EFFECTS OF METHYLMERCURY IN THE DOPAMINE SYNTHESIZING PHEOCHROMOCYTOMA PC12 CELL LINE By Chelsea Tate Tiernan A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Neuroscience – Doctor of Philosophy 2013 ABSTRACT EFFECTS OF METHYLMERCURY IN THE DOPAMINE SYNTHESIZING PHEOCHROMOCYTOMA PC12 CELL LINE By Chelsea Tate Tiernan Methylmercury (MeHg) is a potent bioaccumulative neurotoxicant that targets discrete neuronal populations, including the nigrostriatal dopamine (NSDA) neuronal system. Epidemiological evidence has implicated chronic exposure to MeHg as an environmental risk factor for Parkinson disease (PD), and experimental analyses using in vivo animal and in vitro cell culture models have demonstrated that acute exposure to MeHg alters DA homeostasis, including release, reuptake, and metabolism. The purpose of the studies described in this dissertation is to characterize MeHg-induced changes in DA synthesis, release, reuptake, and metabolism, and to investigate mechanisms by which MeHg exerts its neurotoxic effects in the pheochromotocytoma (PC12) cell line. MeHg causes a concentration- and time-dependent increase in DA release. Higher concentrations (5μM) are associated with an increased incidence of cell death at later time points (60-120 min). However 2μM MeHg induces DA release by 60 min without altering cell viability. Thus this concentration and time-point was selected to examine other indices of DA homeostasis. MeHg-induced DA release is abolished by inhibition of vesicular exocytosis with reserpine, but not inhibition of membrane transport with desipramine. A role for synthesis in MeHg-induced DA release was indicated by an increase in the concentration of intracellular DA and the rate of decline of intracellular DA following acute treatment with the tyrosine hydroxylase (TH) inhibitor α-methyltyrosine (AMT). MeHg stimulates DA synthesis indicated by an increase in DOPA accumulation following treatment with the DOPA decarboxylase inhibitor NSD-1015. This is supported by the observation that MeHg elevates phosphorylation of TH at serine reside 40, without altering the total amount of TH. Moreover, MeHg-induced DA release is dependent upon DA synthesis because pre-treatment with AMT abolishes MeHg-induced DA release. MeHg induces aberrant DA metabolism. Intracellular concentrations of DOPAC are decreased, while intracellular concentrations of the intermediate metabolites DOPAL and DOPET are increased. This metabolomic profile suggests that MeHg inhibits the oxidation of DOPAL to DOPAC and thus inhibits aldehyde dehydrogenase (ALDH). MeHg does not directly impair ALDH activity. Instead, inhibition may be indirect because MeHg inhibits mitochondrial respiration and ATP synthesis, and decreases availability of the ALDH cofactor nicotinamide adenine dinucleotide (NAD). To assess the roles of extracellular and intracellular calcium (Ca2+) in altered DA release and metabolism, undifferentiated PC12 cells were exposed to MeHg in both the absence and presence of extracellular and/or intracellular Ca2+. Removal of intracellular but not extracellular Ca2+ attenuates MeHg-induced DA release. MeHg-impaired DA metabolism is not influenced by chelation of either Ca2+ source. The present findings are consistent with the follow conclusions: 1) MeHg-induced DA release is dependent upon DA synthesis and vesicular exocytosis, 2) MeHg impairs DA metabolism by indirectly inhibiting ALDH, 3) release of Ca2+ from intracellular stores triggers MeHg-induced DA release, and 4) aberrant DA metabolism is not affected by changes in intracellular Ca2+ homeostasis. ACKNOWLEDGEMENTS Completing this dissertation has been a long, arduous journey, and I would not have succeeded without the love, support, and guidance of many people. First and foremost, I express my deepest gratitude to my advisors Drs. Keith Lookingland and John Goudreau. Keith, thank you for always being optimistic and supportive. Thank you for honing my writing and critical thinking skills, and thank you for fostering my passion to teach. Your dedication to your students is inspirational, and your encouragement over the years has been essential to my success. John, thank you for all of your ideas. Your expertise and advice with writing, networking, brainstorming, and presentations have been invaluable. I also owe an immense debt of gratitude to Dr. William Atchison, committee member and coinvestigator. Thank you for providing me with an opportunity to continue my education. Thank you for inspiring me to pursue science outreach and education. And thank you for the many sunny trips to Puerto Rico. Your continued assistance and guidance is inestimable. To my committee members Drs. Kyle Miller and Peter Cobbett, thank for the many meetings, conversations, and assistance. Your presence has enriched my graduate school experience. A number of faculty members at MSU and my previous academic institutions, Western State Colorado University and Roaring Fork High School, have also provided crucial support for my education. I thank Dr. Lynwood Clemens for allowing me to assist in his Integrative Neurobiology course, and to use that experience as part of my Certification in College Teaching. I thank Dr. Charles Patrick Stark for my first research experience and for encouraging me to attend graduate school. May you find peace in your next adventure. I iv thank Mrs. Wendy Boland and Mrs. Laura French who first inspired me to become a lifelong learner, recognized my true talents and passions in science, and quietly challenged me to pursue my dreams. I sincerely thank my fellow GoudLooking laboratory members, past and present. The collaborative, fun, and sometimes shocking environment has been a true pleasure. Special thanks to Dr. Bahareh Behrouz, Dr. Sam Pappas, Dr. Matt Benskey, (almost Dr.) Tyrell Simkins, and Hae-young Hwang. I thank my undergraduate technicians, Ethan Edwin, María Bernard, and Ninotchska Del Valle. Your dedication and hard work has helped make this dissertation a success. I wish you luck in the coming years as you all begin your graduate studies. I thank my many friends who have been so loving and supportive over the years. However there are two people who I would like to thank especially. Chelsea Hutch, I know there is absolutely no way on Earth I would have made it through this gauntlet without you by my side every step of the way. Thank you for the late night runs to Tasty Twist and the marathons of Dexter. Matt Benskey, it is difficult for me to find words to express my gratitude for your love and encouragement. Without your intellectual and emotional support, I would be lost. Thank you for instilling confidence in me. Thank you for not accepting anything less than the very best from me. And most importantly, thank you for helping me learn how to relax and enjoy a beautiful sunset. Last, but certainly not least, I would like to thank my amazing, extended, and incredibly understanding family. To my brother Zeke Tiernan, thank you for reminding me what true endurance is. I can’t wait to cheer you on during your next 100 miles! To my brother Alex Tiernan, thank you for reminding me that life is for living. Your love and v support has always been one of the greatest blessings in my life. And to my parents Edward Tiernan and Carrie Katsis thank you. Dad, I don’t think a father has ever been so proud of his daughter. Your constant love is a treasure I will carry with me always. Thank you for encouraging me to complete this educational endeavor and thank you for taking an active interest in my research. I love you forever, I love you for always, and long as I’m living, your baby I’ll be. Mom, thank you for being patient, even when your life was chaotic, and thank you for being understanding, even when my problems were ridiculous. I am eternally grateful for the love and friendship that we share. Thank for being the eccentric, beautiful, and passionate woman you are. To you both, I am so blessed. I can only hope to be the kinds of parents you are some day. Thank you! vi TABLE OF CONTENTS LIST OF TABLES……………………………………………………………………………………………………………xii LIST OF FIGURES………………………………………………………………………………………………………….xiii KEY TO ABBREVIATIONS…………………….………………………………………………………………………..xvi Chapter 1. General Introduction…………………………………………………………………………………..1 1.1. Statement of Purpose……………………………………………………………………………………..1 1.2. Methylmercury…………..………………………………………………………………………………….3 1.3. Neurotoxicity of MeHg……………………………………………………………………………………6 1.4. Molecular mechanisms of MeHg-mediated neurotoxicity ………………………………..7 1.5. Parkinson’s disease……………………………………………………………………………………...10 1.6. Epidemiological evidence of MeHg as an environmental risk factor for PD……..11 1.7. Dopamine……………………………………………………………………………………………………13 1.7.1. DA synthesis…………………………………………………………………………………..15 1.7.2. DA storage……………………………………………………………………………………..16 1.7.3. DA release and activation of DA receptors……………………………………….17 1.7.4. DA reuptake…………………………………………………………………………………...17 1.7.5. DA metabolism……………………………………………………………………………….18 1.8. Effects of MeHg on DA neuronal homeostasis………………………………………………..20 1.8.1. MeHg targeting of DA synthesis……………………………………………………….20 1.8.2. MeHg targeting of DA release…………………………………………………………..21 1.8.3. MeHg targeting of DA reuptake……………………………………………………….22 1.8.4. MeHg targeting of DA metabolism…………………………………………………...23 1.9. Pheochromocytoma 12 (PC12) cell line………………………………………………………...24 1.9.1. PC12 cell morphology……………………………………………………………………..25 1.9.2. PC12 cell neurochemistry……………………………………………………………….26 1.9.3. PC12 cell VGCC subtype expression…………………………………………………29 1.9.4. Differences between undifferentiated PC12 cells and DA neurons…….30 1.9.5. Effects of MeHg in PC12 cells…………………………………………………………..31 1.10. Summary…………………………………………………………………………………………………...32 1.11. Thesis objective…………………………………………………………………………………………33 Chapter 2. General Materials and Methods.……………………………………………………………….35 2.1. Culture of PC12 cells…………….………………………………………………………………………35 2.2. Chemicals and solutions……………………………………………………………………………….37 2.3. Neurochemical analyses……………………………………………………………………………….41 2.3.1. Calculation of rate constant…..………………………………………………………...41 2.3.2. Synthesis of DOPAL standard…………………………………………………………..42 2.4. Cell viability…………………………………………………………………………………………………46 2.5. Western blot analysis of TH………………………………………………………………………….48 2.6. Mitochondrial bioenergetics…………………………………………………………………………49 vii 2.6.1. XF24 assay medium………………………………………………………………………..49 2.6.2. Test compounds……………………………………………………………………………..49 2.6.3. Extracellular flux (XF) analysis………………………………………………………..51 2.6.4. XF bioenergetics assay…………………………………………………………………....54 2.6.5. XF24 drug exposure………………………………………………………………………..55 2.6.6. XF24 optimization of experimental conditions…………………………………55 2.7. ALDH enzyme kinetics assay………………………………………………………………………...61 2.8. NADH/NAD quantification……………………………………………………………………………64 2.9. Measurement of fura-2 fluorescent change……………………………………………………66 2.10. Statistical analyses……………………………………………………………………………………..67 Chapter 3. The role of de novo catecholamine synthesis in mediating MeHg-induced vesicular DA release…………………………………………………………………………………………………...68 3.1. Introduction………………………………………………………………………………………………...68 3.2. Methods………………………………………………………………………………………………………70 3.2.1. Chemicals……………………………………………………………………………………….70 3.2.2. Chemical treatment...………………...……………………………………………………70 3.2.3. Neurochemistry.……………………………………………………………………………..70 3.2.4. Western blot analysis……………………………………………………………………...71 3.2.5. Cell viability……………………………………………………………………………………71 3.2.6. Statistical analysis…………………………………………………………………………..72 3.3. Results………………………………………………………………………………………………………...72 3.3.1. MeHg increases spontaneous DA release in a concentration- and timedependent manner…………………………………………………………………………………..72 3.3.2. Spontaneous MeHg-mediated DA release requires a functional vesicular, but not membrane, transporter………………………………………………...78 3.3.3. MeHg-mediated DA release is associated with increased intracellular DA concentrations and utilization of DA stores…………………………………………80 3.3.4. MeHg-mediated DA release is associated with de novo DA synthesis and increased TH enzymatic activity………………………………………………………...83 3.4. Discussion…………………………………………………………………………………………………...87 3.4.1. MeHg-induced DA release……………………………………………………………….87 3.4.2. Vesicular transport, but not membrane transport contributes to MeHginduced DA release………………………………………………………………………………….88 3.4.3. Evidence for a role of de novo DA synthesis in MeHg-induced DA release…………………………………………………………………………………………………….90 Chapter 4. The roles of mitochondria and associated catabolic enzyme function in MeHg-impaired DA metabolism…………………………………………………………………………………92 4.1. Introduction………………………………………………………………………………………………...92 4.2. Methods………………………………………………………………………………………………………97 4.2.1. Chemicals……………………………………………………………………………………….97 4.2.2. Chemical treatment …………………………...…………………………………………..97 4.2.3. Neurochemistry……………………………………………………………………………...97 4.2.4. ALDH2 enzyme kinetics assay…………………………………………………………98 4.2.5. Mitochondrial bioenergetics……………………………………………………………98 viii 4.2.6. NADH/NAD quantification………………………………………………………………99 4.2.7 Statistical analysis…………………………………………………………………………...99 4.3. Results………………………………………………………………………………………………………100 4.3.1. MeHg impairs DA metabolism, shunting DA along an alternative metabolic pathway………………………………………………………………………………...100 4.3.2. Selective inhibition of ALDH with daidzin impairs DA metabolism in a manner similar to MeHg………………………………………………………………………...105 4.3.3. Daidzin and rotenone impair DA metabolism in a manner that resembles MeHg……………………………………………………………………………………108 4.3.4. ALDH activity is not directly altered by MeHg………………………………..110 4.3.5. MeHg impairs mitochondrial function and decreases availability of the ALDH cofactor NAD……………………………………………………………………………….113 4.4. Discussion…………………………………………………………………………………………………119 4.4.1. The DA metabolomic profile induced by MeHg………………………………119 4.4.2. Comparative effects of MeHg, daidzin, and rotenone on DA metabolism…………………………………………………………………………………………...121 4.4.3. Evidence for a role of mitochondrial dysfunction rather than direct ALDH inhibition in MeHg-impaired DA metabolism……………………………..….122 4.4.4. Consequences of decreased NAD(H) following MeHg exposure………124 4.4.5. Potential biological effects of MeHg-impaired DA metabolism………...126 4.4.6. Summary……………………………………………………………………………………...127 Chapter 5. The role of extracellular Ca2+ in MeHg-induced alterations in DA release and metabolism………………………………………………………………..……………………………………....129 5.1. Introduction………………………………………………………………………………………………129 5.1.1. Neurotransmitter release and VGCCs…………………………………………….129 5.1.2. Mitochondrial function and DA metabolism…………………………………..131 5.2. Methods…………………………………………………………………………………………………….133 5.2.1. Chemicals……………………………………………………………………………………..133 5.2.2. Chemical treatment……...……………………………………………………………….134 5.2.3. Neurochemistry.…………………………………………………………………………...134 5.2.4. Mitochondrial bioenergetics………………………………………………………….134 5.2.5. Statistical analysis………………………………………………………………………...135 5.3. Results………………………………………………………………………………………………………135 5.3.1. Spontaneous MeHg-mediated DA release is partially dependent upon the presence of extracellular Ca2+, but not Ca2+ influx through VGCCs…….135 5.3.2. Extracellular Ca2+ stimulates mitochondrial bioenergetics, but does not contribute to MeHg-induced mitochondrial dysfunction or aberrant DA metabolism…………………………………………………………………………………………...140 5.4. Discussion…………………………………………………………………………………………………146 5.4.1. Role of extracellular Ca2+ in spontaneous and MeHg-mediated DA release…………………………………………………………………………………………………..147 5.4.2. Role of VGCCs in spontaneous and MeHg-mediated DA release……….148 ix 5.4.3. Role of extracellular Ca2+ in mitochondrial bioenergetics and targeting by MeHg………………………………………………………………………………………………..149 5.4.4. Role of extracellular Ca2+ in basal and MeHg-impaired DA metabolism…………………………………………………………………………………………...152 5.4.5. Summary……………………………………………………………………………………...143 Chapter 6. The role of intracellular Ca2+ in MeHg-induced alterations in DA release and metabolism………………………………………………………………………………………………………..154 6.1. Introduction………………………………………………………………………………………………154 6.1.1. Regulation of intracellular Ca2+…………………………………………………….154 6.1.2. Effects of MeHg on intracellular Ca2+ homeostasis…………………………156 6.1.3. Effects of MeHg on DA release and metabolism………………………………157 6.2. Methods…………………………………………………………………………………………………….158 6.2.1. Chemicals.…………………………………………………………………………………….158 6.2.2. Treatment for neurochemistry and mitochondrial bioenergetics……158 6.2.3. Neurochemistry.…………………………………………………………………………...159 6.2.4. Mitochondrial bioenergetics………………………………………………………….159 6.2.5. Measurement of fura-2 fluorescent change…………………………………….160 6.2.6. Statistical analysis………………………………………………………………………...160 6.3. Results………………………………………………………………………………………………..……..160 6.3.1. MeHg-induced DA release is dependent upon the presence of intracellular Ca2+………………………………………………………………………………......160 6.3.2. Intracellular Ca2+ is essential for the maintenance of mitochondrial bioenergetics…………………………………………………………………………………………166 6.3.3. Aberrant DA metabolism induced by MeHg is not dependent upon intracellular Ca2+……………………………………………………………………………….….170 6.4. Discussion………………………………………………………………………………………………....172 6.4.1. Role of intracellular Ca2+ in spontaneous and MeHg-mediated DA release…………………………………………………………………………………………………..172 6.4.2. Role of intracellular Ca2+ in mitochondrial bioenergetics……………….175 6.4.3. Role of intracellular Ca2+ in basal and MeHg-impaired DA metabolism…………………………………………………………………………………………...176 6.4.4. Summary……………………………………………………………………………………...178 Chapter 7. General Discussion and Concluding Remarks………………………………………...179 7.1. Role of Ca2+ in MeHg-induced activation of DA synthesis…………………………….179 7.2. Role of Ca2+ in MeHg-induced vesicular exocytosis……………………………………..181 7.3. Collaborative relationship between extracellular and intracellular Ca2+ pools………………………………………………………………………………………………………………..182 7.4. Aberrant DA metabolism induced by MeHg…………………………………………………185 7.5. Pathways by which MeHg targets DA neurons……………………………………………..188 x 7.6. Factors influencing sensitivity of DA-synthesizing cells to MeHg………………….191 7.7. Biological and clinical relevance to PD………………………………………………………..193 7.8. Concluding Remarks…………………………………………………………………………………..197 REFERENCES……………………………………………………………………………………………………………...198 xi LIST OF TABLES Table 1.1. Catecholaminergic and cholinergic neurotransmitter content and enzyme activity in undifferentiated and differentiated PC12 cells ………………………………………..……..28 Table 2.1. Summary of drug actions……………………………………………………………………………...40 Table 2.2. Dilutions of test compounds for XF24 measurements of mitochondrial bioenergetics...………...……………………………………………………………………………………………………50 xii LIST OF FIGURES Figure 1.1. Global mercury (Hg) cycle…………………………………………………………………………….5 Figure 1.2. The molecular structure of dopamine (3-hydroxytyramine).………………………...14 Figure 1.3. DA synthesis, release, and metabolism…………………………………………………………19 Figure 2.1. Representative brightfield images of pheochromocytoma (PC12) cells in culture (passage 16) …………………………………………………………………………………………………….36 Figure 2.2. High performance liquid chromatography (HPLC) chromatographs..…………….44 Figure 2.3. Synthesis of DOPAL standard………………………………………………………………………45 Figure 2.4. Hoechst/ propidium iodide fluorescent stain for cell viability in PC12 cells.….47 Figure 2.5. Mitochondrial Stress Test using the XF24 Analyzer………………………………………53 Figure 2.6. XF24 cell density optimization…………………………………………………………………….58 Figure 2.7. XF24 assay medium optimization………………………………………………………………..59 Figure 2.8. XF24 FCCP titration…………………………………………………………………………………….60 Figure 2.9. ALDH assay substrate optimization……………………………………………………………..63 Figure 2.10. Sample volume titration for NADH/NAD quantification……………………………..65 Figure 3.1. Concentration-response and time-course effects of MeHg on extracellular DA concentrations..…………….…………..………………………………………………………………………………….74 Figure 3.2. Concentration-response and time-course effects of MeHg on cell viability…….75 Figure 3.3. Representative images of cell viability……..…………………………………………………..76 Figure 3.4. Effects of DA storage depletion and reuptake transporter inhibition on MeHginduced DA release……………………………………………………………………………………………………….79 Figure 3.5. Effect of MeHg on intracellular DA concentrations..………………………………………81 Figure 3.6. Effect of MeHg on DA storage utilization………………………………………………………82 Figure 3.7. Effect of TH inhibition on MeHg-induced DA release…………………………………….84 xiii Figure 3.8. Effect of DOPA decarboxylase inhibition on TH enzymatic activity………………..85 Figure 3.9. Effect of MeHg on TH expression…………………………………………………………………86 Figure 4.1. Diagram depicting DA metabolic pathways………………………………………………….96 Figure 4.2. MeHg shunts DA metabolism along the reductive metabolic pathway……..…..102 Figure 4.3. Percentage changes in the relative amounts of DA metabolites induced by MeHg………………………………………………………………………………………………………………………….104 Figure 4.4. Concentration-response effects of the ALDH inhibitor daidzin on concentrations of intracellular metabolites. …………………………………………………….…………..106 Figure 4.5. Time-course effects of the ALDH inhibitor daidzin on concentrations of intracellular metabolites……………………………………………………………………………………………..107 Figure 4.6. Comparative effects of MeHg, daidzin, and rotentone on concentrations of intracellular metabolites……………………………………………………………………………………………..109 Figure 4.7. Comparative effects of MeHg, daidzin, and rotenone on ALDH activity.……….111 Figure 4.8. Concentration-response effects of MeHg on ALDH activity ex vitro……………...112 Figure 4.9. Comparative effects of MeHg, daidzin, and rotenone on mitochondrial bioenergetics……………………………………………………………………………………………………………...114 Figure 4.10. Quantification of changes in mitochondrial bioenergetics induced by MeHg, daidzin, or rotenone …………………………………………………………………………………………………...115 Figure 4.11. Effect of MeHg on the production of NAD(H).…………………………………………...117 Figure 5.1. Extracellular Ca2+ does not contribute to MeHg-induced DA release..…………137 Figure 5.2. Spontaneous activation of VGCCs does not contribute to MeHg-induced DA release………………………………………………………………………………………………………………………..139 Figure 5.3. Changes in mitochondrial bioenergetics induced by MeHg in the absence or presence of extracellular Ca2+……………………………………………………………………………………..142 Figure 5.4. Quantification of changes in mitochondrial bioenergetics induced by MeHg in the absence or presence of extracellular Ca2+………………………………………………………………143 xiv Figure 5.5. Effect of MeHg in the absence or presence of extracellular Ca2+ on concentrations of intracellular metabolites. ………………………………………………………………...145 Figure 6.1. Intracellular Ca2+ partially contributes to MeHg-induced DA release………….163 Figure 6.2. Representative traces showing changes in the ratio of fura-2 intensity……….165 Figure 6.3. Changes in mitochondrial bioenergetics induced by MeHg in absence or presence of either intracellular or extracellular Ca2+……………………………………………………167 Figure 6.4. Quantification of changes in mitochondrial bioenergetics induced by MeHg in absence or presence of either intracellular or extracellular Ca2+………………………….……….168 Figure 6.5. Effect of MeHg in the absence or presence of intracellular and/or extracellular Ca2+ on intracellular concentrations of DOPAC and DOPET…………………………………………..171 Figure 7.1. Proposed model identifying molecular targets of MeHg toxicity in DAsynthesizing cells. ……..