37C. Serum samples were added into the wells and incubated for 1-1.5 h at 37C. Plates were washed again and incubated with a 1/1000 dilution of a peroxidase-conjugated goat anti-mouse IgG secondary antibody (Sigma) for 1-1.5 h. Plates were washed and developed with 1 mg/ml ABTS substrate (Roche Applied Science, Branford, CT). Samples were read at 450 nm every minute for 1 hr on kinetic mode using the BioTek Synergy HT plate reader. XVI. Calcium Determinations 7 Splenocytes (2 x 10 cells) were incubated with 2 µM Fluo-3 (Invitrogen) and 5 µM Fura Red (Invitrogen) for 30-60 min at RT in the dark. Cells were analyzed on a FACSCalibur (BD Biosciences) using the kinetic acquisition setting over 5-min intervals. 54 Baseline analysis was conducted for 1 min, after which cells received various treatments (VH or THC or CBD at 1 min, cellular activation at 2 min). Kinetic analyses were performed with FlowJo software. XVII. Low Density Array Mouse Immune Panel RNA was isolated from splenocyte samples using the TRI Reagent method as described above. cDNA was synthesized using the High Capacity cDNA Reverse Transcription Kit with 500 ng of RNA in a total volume of 40 l following the manufacturer’s instructions. TaqMan low density array mouse immune panel (Applied Biosystems) was utilized according to manufacturer’s instructions. In brief, 50 ng of cDNA in a total volume of 100 l was loaded into each row on the plate, followed by centrifugation at 1100 rpm at 1 min twice using a Sorvall Legend T centrifuge (Thermo Fisher, Waltham, MA). Analysis was performed using the Applied Biosystems 7900HT Real-Time PCR system. Thresholds for Ct values were determined by setting baseline fluorescence at 2-5 cycles for the internal control, 18S rRNA gene, and 2-13 cycles for target genes. Fold changes were calculated using the Ct method [226] when comparing CO-pVRCgp120 group to the control CO-pVRC2000, and comparing THC-pVRCgp120 group to the control CO-pVRCgp120. Genes that were differentially regulated by at least 1.5-fold were selected. XVIII. Verification of Genotype Real-time PCR was performed using genomic DNA samples extracted from WT and -/- -/- CB1 CB2 mouse-tails to verify the genotype. Genomic DNA from WT and KO 55 mouse-tails was extracted using the DirectPCR Lysis Reagent according to the manufacturer’s instructions (Viagen Biotech, Los Angeles, CA). One l of isolated genomic DNA was assayed in a total volume of 20 l, containing Taqman Universal Master mix (Applied Biosystems, Foster City, CA) and primers, in a real-time PCR reaction using standard amplification procedures (50C for 2 min, 95C for 10 min, 40 cycles of 95C for 15 sec, and 60C for 1 min) with a 7900 HT Thermocycler (Applied Biosystems). The presence of CB1 and CB2 was determined using commercial primers for cnr1 (Mm00432621-s1) and custom primers for cnr2 (forward 5 ′ CCTGATAGGCTGGAAGAAGTATCTAC-3 , ′ reverse 5 ′ ACATCAGCCTCTGTTTCTGTAACC-3 ′ ) [232] from Applied Biosystems. The average of Ct value for CB1 and CB2 in the WT mice was 24.70  1.22 and 26.84  2.14, -/- -/- respectively. All Ct values for both genes in the CB1 CB2 mice were below the level of detectable product and were reported as undetermined. XIX. Statistical Analysis Statistical analyses were performed using GraphPad Prism version 4.0a (GraphPad Software, La Jolla, CA). The mean  SE is displayed in all bar graphs. Differences between means of each treatment group were determined using parametric ANOVA. When significant differences were detected, treatment groups were compared to the appropriate control using Dunnett’s post-tests. Following a two-way ANOVA, all groups were compared using Bonferroni’s post-tests. For cytokine PCR or low density array data, Grubb’s outlier test was performed for each treatment group using Ct (Cttarget gene 56 - Ct18S). Fold-change values were transformed using natural log (fold-change + 1) for statistical analysis using ANOVA or unpaired t test. P value of < 0.05 was deemed statistically significant. 57 EXPERIMENTAL RESULTS I. In Vitro Cannabinoid-Mediated Effects on T Cell Response to HIVgp120 A. Expression of gp120 mRNA and protein in transduced DC2.4 and EL4 clones DC2.4 and EL4 were transduced with pLexPuro-gp120 and pLexNeo-gp120, respectively (Fig. 5). Importantly, because all DC2.4gp120 cells endogenously expressed cellular proteins are sampled and expressed within the context of MHC, including gp120, any peptide recognized as “nonself” has the potential of eliciting an immune response. To ensure that the immune response being measured during the effector phase of the CTL response was against gp120 but not an antibiotic resistance gene, each cell line (i.e., DC2.4gp120 and EL4gp120) was engineered to express a different antibiotic resistance gene. All transduced DC2.4gp120 r clones (C1, C2, C3, and C4) expressed puro transcript compared with parental DC2.4, as shown by lower Ct values (Fig. 6A). However, only C1 and C4 expressed gp120 transcript but not C2 and C3 (Fig. 6A). The presence of gp120 protein was detected in C1, but not in parental DC2.4 or C4 (Fig. 6B). Two tested EL4gp120 clones, C135 and C149, showed r expression of both gp120 and neo transcripts, compared with parental EL4 (Fig. 6C), and both clones expressed high levels of gp120 protein (Fig. 6D). Subsequent experiments were conducted with DC2.4gp120 C1 for elicitation and EL4gp120 C135 as target cells in the effector phase. Furthermore, the presence of gp120 transcript and protein in DC2.4gp120 C1 58 and EL4gp120 C135 was verified after cells had been passaged several times in culture, confirming stable insertion of the cassette into the genome of clones. The clones will be referred to as DC2.4gp120 and EL4gp120, respectively, in the following sections. B. Elicitation of gp120-specific CD8+ T cell response Following co-culture stimulation with DC2.4gp120, proliferation of CD8 + T cells in + splenocytes was observed, and approximately 80% of total viable CD8 T cells had divided at least once by day 5 (Fig. 7). IFN ELISPOT was performed as a proof-of-principle experiment to demonstrate a gp120-specific CTL response. DC2.4gp120 elicited a greater IFN response against EL4gp120 than against parental EL4 target cells (Fig. 8A) used to assess the contribution to the IFN response from putative antigens derived from BCS, EL4 tumor antigens, or antibiotic resistance peptides. The IFN response was low when there was no antigenic restimulation (CTL only group in Fig. 8A), demonstrating the requirement of target cell restimulation for measuring effector activity. The same trend of gp120-specific CTL response was confirmed by intracellular IFN staining (Fig. 8B). Furthermore, almost + all viable IFN CD8 CD62L low + + T cells had proliferated (Fig. 8C). Finally, the percent of + CD8 T cell (differentiated CD8 T cell) population increased after elicitation with + DC2.4gp120 for 5 days, compared with naïve CD8 T cells from the splenocytes (Fig. 9A), + + low and most IFN CD8 cells were also the CD62L 59 population (Fig. 9B). r Figure 5. Plasmid cassette constructs for expression of gp120 and puro in DC2.4 (top) r or gp120 and neo in EL4 cells (bottom). 60 Figure 6. Expression of gp120 transcript and protein in transduced DC2.4gp120 or EL4gp120 clones. (A, C) Total RNA was extracted from DC2.4gp120 clones (C1, C2, C3, C4), parental DC2.4 cells, EL4gp120 clones (C135, C149), or parental EL4 cells. r r Puro , Neo and gp120 transcript levels were analyzed by Real Time PCR. Data are presented as Ct values. Results shown are from gp120 3’ end Taqman primers, and 5’ end primers gave similar values. (B, D) Cytosolic proteins were isolated from DC2.4gp120 C1 and C4, EL4gp120 C135 and C149. 20 g of each sample was assessed by Western blotting for gp120 expression. Protein extract from parental DC2.4 or EL4 was included as a negative control, and purified gp120 protein (500 ng) was included as a positive control. (A-D) Data are representative of at least three separate experiments. 61 Figure 6 (cont’d) 62 + Figure 7. Proliferation of gp120-specific CD8 T cells was elicited in the in vitro model. 6 CellTrace Violet-labeled splenocytes (1 x 10 cells/well) from C57Bl/6 mice were incubated 5 with irradiated DC2.4gp120 cells (1 x 10 cells/well). On days 1-5 (D1 – D5) post initiation of the co-culture, cells were stained for viability and CD8 and assessed by flow cytometry for + + proliferation of CD8 T cells. Cells were gated on singlet, viable, lymphocyte, and CD8 + populations. The percent of proliferated CD8 T cell population on D5 is indicated in the plot. 63 + Figure 8. gp120-specific IFN production by CD8 T cells was elicited in the in vitro 6 model. (A) Splenocytes (1 x 10 cells/well) were co-cultured with irradiated DC2.4gp120 5 cells (1 x 10 cells/well) for 5 days. Elicited cells were assayed for CTL activity against EL4gp120 or parental EL4 target cells. IFN secretion was measured by ELISPOT. Each well 6 4 received 1 x 10 effectors and 5 x 10 target cells. No target cells were added to the CTL only group. (B-C) Labeled splenocytes were co-cultured with irradiated DC2.4gp120 cells and restimulated with EL4gp120 or parental EL4 target cells at a ratio of 20:1 as in A. Brefeldin A was added during the last 6 h of incubation. Cells were stained for viability, surface CD8, and + intracellular IFN. Cells were gated on singlet, viable, lymphocyte, and CD8 populations. + Cells were analyzed for IFN production and CD8 T cell proliferation in the plots. 64 Figure 8 (cont’d) 65 + Figure 9. Differentiation of gp120-specific CD8 T cells was elicited in the in vitro model. 6 (A) Splenocytes (1 x 10 cells/well) from C57Bl/6 mice were incubated with irradiated 5 DC2.4gp120 cells (1 x 10 cells/well). DC2.4gp120-elicited (gp120, day 5) and naive (NA) low splenocytes were stained for viability, CD8, and CD62L. The percent of CD62L cells + within CD8 T cells is presented in each plot. (B) Elicited cells were restimulated with + EL4gp120 target cells and analyzed for IFN and CD62L expression by CD8 T cells in one plot. (A-B) Three replicates were concatenated for each treatment in the flow cytometric analysis. 66 + C. Differential effects of cannabinoids on CD8 T cell IFN production and proliferation When THC was added at the initiation of the co-culture, IFN release by CTL was suppressed with increasing concentrations of THC (Fig. 10A), suggesting modulation by THC during the elicitation of CTL effectors. Interestingly, enhancement of the CTL IFN response by THC was also observed in some experiments (Fig. 10B). The overall THC effect was modest, but the trend towards suppression or enhancement was clear. An inverse correlation + + between THC effect and the level of IFN response, represented by percent of IFN CD8 T cells in the VH group, was drawn based on the data from eight separate experiments (Fig. 2 10C; p = 0.0272, R = 0.5841). In light of the differential modulation on the CTL IFN response by THC (Fig. 10C) and + the correlation between the IFN response and CD8 T cell proliferation by day 5 (Fig. 8C), + we sought to investigate the effect of THC on CD8 T cell proliferation during the elicitation + phase. These experiments were performed on day 5 because the majority of CD8 T cells exhibited a fully proliferative phenotype by day 5 following elicitation (Fig. 7). Again, an inverse correlation was observed between the effect of THC and the magnitude of stimulation: + THC decreased the percent of proliferating CD8 T cells under conditions in which the IFN + response was suppressed (Fig. 11A), but increased the percent of proliferating CD8 T cells under conditions in which the IFN response was enhanced (Fig. 11B). 2 relationship can be seen in Fig. 11C (p = 0.0099, R = 0.6968). 67 The inverse Figure 10. Differential effects of THC on IFN production. (A-B) Splenocytes (1 x 10 6 5 cells/well) were co-cultured with irradiated DC2.4gp120 cells (1 x 10 cells/well) in the presence of THC (1, 5, 10, 15 M) or VH (0.1% ethanol) or no treatment (NT). On day 5, 6 elicited cells (1 x 10 cells/well) were restimulated with EL4gp120 or parental EL4 target cells 4 (5 x 10 cells/well). Brefeldin A was added during the last 6 h of incubation. Cells were stained for viability, surface CD8, and intracellular IFN. Cells were gated on singlet, viable, + + + lymphocyte, and CD8 populations. The percent of IFN cells within the CD8 T cell population were analyzed, and data from the EL4gp120 target group are presented here. One example of THC-mediated suppression is presented in panel A, and one example of THCmediated enhancement is presented in panel B. (C) The inverse correlation between the magnitude of IFN response in VH and the THC effect on the response was evaluated using data collected from eight independent experiments. 68 Figure 10 (cont’d) 69 Figure 10 (cont’d) 70 + Figure 11. Differential effects of THC on CD8 T cell proliferation. (A-B) CellTrace 6 Violet-labeled splenocytes (1 x 10 cells/well) were co-cultured with irradiated DC2.4gp120 5 cells (1 x 10 cells/well) in the presence of THC (1, 5, 10, 15 M) or VH (0.1% ethanol) or no treatment (NT). Cells were stained for viability and surface CD8. Cells were gated on + + singlet, viable, lymphocyte, and CD8 populations. The percent of the proliferated CD8 T cell population is presented. One example of THC-mediated suppression is presented in panel A, and one example of THC-mediated enhancement is presented in panel B. (C) The inverse correlation between the magnitude of proliferation in VH and the THC effect was evaluated using data collected from eight independent experiments. 71 Figure 11 (cont’d) 72 Figure 11 (cont’d) 73 The role of CB1/CB2 in the THC-modulated differential effects was also investigated -/- -/- using CB1 CB2 (KO) mice. + Both C57Bl/6 wild-type (WT) and KO CD8 exhibited a gp120-specific IFN response, and KO CD8 + T cells T cells showed an enhanced + response when compared with WT CD8 T cells (Fig. 12A). In addition, in three separate experiments, THC produced a concentration-dependent enhancement on gp120-specific IFN + production in WT CD8 T cells, with the effect at 10 and 15 M being significantly different from the VH control (Fig. 12B). On the other hand, THC produced a trend toward + enhancement on gp120-specific IFN response in KO CD8 T cells, but to a much lesser extent when compared with the response in WT CD8 + T cells and the effect was not significant (Fig. 12B), suggesting a CB1/CB2-dependent mechanism. 74 Figure 12. THC-mediated effects on IFN response in a CB1/CB2-dependent manner. -/-/6 (A) Splenocytes (1 x 10 cells/well) from C57Bl/6 wild-type (WT) and CB1 CB2 (KO) 5 mice were co-cultured with irradiated DC2.4gp120 cells (1 x 10 cells/well), respectively. On 6 day 5, elicited cells (1 x 10 cells/well) were restimulated with EL4gp120 or parental EL4 4 target cells (5 x 10 cells/well). Brefeldin A was added during the last 6 h of incubation. Cells were stained for viability, surface CD8, and intracellular IFN. Cells were gated on + + singlet, viable, lymphocyte, and CD8 populations. The percent of IFN cells within the + CD8 T cell population are presented. (B) Splenocytes from WT and KO mice were cocultured with DC2.4gp120 cells as described in A, in the presence of THC (1, 5, 10, 15 M) or VH (0.1% ethanol). The correlation between the magnitude of IFN response in VH and the effect of THC at various concentrations on the response was evaluated using data collected from three independent experiments. * p < 0.05, ** p < 0.01 as compared to respective VH. 75 Figure 12 (cont’d) 76 The differential effects of THC on the IFN response were reminiscent of previous reports in the literature in which both positive and negative modulatory effects of cannabinoids have been observed [144, 159-161, 170, 172, 174, 175, 233]. However, the mechanisms for such differential effects of cannabinoids have produced significant confusion in the literature and are poorly understood. Thus, we sought to determine if the correlation between the magnitude of stimulation and the cannabinoid-mediated effect was broadly observed using two plant-derived cannabinoids (THC and CBD) and readily titratable T cell activators. THC and CBD were compared since they possess different affinities for CB1 and CB2 [101, 102]. We anticipated that the effects of THC and CBD would be similar since we had previously demonstrated that cannabinoid-mediated modulation of T cell function occurred independent of cannabinoid receptors [159, 162, 208]. In response to “optimal” stimulation with either 10x P/I (40 nM/0.5 µM) or immobilized anti-CD3 plus soluble antiCD28 antibodies (iCD3/CD28), IFN was robustly induced, which was suppressed by THC and CBD, similar to previous findings [144, 159, 162] (Fig. 13A, B and C). In contrast, IFN production following “suboptimal” stimulation using 1x P/I (4 nM/0.05 µM) was low, and was enhanced by CBD, but not THC (Fig. 13A, and B). Interestingly, in response to soluble anti-CD3 plus soluble anti-CD28 antibodies (sCD3/CD28), which was previously used as a suboptimal activation for IL-2 [174, 175], IFN production was robustly induced and then suppressed by both THC and CBD (Fig. 13D). 