………………………………………………………………………………………………..190 xv KEY TO ABBREVIATIONS 0-Ca2+ HBS Calcium-free HEPES-buffered saline ACh Acetylcholine ALDH Aldehyde dehydrogenase AMT α-methyltyrosine AR Aldehyde/ aldose reductase ATP Adenosine triphosphate Ca2+ Calcium CaM-MPK Ca2+/calmodulin-dependent multiprotein kinase CAT Choline acetyltransferase DA Dopamine DAT Dopamine transporter DβH Dopamine beta-hydroxylase DDC DOPA decarboxylase DMI Desipramine DOPA 3,4-dihydroxyphenylalanine DOPAC 3,4-dihydroxyphenylacetic acid DOPAL 3,4-dihydroxyphenylacetaldehyde DOPET 3,4-dihydroxyphenylethanol ETC Electron transport chain FCCP Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid xvi HBS HEPES-buffered saline Hg Mercury IP3 Inositol 1,4,5-triphosphate MAO Monoamine oxidase MeHg Methylmercury NAD Nicotinamide adenine dinucleotide, oxidized NADH Nicotinamide adenine dinucleotide, reduced NE Norepinephrine NET Norepinephrine transporter NSDA Nigrostriatal dopamine O2 Oxygen OCR Oxygen consumption rate PCB Polychlorinated biphenyl PD Parkinson disease PKC Ca2+/phospholipid-dependent protein kinase pTH Ser40 Phosphorylated tyrosine hydroxylase at serine 40 ROS Reactive oxygen species SER Smooth endoplasmic reticulum TH Tyrosine hydroxylase VGCC Voltage-gated calcium channel VMAT Vesicular monoamine transporter XF Extracellular flux xvii Chapter 1. General Introduction 1.1. Statement of Purpose Methylmercury (MeHg) is a potent, bioaccumulative neurotoxicant linked to severe neurological dysfunction in humans. Discrete neuronal populations are selectively susceptible to the neurotoxic effects of MeHg, including the nigrostriatal dopamine (NSDA) pathway. Degeneration of NSDA neurons contributes to the onset of Parkinson disease (PD). Epidemiological evidence suggests that prolonged exposure to MeHg in human populations that consume large amounts of MeHg-contaminated marine food increases the incidence of PD. Furthermore, in vitro analyses and animals studies have concluded that MeHg alters DA neuronal homeostasis and induces selective DA cytotoxicity. While it is clear that MeHg impairs DA neuronal function, the mechanisms by which MeHg targets processes in DA-synthesizing cells including synthesis, release, and metabolism remain unclear. Because MeHg cytotoxicity is generally associated with perturbation of intracellular calcium (Ca2+) homeostasis and mitochondrial dysfunction, experiments were performed to identify the contributions of these molecular mechanisms to altered DA homeostasis using DA-synthesizing pheochromocytoma 12 (PC12) cells. The overall aim of the research described in this dissertation is two-fold: 1) to characterize MeHg-induced changes in DA synthesis, release and metabolism in undifferentiated PC12 cells, and 2) examine the contributions of Ca2+ and mitochondrial dysfunction to these changes. The underlying hypothesis of these studies is that MeHginduced increase in intracellular Ca2+ mediates changes in DA synthesis and release, 1 whereas MeHg-induced mitochondrial dysfunction mediates changes in DA metabolism. Evaluation of this hypothesis will elucidate mechanisms by which MeHg targets processes intrinsic to DA neurons thereby predisposing them to degeneration. These studies will provide a platform to develop novel targets of neuroprotective therapies for treating MeHg intoxication and PD. 2 1.2. Methylmercury Methylmercury (MeHg) is composed of a methyl group (CH3) bonded to a mercuric ion (Hg2+), which produces a monovalent charged cation with enhanced lipophilicity. It is produced from elemental mercury through both natural and industrial processes (Figure 1.1; Bhan and Sarkar, 2005). Mercury is released naturally during degassing of the Earth’s crust, volcano emissions, and soil erosion. Human sources of elemental mercury include coal-burning, waste combustion, mercury mining, and mercury-containing fungicide use. Once released, mercury resides in the atmosphere as a vapor, where it is slowly oxidized to inorganic mercury by ozone (Morel et al., 1998). This more water-soluble form of mercury is deposited on the Earth’s surface by rainwater and filters into aquatic ecosystems. The majority of oxidized mercury is reduced back to the vapor state and returns to the atmosphere, but aquatic microorganisms methylate a portion of inorganic mercurous salts to form MeHg (Clarkson, 1998). MeHg biomagnifies in aquatic food chains, meaning that the concentration increases in aquatic organisms of each successive trophic level (i.e. increasing concentrations from algae, to zooplankton, to forager fish, to predatory marine animals). Additionally, aquatic animals accumulate and retain MeHg in their bodies, through a process called bioaccumulation. As a result of these two factors, MeHg reaches high concentrations in upper trophic level marine animals, such as shark, pilot whale, swordfish, and albacore tuna (WHO, 1990). Marine predators are a component of the human diet, and in some cases provide the primary source of protein, vitamins, and minerals. As such, government agencies are 3 concerned about the potential public health risk associated with consumption of MeHgcontaminated marine food. For example, the Environmental Protection Agency (EPA) estimates the average MeHg concentration in marine animals consumed by adults in the general U.S. population is 157 μg/kg (EPA, 2001). The oral reference dose, or the maximum acceptable oral dose, for MeHg is 0.1 μg/kg/day (EPA, 2001). Therefore the consumption of 200 g (7oz.) of fish containing 157 μg/kg MeHg would result in the intake of 31.4 μg MeHg, a concentration well above the recommended daily reference dose. Although studies investigating the adverse consequences of fish-consumption in adult populations have produced largely negative results (Haxton et al., 1979; Turner et al., 1980; Valciukas et al., 1986; Weil et al., 2005), there is concern about chronic, low-concentration consumption (WHO, 1990), as MeHg is known to produce severe neurotoxic effects in humans. Additionally, the issue of superimposition of MeHg-induced neurodegeneration as a function of age is a concern (Weiss et al., 2002; Landrigan et al., 2005; Monnet-Tschudi et al., 2006). 4 Figure 1.1. Global mercury (Hg) cycle. Hg is released from both natural and anthropogenic sources and for the most part enters the atmosphere as a vapor in its elemental state. It is oxidized to a more soluble inorganic form and deposited by rainwater into aquatic ecosystems. Here inorganic Hg can be methylated by microorganisms to form organic Hg, including MeHg. Due to its lipophilicity, organic Hg biomagnifies in aquatic food chains, reaching high concentrations in large predatory fish that are consumed by humans. For interpretation of the references to color in this and all other figures, the reader is referred to the electronic version of this dissertation. (Adapted from Tewalt et al., 2001) 5 1.3. Neurotoxicity of MeHg MeHg pollution and consequent human intoxication was first documented in Minamata Bay in Kumamoto Prefecture, Japan between 1953-1956 (Takeuchi et al., 1962). Two additional large-scale outbreaks occurred in Niigata, Japan in the mid 1960’s (Eto, 1997) and Iraq during the early 1970’s (Bakir et al., 1973). Several other isolated incidences of MeHg intoxication have also been documented in the last half century (Hunter et al., 1940; Hunter and Russell, 1954; Davis et al., 1994; Nierenberg et al., 1998). Despite differences in source, concentration, and duration of exposure, signs and symptoms of MeHg intoxication and targeted brain regions were similar in all cases. Primary signs of MeHg exposure are neurological dysfunctions characterized by ataxia, paresthesia, auditory and visual deficits, and dysarthria (Hunter et al., 1940; AminZaki et al., 1978; Fukuda et al., 1999). Autopsies of brains of affected individuals show a unique and characteristic distribution of pathological features that correlate with clinical signs. In the cerebral cortex, lesions are found in the calcarine cortex of the occipital lobe, primarily along the calcarine fissure (Hunter and Russell, 1954; Takeuchi et al., 1962). Less severe lesions have also been observed in the precentral, postcentral, and temporal cortices (Eto, 1997). In the cerebellum, lesions appear deeper in the hemispheres (Hunter and Russell, 1954). The cerebellar granule cell population is most affected, while the adjacent Purkinje cells are relatively spared (Takeuchi et al., 1962). The basis of selective sensitivity of certain neuronal cell populations to MeHg toxicity is not well understood, and very few comparative analyses have investigated this differential susceptibility. Perhaps the most well studied populations have been the cerebellar granule and adjacent Purkinje cells. As compared to Purkinje cells, granule cells 6 may be more susceptible to MeHg because of an inability to regulate intracellular divalent cations (Edwards et al., 2005). The resistance observed in Purkinje cells could be due to the expression of the Ca2+-binding protein, calbindin D28K (Kadowaki et al., 1993; Amenta et al., 1994), or possibly the differential expression of voltage-gated Ca2+ channel (VGCC) subtypes (Hillman et al., 1991; Randall and Tsien, 1995; Tanaka et al., 1995). 1.4. Molecular mechanisms of MeHg-mediated neurotoxicity Cytotoxicity following MeHg exposure can occur by either apoptosis or necrosis (Kunimoto, 1994; Castoldi et al., 2000). The intensity of the insult is the critical factor in the time-to-onset and the mode of cell death initiated. High concentrations of MeHg (5-10μM) induce immediate necrotic death, whereas low concentrations (≤2.5μM) cause delayed apoptosis (Castoldi et al., 2000). Primary molecular mechanisms responsible for MeHginduced cytotoxicity are still unclear, however disrupted intracellular calcium (Ca2+) homeostasis, impaired mitochondrial function, and inhibition of protein function due the toxicant’s affinity for sulfhydryl groups have all been implicated (Atchison and Hare, 1994; Castoldi et al., 2001; Sanfeliu et al., 2003). MeHg increases intracellular Ca2+ concentrations in several neuronal cell types, including cerebellar neurons (Sarafian, 1993; Marty and Atchison, 1997; Edwards et al., 2005), immature primary neuronal cultures (Mundy and Freudenrich, 2000), and immortalized neuroblastoma cells (Hare et al., 1993; Hare and Atchison, 1995a). The rise in intracellular Ca2+ proceeds in a biphasic manner with temporally and kinetically distinct 7 phases (Denny et al., 1993; Marty and Atchison, 1997). First, Ca2+ is mobilized from intracellular stores, including the smooth endoplasmic reticulum (SER) and mitochondria. MeHg induces an initial release of Ca2+ from an inositol 1,4,5-triphosphate (IP3)-sensitive pool in the SER, which is buffered by the mitochondria, only to be released again after opening of the mitochondrial permeability transition pore (Hare and Atchison, 1995a; Limke and Atchison, 2002; Limke et al., 2003). During the second phase, extracellular Ca2+ enters the cell. This phase is mediated in part by VGCCs, as inhibition of these channels delays the onset of the second phase (Marty and Atchison, 1997). Elevated intracellular Ca2+ is thought to contribute to MeHg-mediated cell death because buffering intracellular Ca2+ with chelators or treatment with VGCC blockers attenuates MeHg-induced cell death in cerebellar granule cells (Marty and Atchison, 1998). Furthermore, VGCC block with the L-type VGCC antagonist flunarizine prevents the appearance of neurological signs in rats treated with MeHg (Sakamoto et al., 1996). MeHg accumulates in mitochondria (Yoshino et al., 1966; Sone et al., 1977) leading to a variety of detrimental effects, including inhibition of mitochondrial respiration and adenosine triphosphate (ATP) production (Sone et al., 1977; Levesque and Atchison, 1991), generation of reactive oxygen species (ROS) (LeBel et al., 1990; Yee and Choi, 1994; Dreiem and Seegal, 2007), and dysregulation of mitochondrial Ca2+ buffering (Levesque and Atchison, 1991; Dreiem and Seegal, 2007). Mitochondrial Ca2+ dysregulation is likely related to MeHg-induced collapse of the mitochondrial membrane potential (Bondy and McKee, 1991; Levesque and Atchison, 1991; Hare and Atchison, 1992; Dreiem et al., 2005). 8 This mitochondrial membrane depolarization results in inhibition of Ca2+ uptake by and induction of Ca2+ efflux from the mitochondria (Levesque and Atchison, 1991; Limke et al., 2003), ultimately contributing to the accumulation of Ca2+ in the cytoplasm. Inhibition of mitochondrial respiration by MeHg is due to direct interaction with respiratory enzyme complexes (Sone et al., 1977; Yee and Choi, 1996; Mori et al., 2011). This is thought to contribute to oxidative damage because ROS formation increases after direct stimulation of the ubiquinol:cytochrome c oxidoreducatase complex (Complex III) of the electron transport chain (ETC) (Yee and Choi, 1996). Furthermore, MeHg-induced inhibition of the mitochondrial ETC and generation of ROS precipitates apoptosis by releasing cytochrome c into the cytoplasm (Mori et al., 2011) that initiates the apoptotic cell death cascade (Cai et al., 1998). MeHg has a high affinity for thiol groups (Hughes, 1957; Sanfeliu et al., 2003). As a result, proteins containing cysteine or methionine residues are acutely susceptible to structural and functional modifications by MeHg. Indeed, MeHg inhibition of PKC in vitro is mimicked by other suflhydryl blocking agents, including 5,5’-dithiobis-2-nitrobenzoic acid and N-ethylmaleimide (Inoue et al., 1988). Furthermore, it is hypothesized that the affinity of MeHg for sulfhydryl groups contributes to oxidative stress provoked by mitochondrial ROS formation. MeHg depletes glutathione (Yee and Choi, 1996; Gatti et al., 2004; Cuello et al., 2010) and reduces the activity of glutathione reductase and glutathione peroxidase (Franco et al., 2009; Cuello et al., 2010), likely by binding thiol groups present in the active sites of these enzymes. The concomitant rise in ROS following MeHg exposure would result in exacerbated oxidative stress and cell death. 9 Because MeHg targets multiple brain regions and many cellular functions, it has been hypothesized that progressive damage, throughout one’s lifetime, contributes to the development of certain neurodegenerative diseases later in life (Monnet-Tschudi et al., 2006). In diseases associated with aging, such as Parkinson disease (PD), the rate of cell loss normally occurring during aging might be accelerated by chronic, low-level exposure to neurotoxicants (Weiss et al., 2002; Weiss, 2011). 1.5. Parkinson’s disease PD is a progressive neurodegenerative disorder affecting more that 850,000 people in the United States alone (Fahn, 2003; Landrigan et al., 2005). Clinical presentation is classically characterized by debilitating motor signs, including resting tremor, rigidity, bradykinesia, and postural instability (Dauer and Przedborski, 2003). The severity of motor symptoms is due to the progressive loss of nigrostriatal dopamine (NSDA) neurons (Fahn, 2003), which originate in the substantia nigra and project to the striatum. Mechanisms of NSDA neuronal loss are multi-factorial, involving both genetic and environmental risk factors (Di Monte, 2003). In the past two decades, genetic approaches to studying neurodegenerative diseases have resulted in the discovery of 28 distinct chromosomal regions associated with PD (Klein and Westenberger, 2012). Most notably, these include the regions that contain the genes that encode α-synuclein, leucine-rich repeat kinase 2, parkin, and DJ-1. Mutations in these genes have been linked to heritable, monogenetic forms of PD. However, these familial cases account for only about 10% of all PD case (Thomas and Beal, 2007). Most patients are diagnosed with sporadic PD, for which causes are multifaceted and relatively unknown. 10 The search for environmental factors that contribute to PD has been more difficult. Toxicant exposure can occur well before clinical manifestation, resulting in a “silent latent period” that can remain undetected for years (Weiss et al., 2002; Landrigan et al., 2005), and reduce the number of neurons below those needed to sustain function in the face of normal aging (Weiss et al., 2002; Landrigan et al., 2005). A more complete understanding of how exposure to particular harmful agents impacts the pathogenesis of PD has far-reaching implications, including determination of gene-environment interactions, as well as the identification of at-risk populations and the development of preventative therapeutic strategies for treatment in these populations. 1.6. Epidemiological evidence of MeHg as an environmental risk factor for PD In the mammalian central nervous system, MeHg accumulates mainly in the cortex, striatum, cerebellum, brain stem, and spinal cord (Møller-Madsen, 1994). Accumulation in the striatum might impair proper basal ganglia function necessary to modulate voluntary movement, and contribute to the presentation of motor signs associated with MeHg intoxication. Additionally, this pattern of accumulation suggests that MeHg exposure plays a pathogenic role in the onset of PD (Monnet-Tschudi et al., 2006). A number of epidemiological studies have been carried out to investigate the relationship between MeHg content and the prevalence of PD. The most well studied population resides in the Faroe Islands, an archipelago in the northern Atlantic Ocean. The Inuit population in the Faroe Islands is stable, demarcated, ethnically homogenous, and geographically isolated, making it well suited for epidemiological studies. Estimated ageadjusted prevalence of idiopathic PD on the Faroe Islands is approximately 183.3 per 11 100,000, which is two-fold higher than that in the island of Als, Denmark or Rogaland, Norway, two geographically proximal locations (Wermuth et al., 1997; 2000; 2008). Genetic mutations are thought to play only a minimal role in the higher prevalence of PD in the Faroe Islands. Of the 100 Faroes patients diagnosed with PD in 2008, only six were diagnosed before the age of 50 (Wermuth et al., 2008). Additionally, only 33% of Faroese patients with PD reported other family members with symptoms of PD or parkinsonism (Wermuth et al., 2008). These statistics suggest that the majority of PD cases in the Faroe Islands are idiopathic. Given the geographical isolation on the islands, it is also plausible that a predisposition to PD could be due to mutations of relevant genes. Mutations in two genes that encode proteins associated with biochemical defense against peroxidation (Collins and Neafsey, 2002), human hemochromatosis gene (HFE; Milman et al., 2005) and the cytochrome P450 gene, CYP2D6 (Halling et al., 2005), occur more frequently in Faroese than other Caucasian populations. However evidence suggests there is no association between mutations in these genes and PD (Halling et al., 2008). The higher prevalence of PD in the Faroe Islands has been associated with chronic, lifetime exposure to neurotoxic contaminants present in pilot whale meat and blubber (Petersen et al., 2008a), including both MeHg and polychlorinated biphenyls (PCBs). Based on this epidemiological analysis it is impossible to discern individual contributions of each of these neurotoxicants. Both MeHg and PCBs affect DA function (Minnema et al., 1989; Kalisch and Racz, 1996; Choksi et al., 1997; Mariussen and Fonnum, 2001; Faro et al., 2002), and there is some evidence to suggest that the contaminants may act in synergy to increase the risk for PD (Bemis and Seegal, 1999). However, a case-control study in Singapore indicates a specific dose-response relationship between blood mercury levels 12 and idiopathic PD (Ngim and Devathasan, 1989), suggesting that MeHg may be capable of contributing to the etiology of PD independently of other neurotoxic contaminants, such as PCBs. Regardless of whether MeHg acts alone or in concert with other chemicals to contribute to the development of PD, in order to understand the synergy of multiple neurotoxicants it is necessary to elucidate more completely how each targets specific neuronal processes individually. Hence, further efforts are needed to elucidate the role of MeHg toxicity in NSDA neuronal loss. To this end, research aimed at investigating mechanisms by which MeHg selectively targets processes within DA neurons (i.e. synthesis, release, reuptake, and metabolism) would contribute to an understanding of how MeHg exposure contributes to the etiology of PD. 1.7. Dopamine DA (3-hydroxytyramine; Figure 1.2) is a catecholamine neurotransmitter synthesized from the dietary amino acid tyrosine. Populations of DA cell bodies originate in discrete cell groups in the olfactory bulb, diencephalon, and mesencephalon, with axons projecting throughout the cortex, striatum, nucleus accumbens, hypothalamus and spinal cord (Björklund and Dunnett, 2007). DA plays roles in many important physiological processes including reward, cognition, movement, pain modulation, and hormone secretion (Graybiel, 1990; Le Moal and Simon, 1991; Spanagel and Weiss, 1999; Lookingland and Moore, 2005; Pappas et al., 2011), as well as serving as the precursor for another brain catecholamine, norepinephrine (NE; Hornykiewicz, 1966). 13 Figure 1.2. The molecular structure of dopamine (3-hydroxytyramine). 14 1.7.1. DA synthesis The maintenance of DA homeostasis in the axon terminal, including synthesis, storage, release, reuptake and metabolism, is depicted in Figure 1.3. DA is synthesized from the dietary amino acid tyrosine, which is transported into the cell by the L-neutral amino acid transporter (Oxender and Christensen, 1963). The rate-limiting enzyme in DA synthesis, tyrosine hydroxylase (TH), converts tyrosine to 3,4-dihydroxyphenylalanine (DOPA; Levitt et al., 1965). Under physiological conditions, DOPA is not detectable in the neuron due to the high affinity and action of DOPA decarboxylase (DDC) that rapidly converts DOPA to DA (Blaschko, 1942). Short-term regulation of DA synthesis is accomplished by changes in the phosphorylation state of TH (Kumer and Vrana, 1996). Phosphorylation of critical serine (Ser) residues (Ser19, -31, and -40) located on the N-terminus of the enzyme increase enzyme efficiency (Haycock, 1990; Daubner et al., 2011). The most important mechanism of TH activation is phosphorylation at Ser40, which decreases end-product feedback inhibition (Daubner et al., 1992; Ribeiro et al., 1992; Ramsey and Fitzpatrick, 1998). Phosphorylation of Ser19 and -31 have modest direct effects on TH activity, but increase the rate of Ser40 phosphorylation, thereby indirectly potentiating TH activity (Bevilaqua, 2001; Lehmann et al., 2006). A number of Ca2+-dependent pathways mediate TH phosphorylation including Ca2+/phospholipid-dependent protein kinase (PKC; Albert et al., 1984; Vulliet et al., 1985) and Ca2+/calmodulin-dependent multiprotein kinase ( CaM-MPK; Yamauchi and Fujisawa, 1981; Vulliet et al., 1984). 15 Dephosphorylation of TH is accomplished by two protein phosphatases (PP), 2A and 2C (Dunkley et al., 2004), both of which are capable of acting at Ser40 (Haavik et al., 1989; Bevilaqua et al., 2003). Studies have only begun to elucidate regulation of these phosphatases and mechanisms by which they dephosphorylate TH. Under physiological conditions, PP2A accounts for ~90% of TH phosphatase activity in rabbit corpus striatum (Haavik et al., 1989), and PP2C activity is approximately one third of this activity (Bevilaqua et al., 2003). However certain physiological conditions may favor PP2C activity, for example high Mg2+ levels (Bevilaqua et al., 2003). Chaperone proteins may also regulate PP2A and PP2C activity. The active form of PP2A requires the co-localization of three subunits, and chaperone proteins play a pivotal role in the formation of the active PP2A heterotrimer (Dobrowsky et al., 1993). One potential chaperone protein is αsynuclein, which shares physical and functional homology with the 14-3-3 family of cytosolic chaperones (Ostrerova et al., 1999). Studies indicate that α-synuclein overexpression in both MN9D and PC12 cells decreased pTH Ser40 phosphorylation by increasing PP2A activity (Peng, 2005). 1.7.2. DA storage The majority of newly synthesized DA is packaged into synaptic vesicles by the vesicular monoamine transporter (VMAT) (Erickson et al., 1992). DA packaging requires energy to store a large amount of neurotransmitter in a small vesicular space at a high concentration. VMAT relies upon the pH and electrochemical gradient generated by a vesicular H+/K+-ATPase. ATP is used to sequester protons creating a low intravesicular pH 16 under physiological conditions (Rudnick, 1998). As a result, VMAT can couple DA uptake to the release of protons down their chemiosmotic pH gradient (Johnson and Scarpa, 1979). 1.7.3. DA release and activation of DA receptors Action potential-induced depolarization of the axon terminal causes DA to be released from the synapse via Ca2+-dependent vesicular exocytosis (Rubin, 1970; Leslie et al., 1985). Released DA can bind either pre- or post-synaptic DA receptors. DA receptors are G-protein coupled receptors categorized as D1- or D2-like (Missale et al., 1998). D1-like receptors are present on the post-synaptic membrane, and elicit an excitatory response by stimulating adenylate cyclase (Monsma et al., 1990). Conversely, D2-like receptors have an inhibitory effect on adenylate cyclase (Onali et al., 1985). They are present pre- and postsynaptically, where they inhibit DA synthesis through end-product feedback inhibition (Nowycky and Roth, 1978), or modulate the excitability of post-synaptic neurons, respectively. 1.7.4. DA reuptake To terminate the action of DA in the synaptic cleft, DA is recaptured by the DA transporter (DAT) located on the presynaptic membrane (Kilty et al., 1991; Shimada et al., 1991; Usdin et al., 1991). The driving force for DA reuptake by DAT is provided by the transport of two Na+ and one Cl- into the cell (Gu et al., 1994; Torres et al., 2003). Recaptured DA may either be re-packaged into synaptic vesicles for subsequent release, or metabolized in an effort to maintain low levels of non-vesicular DA. 17 1.7.5. DA metabolism The major metabolic pathway of DA is oxidative deamination by the mitochondrial enzyme monoamine oxidase (MAO) (Kopin, 1964) to the reactive, transient intermediate 3,4-dihydroxyphenylacetaldehyde (DOPAL). Aldehyde dehydrogenase (ALDH), a nicotinamide adenine dinucleotide (NAD)-dependent enzyme, rapidly oxidizes DOPAL to the inactive metabolite 3,4-dihydroxyphenylacetic acid (DOPAC) (Marchitti et al., 2008). Under some circumstances DOPAL is also reduced to 3,4-dihydroxyphenylethanol (DOPET) by aldehyde/ aldose reductase (AR) (Elsworth and Roth, 1997). Impaired conversion of DA to DOPAC renders cells susceptible to a variety of toxic compounds. Cytosolic DA can be autooxidized, which results in the formation of DAquinones and ROS (Graham, 1978; Graham et al., 1978; LaVoie and Hastings, 1999; 2002). Furthermore, DOPAL itself is a highly reactive intermediate, which readily forms adducts with many cellular components (Marchitti et al., 2007). Because alterations in normal metabolism can impair cellular integrity, it is essential for cells to maintain tight regulation of DA synthesis, storage, release, and metabolism. 18 Figure 1.3. DA synthesis, release, and metabolism. Dietary tyrosine is hydroxylated to DOPA, which is rapidly decarboxylated to DA. DA is packaged into synaptic vesicles by VMAT, and is released from the synapse by depolarization-evoked, Ca2+-dependent exocytosis. Once released, DA can bind pre- and post-synaptic D1 and D2 DA receptors to elicit either excitatory or inhibitory responses, depending on the type of receptor to which it binds and their subsequent effect on adenylate cyclase (AC) to generate cAMP. Termination of DA’s action in the synapse is accomplished by reuptake through DAT. Recaptured DA can either be repackaged into synaptic vesicles or metabolized. DA metabolism proceeds in two steps, orchestrated by two mitochondrial-associated enzymes. First DA is deaminated by MAO to form DOPAL, and then DOPAL is oxidized by ALDH to form DOPAC. Tyr, tyrosine; L-NAAT, L-neutral amino acid transporter; TH, tyrosine hydroxylase; DOPA, dihydroxyphenylalanine; DDC, DOPA decarboxylase; DA, dopamine; VMAT, vesicular monoamine transporter; Ca2+, calcium; D1, excitatory D1-like dopamine receptor; D2, inhibitory D2-like dopamine receptor; AC, adenylate cyclase; cAMP, cyclic adenosine monophosphate; DAT, dopamine transporter; MAO, monoamine oxidase; DOPAL, dihydroxyphenylacetaldehyde; ALDH, aldehyde dehydrogenase; DOPAC, dihydroxyphenylacetic acid. 19 1.8. Effects of MeHg on DA neuronal homeostasis DA neurons are particularly sensitive to the toxic effects of MeHg. Exposure to nanomolar concentrations of the neurotoxicant selectively reduces the number of THpositive neurons in murine embryonic stem cell cultures (Zimmer et al., 2011). Chronic, low-concentration exposure (1-2μM) to MeHg causes a significant loss of DA neurons in the nematode Caenorhabditis elegans (VanDuyn et al., 2010). Furthermore, MeHg impairs a variety of DA-specific bioprocesses in the cell, including synthesis, release, reuptake, and metabolism. 