77 Figure 13. Effects of cannabinoids on IFN production in response to different stimuli. 5 B6C3F1 splenocytes (8 x 10 cells/well) were treated with THC (15 M; A) or CBD (5 M; B) for 30 min, and then activated with P/I (40 nM/0.5 µM, 10x P/I or 4 nM/0.05 µM, 1x P/I) 5 for 18-24 h. (C-D) B6C3F1 splenocytes (8 x 10 cells/well) were treated with THC or CBD at various concentrations for 30 min, and then activated with immobilized anti-CD3 plus soluble anti-CD28 (iCD3/CD28; C) or soluble anti-CD3 plus soluble anti-CD28 (sCD3/CD28; D) for 2 days. (A-D) IFN in the supernatant was quantified by ELISA. Data are presented as the mean units/ml  S. E. of triplicate cultures. * p < 0.05 as compared to respective VH. Data are representative of at least two separate experiments. 78 Figure 13 (cont’d) 79 D. Differential effects of cannabinoids on IL-2 response With the demonstration that THC differentially modulated CD8+ T cell proliferation in response to DC2.4gp120 stimulation, IL-2 production in the presence of P/I or anti-CD3/CD28 and the effect of cannabinoids were evaluated. Optimal stimulation, 10x P/I or iCD3/CD28, induced robust IL-2 production, which was significantly suppressed by THC and CBD, similar to previous reports [144, 159, 162, 175] (Fig. 14A and B). In response to 1x P/I, CBD, but not THC, enhanced IL-2 production (Fig. 14A), consistent with the differential cannabinoid effects on the IFN response. Again, using sCD3/CD28 as a suboptimal stimulation for IL-2, as previously performed [174, 175], IL-2 production was low, and was enhanced by both THC and CBD (Fig. 14C). Additional experiments were conducted in order to decipher the mechanisms by which cannabinoids differentially affected cytokine production with a focus on 1x P/I and CBD. Since P/I likely activated all cells in the splenocyte population, a comparison between splenocytes and purified T cells in response to 1x P/I was performed and demonstrated that the CBD-induced enhancement of the IL-2 response also occurred in purified T cells (Fig. 15). Moreover, FACS analysis of the cytokine-producing cells in splenocytes in response to + 1x P/I plus CBD demonstrated that CD4+ T cells produced most of the IL-2, whereas CD4 + and CD8 T cells and NK cells all contributed to IFN production (Fig. 16). Finally, in order to determine whether the differential effects of cannabinoids also occur in human PBMC, human IL-2 production was quantified. CBD suppressed IL-2 production when using 10x P/I, and enhanced IL-2 production when using 1x P/I (Fig. 17). 80 Figure 14. Effects of cannabinoids on IL-2 production in response to different stimuli. 5 (A) B6C3F1 splenocytes (8 x 10 cells/well) were treated with THC (15 M) or CBD (5 M) for 30 min, and then activated with P/I (40 nM/0.5 µM, 10x P/I or 4 nM/0.05 µM, 1x P/I) for 5 18-24 h. (B-C) B6C3F1 splenocytes (8 x 10 cells/well) were treated with THC or CBD at various concentrations for 30 min, and then activated with immobilized anti-CD3 plus soluble anti-CD28 (iCD3/CD28; B) or soluble anti-CD3 plus soluble anti-CD28 (sCD3/CD28; C) for 2 days. (A-C) IL-2 in the supernatant was quantified by ELISA. Data are presented as the mean units/ml  S. E. of triplicate cultures. * p < 0.05 as compared to respective VH. Data are representative of at least two separate experiments. 81 Figure 14 (cont’d) 82 Figure 15. Enhancement of the IL-2 response by CBD in purified T cells. B6C3F1 5 splenocytes or T cells purified from the splenocytes (SPLC, 8 x 10 cells/well) were treated with 0.2 – 5 M CBD for 30 min, then activated with 1x P/I for 48 h. IL-2 in the supernatant was quantified by ELISA. Data are presented as the mean units/ml  S. E. of triplicate cultures. * p < 0.05 as compared to respective VH. Data are representative of at least two separate experiments. 83 5 Figure 16. Enhancement of cytokine response by CBD in different cellular populations. B6C3F1 splenocytes (8 x 10 cells/well) were treated with 0.5 – 10 M CBD for 30 min, followed by activation with 1x P/I for 18-24 h. Cells were also treated with Brefeldin A 4 h prior to harvest. Cells were stained for viability and extracellular markers, CD4, CD8, CD3 and CD49b, as well as intracellular IL-2 and IFN. 84 Figure 17. Differential effects of CBD on IL-2 response in human PBMC. PBMC were 6 enriched from human leukocyte packs by density gradient centrifugation. PBMC (2 x 10 cells/well) were treated with 0.1 – 10 M CBD for 30 min, then activated with 1x or 10x P/I for 18-24 h. IL-2 in the supernatant was quantified by ELISA. Data are presented as the mean units/ml  S. E. of triplicate cultures. * p < 0.05 as compared to respective VH. Data are representative of at least three separate experiments. 85 E. Differential effects of cannabinoids on cytokine production were correlated with effects on NFAT nuclear translocation The differential effects of cannabinoids on cytokine production was also observed at the steady state mRNA level as CBD suppressed or enhanced IFN and IL-2 mRNA expression in response to 10x or 1x P/I, respectively, at several time points following stimulation (Fig. 18A and 19A). In addition, CBD-induced IFN and IL-2 mRNA expression was increased in a concentration-dependent manner in response to 1x P/I (Fig. 18B and 19B). The concentration-dependent suppression of IFN and IL-2 mRNA expression by CBD on 10x P/I-stimulated splenocytes at 24 hr has been reported previously [159]. These results suggested that CBD-induced modulation of cytokine production might occur at the level of transcription. NFAT family members NFAT1 (NFATc2) and NFAT2 (NFATc1) are the two predominant NFAT proteins expressed by T cells [234], and of the two, NFAT2 is the primary form induced under lower calcium or anergic conditions [235]. Since CBD enhanced cytokine production in response to suboptimal cellular activation, and NFAT plays an important role in IFN and IL-2 transcription [236-239], the effect of CBD on NFAT2 nuclear translocation was examined. 10x P/I induced nuclear translocation of NFAT2, which was modestly suppressed by CBD, whereas 1x P/I-stimulated NFAT2 translocation was enhanced by CBD (Fig. 20). 86 Figure 18. Cannabinoids differentially regulated IFN steady state mRNA expression. 6 (A-B) B6C3F1 splenocytes (5 x 10 cells) were treated with CBD (0.5 – 5 µM) or VH (0.1% ethanol) for 30 min, then activated with P/I (40 nM/0.5 µM, 10x P/I or 4 nM/0.05 µM, 1x P/I) for various times. Total RNA was isolated and reverse transcribed, and Real Time PCR was performed for IFN. Data are expressed as the mean fold change  S. E. of triplicate cultures. * p < 0.05 as compared to respective VH. 87 Figure 19. Cannabinoids differentially regulated IL-2 steady state mRNA expression. 6 (A-B) B6C3F1 splenocytes (5 x 10 cells) were treated with CBD (0.5 – 5 µM) or VH (0.1% ethanol) for 30 min, then activated with P/I (40 nM/0.5 µM, 10x P/I or 4 nM/0.05 µM, 1x P/I) for various times. Total RNA was isolated and reverse transcribed, and Real Time PCR was performed for IL-2. Data are expressed as the mean fold change  S. E. of triplicate cultures. * p < 0.05 as compared to respective VH. 88 Figure 20. Cannabinoids differentially modulated nuclear translocation of NFAT2. 6 B6C3F1 splenocytes (5 x 10 cells) were treated with 0.5 – 5 M CBD or VH (0.1% ethanol) for 30 min, followed by activation with 1x or 10x P/I for 30 min. Nuclear protein was isolated and loaded into an 8% SDS-PAGE gel. NFAT2 is detected as multiple bands in the 100-130 kDa range. 89 F. Cannabinoids enhanced intracellular calcium regardless of the magnitude of cellular activation Intracellular calcium is critical for activation of NFAT [236, 238], therefore we hypothesized that the effect of cannabinoids on intracellular calcium might correlate with the differential regulation of NFAT translocation and cytokine production. THC has been shown to induce intracellular calcium in resting lymphocytes [143, 163] and as seen in Fig. 21A, CBD also induced intracellular calcium in resting splenocytes in a concentration-dependent manner. 1x P/I modestly induced intracellular calcium, and both THC and CBD further elevated the level of intracellular calcium (Fig. 21B). 10x P/I robustly induced intracellular calcium, but unlike the effects of CBD or THC on NFAT nuclear translocation and cytokine production, THC and CBD further enhanced the intracellular calcium above the level induced by P/I alone (Fig. 21B). 90 7 Figure 21. Cannabinoids increased intracellular calcium regardless of activation level. (A-B) B6C3F1 splenocytes (2 x 10 cells) were incubated with Fura Red and Fluo-3 for 30 – 60 min in the dark. Cells were then analyzed by flow cytometry, during which time they received CBD (1 – 10 µM) in panel A or THC (10 µM)/CBD (5 µM)/VH (0.1% ethanol) in panel B for 1 min followed by 1x or 10x P/I or VH (0.1% DMSO) for 3 additional min. Results are representative of at least two separate experiments. 91 Figure 21 (cont’d) 92 II. In Vivo Cannabinoid-Mediated Effects on Immune Responses to HIVgp120 To investigate the effect of THC and the role of CB1/CB2 on gp120 antigen-specific immune responses in vivo, studies were designed such that the gp120-expressing plasmid was -/- -/- injected intramuscularly into WT and CB1 CB2 mice to induce the immune responses and THC was orally given to the mice post each plasmid injection. A. pVRCgp120 induced gp120-specific cytokine production by ELISPOT assay To determine if pVRCgp120 plasmid DNA can elicit gp120-specifc T cell responses in vivo, peptides derived from HIV-1 IIIB envelope protein were split into 53 peptide pools and screened upon peptide restimulation. A robust IFN response by ELISPOT assay was detected when splenocytes from pVRCgp120-injected mice were restimulated with peptide pool 21 compared with the other 52 peptide pools (Fig. 22A). When peptide pool 21 was used to restimulate the splenocytes from pVRC2000-injected mice, the IFN response was minimal (Fig. 22B). This result suggests that pool 21 contains potentially-immunodominant peptide(s) to induce a gp120-specific T cell response. To further identify the immunogenic peptide(s) within pool 21, peptides 81-84 were tested individually. IFN production was significantly higher when peptide 81 (IIGDIRQAHCNISRA) was used for restimulation in the pVRCgp120 group as compared with the other three peptides within pool 21 (p < 0.05), suggesting that peptide 81 is the most immunogenic gp120 peptide (Fig. 23A). Also, an IFN response was not observed in the pVRC2000 control group. A similar profile was observed with IL-2 production in response to peptide 81 restimulation, although less IL-2 was produced compared to IFN (Fig. 23B). 93 6 Figure 22. gp120-specific cytokine production by splenocytes. Splenocytes (1 x 10 cells/well) from pVRCgp120- (A) or pVRC2000- (B) injected C57Bl/6 mice were restimulated overnight with 53 peptide pools (0.2 g/peptide/well) respectively, which are derived from the HIV envelope protein. IFN secretion was measured by ELISPOT. No treatment (NT) or VH (0.01% 6 DMSO) were included as controls. Data are presented as the mean spots/10 cells of 5 samples in each treatment group. 94 Figure 22 (cont’d) 95 Figure 23. Identification of immunodominant gp120 peptide for restimulation. Splenocytes 6 (1 x 10 cells/well) from pVRCgp120- or pVRC2000-injected C57Bl/6 mice were restimulated with the immunodominant peptide 81 (0.2 g/well) overnight. IFN (A) or IL-2 (B) secretion was measured by ELISPOT. No treatment (NT) or VH (0.01% DMSO) were included as 6 controls. Data are presented as the mean spots/10 cells of 5 samples in each treatment group. Data were analyzed using two-way ANOVA with Bonferroni post-tests: *** p < 0.001 as compared with respective VH in each treatment group; ### p < 0.001 as compared with pVRC2000 control for each peptide restimulation. 96 B. pVRCgp120-induced T cell activation With the demonstration that gp120-specifc cytokine production was induced in response to pVRCgp120 antigens in vivo, activation of splenic T cell populations was assessed by measuring + the expression level of the activation marker, CD69. MFI of CD69 on CD8 (Fig. 24A) and + CD4 (Fig. 24B) T cells in the splenocyte populations was significantly increased when peptide 81 was used for restimulation in the pVRCgp120 group, when compared with the other three peptides within pool 21 (p < 0.05), once again confirming that peptide 81 is the immunodominant peptide. In addition, with peptide 81 restimulation, CD69 expression was higher in the pVRCgp120 group compared with the pVRC2000 control, which was statistically different for CD4+ T cells (Fig. 24B, p < 0.05), hence demonstrating a gp120-specific T cell activation. C. pVRCgp120-induced activation of non-T cells In addition to T cells, activation of non-T cell populations using the same in vivo model system was also examined. The magnitude of activation marker expression, including CD69 - - (Fig. 25A), CD80 (Fig. 25B), and MHC II (Fig. 25C), on the non-T cell (CD8 CD4 ) populations in the pVRCgp120 group was significantly increased in response to peptide 81 restimulation (p < 0.05), when compared with the other three peptides. Moreover, the expression levels of all three activation markers in the pVRCgp120 group were significantly higher than in the pVRC2000 control in the presence of peptide 81 restimulation, again demonstrating a gp120-specific response. Peptide 82 was also found to be immunogenic for some endpoints, but to a lesser extent than peptide 81, which is likely due to the sequence overlap between peptide 81 and 82. 97 D. pVRCgp120-induced B cell responses In light of the fact that non-T cells demonstrated a gp120-specific response, B cells were assayed for antibody production in response to pVRCgp120 antigen stimulation in vivo. + Expression levels of CD69 (Fig. 26A) and MHC II (Fig. 26B) on B cells (CD19 ) in the spleen were significantly increased in the pVRCgp120 group as compared with the pVRC2000 control (p < 0.05), in response to peptide 81 restimulation. Furthermore, a gp120-specific IgG response was detected using the serum samples from pVRCgp120-treated mice, which was significantly different from the pVRC2000 control and NA groups (Fig. 26C, p < 0.05), demonstrating the generation of a secondary antibody response. 98 Figure 24. Activation of T cells in response to pVRCgp120 stimulation. (A-B) Splenocytes 6 (1 x 10 cells/well) from pVRCgp120- or pVRC2000-injected C57Bl/6 mice were restimulated overnight with gp120-derived peptides 81, 82, 83, 84, or pool 21, containing all four peptides (0.2 g/peptide/well). Cells were stained for viability, surface CD8, CD4, and CD69, and + + assessed by flow cytometry. Cells were gated on singlet, viable, lymphocyte, and CD8 /CD4 populations. No treatment (NT) or VH (0.01% DMSO) were included as controls. Data are presented as MFI of CD69 from 5 samples in each treatment group. Data were analyzed using two-way ANOVA with Bonferroni post-tests: ** p < 0.01, *** p < 0.001 as compared with respective VH in each treatment group; # p < 0.05, ### p < 0.001 as compared with pVRC2000 control for each peptide restimulation. 99 Figure 25. Activation of non-T cells in response to gp120 antigen stimulation. (A-C) 6 Splenocytes (1 x 10 cells/well) from pVRCgp120- or pVRC2000-injected C57Bl/6 mice were restimulated overnight with gp120-derived peptides 81, 82, 83, 84, or pool 21, containing all four peptides (0.2 g/peptide/well). Cells were stained for viability, surface CD8, CD4, CD69, CD80, and MHC II, and assessed by flow cytometry. Cells were gated on singlet, viable, lymphocyte, and CD8 CD4 populations. No treatment (NT) or VH (0.01% DMSO) were included as controls. Data are presented as MFI of CD69 (A), CD80 (B), or MHC II (C) from 5 samples in each treatment group. Data were analyzed using two-way ANOVA with Bonferroni post-tests: ** p < 0.01, *** p < 0.001 as compared with respective VH in each treatment group; # p < 0.05, ### p < 0.001 as compared with pVRC2000 control for each peptide restimulation. 100 Figure 25 (cont’d) 101 6 Figure 26. B cell responses to pVRCgp120 stimulation. (A-B) Splenocytes (1 x 10 cells/well) from pVRCgp120- or pVRC2000-injected C57Bl/6 mice were restimulated overnight with gp120-derived immunodominant peptide 81 (0.2 g/well). Cells were stained for viability, surface CD19, CD69, and MHC II, and assessed by flow cytometry. Cells were + gated on singlet, viable, lymphocyte, and CD19 populations. Data are presented as MFI of CD69 (A) or MHC II (B) from 5 samples in each treatment group. (C) Serum from trunk blood was collected from pVRCgp120- or pVRC2000-injected C57Bl/6 mice. IgG production in the serum was quantified by ELISA. Data are presented as the mean ng/ml of 5 samples in each treatment group. (A-C) Data were analyzed using two-tailed t test or oneway ANOVA with Dunnett’s post-tests: ** p < 0.01, *** p < 0.001 as compared with the pVRC2000 control. 102 E. THC treatment enhanced pVRCgp120-induced IFN production by lymphocyte populations -/- -/- in the spleens of WT but not CB1 CB2 mice Using the in vivo model system, the effects of THC and the role of CB1/CB2 on gp120-/- -/- specific immune responses in WT and CB1 CB2 mice were investigated. In initial studies, we conducted THC treatment surrounding the DNA injection, similar to our studies examining effects of THC on influenza-induced immune responses [206]. However, given the likely differences in pharmacokinetics between intranasal injection of influenza virus and intramuscular injection of viral plasmid DNA, it was anticipated that initial gp120 immune response in the spleen following intramuscular injection would require some time. Thus, the studies were repeated using the regimen presented in Fig. 4. Overall, the experiment was repeated four times: twice in which THC dosing surrounded DNA injection and twice in which THC dosing was performed 2 days following DNA injection. Results presented are from the latter regimen (Fig. 4), but are representative of all four experiments. + + In CO-treated mice, pVRCgp120 injection induced a higher percentage of CD8 , CD4 , + and NK1.1 cells that produced IFN in response to peptide 81 restimulation when compared -/- -/- with pVRC2000 control in both WT and CB1 CB2 mice (Fig. 27A and 27B). Interestingly, + the percentage of IFN cells in all three lymphocyte populations was further enhanced by THC treatment in WT mice (Fig. 27A). In contrast, there was a trend toward THC-mediated -/- -/- suppression on IFN production in CB1 CB2 103 mice, demonstrating a CB1/CB2-independent mechanism of THC modulation of the IFN response (Fig. 27B). On the other hand, the -/- -/- magnitude of the IFN response in the absence of THC was higher in CB1 CB2 mice than in WT mice, suggesting endocannabinoid regulation of immune responses via CB1 and/or CB2 (Fig. 27B). -/- F. THC treatment enhanced pVRCgp120-induced activation of T cells in WT but not CB1 -/- CB2 mice Next, the effects of THC and the role of CB1/CB2 were studied on the magnitude of T cell activation. The same trend of THC effects was observed on T cell activation as on the IFN response in Fig. 27A and 27B. In the CO-treated mice, pVRCgp120 induced a higher percentage + + + + of CD69 CD8 and CD69 CD4 cells in response to peptide 81 restimulation in both WT and -/- -/- CB1 CB2 /- -/- - mice, which was further increased by THC treatment in WT but not in CB1 CB2 mice (Fig. 28A and 28B). 104 Figure 27. THC-mediated enhancement on gp120-specific IFN response in WT but not -/-/6 CB1 CB2 mice. (A-B) Splenocytes (1 x 10 cells/well) from pVRCgp120- or pVRC2000-/- -/- injected C57Bl/6 (WT, A) and CB1 CB2 (KO, B) mice were restimulated overnight with gp120-derived immunodominant peptide 81 (0.2 g/well). Cells were stained for viability, surface CD8, CD4, NK1.1, and intracellular IFN, and assessed by flow cytometry. Cells were + + + gated on singlet, viable, lymphocyte, and CD8 /CD4 /NK1.1 populations. The percentage of IFN cells within each lymphocyte population is presented in the dot plots. Displayed are concatenated samples (n=5) of each treatment group. 105 Figure 27 (cont’d) 106 -/- -/- Figure 28. THC-mediated enhancement on gp120-specific T cell activation in WT but not CB1 CB2 mice. (A-B) -/-/6 Splenocytes (1 x 10 cells/well) from pVRCgp120- or pVRC2000-injected C57Bl/6 (WT, A) and CB1 CB2 (KO, B) mice were restimulated overnight with gp120-derived immunodominant peptide 81 (0.2 g/well). Cells were stained for viability, surface + + CD8, CD4, and CD69, and assessed by flow cytometry. Cells were gated on singlet, viable, lymphocyte, and CD8 /CD4 + + + populations. The percentage of CD69 cells within CD8 or CD4 populations is presented in the dot plots. Displayed are concatenated samples (n=5) of each treatment group. 107 Figure 28 (cont’d) 108 -/- G. THC treatment enhanced pVRCgp120-induced B cell activation in WT but not CB1 -/- CB2 mice Expression levels of activation marker CD69 on B cells in the presence of THC treatment was also assessed. In response to peptide 81 restimulation, the MFI of CD69 expression on B -/- + -/- cells (CD19 ) was higher in pVRCgp120-treated WT and CB1 CB2 mice as compared with their corresponding control mice (Fig. 29A). Although the THC effect was modest, it -/- -/- clearly showed a trend toward enhancement on CD69 expression in WT but not CB1 CB2 mice (Fig. 29A), which was consistent with its effects on other endpoints reported above. In addition, upon pVRCgp120 stimulation, MHC II expression on B cells was significantly -/- -/- increased in both WT and CB1 CB2 mice as compared with the pVRC2000 control mice (Fig. 29B, p < 0.05). THC also significantly elevated the expression level of MHC II on B -/- -/- cells in pVRCgp120-treated WT mice but not in CB1 CB2 mice (Fig. 29B, p < 0.05) H. THC treatment enhanced pVRCgp120-induced activation of non-T/non-B cells in WT but -/- -/- not CB1 CB2 mice In addition to T cell and B cell populations, the splenic non-T/non-B cell population - - - (CD8 CD4 CD19 ) was also studied is this model system. Both CD69 (Fig. 30A) and MHC II (Fig. 30B) expression levels on the non-T/non-B cells was higher in pVRCgp120-treated -/- -/- WT and CB1 CB2 mice when compared with the pVRC2000 controls, with the increase -/- -/- being statistically significant for MHC II expression in CB1 CB2 109 mice (p < 0.05). The THC treatment group showed a trend toward enhancement in the MFI for CD69 and MHC II -/- -/- in WT mice but not in CB1 CB2 -/- -/- mice. It is also noteworthy that CB1 CB2 mice upon gp120 stimulation exhibited a higher MFI for CD69 and MHC II on non-T/non-B cells, when compared to the WT mice, which was significantly different for MHC II expression (p < 0.05). I. THC modulated mRNA expression of genes induced by pVRCgp120 To further investigate the effects of THC on the anti-gp120 immune responses at gene transcript levels, a 96-gene immune array was employed with spleen RNA from WT and -/- -/- CB1 CB2 mice on day 15 post pVRCgp120 injection. Differential gene expression was -/- -/- observed in treated WT and CB1 CB2 mice. Specifically, in CO-treated mice, mRNA -/- -/- levels of several genes were increased in CB1 CB2 mice in response to pVRCgp120 compared with the pVRC2000 control group, including cytokine Il2, chemokine receptor Ccr4 and Cxcr3, and surface marker Cd40lg (Table 2). THC treatment upregulated expression of surface marker Cd68 and downregulated expression of Ctla4 compared with the CO control group in WT mice. In contrast, THC treatment downregulated expression of cytokines Il17, Il12, and Il2, negative regulators of cytokines Socs1 and Socs2, and -/- -/- chemokine receptor Ccr2 compared with the CO control group in CB1 CB2 110 mice (Table 2). Figure 29. THC-mediated enhancement on gp120-specific B cell activation in WT but -/-/6 not CB1 CB2 mice. (A-B) Splenocytes (1 x 10 cells/well) from pVRCgp120- or -/- -/- pVRC2000-injected C57Bl/6 (WT) and CB1 CB2 (KO) mice were restimulated overnight with gp120-derived immunodominant peptide 81 (0.2 g/well). Cells were stained for viability, surface CD19, CD69, and MHC II, and assessed by flow cytometry. Cells were + gated on singlet, viable, lymphocyte, and CD19 populations. Data are presented as MFI of CD69 (A) and MHC II (B) from 5 samples in each treatment group. Data were analyzed using two-way ANOVA with Bonferroni post-tests: ## p < 0.01, ### p < 0.001 as compared with the respective CO-pVRCgp120 control for each genotype; a p < 0.05, aaa p < 0.001 as compared with the WT control. 111 Figure 30. THC-mediated enhancement on gp120-specific non-T/non-B cell activation in -/-/6 WT but not CB1 CB2 mice. (A-B) Splenocytes (1 x 10 cells/well) from pVRCgp120-/- -/- or pVRC2000-injected C57Bl/6 (WT) and CB1 CB2 (KO) mice were restimulated overnight with gp120-derived immunodominant peptide 81 (0.2 g/well). Cells were stained for viability, surface CD8, CD4, CD19, CD69, and MHC II, and assessed by flow cytometry. Cells were gated on singlet, viable, and CD8 CD4 CD19 populations. Data are presented as MFI of CD69 (A) and MHC II (B) from 5 samples in each treatment group. Data were analyzed using two-way ANOVA with Bonferroni post-tests: # p < 0.05, ## p < 0.01 as compared with the respective CO-pVRCgp120 control for each genotype; a p < 0.05, aa p < 0.01 as compared with the WT control. 112 Table 2. Differential gene expression in response to pVRCgp120 injection and THC -/-/treatment in WT and CB1 CB2 mice. Mice (n=3) were injected with pVRCgp120 or pVRC2000 prior to THC or CO treatment. Spleen RNA was isolated on day 15 and converted to cDNA, and gene expression levels were analyzed using a PCR-based low density immune array. Fold change values were calculated, and shown are genes differentially regulated by at least 1.5-fold when comparing CO-pVRCgp120 to the control CO-pVRC2000 group, and -/-/comparing THC-pVRCgp120 to the control CO-pVRCgp120 group in WT and CB1 CB2 mice, respectively, with a p value of at lease 0.05. Red color indicates upregulation, and green color indicates downregulation. CO-pVRCgp120 vs CO-pVRC2000 KO WT Ccr4 Cxcr3 Il2 Cd40lg THC-pVRCgp120 vs CO-pVRCgp120 WT KO Cd68 Ccr2 Ctla4 Il17 Il12 Il2 Socs1 Socs2 113 DISCUSSION The effect of cannabinoids and the role of CB1/CB2 on HIV viral antigen-specific immune responses were determined throughout this dissertation work, which was divided into two parts. The first part focused on characterizing the effect of cannabinoid on HIVgp120specific T cell responses using an in vitro model. The second part focused on investigating the effects of cannabinoid and the role of CB1/CB2 on gp120-specific immune responses using an in vivo model. I. In vitro cannabinoid-mediated effects on T cell responses to HIVgp120 As described earlier, HIV patients use marijuana to mitigate AIDS-associated adverse effects [94], but there are no suitable mouse models that address the effect of cannabinoids, the active components of marijuana, on initial anti-HIV immune responses. The novel in vitro mouse model established here mimics the T cell response against HIV-associated antigens in the early stages of infection when T cells are critical in eliminating the virus, and was used to investigate the modulation of the T cell response by cannabinoids. Although only one HIV protein, gp120, was studied in our model, broad T cell responses against multiple viral epitopes of this protein are elicited. In this model, different engineered cell lines were employed, which provided the ability to study the response specific to gp120 antigens shared by both DC2.4gp120 and EL4gp120 cells, but not to antigens from BCS, parental cells, or + antibiotic resistance markers. As demonstrated, DC2.4gp120 was capable of eliciting CD8 T 114 cell proliferation, differentiation, and therefore, gp120-specific IFN production from CTL, + indicating that APC DC2.4gp120 cells, CD8 T cells, and target EL4gp120 cells all play functional roles in this model system. The present studies demonstrated that THC differentially modulated IFN production from gp120-specific CTL and proliferation of CD8 + T cells in response to DC2.4gp120 elicitation. This is the first report of the differential effect of cannabinoids on an antigenspecific T cell response. The experimental factors that control the magnitude of cellular activation in this in vitro system are unknown. Across eight independent experiments, a general trend was observed such that the magnitude of the immune response, as measured by + IFN and CD8 T cell proliferation, appeared to be one important determinant of whether THC produced enhancement or suppression. Moreover, the suppression or enhancement + produced by THC on the IFN response was consistent with its effect on CD8 T cell + proliferation, indicating that THC modulated CD8 T cell effector function, in part, via + regulating CD8 T cell proliferation first. Due to the complexity of the factors involved in the gp120-induced response in this cognate model (i.e., magnitude of gp120 expression, gp120 processing and presentation associated with MHC, and initiation of T-cell receptor signaling), more direct activators of T cell effector function were employed to study the differential immune modulation by cannabinoids. The pharmacological activators, P/I and anti-CD3/CD28, facilitated a more readily controlled titration of the stimulus used to induce the immune response. In addition, CBD was also used in several studies in order to demonstrate that the differential effect of 115 cannabinoids was not specific to THC. THC exhibits affinity for CB1 and CB2, whereas CBD does not exhibit high affinity for either cannabinoid receptor [101, 102]. In fact, we anticipated that CBD and THC would produce similar effects since we have previously demonstrated that many effects of cannabinoids on T cells are mediated independently of either CB1 or CB2 [140, 159-162]. With one exception (cannabinoid effect in response to 1x P/I), the effects of THC and CBD were, in fact, similar. The effect of CBD on IFN and IL-2 production was, in part, dictated by the magnitude of stimulation delivered through either P/I or anti-CD3/CD28 treatment. Under conditions of relatively high stimulation (10x P/I, iCD3/CD28 or sCD3/CD28 only for IFN), CBD suppressed cytokine production, whereas under conditions of relatively low stimulation (1x P/I or sCD3/CD28 only for IL-2), CBD enhanced cytokine production. The effect of THC on IFN and IL-2 production followed the same pattern as CBD, with the exception that THC did not enhance cytokine production in response to 1x P/I. It was also unexpected that sCD3/CD28 induced robust IFN production since we previously showed that iCD3/CD28 was a stronger activation stimulus than sCD3/CD28 for IL-2 induction in splenocytes [174, 175], but THC and CBD still produced similar effects on sCD3/CD28-stimulated IFN. It is possible that suboptimal stimulation with sCD3/CD28 leads to T cell anergy or activation-induced cell death (AICD), which requires IFN for the production of caspases and also results in decreases in IL-2 production and proliferation to regulate T cell homeostasis [240-243]. These results are consistent with previous reports that cannabinoids either suppressed or enhanced immune function depending on the experimental conditions, such as cell type, immune stimuli, and