1.8.1. MeHg targeting of DA synthesis Studies investigating effects of MeHg on DA synthesis have primarily measured changes in enzymatic activity of TH. Results are variable and inconsistent. Some studies report decreased TH activity following acute, high dose MeHg exposure in adult rats (Sharma et al., 1982; Kung et al., 1989). Others demonstrate no change in TH activity following either in utero or early postnatal exposure to acute, low doses of MeHg (Taylor and DiStefano, 1976; Bartolome et al., 1984). Omata et al. (1982) have suggested that MeHg stimulates TH activity during an early, asymptomatic phase of exposure to a high dose of MeHg in adult rats (days 5-10 of daily treatment). However, with the onset of neurological signs, 15 days into treatment, TH activity had returned to control rates. Discrepancies between these studies likely result from different exposure paradigms. As an additional caveat to the majority of these studies, TH activity was assessed in whole brain homogenates (Taylor and DiStefano, 1976; Omata et al., 1982; Sharma et al., 1982; Bartolome et al., 1984), which confounds interpretation of how MeHg selectively targets DA synthesis. 20 1.8.2. MeHg targeting of DA release Spontaneous DA release increases following exposure to MeHg. This has been well documented in a variety of model systems, including 3H-DA release from rat striatal synaptosomes (Minnema et al., 1989) and mouse striatal slices (McKay et al., 1986), as well as endogenous DA release from rat striatal synaptosomes (Dreiem et al., 2009), rat striatal punches (Bemis and Seegal, 1999), mouse striatal slices (Kalisch and Racz, 1996), and the striatum of conscious, free-moving rats (Faro et al., 1997; 1998; 2000; 2003; 2007). Mechanisms underlying MeHg-induced DA release, however, are not well characterized. While it is hypothesized that extracellular Ca2+ does not contribute (McKay et al., 1986; Minnema et al., 1989; Kalisch and Racz, 1996; Faro et al., 2002), a role for intracellular Ca2+ in MeHg-induced DA release has not been investigated. As MeHg is known to increase the concentration of cytosolic Ca2+ (Marty and Atchison, 1997) and intracellular Ca2+ contributes to MeHg-induced release of other neurotransmitters (Levesque and Atchison, 1987; 1988), this potential contribution is of interest. Only one study to date has specifically investigated mechanisms of MeHg-induced DA release. Faro et al. (2002) suggest that vesicular release is not involved, but instead attribute a role for DAT. However the authors could not discern whether MeHg was inhibiting DAT uptake or promoting a reversal of the transporter to release DA. Given the paucity of mechanistic studies, further investigation is needed to determine how MeHg targets the DA neuron to induce neurotransmitter release. 21 1.8.3. MeHg targeting of DA reuptake The majority of studies investigating DA reuptake following exposure to MeHg have demonstrated inhibition of DAT activity (Komulainen and Tuomisto, 1982; Rajanna and Hobson, 1985; Bonnet et al., 1994; Dreiem et al., 2009; Zimmer et al., 2011). In these studies, 3H uptake of DA or analogs with high affinity for DAT was assessed in striatal synaptosomes in the presence of MeHg. The IC50, or concentration needed to inhibit 50% of transporter activity, was estimated at 2μM (Komulainen and Tuomisto, 1982; Bonnet et al., 1994; Dreiem et al., 2009). However, one study demonstrated inhibition at a much lower concentration (5nM) (Zimmer et al., 2011), while another found it to be much higher (50μM) (Rajanna and Hobson, 1985). In both cases, the discrepancy is likely due to the comparative sensitivity of the model system, mouse embryonic stem cells and whole brain homogenates, respectively. While decreased DAT activity has been demonstrated in the majority of studies to date, a few have observed alternative effects. Bartolome et al. (1984) demonstrated a significant increase in 3H-DA uptake in whole brain synaptosomes isolated from rat pups treated with MeHg in utero. Conversely, Komulainen et al. (1985) reported no change in 3H-DA uptake in striatal synaptosomes isolated from rats treated with 10 mg/kg MeHg for one or five days in vivo. Interestingly, Komulainen and colleagues had previously demonstrated a decrease in DAT activity following in vitro exposure to MeHg (Komulainen and Tuomisto, 1982). It has been proposed that MeHg directly inhibits DAT by binding sulfhydryl groups present on the transporter (Bonnet et al., 1994; Dreiem and Seegal, 2007). If this is the case, the continued presence of MeHg may be necessary to inhibit DA 22 reuptake. However additional studies are needed to elucidate whether MeHg is inhibiting DAT and to determine how this inhibition may impact measurements of MeHg-induced DA release and metabolism. 1.8.4. MeHg targeting of DA metabolism Effects of MeHg on DA metabolism have been assessed using two methods. Concentrations of the primary DA metabolite, DOPAC, have been measured by highperformance liquid chromatography (HPLC). In vitro analysis of striatal tissue or striatal synaptosomes has demonstrated that MeHg decreases DOPAC, and that this effect is concentration-dependent (Bemis and Seegal, 1999; Dreiem et al., 2009). These data suggest that MeHg inhibits DA metabolism. Complementary to HPLC analyses, the activity of the enzyme that deaminates DA, MAO, has been measured by spectrophotometric or radiochemical assays. Results predominantly suggest that MeHg decreases MAO activity (Taylor and DiStefano, 1976; Chakrabarti et al., 1998; Beyrouty et al., 2006; Castoldi et al., 2006). The most powerful effect was reported by Chakrabarti et al. (1998), who demonstrated that MeHg causes a concentration-dependent decrease in MAO activity of rat synaptosomes isolated from the cortex, striatum, hypothalamus, hippocampus, brain stem, and cerebellum. The authors also observed decreased MAO activity in the same brain regions following in vivo treatment with an acute, high dose, but not an acute, intermediate dose of MeHg. Decreased MAO activity was also reported following in utero or neonatal treatment with MeHg (Taylor and DiStefano, 1976; Beyrouty et al., 2006; Castoldi et al., 2006), however these studies only demonstrate small decreases. Beyrouty et al. (2006) examined MAO activity in both male and female offspring 23 in five brain regions, and only demonstrated a decrease in the brainstem of female offspring of dams treated with a low dose of MeHg during gestation. Following a similar treatment paradigm, Castoldi et al. (2006) only reported a decrease in enzyme activity in the cerebellum of male offspring. Overall, these results support the conclusion that MeHg decreases DA metabolism, and suggest that it does so by targeting MAO. Because MAO is bound to the outer mitochondrial membrane (Schnaitman et al., 1967), MeHg-induced loss of mitochondrial membrane potential (Bondy and McKee, 1991; Hare and Atchison, 1992; InSug et al., 1997; Dreiem et al., 2005) could compromise enzyme activity. However there is also one report of increased MAO activity in brain homogenates isolated from rats treated with 10 mg MeHg /kg/ day for 7 days (Omata et al., 1982). Discrepancies could be due to different experimental conditions, including methods of MeHg administration, animal age, or techniques used to measure MAO activity (e.g. substrate). These contradictory results and the lack of compelling evidence from multiple studies to support the conclusion that MeHg decreases MAO activity necessitate further investigation to determine if and how MeHg decreases DA metabolism. 1.9. Pheochromocytoma 12 (PC12) cell line To investigate MeHg targeting of DA homeostasis, undifferentiated PC12 cells were chosen as the model system in the present series of experiments. The PC12 cell line is an immortalized, clonal cell line derived from a rat adrenal catecholamine-secreting pheochromocytoma (Greene and Tischler, 1976). Cultured under normal conditions, PC12 cells resemble adrenal chromaffin cells in morphology, physiology, and biochemistry. However, when cultured in the presence of nerve growth factor (NGF), PC12 cells exit the 24 cell cycle and differentiate to resemble sympathetic neurons. In both states PC12 cells synthesize, store, and release DA, NE, and acetylcholine (ACh). They also express a variety of ion channels and ligand-gated receptors. As such, PC12 cells represent a useful model for studying a plethora of cellular processes including neuronal differentiation, regulation and function of ion channels and membrane-bound receptors, and synthesis, storage and release of neurotransmitters. Accordingly, this cell line has been used in over 13,000 publications to date. PC12 cells, in both their undifferentiated and differentiated states, are a valuable model system for studying DA neurochemistry and homeostasis (Greene and Rein, 1977a; Malagelada and Greene, 2011). They maintain all of the enzymes necessary for DA synthesis (i.e. TH, DDC) and metabolism (i.e. MAO, ALDH) (Greene and Tischler, 1976; Youdim et al., 1986; Robador et al., 2012), express high-affinity vesicular and membrane catecholamine transporters (Greene and Rein, 1977a; Friedrich and Bönisch, 1986), and release DA by both spontaneous and depolarization-evoked, Ca2+-dependent, vesicular exocytosis (Greene and Tischler, 1976; Greene and Rein, 1977a; Ritchie, 1979; Wagner, 1985). However because of disparities in neurotransmitter content, enzyme activity, and cell morphology, undifferentiated and differentiated PC12 cells each provide unique advantages for studying DA synthesizing cells. 1.9.1. PC12 cell morphology The most discernible difference between undifferentiated and differentiated PC12 cells is morphology. According to the original description of this cell line (Greene and Tischler, 1976), undifferentiated PC12 cells (Figure 2.1) are small, round, and tend to 25 clump in culture. Following treatment with NGF, cell multiplication ceases, and neuronallike processes can be observed by 7 days. Over the course of days 7-22, the number, length, and density of processes increases until ~80% of cells have responded to NGF treatment. These neuronal-like processes, which most closely resemble primary sympathetic neuronal cultures (Chamley et al., 1972; Mains and Patterson, 1973), are fine, long, and branched, contain numerous varacosities, and form fascicles. The neuronal, post-mitotic phenotype of differentiated PC12 cells lends itself to studies of neuronal processes in DA neurons, including neuroprotection by growth factors in PD (Shimoke and Chiba, 2001; Salinas et al., 2003) and alteration of DA neurodevelopment and cell proliferation following exposure to toxicants (Parran et al., 2001; Chen et al., 2011). However, while DA neurons more closely resemble differentiated PC12 cells morphologically, disparities in the neurochemical profile suggest that undifferentiated PC12 cells are better suited for studies pertaining to DA homeostasis. 1.9.2. PC12 cell neurochemistry The major differences induced by PC12 cell differentiation in content and activity of catecholaminergic and cholinergic neurotransmitters and enzymes are summarized in Table 1.1. The principal neurotransmitter in undifferentiated PC12 cells is DA, however measureable concentrations of NE and ACh are also present. Following differentiation, concentrations of DA and NE decrease by approximately 4- and 6-fold respectively (Greene and Tischler, 1976), while the concentration of ACh increases by approximately 4-fold (Greene and Rein, 1977b). Synthetic enzyme activity mirrors neurotransmitter content. The activity of catecholamine enzymes, including TH, DDC, and the NE synthetic enzyme DA 26 β-hydroxylase (DβH), decrease following differentiation (Greene and Tischler, 1976; Badoyannis et al., 1991), whereas the activity of enzymes necessary for ACh synthesis, including choline acetyltransferase (CAT), increase (Greene and Rein, 1977b). Consequently, differentiated PC12 cells are more responsive to ACh, as compared to undifferentiated PC12 cells which are more responsive to DA (Dichter et al., 1977). Because DA content and TH activity decrease profoundly with NGF treatment, DA homeostasis has been primarily characterized in undifferentiated PC12 cells. As in DA neurons, DA release in undifferentiated PC12 cells is coupled to synthesis, whereby newly synthesized DA is preferentially released (Greene and Rein, 1978). Furthermore, DA biosynthesis can be regulated acutely by TH phosphorylation (Tank et al., 1986) and factors such as high K+ and dibutyryl cyclic adenosine 3’,5’ monophosphoric acid (Greene and Rein, 1978), two agents demonstrated to mediate endogenous short-term DA synthesis in catecholaminergic tissue (Harris and Roth, 1971; Patrick and Barchas, 1976). 27 Table 1.1. Catecholaminergic and cholinergic neurotransmitter content and enzyme activity in undifferentiated and differentiated PC12 cells. Undifferentiated Differentiated NEUROTRANSMITTER CONTENT (nmol/mg protein) DA 16.6 ± 1.7 1 4.4 ± 0.4 1 NE 6.1 ± 6 1 1.5 ± 0.2 1 Epinephrine < 0.15 1 < 0.15 1 ACh 1.3 ± 0.07 2 ENZYME ACTIVITY (pmol/min/mg protein) 5.6 ± 0.6 2 TH 39 ± 5 1 10 ± 1 1 DDC 770 ± 99 1 130 ± 15 1 DβH 806 ± 84 1 MAO-A 3.9 ± 0.4 1 161 ± 19 1 N.D. CAT 440 ± 17 2 520 ± 20 2 1 Greene & Tischler. (1976) Proc Natl Acad Sci (USA). 73: 2424-2428. 2 Greene and Rein. (1977b) Nature. 268: 349-351. N.D. not determined. Prior to measuring neurotransmitter content and enzyme activity in differentiated state, PC12 cells were treated with 50mg/mL of NGF for 14 (catecholaminergic properties) or 22 (cholinergic properties) days. 28 1.9.3. PC12 cell VGCC subtype expression One additional phenotypic change induced by NGF contributed to the selection of undifferentiated PC12 cells herein. The expression of VGCC subtypes is altered by PC12 cell differentation (Shafer and Atchison, 1991a). Briefly, two VGCC subtypes have been primarily described in PC12 cells: the L- and N-subtypes (Takahashi et al., 1985; Janigro et al., 1989; Plummer et al., 1989; Reber et al., 1990). Biophysically, L- and N-type VGCCs are similar and activate at relatively depolarized membrane potentials; the L-type depolarizes at approximately -10mV and the N-type at approximately -20mV. These channel subtypes are most readily distinguished by pharmacological properties. The L-type VGCC is inhibited by dihydropyridine antagonists, whereas the N-type VGCC is sensitive to ω-conotoxin GVIA (Lacinová, 2005). Undifferentiated PC12 cells predominately express L-type VGCCs (Janigro et al., 1989; Reber et al., 1990). However following treatment with NGF, there is a substantial increase in the expression of N-type VGCCs without a concomitant decrease in L-type VGCCs (Takahashi et al., 1985; Streit and Lux, 1987; Plummer et al., 1989; Usowicz et al., 1990). This difference is of importance presently because NSDA neurons express a high proportion of L-type VGCCs, which are used to maintain autonomous pacemaking activity (Nedergaard et al., 1993; Bonci et al., 1998; Chan et al., 2007). As such, the expression profile of VGCCs in undifferentiated PC12 cells more closely resembles that in NSDA neurons. 29 1.9.4. Differences between undifferentiated PC12 cells and DA neurons While the neuronal phenotype of differentiated PC12 cells more closely parellels DA neuronal morphology, the neurochemical and protein expression profile of undifferentiated PC12 cells provides a more suitable model of DA synthesizing cells. For these reasons, undifferentiated PC12 cells were chosen for the studies presented in this dissertation. However there are a few important differences between undifferentiated PC12 cells and DA neurons. As described, PC12 cells synthesize and store two additional neurotransmitters, NE and ACh (Greene and Tischler, 1976; Greene and Rein, 1977a; 1977b). NE is stored alongside DA in large dense-core vesicles (Kishimoto et al., 2005), while ACh is stored in small clear-core vesicles that are observed most prominently in the differentiated state (Greene and Tischler, 1976; Liu et al., 2005). Because of the presence of these additional neurotransmitters, it is plausible that changes in DA homeostasis in undifferentiated PC12 cells might not directly reflect changes in DA neurons. However this seems unlikely considering the synthetic and degratory pathways are essentially identical, and DA is the most prominent neurotransmitter in the undifferentiated state. The concentration of NE is much lower than that of DA, primarily because DβH requires reduced ascorbate as a cofactor, and in tissue culture medium the cofactor readily degrades (Greene and Rein, 1978). Additionally, in the undifferentiated state the concentration of ACh is relatively low as compared to DA. Another major difference between undifferentiated PC12 cells and DA neurons is the process of DA reuptake. PC12 cells express the NE transporter (NET) as opposed to DAT (Greene and Rein, 1977a; Friedrich and Bönisch, 1986). However, NET and DAT are both members of a family of Na+- and Cl--dependent carriers, and they have a sequence 30 homology of about 78%, with sequences being most similar in their transmembrane domains (Buck and Amara, 1995; Brüss et al., 1997). The major difference between these carriers is that while DAT requires two Na+ for transport, NET only requires one (Gu et al., 1994). Of importance is the affinity of NET for DA. Using kinetic assays of 3H-DA and 3H-NE uptake in the LLC-PK1 cell line stably transfected with NET or DAT, Gu et al. (1994) found that while DAT and NET transport both catecholamines, DA was the preferred substrate for both transporters. 1.9.5. Effects of MeHg in PC12 cells PC12 cells have been used as a model system to study the neurotoxic effects of MeHg. This work has primarily investigated cytotoxicity and MeHg targeting of VGCCs. In PC12 cells, as in other cell types, MeHg decreases cell viability in a concentrationdependent manner, dissipates the mitochondrial membrane potential, decreases ATP and glutathione production, and increases ROS formation (Parran et al., 2001; Gatti et al., 2004). Additionally, it has been demonstrated that MeHg decreases 45Ca2+ influx in undifferentiated PC12 cells (Shafer et al., 1990). Inhibition of Ca2+ influx is complete at 250μM, and mediated by disruption of VGCCs, not cytotoxicity, because no cell death was observed within the 1hr exposure paradigm. These authors went on to demonstrate that MeHg-mediated inhibition of VGCCs in PC12 cells is voltage-dependent, and occurs regardless of the state of the Ca2+ channel or membrane holding potential (Shafer and Atchison, 1991b). 31 1.10. Summary MeHg is a potent environmental neurotoxicant, which produces severe pathological consequences in humans. Molecular mechanisms of MeHg-mediated neurotoxicity are still elusive, however perturbation of intracellular Ca2+, mitochondrial dysfunction, and an affinity for thiol groups resulting in impaired protein function have been implicated. Despite relatively general neurototoxic effects, MeHg does not affect all cell types equally. DA neurons are one of the few cell types selectively susceptible to MeHg. Factors mediating this sensitivity are unknown, but could be related to unique processes in DA neurons that mediated the neurotransmitters synthesis, release, and metabolism. The experiments in this thesis were designed to investigate the susceptibility of DA neurons to MeHg-mediated toxicity using to the immortalized clonal PC12 cell line. 32 1.11. Thesis Objective The studies described in this dissertation were developed in order to test the central hypothesis that MeHg-induced increase in intracellular Ca2+ mediates changes in DA synthesis and release, whereas MeHg-induced mitochondrial dysfunction mediates changes in DA metabolism. The following specific aims and hypotheses were designed to test this central hypothesis. a) Characterize the relationship between MeHg-induced DA synthesis and release in undifferentiated PC12 cells following exposure to MeHg. Hypothesis: MeHg stimulates both DA synthesis and DA release. Activation of DA synthesis contributes to Ca2+-dependent vesicular exocytosis. b) Characterize the aberrant DA metabolomic profile in undifferentiated PC12 cells induced by MeHg, and examine direct and indirect factors mediating the inhibition of enzymes necessary for DA metabolism. Hypothesis: MeHg impairs DA metabolism by inhibiting mitochondrial-associated enzymes that mediate its oxidative catabolism. These enzymes are inhibited indirectly as a result of mitochondrial dysfunction induced by the toxicant. Furthermore, inhibition of DA metabolism results in formation of aberrant toxic intermediates and compensatory up-regulation of an alternative reductive metabolic pathway. 33 c) Examine the contributions of extracellular and intracellular Ca 2+ changes in DA release and metabolism induced by MeHg Hypothesis: Intracellular, but not extracellular, Ca2+ mediates MeHg-induced DA release, whereas neither participate in MeHg-impaired DA metabolism. The following chapters outline the research performed to address the above specific aims. The reader is referred to Chapter 2 for detailed methodology. Chapters 3-6 describe findings as they related to the central hypothesis and thesis objective. Chapter 7 provides a general discussion on the relevance and importance of this research as it relates to previous findings and information from the literature. 34 Chapter 2. General Materials and Methods 2.1. Culture of PC12 cells Cell culture supplies, including RPMI-1640 medium, horse serum, trypsin, and penicillin-streptomycin, were purchased from GIBCO BRL (Grand Island, NY). Hyclone fetal bovine serum was purchased from Thermo Scientific (Logan, UT). PC12 cells (Gift of Dr. M.L. Contreras) were grown in RPMI-1640 medium supplemented with 10% (v/v) horse serum, 2.5% (v/v) fetal bovine serum, and 1% (v/v) penicillin-streptomycin (pH 7.3). Cultures were maintained in either 25-cm2 or 75-cm2 T-flasks in a humidified environment containing 5% CO2 at 37°C. Culture medium was changed every 2-3 days. Every 4-5 days PC12 cultures were detached from the flasks with 0.25% (v/v) trypsin and sub-cultured at a density of 3x105 cells/mL for a 5 day culture or 4x105 cells/mL for a 4 day culture. All cultures were maintained at 80-90% confluence at the time of subculture. To maintain consistency from experiment to experiment, cells were used between passages 14-19 from our receipt. Experimental conditions were repeated in technical triplicate and experiments were replicated at least three times from separate cultures to minimize the risk of culturespecific confound. Representative brightfield images of PC12 cells (passage 16) in culture are depicted in Figure 2.1. 35 Figure 2.1. Representative brightfield images of pheochromocytoma (PC12) cells in culture (passage 16). Panel A) Low magnification. Scale bar = 200µm. Panel B) High magnification. Scale bar = 100µm. 36 2.2. Chemicals and solutions The standard physiological saline used for extracellular solution was 4-(2hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)-buffered saline (HBS), which contained (mM): 150 NaCl, 5 KCl, 2.4 CaCl2, 1.6 MgSO4, 20 HEPES, and 20 d-glucose (pH 7.3). The Ca2+-free buffer (0-Ca2+ HBS) had the same composition as HBS with the following exceptions (mM): 0 CaCl2 and 0.02 ethylene glycol tetraacetic acid. Methyl mercuric chloride (MeHg) was purchased from ICN Biochemicals Inc. (Aurora, OH). It was prepared as 10mM stock solution and stored for no more than 3 months at 4°C. On the day of each experiment, MeHg was diluted to working concentrations of 1, 2, or 5μM in HBS or 0-Ca2+ HBS. Several pharmacological treatments were used to analyze distinct components of the DA biosynthetic and release pathways, VGCC function, and pathways of intracellular Ca2+ release. Concentrations were calculated as free-base. All chemicals were prepared fresh for each experiment. Table 2.1 summarizes the specific action of each chemical used. α-Methyl-DL-tyrosine methyl ester hydrochloride (AMT; Sigma) was prepared in dH2O as a 10mM stock solution and then diluted to a final concentration of 300µM either in culture medium for a 24 hr or 30 min pretreatment or in HBS for a 60 min co-treatment with 2μM MeHg. 37 Reserpine (Sigma) was dissolved in 100% DMSO as a 10mM stock solution and then diluted so the final concentration of DMSO was less than 0.05% (v/v). The 10mM stock was diluted to a final concentration of 50nM in either culture medium for a 3 hr pretreatment or in HBS for a 60 min co-treatment with 2μM MeHg. Desipramine hydrochloride (DMI; Sigma) was prepared in dH2O as a 10mM stock and then diluted to a final concentration of 1μM either in culture medium for a 15 min pretreatment or in HBS for a 60 min co-treatment with 2μM MeHg. NSD-1015 (3-hydroxybenzylhydrazine; Sigma) was dissolved in dH2O as a 1mM stock and then diluted to a final concentration of 10µM in HBS for a 60 min co-treatment with 2μM MeHg. Daidzin (Sigma) was dissolved in 100% DMSO as stock solutions of 10mM and diluted so that the final concentration of DMSO was no more that 0.05% (v/v). The 10mM stock was diluted to a final concentration of 20μM in HBS for a 45 min treatment. Rotenone (Sigma) was dissolved in 100% DMSO as stock solutions of 10mM and diluted so that the final concentration of DMSO was no more that 0.05% (v/v). The 10mM stock was diluted to a final concentration of 50nM in HBS for a 15 min treatment. 