cannabinoid concentrations [171-175, 233], which were described in the 116 Introduction chapter, but a clear mechanism has not been established. We hypothesize that the increase in intracellular calcium by cannabinoids prior to activation contributes to the downstream effects. That is, the combination of cannabinoids with suboptimal cellular activation provides an appropriate or optimal calcium signal to induce cytokine production, whereas the combination of cannabinoids with an optimal level of cellular activation might provide an anergic or cell death signal to suppress cytokine production, which has been suggested for immune suppression by CBD previously [144, 244]. Thus, it is unexpected that THC does not induce IL-2 or IFN in response to 1x P/I, especially since THC has been reported to increase intracellular calcium in lymphocytes [163, 245, 246]. In these studies, we demonstrated that CBD and THC increased calcium in splenocytes, regardless of the magnitude of activation. It is possible that since THC enhanced the calcium level to a lesser extent than CBD in the presence of 1x P/I, the induction on cytokine production was so modest that it is below the level of detection. It was notable that differential regulation of IL2 production by CBD was also observed in human PBMC, implying that the differential effect of cannabinoids occurs in humans. Further investigations are needed to determine whether similar trends in the cannabinoid-mediated effects also occur in immunocompromised HIV patients. As shown here and in previous studies, cannabinoids increased intracellular calcium in resting and activated splenocytes, suggesting that the differential regulation of NFAT nuclear translocation by cannabinoids must occur at a signal distal to increased intracellular calcium [163, 245, 246]. It is possible that cannabinoids in combination with optimal or suboptimal stimulation might differentially produce oscillatory calcium signals, and depending on the frequency of the signals NFAT activation as well as IL-2 expression were differentially 117 regulated, which have been demonstrated previously [247]. Alternatively, the “switch factor” for cannabinoid-induced modulation of NFAT2 nuclear translocation and subsequent cytokine production that must be distal to the increase in intracellular calcium could be cytoplasmic NFAT regulating factors, such as the calcium-sensing stromal interaction molecule (STIM) or calcium-release-activated calcium (CRAC) channel protein ORAI, or nuclear kinases that induce nuclear export of NFAT, such as dual-specificity tyrosine-phosphorylation regulated kinase 1A (DYRK1A) (reviewed in [248]). Another possibility is that the induction of different transcription factors distal to the increase in intracellular calcium dictates the effects on cytokine production. For example, whether the cells would undergo anergy or AICD may depend on the NFAT:AP-1 ratio, as a high AP-1 level is required for AICD [249, 250]. The putative differential effects of cannabinoids on AP-1 are presently unknown, but AP-1 activity has been shown to be suppressed by cannabinoid treatment under optimal stimulation conditions [140, 161, 227]. Overall, these in vitro studies demonstrated that cannabinoids suppressed or enhanced HIVgp120-specific T cell responses. Such differential effects by cannabinoids also occur in response to other stimuli, such as P/I or anti-CD3/CD28, and demonstrated that cannabinoids differentially regulated NFAT nuclear translocation and cytokine production. Despite these opposing effects, cannabinoids elevated intracellular calcium regardless of the presence or absence, or even the magnitude, of cellular activation. These data indicated that the intracellular calcium level resulting from the combination of cannabinoid treatment and cellular activation, at least in part, determined the overall T cell response. In particular, cannabinoids in combination with suboptimal stimulation led to an optimal calcium level, increased nuclear translocation of NFAT, and therefore enhanced the T cell response. On the 118 other hand, cannabinoids plus optimal stimulation led to excessive intracellular calcium, which reduced NFAT nuclear translocation and resulted in suppression of the T cell response. Together, these data provide one potential mechanism to explain the many dichotomous reports in the literature in which cannabinoids either induce or suppress the same immune response [140, 144, 159-161, 170-172, 174, 175, 233]. Our observations also identify the need for more in-depth investigations on evaluating the effect of cannabinoid use on immune competence of HIV patients. II. In vivo cannabinoid-mediated effects on immune responses to HIVgp120 In light of the demonstration that cannabinoids produced differential effects on T cell responses against HIVgp120 antigen in vitro, the effect of cannabinoids on immune responses to HIV viral antigens in vivo was in turn characterized. In the second part of this dissertation work, a novel HIV mouse model was first developed in which viral DNA injections were used -/- -/- to stimulate gp120-specific responses in WT and CB1 CB2 mice. This is a physiologically relevant model using intact animals to mimic the immune responses against multiple viral epitopes in the early stages of HIV infection. Using this model, a comprehensive immune response was induced, including both cell-mediated and humoral immunity, as evidenced by induction of gp120-specific cytokines, serum IgG production, and activation of various cellular populations. In fact, this is the first report in which the gp120-derived immunodominant peptide (IIGDIRQAHCNISRA) was identified in the C57Bl/6 mouse strain in response to the gp120 protein from the HIV-1 IIIB HXBc2 stain. Interestingly, C57Bl/6 T helper cells were found to respond predominantly to the peptide IIGDIRQAHCNISRE 119 derived from the envelope protein in HIV-1 isolate 1007 [251], which differs from ours by only one amino acid. It is also suggested that this immunodominant peptide is likely located in the variable (V) 3 region of gp120 [252], which is responsible for binding to the chemokine coreceptor CCR5 to allow virus entry into the target cells [253]. -/- -/- In the absence of THC treatment, CB1 CB2 mice produced a greater level of IFN upon gp120 antigen stimulation compared with WT mice (Fig. 27A and 27B). Such hyper-/- -/- reactivity in CB1 CB2 mice has also been reported previously in other studies. For -/- -/- example, in a model of contact hypersensitivity, CB1 CB2 mice showed a pronounced induction of type IV hypersensitivity with exacerbated ear swelling in response to contact sensitizer, 2,4-dinitrofluorobenzene [213]. -/- -/- influenza, CB1 CB2 In addition, in a host resistance model to mice exhibited an exaggerated antiviral immune response to influenza challenge, as shown by less viral load in the lung and increased leukocyte infiltration in the + bronchoalveolar lavage fluid [207]. Furthermore, more robust IFN production by CD8 T -/- -/- cells from CB1 CB2 mice compared with WT mice was also observed in the in vitro model system (Fig. 12A). The mechanisms responsible for these findings are yet to be completely elucidated, but are consistent with the widely held premise that CB1 and CB2 are involved in dampening the immune response. CB1 and CB2 are primarily known as inhibitory heterotrimeric G protein coupled receptors (Gi/o) involved in down-regulating the activity of AC, which decreases the levels of intracellular cAMP [97, 134, 138]. The downstream signaling events are in turn influenced, such as decreased activity of transcription 120 factors, AP-1, NFAT, NF-B, which are involved in regulation of cytokine production as -/- - described earlier [141, 142]. Therefore, the enhanced IFN response observed in CB1 CB2 /- mice might be due to the lack of suppression on signaling events downstream of Gi/o mediated by endogenous cannabinoids via CB1 and CB2. In addition, the CB1 agonist AEA was found to attenuate the KV channel-mediated outward currents in DCs, which resulted in a decrease of surface MHC II expression on DCs and therefore a reduced T cell response, suggesting the modulation of DC function by cannabinoids via CB1 signaling [194-196]. Most recently, THC was found to suppress LPS-induced maturation of bone marrow-derived DCs, as evidenced by reduced MHC II, CD80, and CD86 expression, which in turn decreased IFN production by CD8 + T cells after the co-culture [197]. Consistent with these observations, maturation of the non-T/non-B cell population, which is comprised of APCs, -/- -/- such as DCs, was more pronounced in CB1 CB2 surface markers CD69 and MHC II (Fig. 30). -/- -/- observed in CB1 CB2 mice, as assessed by expression of Hence, the enhanced immune response mice might also be due to the lack of suppression on APC function mediated by endogenous cannabinoids via CB1/CB2. As extensively described in the Introduction chapter, THC is primarily known to suppress immune responses, such as host resistance to pathogens and tumor challenge [197, 204, 206, 254, 255]; however, depending on experimental conditions, including cell type, immune stimuli, and cannabinoid concentrations, enhancement by THC and other cannabinoids on immune function has also been reported in some studies. For example, cells from the spleen 121 or lymph nodes in combination with stimulation using anti-CD3, ConA or phytohemagglutinin could result in differential modulation on cellular proliferation by THC [170]. In addition, the age of animals was also found to be an important factor in determining whether cannabinoids produced immune enhancement or suppression [171]. Moreover, nanomolar concentrations of the synthetic cannabinoid, CP55,940, were demonstrated to increase IL-8 and beta-chemokine monocyte chemotactic protein-1 (MCP-1) production in CB2-transfected HL60 cells [172], and low concentrations of cannabinoid-mediated enhancement were also observed in B cells [173]. Cannabinoid-mediated enhancement on IL2 expression was also observed when EL4 T cells were suboptimally activated with low concentrations of PMA, due to binding of NFAT to a distal site in the IL-2 promoter region [174, 175]. Such enhancement was not abrogated by CB2 antagonist SR144528, possibly suggesting a CB-independent mechanism [174, 175]. In particular, from the in vitro studies reported in the first half of this dissertation, we have also demonstrated that THC suppressed the CTL response specific to gp120 antigens when the magnitude of cellular activation was high, and THC enhanced the response when the magnitude of cellular activation was suboptimal. In the current in vivo studies, which were designed such that THC would be present at the time of immune activation, THC produced an enhanced effect in WT mice on the IFN response and activation of various cellular populations, as assessed by expression of CD69 and MHC II. The mechanisms by which THC produced immune enhancement in WT mice are not clear, but based on the above evidence it is possible that the current experimental conditions in this in vivo model system, including the amount of viral DNA employed and the THC dose, induced a modest antigen-specific responses, hence THC produced immune enhancement. In order to test this possibility, further investigations to boost the response, for 122 example, by increasing the number of epitopes or including an adjuvant, would help to resolve this possibility. In addition, it was clearly demonstrated here that the enhancement produced by THC in WT mice on the IFN response was consistent with its effect on the activation level of T cells, indicating that THC modulated T cell effector function, in part, via regulating T cell activation first. Moreover, although CB1 and CB2 are initially reported to interact with inhibitory Gi/o, they have also been shown to couple with stimulatory heterotrimeric G proteins (Gs) to activate AC and therefore the downstream signaling events [149, 256-258]. However, the binding affinity of CB1 or CB2 to Gs or Gi proteins or the expression levels of either G protein in certain tissues remain elusive. Furthermore, since it was shown that CB1 or CB2 agonists, HU-210 and WIN 55,212-2 inhibited the activity of AC isoforms I, III, V, VI, and VIII, but stimulated the activity of AC isoforms II, IV, and VII [150], involvement of different AC isoforms might be another contributing factor to the differential effects mediated by cannabinoids. Therefore, this in vivo model system, which includes numerous components such as THC, endogenous cannabinoids, CB1, and CB2 might lead to stimulation of different G subunits to cause differential effects on the downstream responses. Although the overall THC effects were modest, the trend toward enhancement in WT mice was consistently observed. In contrast, a trend toward suppression by THC on IFN -/- -/- production was seen in CB1 CB2 mice, but no THC-mediated effect was observed on the other endpoints. These results suggest that depending on the particular immune response, the observed THC-induced activity might be mediated through CB1/CB2-independent or 123 dependent mechanisms. Interestingly, the observation that THC enhanced the relatively low magnitude of IFN response in WT mice, but suppressed the relatively strong IFN response -/- -/- in CB1 CB2 mice was consistent with our findings from in vitro studies in which THC differentially modulated immune function depending on the magnitude of immune response. These results also suggest that the effects of THC could be mediated by receptors other than CB1 and CB2. So far, orphan GPCRs, such as GPR18, GPR55, and GPR119, and several TRP channels, such as TRPC1, TRPA1, and TRPV1-6, have all been implicated as targets for cannabinoid ligands [97, 107, 155, 156], but their functions in cannabinoid-mediated immune modulation remain uncertain. Whether any of these receptors plays a role in this in vivo model system is not known and will require further investigation. At the gene transcript levels, increased cytokine (IL-2) production and expression of -/- -/- costimulatory molecule CD40L on T cells in CB1 CB2 -/- mice but not WT mice, is -/- consistent with a more robust immune response in CB1 CB2 -/- mice compared with WT -/- mice, as evidenced by augmented IFN production in CB1 CB2 mice measured by flow cytometry. In response to THC treatment, the mRNA of inhibitory molecule CTLA-4 was downregulated when compared with the CO-pVRCgp120 control group in WT mice, -/- - suggesting an enhanced effect by THC on immune function. On the other hand in CB1 CB2 /- mice, suppression of cytokine production by THC was observed when compared with the CO-pVRCgp120 control group, as evidenced by downregulation of lymphocyte-specific cytokines (IL-17 and IL-2) and APC-produced cytokine (IL-12) [259]. However, THC also attenuated the suppressive effect on cytokine production by downregulating the transcripts of 124 -/- -/- negative regulators Socs1 and Socs2, suggesting opposing effects by THC in CB1 CB2 mice. It is possible that the suppressive effect was more robust, therefore decreased IFN -/- -/- production by THC was evident in CB1 CB2 mice as measured by flow cytometry. Collectively, the in vivo studies identified an immunodominant peptide derived from HIVgp120, which induced antigen-specific immune responses, including production of cytokines, IFN and IL-2, activation of different cellular populations, and the generation of a secondary antibody response. THC treatment enhanced HIVgp120-induced expression levels -/- -/- of activation markers only in WT but not CB1 CB2 mice, suggesting a CB1/CB2- dependent mechanism of THC-mediated activity, which might be due to the low magnitude of stimulation or activation of different G subunits upon ligation of CB1 and/or CB2. -/- -/- Furthermore, the absence of CB1 and CB2 in CB1 CB2 mice led to a phenotype with enhanced immune responsiveness in some respects, such as cytokine production, suggesting the potential involvement of endogenous CB1 and/or CB2 ligands. Finally, THC enhanced -/- -/- IFN production in WT mice, but suppressed IFN production in CB1 CB2 mice, suggesting a CB1/CB2-independent mechanism and the putative involvement of other receptors. Together, these studies provide an approach to investigate cannabinoid-mediated effects on viral antigen-specific immune responses in vivo, and suggest possible enhancement of immune function by cannabinoids in response to HIV antigens. 125 III. Significance and concluding remarks The studies presented in this dissertation have made significant contributions to the understanding of the effect of cannabinoids in the immune system, in particular on HIV antigen-specific immune responses. By characterizing the immune responses induced by -/- -/- HIVgp120 viral antigens in vitro and in vivo using WT and CB1 CB2 mice, the role of CB1/CB2 was also further elucidated. AIDS is one of the leading causes of high death rates in the US [19]. It is known that HIV patients suffer severe pain, nausea, appetite loss and wasting syndrome associated with weight loss and cachexia. Approximately 25% of HIV/AIDS patients have been using cannabis as an appetite stimulant, anti-emetic and/or analgesic for alleviating some of the symptoms listed above. Use of cannabis has provided relief of anxiety, depression, and pain, improved appetite, and increased pleasure [94]. Since HIV patients are immunocompromised, they are more susceptible to further pathogen infections. At the same time, cannabinoids are primarily known to exert a negative impact on immune function as described in the Introduction chapter, therefore whether marijuana can further impair the immune competence of HIV patients has become a major concern. There is still a lack of research and knowledge on understanding the effect of cannabinoids on the immune function in HIV patients. So far, only a few studies have reported on the effects of short-term or chronic THC treatment on HIV viral load and T cell counts in human patients or nonhuman primate models [214-216]. However, it is important to emphasize that in these studies neither antigen-specific immune responses nor the associated molecular and cellular mechanisms were investigated. Therefore, the first unique contribution of this work is the establishment of two antigen-specific models. 126 Specifically, an in vitro model was first utilized to investigate basic parameters associated with T cell response against HIVgp120, and a subsequent in vivo model using the same stimulation strategy was employed to test this model under physiological conditions of both antiviral immune responses and drug metabolism. This is a stepwise incremental approach to increase the complexity of the model system to make it gradually closer to the real life situation in HIV patients who use marijuana for treating HIV-associated adverse effects. These models are important experimental approaches and can be potentially applied to various cannabinoids or other xenobiotics and other HIV or non-HIV derived viral proteins. In addition, the immunodominant epitope(s) derived from interested viral proteins can be identified using a similar in vivo model system. The second novel finding from these studies is that THC exhibits differential effects on HIVgp120 antigen-specific immune responses depending on the magnitude of the immune response. In vitro, THC was demonstrated to suppress gp120-specific IFN production and + proliferation of CD8 T cells when the magnitude of response was high, but enhance the aforementioned responses when the magnitude of response was low. Although the experimental factors determining the magnitude of the response under the current conditions are not clear, such cannabinoid-mediated differential effects on cytokine (IFN and IL-2) production were also observed when pharmacological activators were used to stimulate the cells under optimal (10x P/I, iCD3/CD28, sCD3/CD28 only for IFN) or suboptimal (1x P/I, sCD3/CD28 only for IL-2) conditions. My findings suggested that the magnitude of cellular activation dictates the cannabinoid-mediated differential T cell response to HIVgp120. Previous reports imply that experimental conditions, including cell type, cannabinoid 127 concentration, and stimuli, influence the effect of cannabinoids on immune responses [171175, 233], and my results provide further insight into the decisive experimental factors. The third important finding is that the modulatory effects of cannabinoids on immune function may be acting on various cellular components simultaneously. The in vitro and in vivo studies performed in this dissertation research are very comprehensive, which + + demonstrate that cannabinoids modulate the functions of CD8 T cells, CD4 T cells, NK cells, B cells, and non-T/non-B cells (consisting primarily of APCs). Due to the fact that + DC2.4 cells express very limited amount of MHC II molecules, the CD4 T cell response was + not detected, so the in vitro model mainly focused on the CD8 T cell response. However, the in vivo studies clearly show that multiple cellular populations are the targets of cannabinoid-mediated modulation. In addition, we demonstrate that each stage in the development of cell-mediated and humoral immunity can be modulated by cannabinoids. In vivo, THC-mediated modulatory effects on T cell and B cell effector function, including cytokines specifically induced by gp120 and IgG production, which correlated with effects on the activation levels of T cells and B cells, as measured by expression levels of CD69, CD80, and MHC II. In vitro, the + activity of THC on cytokine production by CD8 T cells was consistent with its effects on + CD8 T cell proliferation. Therefore, the in vitro and in vivo studies complement each other, and together indicate that cannabinoids modulate activation, proliferation and the effector functions of the differentiated immune cells. Furthermore, mRNA expression of IFN and IL-2 and nuclear translocation of transcription factor NFAT were also differentially regulated by cannabinoids under optimal 128 (10x P/I) or suboptimal (1x P/I) stimulation conditions, suggesting that cannabinoid-mediated modulation also occurs at the transcriptional levels. However, surprisingly, cannabinoids did not differentially regulate but increased intracellular calcium levels regardless of the stimulation condition. Therefore, we have speculated that optimal stimulation in combination with cannabinoids leads to excessive intracellular calcium, which decreases downstream cytokine production, whereas suboptimal stimulation in combination with cannabinoids results in an optimal level of intracellular calcium, which increases cytokine production. These results also imply that cannabinoids are likely to differentially modulate signaling events distal to increased intracellular calcium, which might be NFAT cytoplasmic regulators or nuclear kinases (e.g., STIM, ORAI, and DYRK1A). Although both suppressive and enhancing effects by cannabinoids on immune function have been reported previously by us and others, this is the first time a possible mechanism is suggested. Lastly, the role of CB1/CB2 in cannabinoid-mediated immune modulation has been extensively elucidated mainly using the in vivo model. With the demonstration that THC produced enhancement on activation of various cellular populations in WT mice but no effect -/- -/- in CB1 CB2 mice, a CB1/CB2-dependent mechanism is suggested. On the other hand, -/- -/- IFN production was enhanced by THC in WT mice but suppressed in CB1 CB2 mice, hence suggesting a CB1/CB2-independent mechanism and the existence of additional receptors other than CB1/CB2. Therefore, depending on the response being measured, both CB1/CB2-dependent and -independent mechanisms may be involved in cannabinoid-mediated immune modulation. It is unknown what receptor(s) are responsible for the aforementioned 129 THC-mediated effects. Studies identifying the potential target(s) will provide further information on the mechanisms by which cannabinoids modulate immune responses. It is -/- -/- also noteworthy that consistent with findings from other studies [207, 213], CB1 CB2 mice exhibited a phenotype of enhanced immune responsiveness when compared with WT mice, as shown by more robust IFN production, suggesting the involvement of CB1/CB2 in endocannabinoid-mediated immune suppression. In summary, as described above, this dissertation research advances the knowledge of the effects of cannabinoids and the role of CB1/CB2 on immune regulation to viral antigens in several ways. In addition, our findings further elucidate the mechanisms by which cannabinoids and cannabinoid receptors influence immune modulation. Moreover, these studies provide a basis for developing future studies in which primary human PBMCs from healthy or HIV patients, who may also currently use marijuana for appetite stimulation, can be used to determine correlations between the data presented in this dissertation and human health. This work also begins to provide valuable information for HIV/AIDS patients who are currently using marijuana for therapeutic benefits. 130 APPENDIX 131 INDUCED T CELL CYTOKINE PRODUCTION IS ENHANCED BY ENGINEERED NANOPARTICLES ABSTRACT Engineered nanoparticles are widely used in commercial products and yet due to the paucity of safety information there are concerns surrounding potential adverse health effects, especially from inhaled nanoparticles and their putative contribution to allergic airway disease. The objective of this study was to investigate whether engineered nanoparticles possess adjuvant-like or immune enhancing properties on antigen-specific T cell responses using in vitro models. Ovalbumin (OVA)-derived peptides were presented to T cells by either spleenderived endogenous APCs or a mouse DC line, DC2.4. In all models, IFN and IL-2 production by CD8+ or CD4+ T cells in response to peptide OVA257‐264 or OVA323-339, respectively, was measured by flow cytometry. Silica nanoparticles (SNPs) were modified with alkyne-terminated surfaces and appended with polyethylene glycol (PEG) chains via “click” chemistry. Modified SNPs were not likely to agglomerate in in vitro culture media, suggesting that their effect on leukocyte function was the result of true nanoscale. Under suboptimal T cell stimulation conditions, modified SNPs (up to 10 g/ml) enhanced the proportion of CD8+, but not CD4+, T cells producing IFN and IL-2. Various functional groups (-COOH, -NH2, -OH) on modified SNPs enhanced IFN and IL-2 production to different levels, with -COOH SNPs being the most effective. Furthermore, 51 nm -COOH SNPs exhibited a greater enhancing effect on CD8+ T cell response than other sized particles. Collectively, our results show that modified SNPs can enhance antigen-specific CD8+ T cell responses, suggesting that certain modified SNPs exhibit potential adjuvant-like properties. 132 INTRODUCTION Nanoparticles can occur naturally in the environment or can be artificially engineered for a wide range of applications in industry, such as electronics, medicine, food, clothing and cosmetics [260]. Nanoparticles have potentially beneficial effects; for example, their therapeutic use in vaccine development [261]. By contrast, the same mechanisms that account for their beneficial attributes in vaccine development can contribute to putative adverse effects on human health [262, 263]. Some of these concerns have arisen from animal studies, which have demonstrated that nanoparticles can exhibit adjuvant-like properties and can also contribute to allergic pulmonary disease through increased granulocyte infiltration, production of proinflammatory cytokines and allergen-specific antibodies [264, 265]. Little is presently known concerning the consequences resulting from interactions of engineered nanoparticles with the immune system, especially at the cellular level. The objective of this study was to investigate whether engineered nanoparticles can enhance antigen-driven T cell responses to provide a mechanism of nanoparticle-mediated adjuvant effects, and if so, whether these enhancing properties are influenced by the surface chemistry or specific size of the nanomaterials. In the present investigation SNPs were synthesized, which were modified with alkyneterminated surfaces and appended with PEG azides via “click” chemistry. Pegylated SNPs are dispersed in water and can readily interact with, as well as enter, cells [266, 267]. Preliminary in vivo studies using bronchoalveolar lavage fluid (BALF) and lymph node (LN) cells from OVA/SNP-treated mice have demonstrated the potential adjuvant-like effects of these SNPs on cell counts and activation of lymphocytes as well as other cell populations [268]. However, since in vivo study was limited by the amount of materials and the number 133 of T cells obtained in BALF and LN samples, in vitro models were chosen to delineate the effect of SNPs on T cell functions directly. In addition, these in vitro models could facilitate the study of mechanisms associated with SNP-mediated immune enhancing effect. In this study, antigen-specific T cell responses in vitro were induced. T cells were obtained from spleens of OT-I (CD8+ T cells) or OT-II (CD4+ T cells) mice, which have been engineered to express transgenic T cell receptors specific for OVA-derived peptides OVA257264 (SIINFEKL) or OVA323-339 (ISQAVHAAHAEINEAGR), respectively [269, 270]. In the OT-I model, OVA257-264 peptides that bind to MHC I molecules were presented by either endogenous APCs from the splenic population or a mouse DC line, DC2.4, to activate OT-I CD8+ T cells specifically. By using DC2.4 as APCs, OVA peptides were directly loaded onto DC2.4 cells, which were then co-cultured with splenic cells. In the OT-II model, OVA323-339 peptides that bind to MHC II molecules were presented by endogenous APCs from the splenic population to OT-II CD4+ T cells specifically. Production of cytokines, including IFN and IL-2, were used as a measure of T cell function. The physical and chemical properties of the engineered nanoparticles determine their uptake by cells and therefore their activity in the cells [271-273]. Therefore, SNPs were synthesized in different sizes and with modifications to surface chemistry through the addition of functional groups, including -COOH, -NH2, and -OH, which were not likely to agglomerate in in vitro culture media by performing dynamic light scattering (DLS) analysis. Using the above-mentioned antigen specific models, the objective of this study was to evaluate the effects of modified SNPs on T cell functions using suboptimally-activated CD8+ and CD4+ T cells. 134 METHODS SNPs LUDOX® TM-40 colloidal silica (LTM40, 40 wt% suspension in H2O), 3(triethoxysilyl)propyl isocyanate, propargyl alcohol, sodium ascorbate and sodium azide were purchased from Sigma-Aldrich (St. Louis, MO) and used as received. Colloidal silica SNOWTEX XS (SNTXS, 4-6 nm, 20 wt% suspension in H2O), SNOWTEX 20L (SNT20L, 40-50 nm, 20 wt% suspension in H2O), and SNOWTEX ZL (SNTZL, 71 nm, 40 wt% suspension in H2O) were gifts from Nissan Chemical Industries, Ltd (Pasadena, TX). 