38 Nimodipine (Sigma) is light sensitive, and as such was protected from light during preparation and experimentation. It was dissolved in dH2O as a 50mM stock solution, and then diluted to a final concentration of 2μM either in culture medium for a 15 min pretreatment or HBS for a 60 min co-treatment with 2μM MeHg. Cadmium chloride (Sigma) was prepared in dH2O as a 20mM stock solution and then diluted to a final concentration of 200μM either in culture medium for a 15 min pretreatment or HBS for a 60 min co-treatment with 2μM MeHg. BAPTA/AM (1,2-Bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid tetrakis(acetoxymethyl ester; Calbiochem) was dissolved in 100% DMSO as a 10mM stock solution and diluted to a final concentration of 50μM in HBS for a 30 min pretreatment and then a 60 min co-treatment with 2μM MeHg in either HBS or 0-Ca2+ HBS . 39 Table 2.1. Summary of drug actions. Drugs α-methyl-DLtyrosine Action Reference Source TH inhibitor (Spector et al., 1965) Sigma (Drukarch et al., 1996) Sigma Desipramine NSD-1015 VMAT inhibitor; “vesicular depleter” NET inhibitor DDC inhibitor Sigma Sigma Daidzin ALDH inhibitor Rotenone Complex I inhibitor of the mitochondrial ETC L-type VGCC inhibitor Non-specific VGCC inhibitor (Brüss et al., 1997) (Carlsson et al., 1972) (Keung and Vallee, 1993) (Lamensdorf et al., 2000a) (Neal et al., 2010) (Taylor et al., 2000) (Mahata et al., 2011) Reserpine Nimodipine Cadmium BAPTA/AM Intracellular Ca2+ chelator 40 Sigma Sigma Sigma Sigma Calbiochem 2.3. Neurochemical analyses PC12 cells were seeded in 6-well plates (9.5cm2) coated with poly-D-lysine at a density of 6x105 cells/mL (total volume of 2mL/well) 48 hr prior to pharmacological and/or toxicant treatment. At experiment termination, treatment medium was collected and acidified (1:1) with ice-cold tissue buffer (0.1M phosphate-citrate buffer containing 15% methanol (v/v), pH 2.5) for analysis of extracellular DA content. Cells were rinsed once with 1mL ice-cold phosphate buffered saline, harvested, and pelleted by centrifugation at 12,000 g for 5 min at 4°C. After centrifugation, the supernatant was removed and replaced with 100μL of ice-cold tissue buffer. The content of DA and its metabolites in the treatment medium and supernatant was determined by means of high-pressure liquid chromatography coupled with electrochemical detection (HPLC-ED) using a Water 515 HPLC pump (Waters Corp., Milford, MA) and an ESA Coulochem 5100A electrochemical detector with an oxidation potential of +0.4V. Neurochemical content was quantified by comparing peak height of each sample to peak heights of standards and normalized to mL per sample for extracellular measurements or mg protein for intracellular measurements as determined by the bicinchoninic acid (BCA) protein assay (Sigma). Figure 2.2 depicts HPLC chromatographs of extracellular and intracellular samples as compared to 500pg standards. 2.3.1. Calculation of rate constant Releasable stores of transmitter in catecholamine secreting cells are maintained by end-product feedback regulation of the balance between vesicular release and de novo 41 synthesis dependent replenishment. The slope of decline (or rate constant) of intracellular DA following inhibition of synthesis represents a reliable, indirect measurement of release or DA storage utilization (Brodie et al., 1966). In the present study, the rate constant was calculated from intracellular DA concentrations after 90 min AMT treatment. The slope of the lines representing the difference between cells treated with 2µM MeHg alone or combined with 300µM AMT was calculated, and then compared to HBS-treated cells in the absence or presence of AMT. 2.3.2. Synthesis of DOPAL standard DOPAL is an endogenous aldehyde resulting from the normal oxidative metabolism of DA by MAO. DOPAL standards are not commercially available, therefore in order to measure DOPAL by HPLC-ED it was necessary to synthesize a standard. DA was dissolved in phosphate buffer (140mM NaCl, 8.1mM Na2HPO4, 1.4mM KH2PO4, pH 7.5) to a final concentration of 1mM. Phosphate-buffered DA (750µl) was incubated in two 12x75 culture tubes at 37°C for 15 min. The reaction was started by adding 25µL MAO (10 U) to each of the two culture tubes and gently mixing. In order to determine the optimal incubation duration for experimental analysis of DOPAL, 50µLaliquots were withdrawn from the reaction at different time intervals (10, 15, 30, 45 or 60 min). The reaction in the tubes was stopped by adding 450µLof ice-cold tissue buffer. Samples were transferred to a 1.5mL centrifuge and centrifuged at 12,000 x g for 10 min. Supernatants were removed and placed into a fresh tube for HPLC-ED analysis of DA and DOPAL. Concentrations of DA were compared to 1ng standards of DA. Concentrations of DOPAL were determined by normalizing the DOPAL peak height to the amount of DA lost at each time point. Figure 2.3 42 illustrates the time-dependent synthesis of DOPAL correlated with the time-dependent decline in DA. Based on this analysis, the 45min incubation time-point was selected for experimental analysis of DOPAL concentrations. 43 Figure 2.2. High performance liquid chromatography (HPLC) chromatographs. Extracellular (middle panel) and intracellular (lower panel) PC12 cell neurochemistry was compared to standard mixes (upper panel) containing 500pg NE, DOPET, DOPAC, DOPA, and DA. Comparison of the retention time allowed for determination of which neurochemicals were present in medium and cell samples. Peak height of samples was compared to peak height of standards to determine neurochemical content. Extracellular concentrations were normalized to mL per sample and expressed in ng/mL. Intracellular concentrations were normalized to protein content and expressed in ng/mg protein. 44 Figure 2.3. Synthesis of DOPAL standard. The rate of DOPAL formation (white circles) and DA depletion (black circles) after the incubation of DA (1.063 mM) with 10 U MAO. DA and DOPAL were measured by HPLC-ED. DA concentrations were determined by comparison with 163ng DA standards. Concentrations of DOPAL were determined by comparing DOPAL peak heights to the amount of DA decreased at each time point. Based on this analysis the 45 min incubation time-point was selected for experimental analysis of DOPAL concentrations. 45 2.4. Cell viability PC12 cells were seeded in 96-well plates coated with poly-D-lysine at a density of 4x105 cells/mL 48 hr prior to treatment with HBS, 1, 2, or 5µM MeHg for 15, 30, 60 or 120 min. Hoechst 33342 (10 mg/mL) and propidium iodide (1 mg/mL) were diluted 1:10,000 in existing treatment medium. Cells were incubated in fluorophores at 37˚C for 15 min prior to visualization on a Nikon Ellipse TE2000-U inverted microscope with diascopic illumination pillar (Nikon Instruments, Inc., Melville, NY) equipped with a T-FL EPO fluorescence attachment and X-Cite metal halide illumination system. DAPI (358nm/ 461nm) and rhodamine filters (610nm) were used for visualization. Images were acquired using MetaMorph image acquisition and analysis software (Molecular Devices, Sunnyvale, CA). One image was acquired per well, randomized and counted manually using a grid system. Viable cells had a blue-stained (Hoechst) nucleus, whereas non-viable cells had a high red (propidium iodide) fluorescence (Figure 2.4; Yuan et al., 2009). Each treatment was replicated in triplicate wells in three separate experiments. 46 Figure 2.4. Hoechst/ propidium iodide fluorescent stain for cell viability in PC12 cells. Viable cells are stained with the membrane permeable blue-fluorescent dye, Hoechst 33342. The red-fluorescent dye, propidium iodide, is only permeant to cells with compromised membrane integrity, and thus stains dead cells. Scale bar = 200μm 47 2.5. Western blot analysis of TH PC12 cells were seeded in 6-well plates coated with poly-D-lysine at a density of 6x105 cells/mL 48 hr prior to pharmacological and/or toxicant treatment. Following treatment, medium was aspirated and in some cases used for neurochemical analysis of extracellular DA content. Cells were rinsed once with 1mL ice-cold phosphate buffered saline, harvested, and pelleted by centrifugation at 12,000 g for 5 min at 4°C. Pellets were lysed with 50μL radio-immunoprecipitation assay (RIPA) buffer containing protease inhibitor (1:100) and phosphatase inhibitor (1:100). Protein content was determined by the bicinchoninic acid protein assay (Sigma). Supernatant was prepared for Western blot analysis to quantify total and phosphorylated isoforms of TH. Equal amounts of protein (10-15μg) were separated on precast polyacrylamide gels (10% Bio-Rad, Hercules, CA, USA). Proteins were transferred to 0.45µm Immobulin-FL polyvinylidene fluoride (PVDFL) membranes (Millipore, Billerica, MA), incubated 1 hr in 5% skim milk (w/v) or bovine serum albumin blocking buffer and then reacted overnight with either rabbit anti-TH (1:5000; Chemicon, Billerica, MA), rabbit anti-TH Ser40 (1:1000, Cell Signaling, Danvers, MA), rabbit anti‐TH serine‐31 (1:1000; Chemicon, Temecula, CA, USA), rabbit anti‐TH serine‐19 (1:1000; Chemicon), or mouse anti-β-actin (1:2500; Cell Signaling) diluted in blocking buffer. Membranes were incubated with HRP-conjugated anti-rabbit or anti-mouse (1:5000; Cell Signaling) and bound antibodies were visualized using SuperSignal West Femto Chemiluminescent Substrate (Thermo Scientific, Rockford, IL) and the Odyssey Fc System (LiCor, Lincoln, NE). TH and phosphorylated TH bands were normalized to the β-actin signal to correct for variations in loading. 48 2.6. Mitochondrial bioenergetics 2.6.1. XF24 assay medium RPMI-1640 (Sigma R1383-10X1L) was reconstituted in 1L-distilled water and supplemented with 2.5mM d-glucose. Assay medium was pHed to 7.4 and filter sterilized. 2.6.2. Test compounds Sodium pyruvate, malic acid disodium salt, carbonyl cyanide 4‐(trifluoromethoxy)phenylhydrazone (FCCP), and antimycin A were obtained from SigmaAldrich. Oligomycin A was obtained from Seahorse BioScience (Billerica, MA). Working concentrations of mitochondrial test compound were diluted from stock solutions in XF24 Assay Medium as described in Table 2.2. 49 Table 2.2. Dilutions of test compounds for XF24 measurements of mitochondrial bioenergetics. Compound Pyruvate Oligomycin A FCCP Antimycin A Stock Conc. 1M 2.5mM 1mM 1mM Dilution 20 227.27 41.67 76.9 Working Conc. 50mM 11 μM 24μM 13μM 50 Dilution Final Conc. 10 11 12 13 5mM 1μM 2μM 1μM 2.6.3. Extracellular flux (XF) analysis. The XF24 Extracellular Flux Analyzer (Seahorse Bioscience, Billerica, MA) is a fully integrated 24-well instrument that simultaneously measures the two major energy pathways of a cell—mitochondrial respiration and glycolysis. To examine respiration, the XF analyzer measures the rate at which oxygen (O2) is consumed from the medium. Glycolysis is quantified as the rate at which free protons are released and acidify the medium. Each assay kit contains a cell culture plate and disposable biosensor cartridge. The bottom of the biosensor cartridge contains 24 pairs of two distinct fluorophores, one sensitive to O2 and the other pH. Fiber-optic bundles insert themselves into the sleeves of the sensor cartridge, emit a light that excites the embedded fluorophores (O2= 532nm, pH=470nm), and transmits the fluorophore emission signal (O2=650nm, pH=530nm) to a set of photodetectors. Each cartridge is also equipped with four reagent delivery ports per well for dispensing testing agents at specified times during an assay. The rates of oxygen consumption (OCR) and extracellular acidification are measured within a transient microchamber created in each of the 24 wells when the biosensor cartridge is lowered 200μm above the cells. During this time, analytes are measured every 22s until a 10% change in O2 or pH is detected. The cartridge then rises, which allows the cells to restore medium to baseline. At the beginning of each experiment, the biosensors are independently calibrated using an automated routine in the machine to determine the sensor gain based on sensory 51 output measured in a calibration reagent of known pH and O2 concentration. Testing compounds are preloaded into the four injection ports and subsequently delivered pneumatically into the medium. Because XF measures are nondestructive, the real-time kinetic response of the same cell population can be measured repeatedly over time. Experiments presented in Chapters 4-5 measured mitochondrial respiration under basal and stressed conditions. These conditions were created using an XF24 Mitochondrial Stress Test (Figure 2.5). Injection ports are loaded with oligomycin A, FCCP, and antimycin A. Basal respiration is measured first and then drugs are injected sequentially to measure OCR following inhibition of the ATP synthase, uncoupling of electron transporter and ATP synthesis, and inhibition of electron transport. Figure 2.5 describes how OCR measured under these conditions can be used to calculate four parameters of mitochondrial respiration: basal respiration, ATP turnover, proton leak, and maximum or spare respiration. 52 Figure 2.5. Mitochondrial Stress Test using the XF24 Analyzer. Sequential injections of specific mitochondrial inhibitors through the drug injection ports provide measurements of the 4 fundamental parameters of mitochondrial function: Basal Respiration (blue), ATP Turnover (purple), Proton Leak (orange), and Maximum or Spare Respiration (green and yellow). Oxygen consumption rate (OCR) is first measured under basal conditions (with or without the presence of a substrate, such as glucose or pyruvate), which provides a measurement of basal respiration. Oligomycin A is then injected to inhibit the ATP synthase, which prevents the phosphorylation of ADP to ATP. As a result, OCR decreases and the difference between Basal OCR and OCR following Oligomycin A is calculated as the rate at which ATP is generated. Oligomycin A does not reduce OCR to zero, and the remaining OCR is classified as Proton Leak, or the facilitated diffusion of protons into the mitochondrial matrix through an uncoupling protein present in the inner mitochondrial membrane. Oligomycin A is followed by an injection of the uncoupler FCCP, which acts as a mobile ion carrier, transporting protons across the inner mitochondrial membrane. Because FCCP disconnects electron transport from ATP generation, proton transport occurs at a maximal rate. The difference between OCR following FCCP and Basal OCR is calculated as Spare Respiration. This represents the amount of extra ATP that can be generated by oxidative phosphorylation in the face of a sudden increase in energy demand. Antimycin A is injected to inhibit the ETC at Complex III. This completely inhibits mitochondrial respiration. Remaining OCR is considered non-mitochondrial respiration, and is subtracted from all measurements to calculate accurately the parameters of mitochondrial respiration. Adapted from seahorsebio.com. 53 2.6.4. XF bioenergetics assay PC12 cells were seeded in XF 24-well cell culture microplates (Seahorse Bioscience, Billerica, MA) coated with poly-D-lysine at a density of 7x105 cells in 100μL RPMI culture medium. The cell culture plate was incubated at 37°C/5% CO2 for 1 h before 150μL of additional culture medium was added carefully to each well and cells were incubated at 37°C/5% CO2 for 48 h. The day before the experiment 1mL of XF Calibrant solution (Seahorse Bioscience, Billerica, MA) was added to each well of the biosensor cartridge and incubated at 37°C/ no CO2 overnight. On the day of the experiment cells were treated with MeHg as described below. During cell equilibration, cartridge injection ports were loaded with testing compounds at working concentrations (Table 2.2), and then the biosensor cartridge was placed in the XF24 Flux Analyzer for automated calibration. For the XF assay, the cell culture plate was placed in the calibrated XF24 Flux Analyzer. Baseline OCR (nM/min) and ECAR (milli-pH (mpH) units/min) were measured simultaneously 3 times. Between each rate measurement, assay medium was gently mixed by the XF24 Analyzer for 3-5 min to restore normal O2 tension and pH in the microchamber surrounding the cells. Following baseline measurement, Ports A-D were sequentially injected (75μL) to reach testing compound final concentrations and the 54 medium was mixed for 3-5 min before OCR and ECAR were measured 3 times, as described for baseline measurements. 2.6.5. XF24 drug exposure At the time of treatment, complete medium was aspirated and replaced with HBS or HBS containing 2μM MeHg, 50nM rotenone, or 20μM daidzin. In some experiments, cells were treated with 2μM MeHg in 0-Ca2+ HBS. Cells were incubated at 37°C/ 5% CO2 for 60 min. To terminate drug exposure, treatment medium was aspirated, and cells were prepared for the XF Analyzer measurements. First, cells were rinsed with 1mL warmed XF24 assay medium, and then XF24 assay medium was brought to a final volume of 675μL. Cells were incubated at 37°C/no CO2 for 1 hr to allow medium temperature and pH to equilibrate before the first measurements were made. 2.6.6. XF24 optimization of experimental conditions. In order to measure accurately OCR as described in Methods Section 2.6.4, certain parameters were first optimized. These included cell plating density (Figure 2.6), assay medium composition (Figure 2.7), and FCCP concentration (Figure 2.8). Because the O2 biosensor takes a measurement from one small location within each well, plating densities that are too low or too high result in a variable distribution of cells along the bottom of the well and result in an inaccurate measurement of OCR. For this reason cells were seeded at densities between 40,000-90,000 cells/well and OCR was measured during a Mitochondrial Stress Test. As shown in Figure 2.6, the optimal density 55 was ~70,000 cells/well. At lower densities, basal OCR was too low and the maximum respiratory capacity was not pronounced. Although high densities (80,000-90,000 cells/well) had good basal OCR and a pronounced response to FCCP, there was some indication that once oxygen levels were depleted and measurements were terminated (as normal), oxygen levels were not able to return to baseline. This did not happen at 70,000 cells/well, so this density was chosen for future experiments. Seahorse Biosciences provided best practices and basic protocol instructions for using their equipment with immortalized cell lines. One company suggestion is the use of their proprietary Dulbecco’s Modified Eagle Medium (DMEM) for all XF24 assays. However PC12 cells are cultured in RPMI 1640 medium for growth, which contains different types and concentrations of amino acids, vitamins, and inorganic salts. Changing the medium just prior to intracellular measurements of mitochondrial respiration could dramatically alter cellular and respiratory activity. To determine which assay medium provided the most ideal experimental conditions, RPMI and DMEM assay media were compared (Figure 2.7). In this experiment respiration substrates were also compared. Some RPMI medium was prepared with glucose as a substrate for the ETC, and some RPMI medium was prepared with galactose. Galactose was investigated as a potential substrate because PC12 cells have a very slow doubling time (~92 hrs) (Greene and Tischler, 1976). This suggests that the cells normally do not rely on oxidative phosphorylation, but instead are adapted to generate energy from glycolysis. This has been demonstrated to occur in many immortalized cell lines, and is referred to as the Crabtree effect (Rodríguez-Enríquez et al., 2001). It is possible to stimulate respiration in cancer cell lines by swapping glucose in the medium for galactose (Rossignol, 2004; Marroquin et al., 2007). Oxidation of galactose to 56 pyruvate by way of glycolysis yields no net ATP, which forces the cell to use oxidative phosphorylation to generate energy needed for survival. As shown in Figure 2.7, the optimal medium condition was RPMI 1640 medium with 2.5mM glucose present. Under these conditions, basal respiration was highest and FCCP produced a maximal response. The last optimization experiment performed was an FCCP titration. The optimal concentration of this uncoupler must be determined to ensure maximum stimulation of uncoupled respiration without inhibiting respiration by using a concentration that is too high. Figure 2.8 illustrates the results of this titration; 2μM FCCP was chosen for future assays because this concentration maximally stimulated OCR without negatively altering oxygen consumption. 57 Figure 2.6. XF24 cell density optimization. Undifferentiated PC12 cells were plated at 40,000 (red), 50,000 (orange), 60,000 (light green), 70,000 (dark green), 80,000 (teal), or 90,000 (blue) cells/well. Bioenergetics were assessed using the Mitochondrial Stress Test: A) 50mM pyruvate, B) 1μM oligomycin A, C) 1μM FCCP, and D) 1μM antimycin A. Based on these data, a plating density of 70,000 cells/well was selected for future assays because basal OCR was high and cells responded well to FCCP with adequate oxygen levels within each well. These observations suggest that cells were plated at a density that was high enough to allow cell-cell interaction, but not too high as to result in cells growing on top of each other. 58 Figure 2.7. XF24 assay medium optimization. PC12 cells were seeded at a density of 70,000 cells/well and bioenergetics were assessed using the Mitochondrial Stress Test: A) 50mM pyruvate, B) 1μM oligomycin A, C) 1μM FCCP, and D) 1μM antimycin A. The assay was performed in either DMEM Assay Medium (red), RPMI 1640 Assay Medium (orange), RPMI 1640 + 2.5mM glucose (light green), or RPMI 1640 + 2.5mM galactose (dark green). Results demonstrate that RPMI 1640 + 2.5mM glucose was optimal. 59 Figure 2.8. XF24 FCCP titration. PC12 cells were seeded at a density of 70,000 cells/well and bioenergetics were assessed using the Mitochondrial Stress Test: A) 50mM pyruvate, B) 1μM oligomycin A, and D) 1μM antimycin A. In port C FCCP was injected at concentrations of 0μM (red), 0.25μM (orange), 0.5μM (light green), 1μM (dark green), or 2μM (teal). Because 2μM FCCP produced maximum respiration without negatively altering oxygen consumption, this concentration was chosen for future experiments. 60 2.7. ALDH enzyme kinetics assay PC12 cells were seeded in 6-well plates coated with poly-D-lysine at a density of 7x105 cells/mL and cultured for 24-48 h at 37°C/5% CO2. At the time of treatment, growth medium was aspirated, replaced with HBS or HBS containing 2μM MeHg, 50nM rotenone, or 20μM daidzin. Cell were incubated for 60 min with MeHg, 45 min with daidzin, or 15 min with rotenone at 37°C/5% CO2. Following treatment, plates were centrifuged at 300 g for 5 min at 4°C. Treatment medium was aspirated and replaced with 150μL homogenization buffer (0.1M Tris-HCl, 10mM DTT, 20% glycerol, and 1% Triton-X). Cells were scraped and then pelleted by centrifugation at 13,000 g for 8 min at 4˚C. Protein concentration of supernatant was determined using the BCA protein assay (Sigma). ALDH enzyme activity was determined spectrophotometrically by monitoring the reductive reaction of NAD to NADH at 340 nm using the Tecan Infinite® M1000 Pro microplate reader and Magellan data analysis software (Tecan Systems, Inc., San Jose, CA). Assays were carried out in 96-well plates containing 100mM Tris-HCl buffer (pH 8.0) at 30˚C. Two mM nicotinamide adenosine dinucleotide hydrate (NAD+; Sigma) and 100µg PC12 cell lysate were added to the buffer. In some experiments, 2μM MeHg, 50nM rotenone, or 20μM daidzin was added to the assay wells. To start the reaction, 10mM propionaldehyde (Sigma) was added. Accumulation of NADH was recorded every 30 sec for 10 min. Substrate blanks (NAD + lysate) were run simultaneously and results were corrected for blank reactions. Km was calculated using SigmaPlot 12.0 (SysStat Software, Inc., Point Richmond, CA, USA) and Michaelis–Menten kinetics were plotted to determine 61 the optimal concentration of substrate for future reactions (Figure 2.9). ALDH reaction rates were calculated as µmole NADH per min per mg protein. 62 Figure 2.9. ALDH assay substrate optimization. In ALDH assays, propionaldehyde was used as the substrate. The Km (concentration at which the reaction rate is half Vmax) for propionaldehyde was determined. At concentration above 40mM, propionaldehyde inhibited ALDH activity. Vmax = 2.1μmol/min/mg protein and Km = 6.4mM propionaldehyde. Based on these results, 10mM propionaldehyde was selected for future experiments. 63 2.8. NADH/NAD quantification PC12 cells were seeded in 6-well plates at 7 x 105 cells/mL and incubated at 37°C/5% CO2 for 48 hr. At the time of treatment culture medium was aspirated and replaced with HBS for 60 min, 2μM MeHg in HBS for 60 min, or 50nM rotenone for 15 min. Following pharmacological exposure, treatment medium was aspirated, cells were washed once with cold PBS, and then pelleted by centrifugation at 10,000 g for 5 min at 4°C. NADH/NAD was extracted using 400µLNADH/NAD Extraction Buffer (Sigma) by two freeze/thaw cycles (20 min on dry ice followed by 10 min at room temperature). Samples were then vortexed for 10 sec and centrifuged for 10 min at 13,000 g to remove insoluble material. To quantify NAD, samples were measured as described below. For NADH measurements, 200µLwas incubated at 60°C for 30 min before measurement. A 96-well plate was loaded with 50µLsample (Figure 2.10) and 100µLMaster Reaction Mix (NAD Cycling Buffer + NAD Cycling Enzyme; Sigma). Samples were incubated for 5 min to convert NAD to NADH, and then 10µLNADH Developer was added to each well. Samples were incubated for 1 hr at room temperature. Absorbance at 450nm was measured using spectrophotometry and the Tecan Infinite® M1000 Pro microplate reader with Magellan data analysis software (Tecan Systems, Inc., San Jose, CA). Average absorbance was compared to NADH standards to calculate the concentration of total NAD, total NADH, and the ratio of NAD:NADH. 64 Figure 2.10. Sample volume titration for NADH/NAD quantification. NAD Cycling Enzyme was reacted with 0, 10, 20, 30, 40, of 50µL cell lysate, and total NAD was quantified. Based on this optimization experiment, 50μg lysate was selected as the volume for experimental analysis. 65 2.9. Measurement of fura-2 fluorescent change Changes in the concentration of intracellular Ca2+ were monitored as changes in fluorescence of the ratiometric Ca2+-indicating dye fura-2 using microfluorometric imaging. The perfusion buffer used in these experiments was always 0-Ca2+ HBS. PC12 cells were plated at 7x105 cells/mL on the upper surface of sterilized 25-mm diameter circular glass coverslips (Carolina Biological; Burlington, NC) in Petri dishes 24 hr prior to imaging. Cells were loaded with 2μM fura-2 and 2μM pluronic acid in RPMI culture medium and incubated in the absence or presence of 50μM BAPTA/AM for 30 min at 37°C. After loading, the circular coverslip containing fura-2 loaded cells was mounted on a inverted microscope (Diaphot-TMD, Nikon, Toyko, Japan) and cells were perfused with 0-Ca2+ HBS containing 2μM MeHg at 37°C. Digital fluorescent images were obtained using an IonOptix system (Milton, MA). Fura-2 fluorescence changes were monitored simultaneously in the soma of 10 cells within the same microscopic field. Data from these cells was used to visually analyze the time-to-onset of MeHg-induced changes in intracellular Ca2+. These experiments were conducted to establish BAPTA/AM was chelating intracellular Ca2+. Representative soma were selected to illustrate effects of 0-Ca2+ HBS and BAPTA/AM on MeHg-induced changes in intracellular Ca2+. 66 2.10. Statistical analyses SigmaPlot® software version 12.0 (SysStat Software, Inc., Point Richmond, CA) was used to make statistical comparisons among groups using unpaired t-test, one-, two-way ANOVA, or non-parametric alternatives as appropriate. If a significant difference was detected, post hoc analysis was followed by between-group comparisons using Tukey’s test. Differences with a probability of error of less than 5% were considered statistically significant (p<0.05). 67 Chapter 3. The role of de novo catecholamine synthesis in mediating MeHg-induced vesicular DA release 3.1. Introduction Release of DA occurs predominantly by means of canonical Ca2+-dependent vesicular exocytosis. In PC12 cells, DA is stored in large dense-core vesicles concentrated near the plasma membrane (Bauerfeind et al., 1993; Fornai et al., 2007), and released following cellular depolarization (Greene and Rein, 1977a; Kittner et al., 1987). Released DA is primarily recaptured by the NET in undifferentiated PC12 cells (Zhu and Hexum, 1992; Brüss et al., 1997). However certain cellular conditions, such as exposure to psychostimulant drugs (Sulzer et al., 1995), can reverse the transporter resulting in DA leakage into the synaptic cleft (Leviel, 2011). DA release is, in part, regulated by DA synthesis. Once synthesized, DA inhibits TH activity by binding the catalytic site of the enzyme, and also by interfering with the interaction between TH and its cofactor, tetrahydrobiopterin (Kumer and Vrana, 1996; Dunkley et al., 2004). This end-product feedback inhibition decreases the rate of DA synthesis, and thus down-regulates DA release. Conversely, TH phosphorylation at critical Ser residues by a variety of protein kinases increases enzymatic activity (Daubner et al., 2011). Phosphorylation alters the conformation of DA-inhibited TH, which dissociates catecholamine from enzyme, and allows TH to return to its active state. The most important site of phosphorylation is at Ser residue 40 (Ser40), which directly increases TH activity (Daubner et al., 1992; Dunkley et al., 2004). Phosphorylation of other serine 68 resides, namely Ser19 and Ser31, serves to stabilize phosphorylation at Ser40 (pTH Ser40) (Royo et al., 2005). MeHg stimulates spontaneous neurotransmitter release, but impairs depolarizationevoked transmitter release (Atchison and Narahashi, 1982; Atchison et al., 1984). This effect has been demonstrated in many neurotransmitter systems, including the dopaminergic system (Minnema et al., 1989). MeHg increases spontaneous DA release in a concentration-dependent manner through a presynaptic action (Kalisch and Racz, 1996; Faro et al., 2002; Dreiem et al., 2009; Tiernan et al., 2013). Results of studies investigating the mechanisms underlying the stimulatory effects of MeHg on dopaminergic neuronal transmission have been inconsistent with respect to the underlying mechanisms responsible for MeHg-induced DA release. A role for DAT reversal has been proposed based on the finding that DAT inhibition or stimulation of transporter-mediated release produces an effect similar to that of MeHg alone (Faro et al., 2002). However, MeHg does not target DAT to induce release (Gassó et al., 2000), and may decrease transporter reuptake activity (Bonnet et al., 1994; Dreiem et al., 2009). A role for vesicular exocytosis in MeHg-induced catecholamine release has been documented (Gassó et al., 2000), however non-vesicular DA release has also been described following MeHg exposure (Kalisch and Racz, 1996; Faro et al., 2002). Despite the coupling between neurotransmitter release and synthesis, no reports to date have investigated a potential role for DA synthesis in MeHg-mediated release. Therefore the goals of the present study were to: 1) identify pathways by which MeHg induces spontaneous DA release, differentiating between transporter-mediated and non- 69 transporter mediated release, and 2) examine the coupling between DA synthesis and MeHg-induced DA release. 3.2. Methods 3.2.1. Chemicals MeHg (ICN Biochemicals), reserpine (Sigma), desipramine (DMI; Sigma), αmethyltyrosine (AMT; Sigma), and 3-hydroxybenzylhydrazine (NSD-1015; Sigma) were diluted from stock solutions in HBS. Drug solutions were diluted to final concentrations on the day of each experiment as described in General Methods Section 2.2. 3.2.2. Chemical treatment PC12 cells were seeded at a density of 6x105 cells/mL in 6-well plates coated with poly-D-lysine 48 hr prior to treatment. To begin each experiment, culture medium was aspirated and replaced with HBS or HBS containing pharmacological treatment (with the exception of 24hr AMT pretreatment, which was prepared in RPMI-1640 culture medium). Cells were incubated at 37°C/5% CO2 during treatment as described in General Methods Section 2.2. 3.2.3. Neurochemistry At experiment termination, treatment medium was retained and acidified. Cells were rinsed, harvested, and pelleted by centrifugation. The content of DA and its precursor DOPA in the treatment medium and cell pellet were determined using HPLC-ED as 70 described in General Methods Section 2.3. Neurochemical concentrations were normalized to mL per sample for extracellular measurements or mg of protein for intracellular measurements as determined using the BCA protein assay (Sigma). The rate constant was calculated from intracellular DA concentrations after 90 min AMT treatment as described in General Methods Section 2.3.1. The slope of the lines representing the difference between cells treated with 2 µM MeHg alone or combined with 300 µM AMT was calculated, and then compared to HBS-treated cells in the absence or presence of AMT. 3.2.4. Western blot analysis At experiment termination, treatment medium was collected and analyzed as a positive control. Cells were rinsed once with 1 mL ice-cold phosphate buffered saline, harvested, and pelleted by centrifugation at 12,000 g for 5 min at 4°C. Pellets were lysed with 50µLradio-immunoprecipitation assay (RIPA) buffer containing protease inhibitor (1:100) and phosphatase inhibitor (1:100). Protein content was determined using the BCA protein assay (Sigma). Supernatant was prepared for Western blot analysis to quantify total and phosphorylated isoforms of TH as described in General Methods Section 2.5. TH and pTH Ser40 bands were normalized to the β-actin signal to correct for variations in loading. 3.2.5. Cell viability PC12 cells were seeded in 96-well plates coated with poly-D-lysine at a density of 4x105 cells/mL 48 hr prior to treatment with HBS, 1, 2, or 5 µM MeHg for 15, 30, 60 or 120 71 min. Hoechst 33342 (10 mg/mL) and propidium iodide (1 mg/mL) were diluted 1:10,000 in existing treatment medium. Cells were incubated in fluorophores at 37˚C for 15 min prior to visualization. Image acquisition was as described in General Methods Section 2.4. Viable cells had a blue-stained (Hoechst) nucleus, whereas non-viable cells had a high intensity red (propidium iodide) fluorescence (Yuan et al., 2009). 3.2.6. Statistical analysis SigmaPlot® software version 12.0 (SysStat Software, Inc., Point Richmond, CA) was used to make statistical comparisons among groups using unpaired t-test, one-, two-way ANOVA, or non-parametric alternatives as appropriate. If a significant difference was detected, post hoc between-group comparisons were performed using Tukey’s test. Statistically significance was set at p<0.05. 3.3. Results 3.3.1. MeHg increases spontaneous DA release in a concentration- and time-dependent manner Measurements of DA in the medium reflect the balance between changes in DA release and transporter-mediated reuptake. DA was not detected in treatment medium in the absence of cells and any subsequent treatment-induced change in medium DA was, therefore, due to cellular DA release. In the absence of MeHg, the concentration of extracellular DA stabilized within the first 15 min, and remained at a steady state throughout the 120 min sampling period (Figure 3.1). MeHg caused both a concentrationand time-dependent increase in medium DA. At 1 µM, MeHg did not significantly alter 72 extracellular DA accumulation, whereas 2 µM and 5 µM MeHg significantly increased the concentration of extracellular DA by 60 min and 30 min, respectively. These elevated levels were maintained for the duration of the experiment. The significant increase in extracellular DA concentrations induced by 5 µM MeHg at 60 and 120 min was associated with a significant incidence of cytotoxicity in a parallel set of cultures (Figures 3.2-3.3). Because 2 µM MeHg induced a significant increase in DA release by 60 min without inducing significant levels of cytotoxicity, this concentration and time point was selected for further analysis of release mechanisms. 73 Figure 3.1. Concentration-response and time-course effects of MeHg on extracellular DA concentrations. Undifferentiated PC12 cells were treated with 0 µM (black circles), 1 µM (white circles), 2 µM (black triangles) or 5 µM (white triangles) MeHg in HBS for 15, 30, 60, or 120 min. The asterisk (*) indicates a value significantly different than 0 µM MeHg within a given time point (p<0.05). Values are means + S.E.M. (n = 3-4, 3 replicates per n). 74 Figure 3.2. Concentration-response and time-course effects of MeHg on cell viability. Undifferentiated PC12 cells were exposed to 0µM (black circles), 1µM (white circles), 2µM (black triangles), or 5µM (white triangles) MeHg in HBS for 15, 30, 60, or 120 min. Following toxicant treatment, cultures were stained for viable (Hoechst 33342) and nonviable (propidium iodide) cells. The number of viable and non-viable cells was counted and expressed as percent live cells. The asterisk (*) indicates a value significantly different than 0 µM MeHg within a given time point (p<0.05). Values are means + S.E.M (n = 3, 3 replicates per n). 75 A B Figure 3.3. Representative images of cell viability. Undifferentiated PC12 cells were exposed to 0, 1, 2, or 5 µM MeHg in HBS for 15 (Panel A), 30 (Panel B), 60 (Panel C), or 120 (Panel D) min. Viable cells have a blue-stained (Hoechst) nucleus, whereas non-viable cells have an intense red (propidium iodide) fluorescence. Scale bar = 200 µm 76 Figure 3.3 cont’d C D 77 3.3.2. Spontaneous MeHg-mediated DA release requires a functional vesicular, but not membrane, transporter To investigate the contributions of vesicular release and reuptake transporter activity in spontaneous MeHg-mediated DA release from PC12 cells, reserpine was used to inhibit the VMAT (Schuldiner et al., 1993) and DMI was used to block the NET (Brüss et al., 1997). Reserpine (50 nM) decreased spontaneous basal release and blocked the significant increase in extracellular DA induced by MeHg. In contrast, DMI had no significant effect on extracellular DA concentration (p>0.05), but significantly augmented MeHg-induced DA release (p<0.05; Figure 3.4A). However, while reserpine significantly attenuated the percentage change in extracellular DA induced by MeHg, DMI did not (p>0.05, Figure 3.4B). 78 Figure 3.4. Effects of DA storage depletion and reuptake transporter inhibition on MeHg-induced DA release. Panel A) Concentrations of extracellular DA from undifferentiated PC12 cells preincubated with 50 nM reserpine for 60 min, 1 µM DMI for 15 min, or vehicle in culture medium prior to co-treatment with 0 µM (white bars) or 2 µM (black bars) MeHg in HBS for 60 min. Panel B) The percentage change of extracellular DA after treatment with MeHg in cells treated with HBS, 50 nM reserpine, or 1 µM DMI. The asterisk (*) indicates a value significantly different than HBS (p ≤ 0.05). The hash (#) indicates a value significantly different than 2 µM MeHg-HBS (p ≤ 0.05). Values are means + S.E.M. (n = 3, 3 replicates per n). 79 3.3.3. MeHg-mediated DA release is associated with increased intracellular DA concentrations and utilization of DA stores Spontaneous DA release induced by MeHg was associated with a significant increase in the concentration of intracellular DA (Figure 3.5). To determine if increased DA stores correlated with increased release, DA storage utilization was assessed using AMT (Brodie et al., 1966). The rate constant of decline of intracellular DA was significantly greater in MeHg-treated cells as compared to HBS-treated cells (p<0.05), demonstrating that utilization of DA stores was significantly increased following MeHg exposure (Figure 3.6). 80 Figure 3.5. Effect of MeHg on intracellular DA concentrations. Undifferentiated PC12 cells were treated with either 0 µM (white bars) or 2 µM (black bars) MeHg in HBS for 60 min. The asterisk (*) indicates a significant difference from 0 µM MeHg (p ≤ 0.05). Values are means + S.E.M. (n = 3, 3 replicates per n). 81 Figure 3.6. Effect of MeHg on DA storage utilization. Undifferentiated PC12 cells were preincubated with 300 µM AMT or vehicle in culture medium for 30 min prior to treatment with either 0 µM (white circles) or 2 µM (black circles) MeHg in HBS in the absence or presence of 300 µM AMT for 60 min. The rate constant (k) was calculated as the slope of the line between 0 and 90 min after AMT. The asterisk (*) indicates a value significantly different than 0 µM MeHg within a given time point (p ≤ 0.05). The hash (#) indicates a significant difference between rate constant values (p ≤ 0.05). Values are means ± S.E.M. (n = 3, 3 replicates per n). 82 3.3.4. MeHg-mediated DA release is associated with de novo DA synthesis and increased TH enzymatic activity To examine the role of de novo DA synthesis in MeHg-mediated DA release, PC12 cells were treated with either the competitive TH inhibitor AMT (300 μM) for 24 hr or the inhibitor of DOPA decarboxylase NSD-1015 (10 μM) for 60 min. Complete inhibition of DA synthesis with AMT abolished the MeHg-mediated increase in extracellular DA concentrations, but did not alter basal DA release (Figure 3.7). The DA precursor DOPA was not detectable except in the presence of NSD-1015, after which its concentration was proportional to the rate of TH enzymatic activity. MeHg significantly increased the concentration of intracellular DOPA as compared to untreated cells (Figure 3.8). Western blot analysis of total and phosphorylated TH protein revealed that MeHg significantly increased the amount of pTH Ser40, without altering total TH protein (Figure 3.9). 83 Figure 3.7. Effect of TH inhibition on MeHg-induced DA release. Undifferentiated PC12 cells were preincubated with 300 µM AMT or vehicle in culture medium for 24 hr prior to co-treatment with either 0 µM (white bars) or 2 µM (black bars) MeHg in HBS for 60 min. The asterisk (*) indicates a value significantly different than 0 µM MeHg-HBS (p ≤ 0.05). Values are means + S.E.M. (n = 3, 3 replicates per n). 84 Figure 3.8. Effect of DOPA decarboxylase inhibition on TH enzymatic activity. Undifferentiated PC12 cells were treated with 0 µM (white bars) or 2 µM (black bars) MeHg in HBS for 60 min, in the presence of 10 µM NSD-1015. DOPA accumulation was quantified using HPLC-ED as an index of TH enzymatic activity. The asterisk (*) indicates a significant difference between groups (p ≤ 0.05). Values are means + S.E.M. (n = 4, 3 replicates per n). 85 Figure 3.9. Effect of MeHg on TH expression. Panel A) Undifferentiated PC12 cells were treated with 0 µM (white bars) or 2 µM (black bars) MeHg in HBS for 60 min. Amounts of total TH and pTH Ser40 were quantified by Western blot and normalized to the level of expression of the housekeeping protein β-actin. The asterisk (*) indicates a significant difference between groups (p ≤ 0.05). Values are means + S.E.M. (n = 3, 3 replicates per n). Panel B) Representative Western blot images for Total TH, pTH Ser40, and β-actin in cells treated with 0 µM or 2 µM MeHg. 86 3.4. Discussion The present study contributes significantly to our understanding of mechanisms by which MeHg induces spontaneous DA release, in particular its interaction with the coupling between synthesis and vesicular exocytosis. These data are consistent with the following conclusions: 1) MeHg causes a concentration- and time-dependent increase in DA release from undifferentiated PC12 cells; 2) vesicular exocytosis, but not reuptake transporter activity, is responsible for both basal and MeHg-induced DA release; and 3) MeHg-induced DA release is dependent upon de novo DA synthesis and accelerated TH activity. 3.4.1. MeHg-induced DA release Under basal conditions, DA release from PC12 cells is maintained at a steady state. During the first 15 min DA concentrations increased from 0 ng/mL to ~3 ng/mL, and then did not fluctuate significantly for the remainder of the experiment. The consistent level of extracellular DA in the medium likely reflects coupling between release and reuptake under basal conditions (Schmitz et al., 2003). Increased DA release after exposure to MeHg has been well documented in other systems, including 3H-DA release from rat striatum (Minnema et al., 1989) and endogenous DA release from striatal synaptosomes (Dreiem et al., 2009), mouse striatal slices (Kalisch and Racz, 1996), and the striatum of conscious, free-moving rats (Faro et al., 2002). Results from the present study replicate these observations and demonstrate that MeHg-induced DA release is both concentration- and time-dependent. Low concentrations of MeHg (1 μM) do not alter DA release, however higher concentrations (2-5 μM) increase release. Furthermore, 5 μM MeHg increases DA release more rapidly than did 2 μM. 87 The increase in extracellular DA after exposure to 5 μM MeHg at 60 and 120 min was substantially higher than that observed at lower concentrations, but was more variable, and associated with an increased incidence of cell death. Exposure to 5 μM MeHg has previously been associated with rapid impairment of mitochondrial activity and membrane lysis consistent with necrotic cell death (Castoldi et al., 2000). Therefore it is hypothesized that as PC12 cells lyse, DA would be purged from the cell and increase its extracellular concentration above that expected in intact cells. DA released upon cell lysis would account for the sizable increase in extracellular DA and associated high variability. On the other hand, 2 μM MeHg increased DA release by 60 min, an effect not associated with cell death. The increase in DA release in the absence of cell death demonstrates that MeHg-induced DA release is associated with intracellular mechanisms unrelated to cell lysis. Ca2+ plays an essential role in neurotransmitter release and Ca2+-dependent neurotransmitter release is a known target of MeHg toxicity (Atchison and Narahashi, 1982; Atchison, 1986; Yuan and Atchison, 2007). Therefore MeHg-induced Ca2+ signaling could be a primary site of action linking MeHg exposure to increased DA release. 3.4.2. Vesicular transport, but not membrane transport contributes to MeHg-induced DA release Vesicular exocytosis and transporter-mediated reuptake are essential components of catecholamine release and recycling in PC12 cells (Lorang et al., 1994; Kishimoto et al., 2005; Fornai et al., 2007). In the present study, depletion of storage vesicles decreased basal release, confirming a role for vesicular stores in DA release from undifferentiated 88 PC12 cells. NET inhibition caused a small, although non-significant, increase in extracellular DA, likely due to inhibition of re-uptake of released DA. Depletion of DA stores with reserpine abolishes MeHg-induced DA release. These data suggest that mobilization of vesicular DA stores contributes to the stimulatory effect of MeHg on DA release. A role for vesicular exocytosis in MeHg-induced catecholamine release is consistent with reports in the literature demonstrating that reserpine pretreatment blocks MeHg-induced 3H-NE release from hippocampal slices (Gassó et al., 2000). In the present study inhibition of the PC12 DA transporter NET increased DA release above that induced by MeHg alone. However the percentage change in extracellular DA after MeHg was comparable in control and DMI groups. This suggests that NET is functionally active, and does not contribute to the rise in extracellular DA following exposure to MeHg. In contrast, DAT transporter activity is reportedly decreased following exposure to MeHg (Bonnet et al., 1994; Dreiem et al., 2009), due to either a direct interaction between MeHg and the thiol group on DAT (Bonnet et al., 1994), or indirectly through inhibition of the Na+/K+ ATPase (Berg and Miles, 1979) or reduction of ATP needed to maintain the ion gradient used by the transporter (Torres et al., 2003; Gatti et al., 2004). In the present study it is possible that reuptake by the NET was partially inhibited by MeHg because addition of DMI to MeHg produced a larger increase in extracellular DA than either MeHg or DMI treatment alone. 89 3.4.3. Evidence for a role of de novo DA synthesis in MeHg-induced DA release MeHg-induced DA release was accompanied by an increase in intracellular DA stores and acceleration of DA storage utilization. Increased DA stores could result from either stimulated DA synthesis and/or impaired DA metabolism. MeHg impairs DA metabolism; concentrations of the DA metabolites DOPAC and homovanillic acid are significantly decreased following exposure to MeHg (Chapter 4) (Dreiem et al., 2009). However, the increased rate of storage utilization following MeHg exposure also suggests a role for DA synthesis. Newly synthesized DA is preferentially released (Kopin, 1968), thus MeHg could also accelerate DA synthesis to increase DA release. To elucidate the contribution of the DA biosynthetic pathway to the effects of MeHg, two pharmacological manipulations were employed. Inhibition of TH with AMT completely abolished MeHg-induced DA release. Indirect assessment of TH activity using NSD-1015 to block conversion of DOPA to DA suggests that MeHg stimulated TH activity; i.e., DOPA accumulation was increased following toxicant treatment. Western blot analysis confirmed that MeHg targets TH to increase DA synthesis. This effect is achieved by increasing phosphorylation of TH at Ser40, rather than altering the overall amount of enzyme present. These data demonstrates that MeHg targets TH activity, increasing the amount of newly synthesized DA available for release. The mechanism by which MeHg targets TH phosphorylation to increase DA synthesis is unknown. One possible explanation is related to increased cytosolic Ca2+ following exposure to MeHg (Marty and Atchison, 1997). TH phosphorylation is mediated by Ca2+-dependent protein kinases, including PKC (Albert et al., 1984; Vulliet et al., 1985) 90 and CaM-MPK (Yamauchi and Fujisawa, 1981; Vulliet et al., 1984). Activation of these phosphorylation pathways may be responsible for the observed increase in TH activity following exposure to MeHg. However, increased cytosolic Ca2+ could also inhibit the activity of protein phosphastase 2A or 2C (Bevilaqua et al., 2003), preventing TH dephosphorylation. This is the first report demonstrating a stimulatory effect of acute MeHg exposure on TH activity that culminates in an increase in DA synthesis and consequently DA release. A previous report documented elevated TH activity in brain homogenates isolated from rats treated with MeHg for seven days (Omata et al., 1982), however the downstream consequences of MeHg-activated TH activity on catecholamine release were not studied. Results presented herein describe how MeHg targeting of the DA biosynthetic pathway contribute to MeHg-mediated DA release. This could be one mechanism by which MeHg induces cell-specific neurotoxicity. Decoupling of the regulation between biosynthesis and release, leading to an abnormally high level of extracellular DA, could disrupt the signal-tonoise ratio associated with normal neuronal activity-evoked release at the DA synapse. 