1azido-2-(2-(2-(2-methoxyethoxy)ethoxy)ethoxyethane (N34PEG) was synthesized according to a literature procedure [274]. Dialysis membranes (Spectra/Por® Biotech Cellulose ester, MWCO: 300K, 12-14K) were purchased from Spectrum Laboratories Inc. (Rancho Dominguez, CA). Ion exchange resin (Amberlite® IRC-748) was purchased from Alfa Aesar (Ward Hill, MA). Instruments and SNP characterization 1 H Nuclear magnetic resonance (NMR) and 13C NMR spectra were recorded in CDCl3 on a Varian 300 MHz or VXR-500 MHz instrument (Agilent Technologies, Santa Clara, CA). The CDCl3 resonance was used as the internal standard for 13 C NMR ( = 77.0 ppm) and residual CHCl3 for 1H NMR ( = 7.24 ppm). Fourier Transform Infrared spectroscopy (FTIR) spectra were recorded on a Mattson Galaxy series FTIR 3000 (Mattson Instruments, Inc., Madison, WI). Thermogravimetric analyses (TGA) were obtained in air from a Perkin-Elmer TGA 7 (Perkin Elmer Corporation, Norwalk, CT). Samples were held at 120 °C for 30 min to 135 remove adsorbed water from the particle surfaces, and then heated to 850 °C at a rate of 10 °C/min. All FTIR and TGA samples were dried under vacuum at room temperature for 24 h. DLS data were obtained with a Malvern NanoZS ZetaSizer (Malvern Instruments Inc., Westborough, MA) with 178° backscattering detection. Intensity and average diameters were calculated from the autocorrelation function using Malvern’s Zetasizer Software 6.12. Samples for DLS analyses were sonicated at 60 C prior to measuring particle sizes at 25 C (FS20H sonicator, Fisher Scientific, Pittsburgh, PA). For nanoparticle stability studies, particles were added to fresh in vitro culture media (see below), dispersed by vortex mixing, and characterized by DLS at 0, 4, 8, 21, 42, 68 h at 25 C. Particles only and media only were included as experimental controls. Synthesis of reagents for modifying nanoparticles and click chemistry Synthesis of ((2-propynylcarbamate)propyl)triethoxysilane (PPTrEOS) (Fig. 31A). Propargyl alcohol (12 g, 0.21 mol) and triethylamine (19 g, 0.19 mol) were dissolved in 120 mL of dry CH2Cl2. The mixture was cooled in an ice bath, and under nitrogen, a solution of 3-(triethoxysilyl)propyl isocyanate (IPTEOS) (46 g, 0.18 mol) in 60 mL of dry CH2Cl2, was added dropwise to the flask. The reaction mixture was stirred for 24 h at room temperature and then filtered. CH2Cl2, excess propargyl alcohol, and NEt3 were evaporated under reduced pressure to afford 50 g of PPTrEOS as pale-yellow oil. 1H NMR (500 MHz, CDCl3) 0.52 (t, 2H, J = 8Hz), 1.11 (t, 9H, J = 8 Hz), 1.52 (m, 2H), 2.37 (s, 1H), 3.08 (m, 2H), 3.71 (q, 6H, J = 7 Hz), 4.55 (s, 2H). 136 Synthesis of 2-[2-[2-[2-azidoethoxy]ethoxy]ethoxy]ethanol (N34PEGOH) (Fig.31B). A solution of TsCl (14 g, 0.071 mol) in 250 mL of tetrahydrofuran (THF) was added dropwise to a solution of tetraethylene glycol (34 g, 0.18 mol), triethylamine (14 mL) and THF (250 mL). After stirring the resulting solution for 36 h at room temperature, THF was removed by rotary evaporation and the crude product was dissolved in 200 mL of CH2Cl2. The organic layer was washed with water (3 × 150 mL), aqueous NaHCO3 (200 mL), and brine (200 mL). After drying over Na2SO4, the solution was concentrated and dried under vacuum to give 18 g of a light yellow liquid comprised of a mixture of mono-tosylate (85%) and di-tosylate (15%), which corresponded to 14.3 g of mono-tosylate (58% yield). The product was used in the next step without further purification. 1H NMR (500 MHz, CDCl3) δ 7.77 (m, 2H), 7.32 (m, 2H), 4.29 (m, 2H), 3.69-3.50 (m, 14H), 2.5 (br, 1H), 2.42 (s, 3H). The tosylates were dissolved in dimethylformamide (DMF) (210 mL), treated with NaN3 (10.4 g, 15 mmol), and then heated to 60 °C for 24 h. Then the mixture was concentrated, re-dissolved in ethyl acetate, filtered to remove the solid sodium tosylate, and concentrated under reduced pressure. The resulting oily product was dissolved in water and washed with hexane until ,-diazido tetraethylene glycol was not detected in the aqueous layer. Drying under vacuum gave the product as a light yellow liquid (8.0 g, 72%). 1H NMR (500 MHz, CDCl3) δ 3.69 (m, 2H), 3.64 (m, 10H), 3.58 (m, 2H), 3.36 (t, 2H, J = 5.5 Hz), 2.8 (br, 1H); 13 C NMR (125 MHz, CDCl3) 72.41, 70.55, 70.51, 70.43, 70.18, 69.90, 61.56, 50.54. Synthesis of 2-[2-[2-[2-azidoethoxy]ethoxy]ethoxy]ethyl amine (N34PEGNH2) (Fig. 31C). A solution of TsCl (39.9 g, 0.21 mol) in THF (150 mL) was added dropwise into an ice-cold mixture of tetraethylene glycol (13.5 g, 0.07 mol), aqueous KOH (100 g, 47%, w/w), 137 and THF (100 mL). After stirring the solution for 36 h at room temperature, THF was removed by rotary evaporation, and then CH2Cl2 (200 mL) was added to the residue. The organic layer was washed with water (3 × 150 mL), aqueous NaHCO3 (200 mL), and brine (200 mL). After drying the organic layer over Na2SO4, the solution was concentrated and dried under vacuum to give 32 g of a light yellow liquid (91%), which was used in the next step without further purification. 1H NMR (300 MHz, CDCl3) δ 7.81(m, 4H), 7.36 (m, 4H), 4.20 (m, 4H), 3.70-3.65 (m, 12H), 2.47 (s, 6H). The di-tosylate of tetraethylene glycol was treated with NaN3 (11.7 g, 18 mmol) in DMF (300 mL) at 90 °C for 24 h. The mixture was concentrated, dissolved in water, and extracted with CH2Cl2 (4 × 100 mL). The solution was dried over anhydrous Na2SO4 and concentrated under reduced pressure to give 12 g of a yellow liquid. Aqueous HCl (5%, 100 mL) was added to the yellow liquid, and under vigorous stirring at room temperature, a solution of PPh3 (11.4 g, 0.041 mol) in 75 mL of ether was added dropwise. After stirring for 24 h, the ether was removed by rotary evaporation and the aqueous layer was extracted with CH2Cl2 until triphenylphosphine oxide was not detected in the aqueous layer. The aqueous layer was adjusted to pH=12, and amine was extracted from the aqueous layer with CH2Cl2 (6 × 50 mL). Drying under vacuum gave the product as a light yellow liquid (9.0 g 68%). 1H NMR (500 MHz, CDCl3) δ 3.72-3.58 (m, 10H), 3.48 (t, 2H, J = 5 Hz), 3.35 (t, 2H, J = 5 Hz), 2.84 (t, 2H, J = 5Hz), 2.24 (s, 2H); 13C NMR (125 MHz, CDCl3) 72.79, 70.50, 70.47, 70.43, 70.07, 69.85, 50.50, 41.44. Synthesis of 2-[2-[2-[2-azidoethoxy]ethoxy]ethoxy]acetic acid (N34PEGCOOH) (Fig. 31D). A solution of [2-[2-[2-chloroethoxy]ethoxy]ethanol (16.8 g, 0.1 mol) and NaN3 (13 g, 0.2 mol) in 75 mL of water was heated at 75 °C with stirring for 24 h. Water was removed by 138 rotary evaporation and the crude product was extracted from the salts with 200 mL of CH2Cl2. Concentration and drying under vacuum gave the product as a light yellow liquid (17 g, 97%), which was used without further purification. 1 H NMR (500 MHz, CDCl3) δ 3.72-3.62 (m, 8H), 3.58 (m, 2H), 3.37 (t, 2H, J = 5 Hz). The light yellow liquid product (14 g, 0.08 mol), prepared above, was added to NaH (0.1 mol) in 350 mL of dry THF cooled in an ice bath. After stirring for 1 h at room temperature, ethyl bromoacetate (17.7 mL, 0.16 mol) was added dropwise to the solution and then stirred for 48 h at room temperature. The THF was removed by rotary evaporation, and then 150 mL of water was added. The aqueous layer was extracted with CH2Cl2 (3 × 50 mL), dried over anhydrous Na2SO4, and concentrated under reduced pressure at 55 °C to give an amber liquid (26 g), which was used without further purification. 1H NMR (500 MHz, CDCl3) δ 4.24 (q, 2H, J = 10 Hz), 4.17 (s, 2H), 3.77-3.65 (m, 10H), 3.41 (t, 2H, J = 5 Hz). The ester was hydrolyzed with 53 mL of 3M NaOH at room temperature for 18 h. After extracting the aqueous layer with CH2Cl2 (10 × 30 mL) to remove mineral oil and other impurities, the aqueous layer was acidified to pH=1 and extracted with EtOAc (1 × 80 mL, 7 × 30 mL). After drying over anhydrous Na2SO4, the residue was concentrated to a pale red liquid (14 g, 75%). 3.78-3.67 (m, 10H), 3.41 (t, 2H, J = 5 Hz); 1 H NMR (500 MHz, CDCl3) δ 4.18 (s, 2H), 13 C NMR (125 MHz, CDCl3) 177.88, 76.71, 75.95, 75.69, 75.48, 75.31, 74.02, 55.9. Surface modification of SNPs Synthesis of XXXXPTr: One-step alkyne modification of SNPs (Fig. 31E). XXXX represented different commercial SNPs, including LTM40, SNTXS, SNT20L, and SNTZL (9 g, 40% suspension in water), which were diluted with 75 mL of 1:1 EtOH-H2O and sonicated. 139 Particle size was measured by DLS prior to modification. pH of the silica suspension was adjusted to ~9 by adding drops of aqueous ammonia. At room temperature, a solution of PPTrEOS (3 g, 10 mmol) in 10 mL of EtOH was added dropwise to the stirred silica suspension. After stirring for 3 days, the suspension was diluted with EtOH until it was homogeneous. The modified particles were purified by dialysis in EtOH and water (9:1) using dialysis tubing (MWCO: 300K). The surface coverage of the alkyne functional group was evaluated by TGA. The particle concentration (mg/g) was determined by evaporating 2 4 mL of purified particle solutions. Synthesis of XXXXPTrgR’ with different functional groups: “Click” chemistry of SNPs (Fig. 31E). Click chemistry between alkyne-modified SNPs and clicking reagents (R’, including N34PEG, N34PEGOH, N34PEGNH2, and N34PEGCOOH) is described using the synthesis of LTM40PTrg4PEG as an example. LTM40PTr (100 mg, 0.036 mmol of alkyne functional groups) and N34PEG (41.9 mg, 0.18 mmol, 5 equiv.) were added to 13.8 g of EtOH and water (9:1). Then this mixture was diluted in 3:1 DMF/water until the solution was clear. After adding sodium ascorbate (14.3 mg, 2 equiv.) to the solution, trace O2 was removed by three freeze-pump-thaw cycles. Aqueous CuSO4 (72 μL, 0.1 M, 0.2 equiv.) was added to the solution followed by a final degassing process, and the solution changed to light brown-yellow. After stirring at room temperature for 36 h, 1 mL of aqueous EDTA (0.1M) was added and the solution was stirred for another 6 h. The resulting particles were dialyzed against (MWCO:12-14K) EtOH/water (9:1, 5 × 2 L) and against distilled water to afford purified surface-modified SNPs with different surface chemistry. For the synthesis of LTM40PTrg4PEGNH2, EDTA was not added after click chemistry, but was dialyzed directly 140 as stated above, and then the solution of LTM40PTrg4PEGNH2 was sonicated to be homogeneous. Ion exchange resin (2 g, Amberlite® IRC-748) was added to remove the Cu2+, and it was removed via filtration after 8 h. The modified SNPs, including LTM40PTrg4PEG, LTM40PTrg4PEGOH, LTM40PTrg4PEGNH2, LTM40PTrg4PEGCOOH, SNTXSPTrg4PEGCOOH, SNT20LPTrg4PEGCOOH, and SNTZLPTrg4PEGCOOH, were kept in distilled water until further characterization. The surface coverage of surface functional groups after each step modification was evaluated by TGA. g indicates that the clicking reagents were grafted to the particles via “click” chemistry. DLS samples were prepared by diluting or concentrating solutions to 0.1 or 1 mg/mL. The particle concentration (mg/g) was determined by evaporating 2-4 mL of purified particle solutions. Mice Female OT-I and OT-II mice were purchased from Jackson Laboratory (Bar Harbor, ME, USA) and used at 8 -12 weeks of age. Mice were randomized at no more than 5 animals per cage and provided rodent chow (Harlan Teklad, Madison, WI) and water ad libitum. Rooms were kept on a 12-hr light/dark cycle at 21-24°C and 40-60% humidity. All experiments were performed following the guidelines set forth by Michigan State University Institutional Animal Care and Use Committee. Cell culture conditions DC2.4 cells (a kind gift from Dr. Kenneth Rock at University of Massachusetts) were maintained in complete RPMI media (10% BCS, 100 units/ml penicillin and 100 µg/ml streptomycin [pen/strep]). 141 Figure 31. Synthesis of different SNPs. (A-D) Synthesis steps are illustrated for PPTrEOS (A), N34PEGOH (B), N34PEGNH2(C), and N34PEGCOOH (D). (E) Particle synthesis procedure is illustrated for modified SNPs. XXXX represents different commercial SNPs, including LTM40, SNTXS, SNT20L, and SNTZL. PTr represents particles modified with alkyne functional groups. R’ represents different clicking reagents, including N34PEG, N34PEGOH, N34PEGNH2, and N34PEGCOOH, which are PEG chains of 4 repeating units terminated with -OMe, -OH, -NH2, and -COOH functional groups, respectively. g represents that the clicking reagents were grafted to the particles via “click” chemistry. 142 Activation of OT-I or OT-II T cells with OVA peptides in vitro Splenocytes were isolated aseptically from OT-I or OT-II mouse spleens and made into single-cell suspensions. Splenocytes (8 x 105 cells/well) were treated with various concentrations (10-8 – 10-2 g/ml) of OVA257-264 peptide SIINFEKL or OVA323-339 peptide ISQAVHAAHAEINEAGR (Anaspec, San Jose, CA) in 48-well plates for 2 days in 2% BCS RPMI media (2% BCS, pen/strep, and 50 µM 2-mercaptoethanol). When DC2.4 cells were used as APCs, they were plated at 2 x 106 cell/well in 6-well pates and pretreated with 1 g/ml LPS (Salmonella typhosa, Sigma-Aldrich) in 2% BCS RPMI media overnight. DC2.4 cells were then incubated with various concentrations of SIINFEKL (10-2 – 10 g/ml) for 3 – 4 h, X-ray irradiated at 35 gray to prevent their proliferation, washed 3 times to remove the extra SIINFEKL, and plated 2 x 105 cell/well in 96 U-Bottom plates with 1 x 106 cells/well OT-I splenocytes for 4 days in 5% BCS RPMI media (5% BCS, pen/strep, and 50 µM 2mercaptoethanol). Peptide concentration for suboptimal stimulation of T cell response was determined and used to further study the influence of modified SNPs on T cell function. Modified SNPs treatment in cell culture Once the suboptimal stimulation condition was determined, splenocytes were treated with modified SNPs with or without different surface functional groups or in different sizes, including LTM40PTrg4PEG, LTM40PTrg4PEGCOOH, LTM40PTrg4PEGOH, SNTXSPTrg4PEGCOOH, LTM40PTrg4PEGNH2, SNT20LPTrg4PEGCOOH and SNTZLPTrg4PEGCOOH. These modified SNPs were sonicated at 60C for 35 min and then added to the splenocyte suspension at various concentrations (10-2 – 10 g/ml) prior to the OVA peptide stimulation as describe above. As reviewed by Ganwal et al. [275], 10 g/ml 143 concentration does not fall into the high-end concentrations (~30 – 400 µg/ml) of nanoparticles for in vitro assay testing based on estimates of lung deposition after occupational exposure. Flow cytometry analysis On day 2 post the initiation of peptide stimulation, brefeldin A (Biolegend, San Diego, CA) was added to the culture during the last 6 h of incubation to prevent cytokine release and allow for the identification of cytokine producing T cells. Cells were washed with 1 x HBSS buffer (Invitrogen, Carlsbad, CA) and incubated with LIVE/DEAD Fixable Near-IR Dead Cell Stain Kit (Invitrogen) to assess cell viability according to manufacturer’s instructions. For surface molecule staining, cells were blocked for Fc receptors with purified rat antimouse CD16/CD32 (“Fc block”, BD Pharmingen, San Diego, CA), then incubated with PE/Cy7-conjugated rat anti-mouse antibodies against CD8 (clone 53-6.7, Biolegend,) or CD4 (clone RM4-5, Biolegend), and fixed with Cytofix (BD Pharmingen). On the day of flow cytometric analysis, cells were permeabilized with 1 x Perm/Wash (BD Pharmingen) and stained with PE-conjugated anti-mouse IFN antibody (clone XMG1.2, Biolegend) and APCconjugated anti-mouse IL-2 antibody (clone JES6-5H4, Biolegend). Single stain controls were included in all experiments to compensate for fluorescence interference between detectors. Cells were washed with 1 x Perm/Wash, resuspended in FACS buffer (1 x HBSS containing 1% BSA and 0.1% sodium azide), assessed on a FACS Canto II flow cytometer (BD Biosciences, San Jose, CA), and analyzed with FlowJo v8.8.6 (Tree Star, Ashland, OR) or Kaluza 1.1 (Beckman Coulter, Miami, FL) software. In the experiments where DC2.4 cells were used for elicitation, on day 4 post the initiation of the co-culture, cells were restimulated 144 with SIINFEKL (10 g/ml) in the presence of brefeldin A for 6 h in 2% BCS RPMI media. Cells were then stained for viability, surface CD8 expression, and intracellular cytokine production as described above. Statistical analysis Graphing and statistical analyses were performed using GraphPad Prism v4.0 (Graphpad Software, San Diego, CA). The mean  S.E. is presented in all bar graphs. To determine the statistically significant changes between peptide stimulated and unstimulated cells, two-way ANOVA for parametric data was performed. Bonferroni post-test was used to compare samples to their control group. Differences between means from each treatment group were determined using one-way ANOVA. When significant differences were detected, treatment groups were compared to the appropriate control using Dunnett’s test. 145 RESULTS Surface chemistry of modified LTM40PTrg4PEG SNPs PPTrEOS was synthesized as a pure material with a high yield. Characterization by 1H NMR and 13 C NMR indicated that further purification was not required [276, 277]. Commercial SNPs were condensed with PPTrEOS in basic EtOH-H2O mixture for 3 days, as outlined in Fig. 31E. After condensation, alkyne-modified SNPs were purified via dialysis, using tubing with molecular weight cut off (MWCO) values appropriate for the sizes of the particles. Purification continued until no