91 Chapter 4. The roles of mitochondria and associated catabolic enzyme function in MeHg-impaired DA metabolism 4.1. Introduction DA is metabolized in a two-step process. First, MAO catalyzes the oxidative deamination of DA to DOPAL. Second, this reactive intermediate metabolite either undergoes further oxidation mediated by ALDH to form the carboxylic acid DOPAC, or it is reduced by aldehyde/aldose reductase (AR) to form the alcohol DOPET (Eisenhofer, 2004; Meiser et al., 2013). These oxidative and reductive pathways of DA metabolism are illustrated in Figure 4.1. DA is predominately degraded along the oxidative metabolic pathway due to the absence of a β-hydroxyl group. This is contrast to the other catecholamines, NE and epinephrine, which contain a β-hydroxyl group, and are preferentially degraded along the reductive pathway (Duncan and Sourkes, 1974; Turner et al., 1974; Kawamura et al., 2002). Tight regulation of DA metabolism is necessary because both cytosolic DA and DOPAL can be cytotoxic (Graham, 1978; Graham et al., 1978; Rees et al., 2009). Cytosolic DA undergoes autooxidation in the presence of molecular oxygen to form either a quinone or semiquinone (Graham, 1978; Kalyanaraman et al., 1985; Lévay and Bodell, 1993). These reactive species can covalently bind cellular proteins and DNA (Graham et al., 1978). Similarly, DOPAL has been reported to form adducts with protein amino acids, and to cross-link proteins and DNA (Nair et al., 1986; Nilsson and Tottmar, 1987; Nadkarni and Sayre, 1995; Brooks and Theruvathu, 2005; Rees et al., 2009). Subsequent biological effects of DA and aldehyde adducts could contribute to inactivation of enzymes (Chio and Tappel, 92 1969), depletion of antioxidant systems (Esterbauer et al., 1975), and altered signal transduction (Leonarduzzi et al., 2004), gene expression (Kumagai et al., 2000), and DNA repair (Feng et al., 2004). Indeed, the specific vulnerability and death of DA neurons in PD is hypothesized to involve the accumulation of these toxic compounds that are selectively produced in DA neurons (Li et al., 2001; Burke et al., 2003). Dysfunction of oxidative DA metabolism at the level of either MAO or ALDH could contribute to oxidative stress and cytotoxicity associated with DA autooxidation and DOPAL adduct formation. MAO is localized on the outer mitochondrial membrane (Schnaitman et al., 1967), and is covalently bound to a flavin adenine dinucleotide molecule, which is used as a cofactor (Youdim et al., 2006). MAO catalyzes the oxidative deamination of a number of biogenic amines, including DA, NE, epinephrine, and serotonin (Shih et al., 1999; Nagatsu, 2004). In the case of DA, the reaction produces the corresponding aldehyde (i.e., DOPAL), hydrogen peroxide, and ammonia. The ALDH superfamily includes enzymes that catalyze the NAD(P)+-dependent oxidation of a wide variety of aldehydes into their corresponding carboxylic acids. There are 19 known human ALDH proteins found in all subcellular regions, including the cytosol, mitochondria, endoplasmic reticulum, and nucleus (Marchitti et al., 2008). While ALDH enzymes are distributed widely throughout different tissues in the body, they display distinct substrate selectivity. In DA metabolism, two isozymes, ALDH1A1 and ALDH2, have demonstrated roles. ALDH1A1 is highly expressed in the cytosol of DA neurons (Galter et al., 2003; Anderson et al., 2011b), and its gene expression is down-regulated in PD patients (Mandel et al., 2005). ALDH2 is a mitochondrial matrix protein constitutively expressed in a variety of tissues and cell types, including the brain (Goedde and Agarwal, 1990) and 93 PC12 cells (Robador et al., 2012). Both isozymes are thought to play critical roles in maintaining low intracellular levels of DOPAL (Maring et al., 1985; Galter et al., 2003; Marchitti et al., 2007). MeHg has demonstrated effects on DA metabolism. The toxicant decreases the concentration of intracellular DOPAC (Bemis and Seegal, 1999; Dreiem et al., 2009), which may result from an observed decrease in MAO activity (Taylor and DiStefano, 1976; Chakrabarti et al., 1998; Beyrouty et al., 2006; Castoldi et al., 2006). MeHg could inhibit MAO by two potential mechanisms. Because MAO is localized to the outer mitochondrial membrane (Schnaitman et al., 1967), MeHg-induced loss of mitochondrial membrane potential (Bondy and McKee, 1991; Hare and Atchison, 1992; Limke and Atchison, 2002) could compromise enzyme activity. Additionally, several cysteine residues are required for proper MAO enzymatic function (Hubalek et al., 2003). MeHg has a high affinity for sulfhydryl groups (Hughes, 1957; Sanfeliu et al., 2003), and could bind these cysteine residue to impair MAO activity. MeHg targeting of mitochondrial function and protein thiols to impair MAO activity might also contribute to inhibition of ALDH function. Mitochondrial-associated ALDH mediates the catabolism of DOPAL to DOPAC (Marchitti et al., 2007), and thus mitochondrial dysfunction induced by MeHg could impair ALDH enzymatic activity. Furthermore, the active site of ALDH contains two cysteine residues (Hempel et al., 1985), which MeHg could readily bind to inhibit enzymatic activity. Impaired MAO or ALDH activity would be indicated by changes in the metabolomic profile of DA. Alterations in the ratios of DOPAC, DOPAL and DOPET would denote the level of enzymatic inhibition. While DA is predominately metabolized to DOPAC, the reductive 94 metabolic pathway has been proposed to compensate for DOPAL metabolism in the event of ALDH inhibition (Turner et al., 1974). Indeed, inhibition of the oxidative pathway decreases DOPAC formation and increases DOPET formation in PC12 cells (Lamensdorf et al., 2000b), indicating that when ALDH is blocked, a major portion of DOPAL is reduced by AR to DOPET. Therefore inhibition of MAO activity would be reflected as a decrease in both DOPAL and DOPAC, with a concomitant increase in DOPET, whereas inhibition of ALDH activity would be associated with a decrease in DOPAC and an increase in DOPAL and DOPET. The goals of the present study were to 1) characterize the DA metabolomic profile following exposure to MeHg in undifferentiated PC12 cells, and 2) examine the specific contributions of direct and indirect ALDH inhibition to MeHg-impaired DA metabolism. To this end, the effects of MeHg on DA metabolism, ALDH activity, and mitochondrial function were compared with those of the reversible ALDH2 inhibitor daidzin (Keung and Vallee, 1993), and the reversible mitochondrial ETC Complex I inhibitor rotenone (Schuler and Casida, 2001). 95 Figure 4.1. Diagram depicting DA metabolic pathways. First DA is deaminated to DOPAL by MAO. The majority of DA is degraded along the “oxidative pathway”, where DOPAL is oxidized to DOPAC by ALDH. A small portion of DOPAL is reduced by AR to form DOPET along a “reductive pathway”. MAO and ALDH are mitochondrial-associated enzymes. MAO is localized to the outer mitochondrial membrane, and ALDH is present in the mitochondrial matrix. Additionally, both enzymes rely on cofactors provided by the mitochondria, FAD is used by MAO and NAD is used by ALDH. (Adapted from Lamensdorf et al., 2000a) 96 4.2. Methods 4.2.1. Chemicals MeHg (ICN Biochemicals), daidzin (Sigma), and rotenone (Sigma) were diluted from stock solutions in HBS. Drug solutions were diluted to final concentrations on the day of each experiment as described in General Methods Section 2.2. 4.2.2. Chemical treatment PC12 cells were seeded at a density of 6-7x105 cells/mL in 6-well plates or XF24well plates coated with poly-D-lysine 24-48 hr prior to treatment. To begin each experiment, culture medium was aspirated and replaced with HBS in the absence or presence of pharmacological and toxicant treatments. Cells were incubated at 37°C/5% CO2 during treatment as described in General Methods Section 2.2. 4.2.3. Neurochemistry At experiment termination, treatment medium was retained and acidified. Cells were rinsed, harvested, and pelleted by centrifugation. The content of DA and its metabolites in the treatment medium and cell pellet was determined by HPLC-ED as described in General Methods Section 2.3. Neurochemical concentrations were normalized to mL per sample for extracellular measurements or mg of protein for intracellular measurements as determined by the BCA protein assay (Sigma). To measure DOPAL, a standard was synthesized as described in General Methods Section 2.3.2. Briefly, 750µL of 1.063mM phosphate-buffered DA was incubated at 37°C for 97 15 min. The reaction was started by adding 25µL MAO (10 U; BD Biosciences, San Jose, CA) to each of the two culture tubes and gently mixing. At 45min, a 50µL aliquot was withdrawn from the reaction and added to 450µL of ice-cold tissue buffer. Following centrifugation, supernatants were removed and placed into a fresh tube for HPLC-ED analysis of DA and DOPAL. Concentrations of DA were compared to 1ng standards of DA, and concentrations of DOPAL were determined by normalizing the DOPAL peak height to the amount of DA lost. DOPAL standards are unstable and degrade within a few days of synthesis. As such a fresh standard was synthesized each time measurements of DOPAL were made. 4.2.4. ALDH2 enzyme kinetics assay To terminate the experiment, treatment medium was aspirated and replaced with 150µL homogenization buffer (0.1M Tris-HCl, 10mM DTT, 20% glycerol, and 1% Triton-X). Cells were scraped and pelleted by centrifugation at 13,000 g for 8 min at 4˚C. Protein concentration of supernatant was determined by BCA protein assay (Sigma). ALDH2 enzyme activity was determined spectrophotometrically by monitoring the reductive reaction of NAD to NADH at 340 nm as described in General Methods Section 2.7. ALDH2 reaction rates were calculated as µmole NADH per min per mg protein. 4.2.5. Mitochondrial bioenergetics Following pharmacological and toxicant exposure, treatment medium was aspirated. Cells were washed once with XF24 RPMI Assay Medium and then incubated at 37°C/no CO2 for 1 hr to allow medium temperature and pH to equilibrate before the first 98 measurements were made. The biosensor cartridge was hydrated overnight and injection ports were loaded with test compounds as described in General Methods Section 2.6. OCR was measured under basal conditions and following injection of test compounds using the XF24 Extracellular Flux Analyzer. Basal respiration, ATP generation, and spare respiration were calculated and compared between pharmacological and toxicant treatment groups. 4.2.6. NADH/NAD quantification To terminate the experiment, treatment medium was aspirated and cells were pelleted by centrifugation. NAD and NADH were extracted and measured by spectrophotometry as described in General Methods Section 2.8. Absorbance was compared to NADH standards to calculate the concentration of total NAD, total NADH, and the ratio of NAD:NADH. 4.2.7. Statistical analysis SigmaPlot® software version 12.0 (SysStat Software, Inc., Point Richmond, CA) was used to make statistical comparisons among groups using unpaired t-test, one-way ANOVA, or non-parametric alternatives as appropriate. If a significant difference was detected, post hoc between-group comparisons were performed using Tukey’s test. Statistical significance was set at p≤0.05. 99 4.3. Results 4.3.1. MeHg impairs DA metabolism, shunting DA along an alternative metabolic pathway As demonstrated in Figure 4.2A, treatment with 2μM MeHg for 60 min decreases the intracellular concentration of DOPAC, the primary DA metabolite (p<0.05). Because MeHg, at this same concentration and time-point, also increases intracellular DA (Figure 3.5), the observation that MeHg impairs canonical DA metabolism, suggests that MeHg inhibits some process specific to DOPAC formation. Figures 4.2B-C illustrate that in contrast to the effect on DOPAC, treatment with 2μM MeHg for 60 min increases intracellular concentrations of DOPAL and DOPET (p<0.05), the intermediate DA metabolite and alternative DA metabolic product, respectively. Comparison of the DA metabolomic profile in control and MeHg-treated PC12 cells (Figure 4.3) revealed that treatment with 2μM MeHg for 60 min shifts the distribution of DA metabolites. Under basal conditions, DOPAC comprises 56% of total metabolites, while DOPAL and DOPET account for 31% and 13% respectively. Following exposure to MeHg, the concentration of DOPAC decreases to account for only 14.5% of the total metabolites, whereas DOPAL and DOPET concentrations increase to accounts for 53.5% and 32%, respectively. Additionally, MeHg significantly increases the total concentration of DA metabolites (i.e. DOPAC, DOPAL, and DOPET) from 257.9 ± 31.4 to 398.1 ± 40.2 (p<0.05, HBS vs. MeHg). These data together suggest that MeHg impairs DOPAC formation at the level of the second DA metabolic enzyme, ALDH. Inhibition of MAO activity would have been reflected as a decrease in both DOPAL and DOPAC formation. Based on these data additional experiments were designed to investigate whether MeHg directly or indirectly impairs 100 ALDH activity. Furthermore, given that changes in DOPAL directly correspond to changes in DOPET, only DOPET was analyzed in the remaining studies. 101 Figure 4.2. MeHg shunts DA metabolism along the reductive metabolic pathway. Intracellular concentrations of DOPAC (Panel A), DOPAL (Panel B), and DOPET (Panel C). Undifferentiated PC12 cells were treated with 0µM (white bars) or 2µM (black bars) MeHg in HBS for 60 min. The asterisk (*) indicates a value significantly different than 0µM MeHg (p<0.05). Values are means + S.E.M. (n = 3, 3 replicates per n). 102 Figure 4.2 cont’d 103 Figure 4.3. Percentage changes in the relative amounts of DA metabolites induced by MeHg. The DA metabolomic profile was expressed as the percentage of DOPAC (white), DOPAL (gray), or DOPET (hatched) as a proportion of all three metabolites in undifferentiated PC12 cells treated with 0µM and 2µM MeHg for 60min. 104 4.3.2. Selective inhibition of ALDH with daidzin impairs DA metabolism in a manner similar to MeHg MeHg could impair ALDH activity directly by interacting with sulfhydryl groups present in the active site of the enzyme (Hempel et al., 1985). It could also indirectly decrease ALDH activity by decreasing availability of the ALDH cofactor, NAD, which is provided by Complex I of the mitochondrial ETC (Lamensdorf et al., 2000a; 2000b). In order to determine whether MeHg directly or indirectly inhibits ALDH activity, effects of MeHg on DA metabolism were compared to those of the specific ALDH2 inhibitor daidzin and the ETC Complex I inhibitor rotenone. Rotenone alters DA metabolism in PC12 cells (Lamensdorf et al., 2000a). Treatment with 50nM rotenone for 15 min decreases the combined concentrations of intracellular and extracellular DOPAC, while increasing DOPET concentrations. Effects of the specific ALDH2 inhibitor daidzin (Keung and Vallee, 1998) on DA metabolism are less well characterized. Therefore preliminary concentration-response and time-course experiments were conducted to characterize how daidzin alters intracellular concentrations of DOPAC and DOPET. As demonstrated in Figure 4.4, treatment with daidzin for 45 min significantly increased DOPET formation in undifferentiated PC12 cells only at the highest concentration investigated, 20μM (p<0.05). Intracellular DOPAC was not significantly altered by any concentration of daidzin. To examine time-course effects of daidzin, undifferentiated PC12 cells were exposed to 20μM daidzin for 0.75-24 hr (Figure 4.5). At all time points investigated, intracellular DOPAC was significantly decreased and intracellular DOPET was significantly increased (p<0.05). Based on these results, treatment with 20μM daidzin for 45 min was selected for comparison with MeHg. 105 Figure 4.4. Concentration-response effects of the ALDH inhibitor daidzin on concentrations of intracellular metabolites. Undifferentiated PC12 cells were treated with 0-20μM daidzin for 45 min, and concentrations of DOPAC (Panel A) and DOPET (Panel B) were measured. The asterisk (*) indicates a value significantly different than control values (p<0.05). Values are means ± S.E.M. (n = 1, 3 replicates per n). 106 Figure 4.5. Time-course effects of the ALDH inhibitor daidzin on concentrations of intracellular metabolites. Undifferentiated PC12 cells were treated with 20μM daidzin for 0-24 hr, and concentrations of DOPAC (Panel A) and DOPET (Panel B) were measured. The asterisk (*) indicates a value significantly different than control values (p<0.05). Values are means ± S.E.M. (n = 1, 3 replicates per n). 107 4.3.3. Daidzin and rotenone impair DA metabolism in a manner that resembles MeHg The comparative effects of MeHg, rotenone, and daidzin on intracellular DOPAC and DOPET concentrations are presented in Figure 4.6. Treatment with all three chemicals decreased intracellular DOPAC concentrations by approximately 60% as compared to cells treated with HBS (p<0.05). MeHg and daidzin, but not rotenone, increased DOPET concentrations (p<0.05). Furthermore, the increase induced by MeHg was more pronounced. While MeHg increased DOPET by over 300% as compared to HBS, daidzin only increased DOPET by ~200%. Rotenone did increase DOPET concentrations by ~100%, however this effect was not significantly different than HBS. 108 Figure 4.6. Comparative effects of MeHg, daidzin, and rotentone on concentrations of intracellular metabolites. Undifferentiated PC12 cells were treated with HBS for 60 min (white bars), 2μM MeHg for 60 min (black bars), 20μM daidzin for 45 min (gray bars), or 50nM rotenone for 15 min (hatched bars), and intracellular concentrations of DOPAC (Panel A) and DOPET (Panel B) were measured. The asterisk (*) indicates a value significantly different than HBS (p<0.05). Values are means + S.E.M. (n = 3, 2-3 replicates per n). 109 4.3.4. ALDH activity is not directly altered by MeHg To determine if MeHg has a direct effect on ALDH, enzymatic activity was measured as the production of NADH over time. ALDH uses NAD as a cofactor during substrate oxidation (Marchitti et al., 2008), and as a by-product, NADH is formed (Figure 4.1). NADH production was measured in PC12 cell lysate either following pre-treated with or in the presence of HBS, 2μM MeHg, 20μM daidzin, or 50nM rotenone. Following pre-treatment, only rotenone increased NADH production (Figure 4.7A; p<0.05). However when ALDH activity was measured in the presence of each chemical, daidzin decreased and rotenone increased NADH production (Figure 4.7B; p<0.05). MeHg had no effect on NADH production under either experimental condition. In order to determine if MeHg impaired ALDH activity at higher concentrations, a concentration-response experiment was performed (Figure 4.8). NADH production was measured in the presence of 1-100μM MeHg. At no concentration did MeHg affect NADH production (p>0.05). 110 Figure 4.7. Comparative effects of MeHg, daidzin, and rotenone on ALDH activity. Panel A) Undifferentiated PC12 cells were treated with HBS for 60 min (white bars), 2μM MeHg for 60 min (black bars), 20μM daidzin for 45 min (gray bars), or 50nM rotenone for 15 min (hatched bars). Cells were then harvested and ALDH activity was measured in each treatment group. Drugs were not present during enzyme activity assay. Panel B) Undifferentiated PC12 cells were harvested and ALDH activity was measured in the presence of HBS (white bars), 2μM MeHg (black bars), 20μM daidzin (gray bars), or 50nM rotenone (hatched bars). The asterisk (*) indicates a value significantly different than HBS (p<0.05). Values are means + S.E.M. (n = 3, 6 replicates per n). 111 Figure 4.8. Concentration-response effects of MeHg on ALDH activity ex vitro. ALDH activity was measured in undifferentiated PC12 lysate in the presence of 0-100μM MeHg. Values are means + S.E.M. (n = 1, 6 replicates per n). 112 4.3.5. MeHg impairs mitochondrial function and decreases availability of the ALDH cofactor NAD The indirect effects of MeHg on ALDH activity were investigated by measuring mitochondrial function and ALDH cofactor production. Mitochondrial respiration was measured in undifferentiated PC12 cells pre-treated with HBS for 60 min, 2μM MeHg for 60 min, 20μM daidzin for 45 min, or 50nM rotenone for 15 min. Following pre-treatment, OCR was measured under basal conditions and following treatment with 5mM pyruvate, 1μM oligomycin A, 2μM FCCP, and 1μM antimycin A. Treatment with these compounds is used to measure basal respiration, ATP generation, maximum respiration, and non-mitochondrial respiration, respectively. The effects of MeHg, daidzin, and rotenone on OCR is presented in Figure 4.9. The quantification of these effects revealed that MeHg attenuated basal and spare respiration, as well as ATP generation (p<0.05). Rotenone only impaired spare respiration, and daidzin had no effect on mitochondrial function (Figure 4.10; p<0.05). Because Complex I of the mitochondrial ETC accepts electrons from NADH thereby forming NAD, and because ALDH uses NAD provided by Complex I, impaired mitochondrial respiration could detrimentally affect the production of NAD. In order to determine if MeHg alters the production of NAD, the amount of the coenzyme present in undifferentiated PC12 cells treated with either HBS or 2μM MeHg for 60 min was quantified. As illustrated in Figure 4.11, MeHg caused similar decreases in the amount of both NAD and NADH (p<0.05) such that there was no change in the NAD/NADH ratio. 113 Figure 4.9. Comparative effects of MeHg, daidzin, and rotenone on mitochondrial bioenergetics. Undifferentiated PC12 cells were pretreated with HBS for 60 min (red), 2μM MeHg for 60 min (orange), 20μM daidzin for 45 min (yellow), or 50nM rotenone for 15 min (green). Following pre-treatment, OCR (pMoles/min) was measured under basal conditions and following injections of 5mM pyruvate (A), 1μM oligomycin (B), 2μM FCCP (C), and 1μM antimycin A (D). Values are means ± S.E.M. (n = 3, 4-5 replicates per n). 114 Figure 4.10. Quantification of changes in mitochondrial bioenergetics induced by MeHg, daidzin, or rotenone. Undifferentiated PC12 cells were treated with HBS for 60 min (white bars), 2μM MeHg for 60 min (black bars), 20μM daidzin for 45 min (gray bars), or 50nM rotenone for 15 min (hatched bars). Panel A) Basal respiration (OCR) was measured in the presence of 25mM glucose and 5mM pyruvate/2.5mM malate. Panel B) Spare respiration was calculated as the difference between maximum respiration (following treatment with 2μM FCCP) and basal respiration. Panel C) ATP generation was calculated as the difference between basal respiration and respiration following treatment with 1μM oligomycin A. The asterisk (*) indicates a value significantly different than HBS (p<0.05). Values are means + S.E.M. (n = 3, 4-5 replicates per n). 115 Figure 4.10 cont’d 116 Figure 4.11. Effect of MeHg on the production of NAD(H). Undifferentiated PC12 cells were treated with 0μM (white bars) or 2μM (black bars) MeHg for 60 min, and then the amount of NAD (Panel A) and NADH (Panel B) were measured by spectrophotometry using an NAD/NADH Quantification Kit (Sigma). The ratio of NAD/NADH (Panel C) was calculated as (NAD-NADH)/NADH. The asterisk (*) indicates a value significantly different than 0μM MeHg (p<0.05). Values are means + S.E.M. (n = 4, 4 replicates per n). 117 Figure 4.11 cont’d 118 4.4. Discussion The present study provides evidence that MeHg impairs DA metabolism by indirectly inhibiting ALDH activity. These data are consistent with the following conclusions: 1) MeHg impairs conventional DA metabolism and shunts DA along an alternative metabolic pathway, 2) MeHg does not directly impair ALDH activity to alter DA metabolism, 3) mitochondrial dysfunction induced by MeHg decreases basal respiration, as well as ATP generation and reserve respiratory capacity, and 4) MeHg decreases availability of the enzyme cofactor NAD(H), which could indirectly impair ALDH activity. 4.4.1. The DA metabolomic profile induced by MeHg MeHg inhibits DA metabolism. Previous reports demonstrate a concentrationdependent decrease in intracellular DOPAC in the striatum (Bemis and Seegal, 1999; Dreiem et al., 2009), and decreased striatal release of DOPAC (Faro et al., 2003; 2007). Results from the present study support and extend these findings. Intracellular DOPAC was significantly decreased by exposure to MeHg, and this effect was accompanied by a significant increase in intracellular concentrations of DOPAL and DOPET. Analysis of the percentage changes in these three metabolites reveals they are proportional; DOPAC decreases by approximately 60%, while DOPAL and DOPET increase by approximately 65% and 70% respectively. These data suggest that MeHg impairs DA metabolism by inhibiting the oxidation of DOPAL to DOPAC. MAO is functional and its activity is intact because the concentration of DOPAL did not decrease. Conversely, the concentration of DOPAL increased, suggesting that ALDH activity is impaired. As a result, DOPAL metabolism shifts to the lower-affinity 119 reductive pathway mediated by AR, which enhances DOPET accumulation (Turner et al., 1974; Lamensdorf et al., 2000b). The present analysis suggests that MeHg does not stimulate AR activity. DOPAL and DOPET increased in proportion to one another. If AR were stimulated, a profound increase in DOPET without a concomitant increase in DOPAL would be expected. Furthermore, while DOPET accumulation was moderately enhanced, this was expected as inhibition of the oxidative pathway has been previously demonstrated to increase DOPET formation (Lamensdorf et al., 2000b). However based on the present analysis, there is no conclusive evidence that MeHg does not stimulated AR, and further investigate would be needed to address this point. The conclusion that MeHg inhibits ALDH is novel, however somewhat inconsistent with reports in the literature. Previous studies investigating the effects of MeHg on DA metabolism have primarily indicated that MeHg inhibits MAO activity (Taylor and DiStefano, 1976; Chakrabarti et al., 1998; Beyrouty et al., 2006; Castoldi et al., 2006). In these studies MAO activity was measured in either whole or regional brain homogenates. Therefore the analysis of MAO activity is these studies was not specific to DA neurons, and the observed decrease in MAO activity in these studies could have been due to collective changes in multiple types of neurons. It is also plausible that the lack of change in MAO activity observed presently is specific to the undifferentiated PC12 cell line. For example, expression of different MAO and ALDH isozymes in NSDA neurons could mediate alterative changes in the aberrant DA metabolomic profile induced by exposure to MeHg. Two isozymes of MAO, designated A and B, have been proposed based on substrate selectivity and inhibitor sensitivity (Cai et 120 al., 1998). MAO A has a higher affinity for DA, and is predominately found in rat catecholamine neurons and PC12 cells (Naoi et al., 1987; Jahng et al., 1997). However, in most species both forms can oxidize DA (O'Carroll et al., 1983), and in humans MAO B is primarily responsible for DA oxidation (Glover et al., 1977; Riederer and Jellinger, 1982). Similarly, two isozymes of ALDH, ALDH2 and ALDH1A1, can mediate DA metabolism (Marchitti et al., 2008). In PC12 cells, ALDH2 catalyzes the oxidation of DOPAL (Marchitti et al., 2007; Robador et al., 2012), whereas in NSDA neurons both ALDH2 and ALDH1A1 can participate (Galter et al., 2003; Anderson et al., 2011b). Given these differences, an investigation of the aberrant DA metabolomic profile induced by MeHg in NSDA neuronal cultures is warranted. However, the observed changes in DA metabolites suggest that mechanisms by which MeHg targets DA metabolism are more complex that previous assumed. 