organic chemicals were detected in the dialysate residue by 1H NMR analysis, usually lasting between 1.5 to 8 days. After dialysis was completed, particles were recovered by evaporating the dialysis solvent for further characterization. ‘Click’ chemistry on SNP surfaces was conducted to afford SNPs with different functional group moieties. These modified SNPs were purified by dialysis and used for further surface characterization, as well as in in vitro culture studies. The changes in nanoparticle surface chemistry were tracked by FTIR. Fig. 32A showed normalized FTIR spectra of LTM40 and its modified SNPs as described in Materials and Methods. The band at 806 cm-1, characteristic of bulk silica, allowed normalization of the IR data and semi-quantitative comparison of the nanoparticles. For example, LTM40PTr showed new absorption at 2900-3300 cm-1, 2123 cm-1, 1700 cm-1 and 1430 cm-1, which corresponded to the alkyl group, triple bond, carbonyl group, and another alkyl group, respectively, and confirmed the functionalization of particle surface with alkynes. After “click” modification of the particles, bands at 2930 and 1640 cm-1 increased and were consistent with adding methylene groups from PEG units and the triazole ring (top 4 traces, Fig. 32A). FTIR was limited for characterizing pegylation, since the characteristic C-O bands of PEG were buried 146 under intense IR bands from the SNPs. However, C-H bending and stretching absorption bands were consistent with the PEG chain in LTM40PTrg4PEG (-OMe terminated) particles, as was seen in all other pegylated SNPs (-OH, -NH2, and -COOH terminated particles). TGA data in Fig. 32B and Table 3 confirmed the step-by-step modification of SNPs with different functional groups on the surface. The grafting density on particle surfaces was calculated from TGA mass loss in air, which corresponded to oxidation of the organic layer on the nanoparticle surfaces. Table 3 showed a 5.1% weight loss for alkyne-modified nanoparticles (LTM40PTr), which was calculated to a grafting density of 1.54 alkyne groups/nm2. After clicking pegylated functional groups to LTM40PTr, TGA analyses showed 7.5-8.7% weight loss for all SNPs. The mass losses corresponded to a grafting density of ~1.5 chain/nm2 and >90% high click conversion, indicating successful synthesis of waterdispersed nanoparticles. DLS measurements were conducted with LTM40, LTM40PTr, and the four click products, LTM40PTrg4PEG (-OMe), LTM40PTrg4PEGOH (-OH), LTM40PTrg4PEGNH2 (NH2), and LTM40PTrg4PEGCOOH (-COOH) (Table 3 and Fig. 32C). All surface-modified nanoparticles had monomodal distributions. After modifying the surface of LTM40 with alkynyl groups to make LTM40PTr, its size increased from 30 nm to 37 nm with a zeta potential of -35mV measured in zwitterionic buffer (100mM KCl/10 mM HEPES). The sizes of the four products obtained from LTM40PTr using click chemistry increased to 56, 50, 66 and 51 nm for LTM40PTrg4PEG, LTM40PTrg4PEGOH, LTM40PTrg4PEGNH2, and LTM40PTrg4PEGCOOH, respectively, and zeta potentials were changed to -10mV, -13mV, 7mV, and -17mV, respectively. The DLS size of the click products slightly depended on the surface chemistry and their steric stabilization. 147 Surface chemistry of modified -COOH SNPs at different sizes Grafting data from different sized -COOH SNPs are summarized in Table 4. The initial particle diameters ranged from 20 to 120 nm. Fig. 32D showed normalized FTIR spectra for SNPs terminated with -COOH groups, including LTM40PTrg4PEGCOOH, SNTXSPTrg4PEGCOOH, SNT20LPTrg4PEGCOOH, and SNTZLPTrg4PEGCOOH. Again, the IR band at 806 cm-1 (silica) was used to normalize the spectra and allowed semiquantitative comparisons of the nanoparticles. All spectra showed IR absorption bands at 2900-3300 cm-1 and 1700 cm-1, which corresponded to C-H stretching and a carbonyl group, respectively, confirming successful modification of the particle surface. Since the specific surface areas of small SNPs are higher than large particles, small particles appear to have higher grafted layers than larger particles. Therefore, the IR bands in Fig. 32D decreased in intensity from top to bottom (smallest particle to largest). Table 4 and Fig. 32E showed TGA results for -COOH terminated SNPs. With increasing particle sizes (33 nm to 137 nm), TGA weight losses decreased from 15.9% to 2.2%, as expected due to the decreasing surface to mass ratios. The click conversion calculated from TGA data showed high conversion for all particles (>80%). The DLS traces (Fig. 32F) and the data from SNPs modified with -COOH groups are presented in Table 4. All modified nanoparticles had monomodal distributions (intensity distribution), a result of the steric stabilization of surface modified particles. 148 Figure 32. Surface chemistry of modified SNPs with different functional groups and at different sizes. (A and D) FTIR spectra were obtained by scanning the pellets of different SNPs and KBr from 500 to 4000 cm-1, and the absorbance was recorded. FTIR spectra were normalized using the band intensity at 806 cm-1 for semi-quantitative comparison of the SNPs. (B and E) TGA traces of different SNPs were measured by burning the SNPs in air at a rate of 10 C/min, and the weight retention of SNPs at different temperatures (120 to 850 C) was recorded. (C and F) Different SNPs (100 μg/ml) were sonicated, and DLS data (size distribution by intensity) were measured at 25C. 149 Table 3. Grafting amount and size of SNPs. Sample Grafting Grafting Click Amount Density Conversiona (mmol/g)a (group/nm2)a (%) Weight Loss (%)a Size (nm)b LTM40 -- -- -- -- 30 LTM40PTr 5.1 0.36 1.5 -- 37c LTM40PTrg4PEG 7.6 0.33 1.4 92 56 LTM40PTrg4PEGOH 7.5 0.34 1.4 94 50 LTM40PTrg4PEGNH2 7.2 0.33 1.4 90 66 LTM40PTrg4PEGCOOH 8.7 0.37 1.5 97 51 a Weight loss, grafting amount, density and click conversion were determined after each modification step. b Size measured by DLS after sonication for 30 min in H2O. 150 Table 4. Grafting data from different sized SNPs terminated with carboxylic acids. Weight Grafting Grafting Click Loss Amount Density Conversiona (%)a (mmol/g)a (group/nm2)a (%) SNTXS -- -- -- -- 20b SNTXSPTr 10.4 0.74 0.8 -- 19c SNTXSPTrg4PEGCOOH 15.9 0.68 0.7 91.9 33c LTM40 -- -- -- -- 30b LTM40PTr 5.1 0.36 1.5 -- 37c LTM40PTrg4PEGCOOH 8.7 0.37 1.5 100 51b SNT20L -- -- -- -- 80b SNT20LPTr 1.9 0.14 1.4 -- 90c SNT20LPTrg4PEGCOOH 2.8 0.12 1.2 84.3 82b SNTZL -- -- -- -- 150b SNTZLPTr 1.6 0.12 2.1 -- 117b SNTZLPTrg4PEGCOOH 2.2 0.10 1.7 79.2 137b Sample a Size (nm)b Weight loss, grafting amount, density and click conversion were determined after each modification step. b Size measured in H2O by DLS after sonication. c Size measured in EtOH. 151 Aqueous media dispersion stability of modified SNPs DLS was used to determine whether the modified SNPs remained dispersed in cell culture media (RPMI containing 2% BCS) as well as their stability over time. Four SNPs with different surface chemistry (-OMe, -OH, -NH2, and -COOH) were measured by DLS, and all were stabile for 2-3 days in the in vitro culture media without aggregation. For example, Fig. 33 showed data for LTM40PTrg4PEG (-OMe) in in vitro media. At 0 h in water, the distribution of the nanoparticles was monomodal with a hydrodynamic size of ~60 nm (black curve). The results from in vitro culture media showed two peaks, a small narrow peak at ~10 nm, and a broad particle population from >50 nm to <500 nm (red curve). After combining the particles and culture media (blue curve), the small peak at ~10 nm, seen in the culture media, shifted slightly to lower diameters (<10 nm), and the combined scattering from the nanoparticle and the media formed a single peak, which broadened toward higher particle sizes. On the high size of the distribution, the scattering was comprised of the sum of the particles and the media. Importantly, the onset for scattering from the particles, the nearconstant slope of the low side of the distribution, and the peak of the distribution did not change as a function of time. We concluded that the pegylated SNPs in in vitro culture media showed no significant aggregation, and the main effect of the media was to broaden the distribution. The other three SNPs showed similar DLS behavior when added to in vitro culture media as presented in Fig. 34-36. 152 Figure 33. Aqueous dispersion stability of LTM40PTrg4PEG SNPs in in vitro culture media. LTM40PTrg4PEG SNPs were sonicated, and dispersed in culture media by vortexing. DLS data (size distribution by intensity) from LTM40PTrg4PEG SNPs alone (100 μg/ml, black curve), culture media alone (red curve), and LTM40PTrg4PEG SNPs in culture media (blue curve) at different time points (indicated on the graphs) were recorded. 153 Figure 34. Aqueous dispersion stability of SNPs with -OH functional group in in vitro culture media. LTM40PTrg4PEGOH SNPs were sonicated, and dispersed in culture media by vortexing. DLS data (size distribution by intensity) from SNPs alone (100 μg/ml, black curve), culture media alone (red curve), and SNPs in culture media (blue curve) at different time points (indicated on the graphs) were recorded. 154 Figure 35. Aqueous dispersion stability of SNPs with –NH2 functional group in in vitro culture media. LTM40PTrg4PEGNH2 SNPs were sonicated, and dispersed in culture media by vortexing. DLS data (size distribution by intensity) from SNPs alone (100 μg/ml, black curve), culture media alone (red curve), and SNPs in culture media (blue curve) at different time points (indicated on the graphs) were recorded. 155 Figure 36. Aqueous dispersion stability of SNPs with -COOH functional group in in vitro culture media. LTM40PTrg4PEGCOOH SNPs were sonicated, and dispersed in culture media by vortexing. DLS data (size distribution by intensity) from SNPs alone (100 μg/ml, black curve), culture media alone (red curve), and SNPs in culture media (blue curve) at different time points (indicated on the graphs) were recorded. 156 OVA257-264 (SIINFEKL)-stimulated cytokine production by CD8+ T cells Using the OT-I model in which SIINFEKL was presented by endogenous APCs to CD8+ T cells, the percent of viable CD8+ T cells that produced IFN (Fig. 37A, upper panel) or IL-2 (Fig. 37A, bottom panel) increased with increasing concentrations of SIINFEKL (S; 10-8 to 10-2 g/ml). There was little background cytokine production in the absence of SIINFEKL stimulation (No S). In addition, the fluorescence intensity of IFN was higher than IL-2 (Fig. 37A). SIINFEKL (10-4 g/ml) produced suboptimal, but measurable, stimulation of the CD8+ T cell response and was used in subsequent experiments. Cytokine production was also measured using the model in which DC2.4 cells were used as APCs to present SIINFEKL to OT-I CD8+ T cells. DC maturation can be induced by various stimuli, including lipopolysaccharide (LPS) [278]. However, in this model under current conditions, LPS pretreatment of DC2.4 cells did not further increase IFN (Fig. 37B, left panel) or IL-2 (Fig. 37B, right panel) production by SIINFEKL-activated viable CD8+ T cells. Thus, LPS pretreatment was not utilized in subsequent experiments. Production of IFN (Fig. 37C, upper panel) or IL-2 (Fig. 37C, bottom panel) by viable CD8+ T cells was increased with increasing concentrations of SIINFEKL (0.01 – 1 g/ml) loaded onto MHC I of DC2.4 cells. Also, there was more IFN production than IL-2, which was consistent with the trend observed in Fig. 37A. These studies showed that the concentration of SIINFEKL for MHC I loading on DC2.4, in order to achieve suboptimal activation of CD8+ T cells, was 0.1 g/ml. 157 Figure 37. SIINFEKL concentration-dependent activation of CD8+ T cell responses. (A) OT-I splenocytes (CD8+ T cells, 8 x 105 cells/ well) were incubated with various concentrations of SIINFEKL (S) peptide (10-8 – 10-2 g/ml) for 2 days. Brefeldin A was added during the last 6 h of incubation. Splenocytes were stained for viability, surface CD8 and intracellular IFN or IL-2 expression. The percent of IFN+ or IL-2+ cells within CD8+ T cells is presented in the quadrants of dot plots, with X axis and Y axis representing the fluorescence intensity for CD8 and IFN or IL-2, respectively. (B) DC2.4 cells were pretreated with LPS (1 g/ml) overnight, loaded with SIINFEKL peptide (1 g/ml) for 3 – 4 h, irradiated, and plated at 2 x 105 cells/well with 1 x 106 cells/well OT-I splenocytes for 4 days. Splenocytes were restimulated with SIINFEKL (10 g/ml) in the presence of brefeldin A for 6 h. Splenocytes were stained for viability, surface CD8 and intracellular IFN or IL-2 expression. The percent of IFN+ or IL-2+ cells within CD8+ T cells is presented in histograms, with X axis representing fluorescence intensity for IFN or IL-2 and Y axis representing percent of CD8+ T cells. (C) DC2.4 cells were not pretreated with LPS and incubated with various concentrations of SIINFEKL (10-2 – 1 g/ml), co-cultured with OT-I splenocytes as described in (B). The dot plots are presented as described in (A). (A-C) Three replicates were concatenated for each treatment in the flow cytometric analysis. Data are representative of at least two separate experiments. 158 Figure 37 (cont’d) 159 Modified SNPs enhanced SIINFEKL-activated CD8+ T cell responses LTM40PTrg4PEG SNPs alone showed minimal effects on IFN and IL-2 production by viable CD8+ T cells; whereas LTM40PTrg4PEG enhanced the production of both cytokines by CD8+ T cells that were suboptimally activated using SIINFEKL peptide directly (Fig. 38A and 38B). The effect was deemed statistically significant at multiple LTM40PTrg4PEG concentrations, including 0.01, 1, and 10 g/ml for IFN, and all concentrations for IL-2. In the co-culture OT-I model in which irradiated DC2.4 cells were used as APCs, LTM40PTrg4PEG SNPs showed a similar trend of effect on CD8+ T cell responses. LTM40PTrg4PEG had no effect in the absence of SIINFEKL activation; whereas in the presence of a suboptimal concentration of SIINFEKL loaded onto DC2.4 cells, LTM40PTrg4PEG at 10 g/ml further enhanced IFN (Fig. 38C) and IL-2 (Fig. 38D) production by viable CD8+ T cells, with the latter being significantly different from the no particle treatment control. This suggests that SNP-mediated enhancement is likely to act directly on CD8+ T cell responses. Different functional groups on SNPs influenced the immune enhancing effect to different levels The effect of engineered SNPs on T cell responses was compared between particles with different surface LTM40PTrg4PEGOH functional (-OH), LTM40PTrg4PEGNH2 (-NH2). groups, including LTM40PTrg4PEG LTM40PTrg4PEGCOOH (-COOH), (-OMe), and Similar to the above observations, in the absence of SIINFEKL stimulation, SNPs produced minimal effects on the cytokine response, especially for IL-2 (Fig. 38E and 38F). When OT-I CD8+ T cells were suboptimally activated with 10-4 160 g/ml of SIINFEKL, all tested SNPs showed significant enhancement of IFN (Fig. 38E) and IL-2 (Fig. 38F) production by CD8+ T cells. In particular, -COOH and -NH2 particles exhibited concentration-dependent enhancement on cytokine production. Absence of immune enhancement by SNPs on OVA323-339-induced CD4+ T cell responses In the OT-II model in which OVA323-339 is presented by MHC II molecules on endogenous APCs to OT-II CD4+ T cells, production of IFN and IL-2 was minimal in the absence of OVA323-339 peptide activation and increased with increasing concentrations of OVA323-339 (0.01 – 0.1 g/ml) (Fig. 39A). In contrast to the OT-I model, there was more IL-2 production than IFN in the OT-II model. The OVA323-339 concentration for suboptimal CD4+ T cell activation was determined to be 0.05 g/ml. Next, effects of SNPs with different functional groups on CD4+ T cell responses were characterized. In contrast to their enhancing effects on CD8+ T cell responses, none of the particles showed enhancing activity on IFN and IL-2 production by CD4+ T cells at any of the tested concentrations (Fig. 39B and 39C). Enhancing effect of SNPs on CD8+ T cell responses is size-dependent Next, we investigated the role of particle size on the enhancement of CD8+ T cell responses. Particles with the -COOH functional group at different sizes, including SNTXSPTrg4PEGCOOH (33 nm), LTM40PTrg4PEGCOOH (51 nm), SNT20LPTrg4PEGCOOH (82 nm), and SNTZLPTrg4PEGCOOH (137 nm), were compared. Interestingly, only the 51 nm -COOH particles at 10 µg/ml produced a significantly greater enhancement of IFN and IL-2 production by CD8+ T cells activated with a suboptimal 161 concentration of SIINFEKL. SNPs either smaller or larger in size than 51 nm particles lacked immune enhancing activity as assessed by IL-2 and IFN production (Fig. 40A and 40B). 