4.4.2. Comparative effects of MeHg, daidzin, and rotenone on DA metabolism The observed neurochemical profile suggests that MeHg impairs ALDH activity, but does not provide evidence to distinguish between direct versus indirect effects. Directly, MeHg could bind cysteine residues present in the active site of the enzyme (Weiner and Wang, 1994). Indirectly, impaired mitochondrial respiration induced by MeHg could decrease availability of the ALDH cofactor NAD, which is produced when NADH donates electrons to Complex I of the ETC (Lamensdorf et al., 2000a). To identify direct or indirect mechanisms by which MeHg may target ALDH, the effect of the toxicant was compared with two other pharmacological inhibitors, daidzin and rotenone. Daidzin is a reversible, selective ALDH2 inhibitor (Keung and Vallee, 1993), and rotenone is a reversible inhibitor 121 of NADH:ubiquinone oxidoreducase (Complex I of the mitochondrial ETC; Schuler and Casida, 2001). Rotenone and daidzin reduced the intracellular concentration of DOPAC to a similar extent as compare to MeHg. However the increase in DOPET induced by MeHg and daidzin was more pronounced than that induced by rotenone. These results suggest that all three compounds inhibit the oxidative pathway of DA metabolism; daidzin by directly inhibiting ALDH (Keung and Vallee, 1993), rotenone by decreasing availability of the ALDH cofactor NAD (Schuler and Casida, 2001), and MeHg by a yet to be elucidated mechanism. However, the three compounds stimulate the reductive pathway of DA metabolism to different degrees. 4.4.3. Evidence for a role of mitochondrial dysfunction rather than direct ALDH inhibition in MeHg-impaired DA metabolism To investigate the direct effect of MeHg on ALDH, enzymatic activity was measured as the accumulation of NADH, a by-product of reactions catalyzed by ALDH (Guru and Shetty, 1990). The activity of ALDH was measured either following pre-treatment or in the presence of HBS, MeHg, daidzin, or rotenone. As expected, daidzin decreased NADH production, but only during co-treatment. Because daidzin is a reversible ALDH2 inhibitor, its presence is required to inhibit ALDH enzymatic activity. Rotenone increased NADH production. This observation was likely a false positive, resulting from the action of rotenone on NADH:ubiquinone oxidoreducase (Schuler and Casida, 2001). Inhibition of this enzyme complex by rotenone resulted in an artificially high accumulation of NADH in the enzyme assay. MeHg, however, had no observable effect on NADH production. This lack of 122 effect was demonstrated not only following pretreatment with 2μM for 60 min, but also in the presence of 1-100μM MeHg. Based on these results, it can be concluded that MeHg does not directly inhibit ALDH. MeHg could indirectly inhibit ALDH activity by decreasing availability of its cofactor NAD. Mitochondrial dysfunction resulting in inhibition of the mitochondrial ETC and decreased respiration would contribute to decreased production of NAD. To investigate this hypothesis, mitochondrial respiration and NAD(H)+ were measured. In comparison to pre-treatment with daidzin and rotenone, MeHg not only decreased basal respiration and ATP generation, as demonstrated previously (Sone et al., 1977; Yee and Choi, 1994), but it also decreased spare respiratory capacity. Daidzin had no effect on mitochondrial function, as anticipated, and rotenone only impaired spare respiration. The observed effects of rotenone were less pronounced than expected, but likely resulted because rotenone is a reversible inhibitor. Spare respiratory capacity is the ability of a cell to respond to an energy crisis (Nicholls et al., 2007). It is dependent upon increased usage of substrate supply and electron transport. Maintenance of an energy reserve is important in the face of oxidative stress (Vesce, 2005) or restriction of Complex I and/or II activity (Yadava and Nicholls, 2007; Choi et al., 2009). In cells with variable ATP requirements, such as neurons, spare respiratory capacity is a powerful diagnostic of cellular bioenergetics (Brand and Nicholls, 2011). Indeed, Nicholls et al., (2007) argue that the maintenance of some spare respiratory capacity is a major factor in survival of neurons. The ability of MeHg to abolish completely spare respiratory capacity in undifferentiated PC12 cells is a novel finding with significant implications. These data 123 suggest that in the face of MeHg exposure, cells are operating at the upper limit of their bioenergetic capacity. As a result, they would be unable to combat subsequent oxidative stress and excitotoxicity induced by MeHg (Dreiem and Seegal, 2007; Franco et al., 2007; Mori et al., 2007; Liu et al., 2012). Abolition of spare respiratory capacity could be a critical early factor in the initiation of cell death (Nicholls et al., 2007). These data are also indicative of impaired electron transport. The exact level at which MeHg impairs the ETC was not determined presently, however previous reports suggest that the toxicant inhibits both Complexes II and III (Sone et al., 1977; Yee and Choi, 1994; Mori et al., 2011). Inhibition at these positions along the respiratory chain would down-regulate the activity of Complex I (DiMauro and Schon, 2008; Cannino et al., 2012), and oxidation of NADH to NAD would diminish. 4.4.4. Consequences of decreased NAD(H) following MeHg exposure Quantification of NAD(H) revealed a decrease in both total NAD and NADH, without altering their ratio. These data suggest that MeHg decreases the total amount of NAD(H) in either its oxidized or reduced form. This finding supports the conclusion that decreased availability of the ALDH cofactor contributes to the impairment of DA metabolism following exposure to MeHg. Mitochondrial dysfunction induced by MeHg was hypothesized to contribute to the decreased availability of NAD. The present set of experiments does not provide conclusive evidence to support this hypothesis. Indeed, if MeHg did impair mitochondrial function to indirectly inhibit ALDH, the DA neurochemical profile would mimic that induced by rotenone. Instead MeHg increased the concentration of DOPET to a much greater extent 124 than did rotenone. Additionally, although MeHg profoundly impaired mitochondrial respiration, there is no evidence to correlate that dysfunction with the observed decrease in NAD. Lastly, because MeHg decreased the total amount of NAD(H), inhibition of the ETC, particularly Complex I, cannot completely account for the observed alterations in DA metabolism, as inhibition of Complex I would decrease the oxidation of NADH (Kang et al., 1997). Yet unidentified intracellular factors must contribute to the observed decrease in NAD(H). Two possibilities are MeHg-induced inhibition of the citric acid cycle and/or activation of a protective metabolic network dedicated to the conversion of NADH to NADPH. The citric acid cycle is ubiquitous and comprises an essential series of chemical reactions responsible for generating cellular energy in aerobic organisms. Additionally, the cycle provides NADH, which can be used in numerous biochemical reactions, including the mitochondrial ETC. There is some evidence to suggest that MeHg inhibits the citric acid cycle. The formation of substrates for the cycle, including pyruvate, are increased in guinea pig cerebral cortical slices treated with 10-50μM MeHg (Fox et al., 1975). Furthermore MeHg is demonstrated to inhibit the activity of specific enzymes in the cycle, including succinate dehydrogenase (Yoshino et al., 1966) and malate dehydrogenase (Hamdy and Noyes, 1977). Resulting changes in substrate availability would regulate the activity of the citric acid cycle, and decrease production of NADH. Another possible explanation for decreased availability of NAD(H) is conversion of NADH to NADPH. NADPH is an essential reducing agent, and contributes to antioxidant defense mechanisms by sustaining the activity of enzymes such as catalase, superoxide dismutase, and glutathione peroxidase (Minard, 2005). There is evidence to suggest that 125 under conditions of oxidative stress, NADH is converted to NADPH by a compensatory metabolic network involving NADH kinase and malic enzyme (Singh et al., 2008). Activation of this pathway could potentially contribute to antioxidant defense strategies in the face oxidative stress induced by MeHg (LeBel et al., 1990; Sarafian et al., 1994; Gatti et al., 2004). However because MeHg inhibits many antioxidant enzymes, including glutathione peroxidase (Kaur et al., 2006; Franco et al., 2009) and catalase (Cuello et al., 2010), the actual protection offered by activation of this network could be negligible. It is unknown if and how NADPH levels are modulated following exposure to MeHg, however a potential increase in NADPH could contribute to AR stimulation and DOPET accumulation observed in these studies. 4.4.5. Potential biological effects of MeHg-impaired DA metabolism The observation that MeHg increases DOPAL accumulation is of significant consequence. Increased production of DOPAL is one mechanism by which MeHg may exert toxicity in DA synthesizing cells. DOPAL has demonstrated toxic effects in DA neurons both in vivo and in vitro. Microinjections of DOPAL in the substantia nigra cause focal, generalized lesions associated with a loss of TH immunoreactivity and other phenotypic neuronal markers, as well as marked gliosis (Burke et al., 2003). In differentiated PC12 cells and striatal synaptosomes, exposure to micromolar concentrations of DOPAL increases the incidence of cell death (Mattammal et al., 1995). In addition to increasing toxicity itself, DOPAL in combination with glucose deprivation and mitochondrial respiratory chain inhibition enhances toxicity (Lamensdorf et al., 2000a). Hence, inhibition of DOPAL oxidation potentiates toxicity induced by 126 metabolic stress and mitochondrial dysfunction. As demonstrated presently, MeHg impairs mitochondrial respiration and abolishes reserve respiratory capacity in cells. Therefore, the selective sensitivity of DA synthesizing cells to MeHg may result from a combination of the generalized molecular mechanism by which the toxicant exerts its effects and the distinct cellular environment created by the presence of DA and the enzymes necessary for its metabolism. Furthermore, the finding that MeHg increases DOPAL activity as a result of indirect ALDH inhibition provides evidence in support of the hypothesis that exposure to MeHg contributes to the etiology of PD. A link between ALDH inhibition and PD has previously been established following occupational exposure to another environmental toxicant. Fitzmaurice et al. (2013) demonstrate that the fungicide benomyl causes concentrationdependent inhibition of ALDH, which is associated with decreased production of DOPAC in primary DA cultures and decreased VMAT positive neurons in zebrafish. In humans, occupational exposure to benomyl increases the incidence of PD by 67% (Fitzmaurice et al., 2013). Therefore environmental toxicants, including benomyl and MeHg, inhibit ALDH sufficiently to damage DA neurons and increase the risk of PD in exposed humans. 4.4.6. Summary In conclusions, MeHg alters the DA metabolic profile in undifferentiated PC12 cells. Results suggest that ALDH activity is inhibited indirectly by mitochondrial dysfunction and decreased availability of the ALDH cofactor NAD. Consequences of impaired DA metabolism contribute to accumulation of the toxic DA metabolic intermediate, DOPAL. These data 127 provide evidence for a mechanism by which DA neurons may be selectively sensitive to the toxic effects of MeHg. 128 Chapter 5. The role of extracellular Ca2+ in MeHg-induced alterations in DA release and metabolism 5.1. Introduction 5.1.1. Neurotransmitter release and VGCCs Extracellular Ca2+ influx through VGCCs is the primary event responsible for initiating synaptic transmission (Olivera et al., 1994; Dunlap et al., 1995). Once Ca2+ enters the cell, it binds the Ca2+ binding protein on the vesicle membrane, synaptotagmin, and activates proteins involved in vesicle docking and neurotransmitter release (Bommert et al., 1993; Elferink et al., 1993; Bajjalieh and Scheller, 1995; Catterall and Few, 2008). In the mammalian central nervous system, the P/Q-type VGCC plays the major role mediating vesicular exocytosis at most synapses (Sheng et al., 1998). However N- and L-type VGCCs have also been implicated in specific subsets of neurons, including inhibitory interneurons in the hippocampus (Poncer et al., 1997) and neuroendocrine cells (Yamada et al., 1996). MeHg causes a concentration- and time-dependent increase in intracellular Ca2+ (Kauppinen et al., 1989). The first phase is slow, gradual, and mediated by the release of Ca2+ from intracellular stores (Hare and Atchison, 1995a; Limke et al., 2003). The second phase is rapid and occurs at a magnitude that overwhelms the cellular buffering capacity (Marty and Atchison, 1997). Removal of extracellular Ca2+ abolishes the second phase of intracellular Ca2+ elevation induced by MeHg, demonstrating a strict dependence on 129 extracellular Ca2+ (Denny et al., 1993). To date, the pathways mediating second phase Ca2+ influx have yet to be elucidated, however influx through L-, N-, or P/Q-type VGCCs has been implicated because their inhibition delays the time-to-onset (Marty and Atchison, 1997). Effects of MeHg on VGCCs are well documented although somewhat enigmatic. Immediately following application, MeHg decreases VGCC-mediated 45Ca2+ uptake in forebrain synaptosomes, and decreases specific binding of L- and N-type VGCC antagonists in PC12 cells (Shafer et al., 1990). Additionally, whole cell currents carried by Ba2+, a Ca2+ surrogate, through L- and N-type VGCCs is rapidly and completely blocked by MeHg (Shafer and Atchison, 1991b). Based on these findings it has been predominantly concluded that MeHg interacts directly with channels to irreversibly block Ca2+ flux. However, MeHg does not reduce mean Ca2+ current amplitude when applied to the cytoplasmic side of the membrane (Shafer, 1998). One possible explanation for these paradoxical findings is that MeHg uses VGCCs to enter the cell, which competitively but temporarily blocks the channel. Once on the cytoplasmic side, the toxicant interacts with the channel to enhance extracellular Ca2+ influx. Delay of the second Ca2+ phase by VGCC inhibition supports this hypothesis (Marty and Atchison, 1997). In this instance, VGCC antagonism could actually delay the interaction of MeHg with target sites or delay its entry into the cell. Because of the crucial role of Ca2+ influx through VGCCs in neurotransmitter release, it follows that extracellular Ca2+ influx induced by MeHg may act as a primary 130 mechanism mediating MeHg-induced DA release (Chapter 3). A role for extracellular Ca2+ in MeHg-induced catecholamine release has been documented (Gassó et al., 2000), however extracellular Ca2+-independent DA release has also been described following MeHg exposure (Kalisch and Racz, 1996; Faro et al., 2002). As such, the present study sought to elucidate the role of extracellular Ca2+ and the potential contribution of VGCCs to MeHg-induced DA release. 5.1.2. Mitochondrial function and DA metabolism Tight regulation of intracellular Ca2+ homeostasis is of critical importance because Ca2+ plays key roles in metabolic regulation and intracellular signaling cascades. Mitochondria are one of two essential Ca2+ buffering organelles responsible for maintaining a low concentration of intracellular Ca2+ (Blaustein, 1988; Meldolesi et al., 1992). Generally, a powerful stimulus of high intracellular Ca2+ is required for mitochondria to actively buffer Ca2+ from the cytosol into the mitochondrial matrix (Nicholls and Scott, 1980; Stockum et al., 2011). The Ca2+ uniporter mediates Ca2+ uptake (Pan et al., 2011), whereas the Na+/Ca2+ exchanger primarily mediates Ca2+ efflux (Deryabina et al., 2004). However Ca2+ can also be released from the mitochondria following reversal of the Ca2+ uniporter, collapse of the mitochondrial membrane potential, or opening of the mitochondrial permeability transition pore (Contreras et al., 2010; Assaly 131 et al., 2012; Roos et al., 2012). These pathological conditions occur as a result of Ca2+ overloading and indicate mitochondrial dysfunction, which can trigger apoptosis (Ravagnan et al., 2002; Jordan et al., 2003). MeHg induces a mitochondrial state consistent with Ca2+ overloading (Roos et al., 2012). Respiration and ATP production are depressed (Kauppinen et al., 1989; Chapter 4; Gatti et al., 2004), Complexes II and III of the ETC are inhibited (Sone et al., 1977; Yee and Choi, 1996; Mori et al., 2011), and the mitochondrial membrane potential is dissipated (Bondy and McKee, 1991; Hare and Atchison, 1992; Limke and Atchison, 2002). Subsequent cell death is thought to result from opening of the mitochondrial permeability transition pore and release of pro-apoptotic factors, including apoptosis inducing factor (Fonfria et al., 2002; Limke and Atchison, 2002). MeHg affects mitochondrial Ca2+ regulation by at least two mechanisms. Directly, MeHg inhibits ETC respiratory proteins (Verity et al., 1975) and causes a loss of mitochondrial membrane potential (Bondy and McKee, 1991; Hare and Atchison, 1992; Limke and Atchison, 2002). Indirectly, MeHginduced increase in intracellular Ca2+ initiates excessive uptake of Ca2+ by the mitochondria (Limke et al., 2003), ultimately depolarizing the inner mitochondrial membrane, inhibiting ATP production, and opening the mitochondrial permeability transition pore. Considering that mitochondria play an essential role in buffering excessive intracellular Ca2+ (Blaustein, 1988; Meldolesi et al., 1992) and extracellular Ca2+ influx induced by MeHg contributes to deleteriously elevated intracellular Ca2+ (Denny et al., 132 1993; Marty and Atchison, 1997), the role of MeHg-induced extracellular Ca2+ influx in mitochondrial dysfunction was investigated. Although removal of extracellular Ca2+ does not alter MeHg-induced ROS formation (Dreiem and Seegal, 2007), other aspects of mitochondrial function, including respiration and ATP production, have yet to be investigated. Since MeHg-impaired DA metabolism is due in part to mitochondrial dysfunction (Chapter 4), the role of extracellular Ca2+ in MeHg-impaired DA metabolism was also investigated. The goals of the present study were to identify the contribution of extracellular Ca2+ to MeHg-induced changes in DA release, DA metabolism, and mitochondrial function. 5.2. Methods 5.2.1. Chemicals The standard physiological saline used for extracellular solution was HBS, which contained (mM): 150 NaCl, 5 KCl, 2.4 CaCl2, 1.6 MgSO4, 20 HEPES, and 20 d-glucose (pH 7.3). In some experiments cells were treated in 0-Ca2+ HBS, which had the same composition as HBS with the following exceptions (mM): 0 CaCl2 and 0.02 ethylene glycol tetraacetic acid. MeHg (ICN Biochemicals), nimodipine (Sigma), and cadmium chloride (Cd2+; Sigma) were diluted from stock solutions in HBS or 0-Ca2+ HBS. Nimodipine was protected from light during preparation and experimentation. Drug solutions were diluted to final concentrations on the day of each experiment as described in General Methods Section 2.2. 133 5.2.2. Chemical treatment PC12 cells were seeded at a density of 6-7x105 cells/mL in 6-well plates or XF24well plates coated with poly-D-lysine 24-48 hr prior to treatment. To begin each experiment, culture medium was aspirated and replaced with HBS or 0-Ca2+ HBS in the absence or presence of pharmacological and toxicant treatments. Cells were incubated at 37°C/5% CO2 during treatment as described in General Methods Section 2.2. 5.2.3. Neurochemistry At experiment termination, treatment medium was retained and acidified. Cells were rinsed, harvested, and pelleted by centrifugation. The content of DA in the treatment medium and its metabolites in the cell pellet were determined by HPLC-ED as described in General Methods Section 2.3. Neurochemical concentrations were normalized to mL per sample for extracellular measurements or mg of protein for intracellular measurements as determined by the BCA protein assay (Sigma). 5.2.4. Mitochondrial bioenergetics Following pharmacological and toxicant exposure, treatment medium was aspirated. Cells were washed once with XF24 RPMI Assay Medium and then incubated at 37°C/no CO2 for 1 hr to allow medium temperature and pH to equilibrate before the first measurements were made. The biosensor cartridge was hydrated overnight and injection ports were loaded with test compounds as described in General Methods Section 2.6. OCR 134 was measured under basal conditions and following injection of test compounds using the XF24 Extracellular Flux Analyzer. Basal respiration, ATP generation, and spare respiration were calculated and compared between treatment groups. 5.2.5. Statistical analysis SigmaPlot® software version 12.0 (SysStat Software, Inc., Point Richmond, CA) was used to make statistical comparisons among groups using unpaired t-test, one-, and twoway ANOVA, or non-parametric alternatives as appropriate. If a significant difference was detected, post hoc between-group comparisons were performed using Tukey’s test. Statistical significance was set at p≤0.05. 5.3. Results 5.3.1. Spontaneous MeHg-mediated DA release is partially dependent upon the presence of extracellular Ca2+, but not Ca2+ influx through VGCCs DA is released from PC12 cells through Ca2+-dependent exocytosis (Kishimoto et al., 2005). Because MeHg induces extracellular Ca2+ influx (Marty and Atchison, 1997), a role for extracellular Ca2+ in MeHg-mediated DA release from undifferentiated PC12 cells was evaluated by measuring extracellular DA concentrations after exposure to MeHg in a Ca2+free solution. As demonstrated previously (Chapter 3), in the presence of extracellular Ca2+, treatment with 2μM MeHg for 60 min increased the concentration of extracellular DA (Figure 5.1A; p<0.05). There was a slight non-significant decrease in spontaneous DA 135 release in cells incubated in Ca2+-free HBS (Figure 5.1A, C; p>0.05). In the absence of extracellular Ca2+, MeHg-induced DA release from PC12 cells was significantly attenuated as compared with HBS-treated cells (p<0.05). However, there was still a dramatic increase in DA released by MeHg in the absence of extracellular Ca2+. When evaluated as the percentage change from medium control, DA release induced by MeHg in the absence of extracellular Ca2+ was not significantly different from DA released by MeHg in the presence of extracellular Ca2+ (Figure 5.1B; p>0.05). VGCCs contribute to the perturbation of intracellular Ca2+ induced by MeHg (Hare and Atchison, 1995b; Marty and Atchison, 1997). Undifferentiated PC12 cells primarily express the L-type VGCC, and Ca2+ current through these channels is blocked by dihydropyridine antagonists, such as nimodipine (Kongsamut and Miller, 1986; Janigro et al., 1989; Avidor et al., 1994). Therefore experiments complementary to those evaluating MeHg-induced DA release in a Ca2+-free buffer evaluated release following inhibition of VGCCs (Figure 5.2). Pretreatment with 2μM nimodipine, a selective L-type VGCC inhibitor, failed to attenuate DA release induced by 2μM MeHg for 60 min. Likewise, the nonspecific VGCC inhibitor Cd2+ (200μM) had no effect (p>0.05). 136 Figure 5.1. Extracellular Ca2+ does not contribute to MeHg-induced DA release. Undifferentiated PC12 cells were treated with 0µM (white bars) or 2µM (black bars) MeHg in the absence (0-Ca2+ HBS) or presence (HBS) of extracellular Ca2+ for 60 min. Panel A) Concentrations of extracellular DA. Panel B) The percentage change of extracellular DA after treatment with MeHg in cells treated with either HBS or 0-Ca2+ HBS. Panel C) The percentage reduction of extracellular DA by treatment with 0-Ca2+ HBS in cells treated with either 0µM or 2µM MeHg. The asterisk (*) indicates a value significantly different than HBS (p ≤ 0.05). The hash (#) indicates a value significantly difference from both HBSControl and HBS-MeHg (p ≤ 0.05). Values are means + SEM (n = 4, 3 replicates per n). 137 Figure 5.1 cont’d 138 Figure 5.2. Spontaneous activation of VGCCs does not contribute to MeHg-induced DA release. Undifferentiated PC12 cells were treated with either 2μM nimodipine or 200μM Cd2+ in culture medium for 15min prior to co-treatment with 0µM (white bars) or 2µM (black bars) MeHg for 60 min. Panel A) Extracellular DA concentrations. Panel B) The percentage change of extracellular DA after treatment with MeHg in each treatment group. The asterisk (*) indicates a significant difference from HBS within treatment group (p ≤ 0.05). Values are means + SEM (n = 3, 3 replicates per n). 