162 Figure 38. Enhancement by modified LTM40PTrg4PEG on CD8+ T cell responses induced by a suboptimal concentration of SIINFEKL. (A-B) LTM40PTrg4PEG SNPs were sonicated and added to OT-I splenocytes (CD8+ T cells, 8 x 105 cells/ well) at various concentrations (10-2 – 10 g/ml) prior to SIINFEKL (S) stimulation (10-4 g/ml) for 2 days. (C-D) SIINFEKL (S, 0.1 g/ml)-loaded DC2.4 cells (2 x 106 cells/well) were irradiated and plated at 2 x 105 cells/well. OT-I splenocytes (1 x 106 cells/well) were treated with LTM40PTrg4PEG SNPs (10 g/ml) and then co-cultured with SIINFEKL-loaded DC2.4 cells for 4 days. Splenocytes were restimulated with SIINFEKL (10 g/ml) in the presence of brefeldin A. (E-F) SNPs with different functional groups, including LTM40PTrg4PEG (OMe), LTM40PTrg4PEGOH (-OH), LTM40PTrg4PEGCOOH (-COOH), and LTM40PTrg4PEGNH2 (-NH2), were sonicated and then added to OT-I splenocytes (8 x 105 cells/ well) at various concentrations (10-1 – 10 g/ml) prior to SIINFEKL (S) stimulation (104 g/ml) for 2 days. (A-F) Brefeldin A was added during the last 6 h of incubation. Splenocytes were stained for viability, surface CD8 and intracellular IFN or IL-2 expression. The percent of IFN+ or IL-2+ cells within CD8+ T cells is presented as the mean %  S. E. of triplicate cultures. * p < 0.05 as compared to respective no SNP treatment control. Data are representative of at least two separate experiments. 163 Figure 38 (cont’d) 164 Figure 39. Absence of enhancement by SNPs with different functional groups on CD4+ T cell responses under suboptimal OVA323-339 stimulation. (A) OT-II splenocytes (CD4+ T cells, 8 x 105 cells/well) were incubated with various concentrations of OVA323-339 peptide (10-2 – 0.1 g/ml) for 2 days. (B-C) SNPs with different functional groups, including LTM40PTrg4PEG (-OMe), LTM40PTrg4PEGOH (-OH), LTM40PTrg4PEGCOOH (COOH), and LTM40PTrg4PEGNH2 (-NH2), were sonicated and added to OT-II splenocytes at various concentrations (10-1 – 10 g/ml) prior to OVA323-339 peptide stimulation (0.05 g/ml) for 2 days. (A-C) Brefeldin A was added during the last 6 h of incubation. Splenocytes were stained for viability, surface CD4 and intracellular IFN or IL-2 expression. The percent of IFN+ or IL-2+ cells within CD4+ T cells is presented in the quadrants in dot plots (A), with X axis and Y axis representing fluorescence intensity for CD4 and IFN or IL2, respectively. Three replicates were concatenated for each treatment in the flow cytometry analysis (A). The percent of IFN+ (B) or IL-2+ (C) cells within CD4+ T cells is presented as the mean %  S. E. of triplicate cultures. * p < 0.05 as compared to respective no SNP treatment control. Data are representative of at least two separate experiments. 165 Figure 39 (cont’d) 166 Figure 40. Effect of -COOH SNPs at different sizes on CD8+ T cell responses under suboptimal induction. (A-B) SNPs with -COOH functional group at different sizes, including SNTXSPTrg4PEGCOOH (33 nm), LTM40PTrg4PEGCOOH (51 nm), SNT20LPTrg4PEGCOOH (82 nm), and SNTZLPTrg4PEGCOOH (137 nm) were sonicated and then added to OT-I splenocytes (8 x 105 cells/ well) at various concentrations (10-1 – 10 g/ml) prior to SIINFEKL (S) stimulation (10-4 g/ml) for 2 days. Brefeldin A was added during the last 6 h of incubation. Splenocytes were stained for viability, surface CD8 and intracellular IFN or IL-2 expression. The percent of IFN+ (A) or IL-2+ (B) cells within CD8+ T cells is presented as the mean %  S. E. of triplicate cultures. * p < 0.05 as compared to respective no SNP treatment control. Data are representative of at least two separate experiments. 167 DISCUSSION As the commercial use of engineered nanoparticles has grown, their potential for causing adverse health issues has also received increasing attention. To date, most investigations focusing on the potential toxicity of engineered nanoparticles have been conducted in laboratory animals. Some of these studies have demonstrated that certain engineered nanoparticles possess adjuvant-like properties, especially on pulmonary inflammation and lung disease. Although inhalation is one of the principal routes of particle exposure [262-265, 279], little is still known concerning the effect of engineered nanoparticles on immune responses in airways or on the immune system in general. Here, we utilized unique in vitro mouse models to stimulate antigen-specific CD8+ and CD4+ T cell responses in order to investigate the effects of engineered nanoparticles on the functions of certain leukocyte populations. It is worth mentioning that two related but distinct OT-I models possessing either endogenous splenic APCs containing multiple cellular populations or the cell line, DC2.4, for antigen presentation of SIINFEKL showed similar trends, as evidenced by enhancement of antigen-specific CD8+ T cell responses by engineered nanoparticles. As reviewed by Banchereau and co-workers, DCs are highly efficient antigen presenting cells [278]. DC2.4 cells were irradiated before co-culturing with splenocytes to arrest their proliferation, so they would not deplete the media of nutrients and/or outgrow naïve T cells; however, their ability to activate naïve T cells in the context of antigen-MHC complexes is not affected by irradiation [280, 281]. By using DC2.4 as the only APC population in the second OT-I model, OVA peptides were directly loaded onto DC2.4, therefore the effects of nanoparticles on antigen processing via APCs as a potential confounder were eliminated, allowing the direct assessment of their potential effects on T cell responses. With two OT-I 168 models, we demonstrated that SNPs produced their immune enhancing effects most likely by affecting T cell function directly. The aqueous-dispersed engineered nanoparticles used in the present study underwent chemical modification by varying the surface functional group moieties and overall particle sizes. An important as well as rare physico-chemical property of the engineered nanoparticles employed in this study was that these intermediate-sized particles are resistant to undergoing aggregation even in the culture media, which allowed them to be used in in vitro assay systems to investigate their influence on leukocyte function in their true nanoscale context. Although the size distribution peak for the -NH2 SNPs in culture media appeared to shift to the right, around 100 nm, distinct from the peaks for other particles, which appeared close to 50 nm, we believe they are unlikely to agglomerate in the culture media. Because if modest changes in the size of -NH2 SNPs led to significant shifts between no agglomeration and agglomeration, one would expect to observe significant peaks at markedly larger sizes than 100 nm (i.e., which would be representative of association between 2 particles). Alternatively, the change in size could reflect an interaction between SNPs and serum proteins as previously suggested [282, 283]. However, if the nanoparticles interacted with protein components in the serum, the particle size would increase dramatically due to formation of agglomerates and keep increasing over time as reported in the literature [284], which were not detected by DLS here. In addition, a relatively low serum concentration in the RPMI media was used in our studies, which would not introduce a big amount of serum proteins, therefore keeping the zeta potentials relatively stable as compared to no serum condition. It is also worth mentioning that the average hydrodynamic diameters of the bare SNPs, measured by DLS in water, were larger than the sizes claimed by manufacturers, which 169 is likely due to the differences in methodologies for particle size measurement, such as using transmission electron microscopy (TEM) or scanning electron microscopy (SEM) images. The zeta potential of all coated SNPs displayed higher values than the bare SNP, indicating that the silanol groups were covered by PEG layers. NH2-modified SNPs showed the highest zeta potential, which indicates the zeta potential of bare SNP was compensated by both the PEG layer and the NH2 (+) surface charge. COOH-modified SNPs with the lowest zeta potential, which was higher than bare SNP, is due to the PEG layer and carboxylate group in the buffer. PEG and OH-modified SNPs showed medium zeta potential, which is due to the neutral surface function and PEG layers. It has been reported recently that surface modified SNPs have a negative zeta potential for neutral surface layers and +0.6mV only for amine modified SNPs in PBS buffer [285, 286]. Our negative zeta potential is most likely due, at least in part, to the remaining negatively charged silanol groups. Since surface chemistry of nanoparticles directly influences cellular uptake and thereby their activity as well as interaction with cellular compartments in the cells [273], three surface functional groups, including -COOH, -NH2, and -OH, were engineered to modify the pegylated SNPs individually. Among these three functional groups, -COOH SNPs were chosen for further characterization, although both -COOH and -NH2 SNPs showed concentration-dependent enhancement of the CD8+ T cell response when compared with -OH SNPs in our assays (Fig. 38E and 38F). It has been suggested that -COOH particles help to facilitate cellular uptake and transport into intracellular compartments with less aggregation inside the cells [287]. Also, since -COOH, -NH2, and -OH particles have different surface charges at physiological pH, cationic charge of the cell membrane due to interaction with NH2 particles may cause excessive proton transport and accumulate inside the cells, therefore 170 generating reactive oxygen species (ROS), which may injury mitochondria and eventually disrupt the cellular function [288, 289]. In addition, no agglomeration in culture media was observed with -COOH particles. Therefore, -COOH particles were selected for the subsequent size-dependent studies. In this assay system, cytokine production by CD8+, but not CD4+, T cells was significantly increased by modified SNPs, suggesting that the particles selectively target CD8+ T cell responses. It has been suggested that certain physico-chemical properties of modified SNPs may induce upregulation of MHC I expression on DCs by increasing phosphorylation of mitogen-activated protein kinases (MAPK), such as p38 kinase, c-Jun Nterminal kinases (JNK), and extracellular signal-regulated kinases (ERK), which are involved in DC activation signaling pathways, and thereby indirectly enhance CD8+ T cell responses [290]. However, because DC2.4 cells were irradiated for experiments employing the coculture model, MHC I expression on these cells should not be further regulated by SNPs, hence suggesting that SNPs did not enhance CD8+ T cell responses via regulating APCs. Therefore, mechanisms underlining the observed enhancement on CD8+ T cell responses remain to be elucidated. It is possible that engineered nanoparticles in our models specifically modulate CD8+ T cell receptor signaling or directly interact with surface receptors on CD8+ T cells, e.g. low-density lipoprotein receptor [291], and thereby increase particle cellular uptake to enhance CD8+ T cell responses, but further investigations will be needed. In addition, there was no direct cytotoxicity from SNPs on the T cells, because all the flow cytometry analyses were performed using viable cells. We have demonstrated that only particles at a size of 51 nm significantly enhanced CD8+ T cell responses, suggesting that the nanoparticle-mediated enhancement of CD8+ T cell 171 responses is size-dependent. Similar nanoparticle size-dependent effect has been demonstrated previously, indicating that within the size range of 2 - 100 nm, 50 nm gold nanoparticles showed the greatest effect on membrane receptor internalization, which in turn affected the downstream cellular proliferation and survival responses [292]. As suggested by others using gold naoparticles in cells or theoretical models, 50 nm particles might fall into the critical size range for receptor-mediated internalization. Smaller particles might dissociate from the receptors on the membrane before being endocytosed due to their small avidity; whereas large particles might cause decreased membrane wrapping efficiency, which would directly impair the nanoparticle internalization process [292-294]. Collectively, we demonstrated immune enhancing properties by engineered SNPs on DCinduced antigen-specific CD8+ T cell responses using an OT-I model in vitro, suggesting the potential adjuvant-like effects of these engineered SNPs on immune function at the cellular level. We also show that by modifying the chemical and physical properties of the particles, such as the surface chemistry and size, the immunomodulatory properties of these particles can be differentially affected. Further studies at the organismal, cellular and subcellular level with special emphasis on engineered nanoparticle-membrane interactions are needed to provide more in-depth understanding of the mechanisms responsible for immune modulation by SNPs. 172 CONCLUSIONS In summary, our results demonstrated that the engineered SNPs were successfully modified with different functional group moieties on the surface and in different sizes, which allowed them to be used to characterize the effect of diverse physical and chemical properties of SNPs on T cell function. In addition, modified SNPs, except for -NH2 SNP, which might interact with proteins in the media, did not agglomerate in in vitro culture condition, which was critical for them to maintain their aqueous distribution at the nanoscale and was an essential prerequisite for studying their immunological effects (Fig. 33-36). Conditions for suboptimal stimulation of T cell responses were determined using OT-I and OT-II models (Fig. 37 and 39A) in order to measure the immune enhancing effects of the SNPs. Modified SNPs (up to 10 g/ml) enhanced OVA-specific IFN and IL-2 production by CD8+, but not CD4+, T cells, suggesting a cell-type selective effect by engineered SNPs on immune function, in particular on antigen-specific CD8+ T cell responses. Enhancement of CD8+ T responses by SNPs was differentially affected by the surface functional groups placed on the SNPs, which is consistent with findings of others [273, 287]. SNPs with -COOH moieties were chosen for subsequent studies, since they exhibited concentration-dependent enhancement of CD8+ T responses in the current assay system as well as advantages over SNPs with other functional groups as suggested in the literature [287-289], such as enhanced cellular uptake, less agglomeration, and limited cellular toxicity. Lastly, 51 nm sized SNPs showed greater enhancement of CD8+ T cell responses than the other sized SNPs, which might be dictated by the nanoparticle size-dependent internalization processes as suggested by others [292-294]. Our findings provide strong evidence that nanoparticles have potential adjuvant-like properties by enhancing immune function, which will cause adverse effects on allergic airway 173 disease in humans. In addition, our studies have significant implications in understanding the role of particle size and other surface chemistry on regulating the biological responses and providing guidelines for the design of nanoproducts, with great potential for the development of novel biomedical applications in diagnostics and therapeutics. 174 BIBLIOGRAPHY 175 BIBLIOGRAPHY 1. Gilbert, P.B., McKeague, I.W., Eisen, G., Mullins, C., Gueye, N.A., Mboup, S., Kanki, P.J. (2003) Comparison of HIV-1 and HIV-2 infectivity from a prospective cohort study in Senegal. Stat Med 22, 573-93. 2. 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