139 5.3.2. Extracellular Ca2+ stimulates mitochondrial bioenergetics, but does not contribute to MeHg-induced mitochondrial dysfunction or aberrant DA metabolism As demonstrated in Chapter 4, MeHg impairs both mitochondrial bioenergetics and DA metabolism. Results suggest MeHg may indirectly impair the oxidation of DOPAL to DOPAC by decreasing availability of the ALDH cofactor NAD, the majority of which is provided by Complex I of the mitochondrial ETC (Weiner and Wang, 1994). As such, MeHginduced mitochondrial dysfunction is hypothesized to contribute to MeHg-impaired DA metabolism. To evaluate a role for extracellular Ca2+ in MeHg targeting of DA metabolism, parameters of mitochondrial bioenergetics and concentrations of intracellular DA metabolites were measured after exposure to MeHg in a Ca2+-free buffer. Mitochondrial respiration was measured in undifferentiated PC12 cells pre-treated with 2μM MeHg for 60 min in the absence or presence of extracellular Ca2+. Following pre-treatment, OCR was measured with the XF24 respirometer under basal conditions and following treatment with 5mM pyruvate, 1μM oligomycin A, 2μM FCCP, and 1μM antimycin A. Figure 5.3 depicts changes in OCR following treatment with these compounds. The quantification of these effects revealed that in the absence of extracellular Ca2+, basal respiration, spare respiratory capacity, and ATP generation were all enhanced (Figure 5.4; p<0.05). Treatment with 2μM MeHg for 60 min decreased all three parameters to a similar extent, regardless of the absence or presence of extracellular Ca2+ in the treatment medium (p<0.05). 140 In a complementary set of cultures, concentrations of DOPAC and DOPET were assessed following treatment with 2μM MeHg for 60 min in 0-Ca2+ HBS. MeHg decreased intracellular DOPAC and increased intracellular DOPET in undifferentiated PC12 cells, as previously observed (Chapter 4). Removal of extracellular Ca2+ from the treatment buffer did not alter the effect of MeHg on either metabolite (Figure 5.5; p>0.05). 141 Figure 5.3. Changes in mitochondrial bioenergetics induced by MeHg in the absence or presence of extracellular Ca2+. Undifferentiated PC12 cells were pretreated for 60 min with 0μM MeHg in the absence (yellow) or presence (red) of extracellular Ca2+ or 2μM MeHg in the absence (green) or presence (orange) of extracellular Ca2+. Following pre-treatment, OCR (pMoles/min) was measured under basal conditions and following injections of 5mM pyruvate (A), 1μM oligomycin (B), 2μM FCCP (C), and 1μM antimycin A (D). Values are means ± S.E.M. (n = 3, 4-5 replicates per n). 142 Figure 5.4. Quantification of changes in mitochondrial bioenergetics induced by MeHg in the absence or presence of extracellular Ca2+. Panel A) Basal respiration (OCR) was measured in the presence of 25mM glucose and 5mM pyruvate. Panel B) Spare respiration was calculated as the difference between maximum respiration (following treatment with 2μM FCCP) and basal respiration. Panel C) ATP generation was calculated as the difference between basal respiration and respiration following treatment with 1μM oligomycin A. The asterisk (*) indicates a value significantly different than vehicle control (p<0.05). The hash (#) indicates a value significantly different than HBS-Control (p ≤ 0.05). Values are means + S.E.M. (n = 3, 4-5 replicates per n). 143 Figure 5.4 cont’d 144 Figure 5.5. Effect of MeHg in the absence or presence of extracellular Ca2+ on concentrations of intracellular metabolites. Undifferentiated PC12 cells were treated with 0µM (white bars) or 2µM (black bars) MeHg in the absence (0- Ca2+ HBS) or presence (HBS) of extracellular Ca2+ for 60 min, and concentrations of DOPAC (Panel A) and DOPET (Panel B) were measured. The asterisk (*) indicates a value significantly different from vehicle medium (p ≤ 0.05). Values are means + SEM (n = 4, 3 replicates per n). 145 5.4. Discussion Ca2+ plays essential roles in many cellular functions including neurotransmitter release and intracellular signaling (Olivera et al., 1994; Dunlap et al., 1995). Because high concentrations of cytosolic Ca2+ can initiate cell death cascades, the concentration of intracellular Ca2+ must be tightly regulated (Kass and Orrenius, 1999). A nonphysiologically high concentration of Ca2+ in the cytosol is a marker of pathology. MeHg increases the concentration of intracellular Ca2+, which contributes to its toxicity (Kauppinen et al., 1989; Marty and Atchison, 1998). Both extracellular and intracellular sources of Ca2+ have been implicated in MeHg-induced elevation of intracellular Ca2+ (Denny et al., 1993; Marty and Atchison, 1997). The present study sought to elucidate the role of extracellular Ca2+ as a primary mechanism mediating MeHg targeting of DA release and metabolism in PC12 cells, as well as impaired mitochondrial respiration and ATP generation induced by the toxicant. The principal findings from the present work are: 1) extracellular Ca2+ influx through VGCCs at rest does not contribute to either basal or MeHg-mediated DA release, 2) lack of extracellular Ca2+ enhances mitochondrial respiration and other parameters of bioenergetics including ATP generation and spare respiratory capacity, 3) mitochondrial dysfunction induced by MeHg is not dependent on extracellular Ca2+, and 4) basal DA metabolism and changes induced by MeHg are not dependent upon the presence of extracellular Ca2+. 146 5.4.1. Role of extracellular Ca2+ in spontaneous and MeHg-mediated DA release Spontaneous basal DA release from PC12 cells was independent of extracellular Ca2+ because removal of Ca2+ from the extracellular medium, in the presence of a chelating agent, did not reduce the concentration of extracellular DA. Although DA release from PC12 cells is classically described as Ca2+-dependent (Greene and Rein, 1977a), the source of Ca2+ can be intracellular (Kishimoto et al., 2005). Therefore, spontaneous DA release from PC12 cells is most likely modulated by changes in cytosolic Ca2+ independent of extracellular Ca2+ influx. MeHg induces a biphasic rise in intracellular Ca2+. Initially Ca2+ is released from intracellular stores followed by Ca2+ entry from the extracellular space (Marty and Atchison, 1997). Removal of Ca2+ from the medium attenuates the rise in extracellular DA following exposure to MeHg, demonstrating a partial dependence on extracellular Ca2+. This observation suggests that second phase Ca2+ influx following exposure to the toxicant contributes to DA release. However, further analysis of these data reveal that the contribution of extracellular Ca2+ is minimal; MeHg induced comparable increases in extracellular DA in the absence and presence of extracellular Ca2+. These data are consistent with previous reports in the literature demonstrating that spontaneous DA release is diminished, but not abolished, in a Ca2+-free superfusate 147 (Kalisch and Racz, 1996). A large contribution of extracellular Ca2+ to the observed MeHginduced DA release was not expected, considering that release was not evoked by depolarization in the present study. Rather, the phenomenon is more likely due to release of Ca2+ from intracellular stores that results during first phase Ca2+ influx following exposure to MeHg (Marty and Atchison, 1997). Alternatively, inhibition of the Na+/K+ ATPase by MeHg produces an increase in intracellular Ca2+ that could also contribute to DA release (Berg and Miles, 1979). 5.4.2. Role of VGCCs in spontaneous and MeHg-mediated DA release Neurotransmitter release occurs is two phases, fast synchronous and slow asynchronous (Varma et al., 2003). The synchronous phase is driven by a precisely timed presynaptic Ca2+ current initiated by the arrival of an action potential, and this mediates a large, fast post-synaptic response (Llinas et al., 1981; Sabatini and Regehr, 1996). The asynchronous phase provides tonic neurotransmitter release at most synapses and results from residual Ca2+ remaining in the axon terminal after an action potential (Atluri and Regehr, 1998; Lu and Trussell, 2000). The observation that inhibition of VGCCs did not impair basal DA release in the present study is aligned with this hypothesis. Under physiological conditions spontaneous DA release in undifferentiated PC12 cells is primarily mediated by residual Ca2+ present in the cell, and not VGCC-mediated Ca2+ influx. 148 Treatment with nimodipine or Cd2+ did not attenuated MeHg-induced DA release. Hence, VGCCs do not contribute to catecholamine release induced by MeHg. These data support the conclusion that extracellular Ca2+ does not contribute to MeHg-induced neurotransmitter release. Although MeHg interacts with and impairs VGCC function (Shafer et al., 1990; Shafer and Atchison, 1991b; Sirois and Atchison, 2000; Hajela et al., 2003), there is little evidence to suggest MeHg is mediating extracellular Ca2+ influx through VGCCs. Instead, DA release induced by MeHg may result from the release of Ca2+ from intracellular stores. However it is also possible that there is an increase in Ca2+ influx, but the level of Ca2+ is not high enough to activate pathways responsible for vesicular release (Atchison, 1986; 1987). 5.4.3. Role of extracellular Ca2+ in mitochondrial bioenergetics and targeting by MeHg The relationship between extracellular Ca2+ and mitochondrial respiration has only recently been established by one report in the literature. In an elegant study, Zhdanov et al. (2010) demonstrated a marked, transient activation of respiration in differentiated PC12 cells in response to depletion of extracellular Ca2+. The authors established that depletion of extracellular Ca2+ with the chelator EGTA slightly decreased mitochondrial membrane potential, and decreased concentrations of Ca2+ in mitochondria and the cytosol. Furthermore treatment with the uncoupler FCCP increased the respiratory response to 149 EGTA by 80-90% (Zhdanov et al., 2010). In agreement, the present results support and extend these findings. While Zdhanov et al. reported an immediate, yet transient increase in oxygen consumption following exposure to EGTA, the present results suggest that sustained elevation of mitochondrial respiration can be induced by prolonged incubation in a Ca2+-free solution. Additionally, the present work demonstrates that depletion of extracellular Ca2+ enhances other parameters of bioenergetics, including ATP generation and spare respiratory capacity. In addition to establishing a prominent respiratory response to extracellular Ca2+ depletion, Zdhanov et al. (2010) also investigated pathways mediating the effect. The authors demonstrate dependence on extracellular Na+ influx through VGCC, which activates mitochondrial Na+/Ca2+ exchange and causes the observed partial decrease in mitochondrial Ca2+. The excessive exchange of Na+ into the mitochondrial matrix would activate mitochondrial Na+/H+ exchange, and result in increased outward pumping of protons, electron transport, and oxygen consumption. In support of this model, the marked respiratory spike induced by EGTA was accompanied by acidification of the mitochondrial matrix, and inhibition of either the mitochondrial Na+/Ca2+ or Na+/H+ exchanger attenuated the response to extracellular Ca2+ depletion by ~50% (Zhdanov et al., 2010). Enhancement of ATP generation and spare respiratory capacity observed in the present study supports the general model proposed by these authors. Stimulated electron transport and respiration would activate the mitochondrial ATPase, and activation of 150 Na+/H+ exchange would provide the proton-motive force necessary for ATP synthesis. The spare respiratory capacity would result naturally from the galvanized system. MeHg abolishes the activation of respiration induced by depletion of extracellular Ca2+. Although the underlying mechanism of this effect is not known, inhibition of VGCCs by MeHg (Shafer and Atchison, 1991b) could prevent Na+ influx. Furthermore, direct interaction of MeHg with the mitochondria would inhibit respiratory proteins (Verity et al., 1975) and dissipate the mitochondrial membrane potential (Bondy and McKee, 1991; Hare and Atchison, 1992; Limke and Atchison, 2002). As a result, even if Na+ influx occurred, cation exchange required for activation of respiration would be impeded as postulated by Zhdanov et al. (2010). The observation that extracellular Ca2+ does not contribute to MeHg-induced mitochondrial dysfunction verifies other reports in the literature. ROS formation mediated by MeHg exposure is not affected by removal of extracellular Ca2+ (Dreiem and Seegal, 2007). These results support the conclusion that MeHg targeting of mitochondria is a primary event in cellular toxicity, and not secondary to perturbation of intracellular Ca2+ homeostasis. However because preventing the increase in intracellular Ca2+ can attenuate or delay cell toxicity (Marty and Atchison, 1998; Gassó et al., 2001), release of Ca2+ from intracellular stores could contribute to MeHg-induced mitochondrial damage. 151 5.4.4. Role of extracellular Ca2+ in basal and MeHg-impaired DA metabolism Extracellular Ca2+ does not contribute to basal DA metabolism. To date, there are no reports investigating this relationship. Although the present conclusion was anticipated, a role for extracellular Ca2+ in DA metabolism may have been uncovered. Ca2+ could either directly or indirectly modify metabolism by interacting with MAO and/or ALDH. Ca2+ dependent phosphorylation or de-phosphorylation of these enzymes could regulate enzymatic activity. Indeed, protein kinase C, which can be activated by increased concentrations of Ca2+ (Farah and Sossin, 2012), translocates to the mitochondrial matrix and activates ALDH under conditions of oxidative stress (Churchill et al., 2009). However there is no demonstrated role for Ca2+-dependent phosphorylation or de-phosphorylation in MAO or ALDH regulation. It is also possible that because the mitochondria play an integral role in both MAO and ALDH function (Schnaitman et al., 1967; Marchitti et al., 2008), changes in Ca2+ buffering induced by removing extracellular Ca2+ could alter enzyme function due to changes in mitochondrial membrane potential. While chelation of extracellular Ca2+ does alter mitochondrial membrane potential (Zhdanov et al., 2010), the magnitude of this decrease is only ~5mV. Therefore, it is likely that more substantial changes in mitochondrial function would be necessary to induce correlative changes in MAO or ALDH activity. 152 Extracellular Ca2+ does not contribute to altered DA metabolism induced by MeHg. Because MeHg-induced mitochondrial dysfunction was also unaltered by the absence of extracellular Ca2+, these observations support the hypothesis that mitochondrial dysfunction is a primary mechanism responsible for MeHg-impaired DA metabolism. However, additional investigation is needed to confirm a relationship between the two factors, and to determine if Ca2+ released from intracellular stores contributes to changes in DA metabolism following exposure to MeHg. 5.4.5. Summary The present work contributes to the understanding of MeHg targeting of DA homeostasis by establishing that extracellular Ca2+ does not play a role in either MeHginduced DA release or MeHg-impaired DA metabolism. While removal of extracellular Ca2+ can stimulate mitochondrial respiration, MeHg targeting of the organelle abolishes this activation. Because MeHg increases intracellular Ca2+ by influx of extracellular Ca2+, as well as release from intracellular stores, additional studies are needed to examine the contribution of intracellular Ca2+ to MeHg-induced DA release, and to determine in mitochondrial dysfunction or perturbation of intracellular Ca2+ homeostasis is a primary event in MeHg-impaired DA metabolism. 153 Chapter 6. The role of intracellular Ca2+ in MeHg-induced alterations in DA release and metabolism 6.1. Introduction 6.1.1. Regulation of intracellular Ca2+ Free intracellular Ca2+ plays many important physiological roles in neurons, including protein cofactor, electric charge carrier, and diffusible intracellular messenger (Carafoli, 1987; Brini et al., 2013). The ability of Ca2+ to contribute to these processes results partly from its differential distribution across the plasma membrane. On the extracellular side, Ca2+ concentrations range from 1-2mM, whereas free cytosolic Ca2+ is maintained at ~100nM (Kass and Orrenius, 1999). This distribution results in a steep electrochemical gradient that is maintained by a sophisticated network of transport and sequestration mechanisms that balance the elevation and diminution of cytosolic Ca2+ (Tsien and Tsien, 1990; Pozzan et al., 1994). Processes that increase intracellular Ca2+ include the influx of extracellular Ca2+ and the release of Ca2+ sequestered in organelles. Processes that reduce intracellular Ca2+ involve extrusion and sequestration. The plasma membrane contains channels, transporters, and exchangers that mediate Ca2+ flux. Ca2+ enters excitable cells primarily through VGCCs, however ligandgated channels permeable to Ca2+, such as the nicotinic ACh receptor and the N-methyl-D- 154 aspartate receptor, also participate (Tsien and Tsien, 1990; Llinas et al., 1992; Denny and Atchison, 1996). Influx occurs readily and does not require energy input because Ca2+ diffuses down its concentration gradient. Extrusion pathways include the Ca2+-ATPase and the Na+/ Ca2+ exchanger (Michaelis et al., 1987; Sanchez-Armass and Blaustein, 1987; Carafoli et al., 1996). Both of these processes require energy. The Ca2+-ATPase uses ATP to pump Ca2+ against its gradient and out of the cell. The Na+/ Ca2+ exchanger uses energy associated with the influx of Na+ down its gradient to drive Ca2+ efflux, but is also dependent upon ATP because the Na+/K+ ATPase maintains the Na+ gradient. Intracellular Ca2+ buffering organelles are also responsible for maintaining Ca2+ homeostasis. The most prominent Ca2+ buffering organelles are the SER and the mitochondria (Somlyo et al., 1985; Richter and Kass, 1991; Pozzan et al., 1994). The SER is a high-affinity, low capacity sequestration site, and participates in Ca2+ buffering within a physiological range. A Ca2+-ATPase present on the SER membrane transports Ca2+ from the cytosol into the lumen of the SER, whereas IP3 receptors and Ca2+-induced Ca2+release from ryanodine receptors mediate Ca2+ release from the SER (McPherson and Campbell, 1993; Pozzan et al., 1994). Mitochondria represent a low-affinity, high-capacity sequestration organelle, and require a powerful, pathologically high intracellular Ca2+ stimulus to begin buffering Ca2+ (Nicholls and Akerman, 1982). The Ca2+ uniporter is 155 responsible for the uptake of Ca2+ into the mitochondrial matrix, whereas release is accomplished by either reversal of the uniporter, Na+/Ca2+ exchanger, or opening of the mitochondrial permeability transition pore (Nicholls and Scott, 1980; Deryabina et al., 2004). 6.1.2. Effects of MeHg on intracellular Ca2+ homeostasis MeHg disrupts intracellular Ca2+ homeostasis. This effect was first indicated by studies of the neuromuscular junction, which demonstrated an increase in spontaneous ACh release following application of MeHg (Atchison and Narahashi, 1982; Atchison, 1986). Imaging studies using the fluorophore fura-2have confirmed that MeHg perturbs intracellular Ca2+ concentrations (Hare et al., 1993; Marty and Atchison, 1997). MeHg induces a biphasic rise in intracellular Ca2+, indicated by alterations in the ratio of fura-2 bound to intracellular Ca2+. This has been observed in many cell types, including neuroblastoma cells (Hare et al., 1993; Hare and Atchison, 1995b), rat forebrain synaptosomes (Komulainen and Bondy, 1987; Denny et al., 1993), and rat cerebellar neurons (Sarafian et al., 1994; Marty and Atchison, 1997; Mundy and Freudenrich, 2000; Edwards et al., 2005). First, Ca2+ is released from intracellular stores. This produces a slow, gradual rise in cytosolic Ca2+ that is within the cellular buffering capacity (Marty and Atchison, 1997). A second Ca2+ phase follows whereby extracellular Ca2+ influx overwhelms the cellular buffering capacity, which contributes to the initiation of cytotoxic 156 events (Kass and Orrenius, 1999). Indeed, inhibition of VGCC channels not only delays the onset of MeHg-induced intracellular Ca2+ elevation, but also decreases the incidence of cell death in cerebellar granule cells (Marty and Atchison, 1998) and the appearance of neurological symptoms associated with MeHg intoxication in rats (Sakamoto et al., 1996). 6.1.3. Effects of MeHg on DA release and metabolism Ca2+ plays an essential signaling role in vesicle docking and neurotransmitter release (Llinas et al., 1981; Sabatini and Regehr, 1996). While influx through VGCCs is the key event in depolarization-evoked vesicular exocytosis (Atluri and Regehr, 1998; Lu and Trussell, 2000), residual cytosolic Ca2+ can participate in spontaneous neurotransmitter release (Chapter 5; Tiernan et al., 2013). As demonstrated previously, spontaneous MeHginduced DA release is dependent upon vesicular exocytosis (Chapter 3), and extracellular Ca2+ does not mediate this effect (Levesque and Atchison, 1987; 1988). Thus intracellular Ca2+ may be the key signaling mechanism responsible for MeHg-induced DA release. Studies at the neuromuscular junction have characterized a role for intracellular Ca2+ in MeHg-induced ACh release (Levesque and Atchison, 1988). Specifically, agents that disrupt Ca2+ transport mechanisms in the mitochondria preclude MeHg from inducing ACh release, whereas agents that either disrupt Ca2+ buffering by the SER or Na+-dependent Ca2+ mobilization in the nerve terminal are ineffective (Levesque and Atchison, 1988). A role for intracellular Ca2+ in the spontaneous release of other neurotransmitters has yet to 157 be defined. Furthermore, because mitochondrial dysfunction contributes to ACh release (Tsien, 1981), and MeHg-impaired DA metabolism (Chapter 4), the relationships between intracellular Ca2+, aberrant DA metabolism, and mitochondrial dysfunction were also investigated in the present study. 6.2. Methods 6.2.1. Chemicals The standard physiological saline used for extracellular solution was HBS, which contained (mM): 150 NaCl, 5 KCl, 2.4 CaCl2, 1.6 MgSO4, 20 HEPES, and 20 d-glucose (pH 7.3). In some experiments cells were treated in 0-Ca2+ HBS, which had the same composition as HBS with the following exceptions (mM): 0 CaCl2 and 0.02 ethylene glycol tetraacetic acid. MeHg (ICN Biochemicals) and BAPTA/AM (Calbiochem) were diluted from stock solutions in HBS or 0-Ca2+ HBS. Drug solutions were diluted to final concentrations on the day of each experiment as described in General Methods Section 2.2. 6.2.2. Treatment for neurochemistry and mitochondrial bioenergetics PC12 cells were seeded at a density of 6-7x105 cells/mL in 6-well plates or XF24well plates coated with poly-D-lysine 24-48 hr prior to treatment. To begin each experiment, culture medium was aspirated and replaced with HBS in the absence or presence of 50μM BAPTA/AM for 30 min, and then 0μM or 2μM MeHg in the absence or 158 presence of 50μM BAPTA/AM in HBS or 0-Ca2+ HBS for 60 min. Cells were incubated at 37°C/5% CO2 during treatment as described in General Methods Section 2.2. 6.2.3. Neurochemistry At experiment termination, treatment medium was retained and acidified. Cells were rinsed, harvested, and pelleted by centrifugation. The content of DA in the treatment medium and its metabolites in the cell pellet were determined by HPLC-ED as described in General Methods Section 2.3. Neurochemical concentrations were normalized to mL per sample for extracellular measurements or mg of protein for intracellular measurements as determined by the BCA protein assay (Sigma). 6.2.4. Mitochondrial bioenergetics Following intracellular Ca2+ chelation and toxicant exposure, treatment medium was aspirated. Cells were washed once with XF24 RPMI Assay Medium and then incubated at 37°C/no CO2 for 1 hr to allow medium temperature and pH to equilibrate before the first measurements were made. The biosensor cartridge was hydrated overnight and injection ports were loaded with test compounds as described in General Methods Section 2.6. OCR was measured under basal conditions and following injection of test compounds using the XF24 Extracellular Flux Analyzer. Basal respiration, ATP generation, and spare respiration were calculated and compared between treatment groups. 159 6.2.5. Measurement of fura-2 fluorescent change PC12 cells were plated at 7x105 cells/mL on circular glass coverslips 24 hr prior to analysis. Cells were preloaded with 2μM fura-2 in the presence of 50μM BAPTA/AM. Following fura-2 loading and intracellular Ca2+ chelation, coverslips containing PC12 cells were mounted on an inverted microscope (Diaphot-TMD, Nikon, Toyko, Japan) and perfused with 0-Ca2+ HBS at 37°C. Digital fluorescent images were obtained using an IonOptix system (Milton, MA) and fura-2 fluorescence changes were monitored as described in General Methods Section 2.9. 6.2.6. Statistical analysis SigmaPlot® software version 12.0 (SysStat Software, Inc., Point Richmond, CA) was used to make statistical comparisons among groups using unpaired t-test, one-, and twoway ANOVA, or non-parametric alternatives as appropriate. If a significant difference was detected, post hoc between-group comparisons were performed using Tukey’s test. Statistical significance was set at p≤0.05. 6.3. Results 6.3.1. MeHg-induced DA release is dependent upon the presence of intracellular Ca 2+ BAPTA/AM is a selective intracellular Ca2+ chelator. The presence of an acetoxymethyl ester group (AM) allows the molecule to traverse biological membranes. Once inside the cell, the ester is cleaved by endogenous esterases, preventing BAPTA from 160 exiting the cell (Limke and Atchison, 2009). To investigate the role of intracellular Ca2+ in MeHg-induced DA release, undifferentiated PC12 cells were loaded with 50μM BAPTA/AM to chelate intracellular Ca2+, and then treated with 2μM MeHg for 60 min. This experiment was performed in both the absence and presence of extracellular Ca2+ to assess the interplay between extracellular and intracellular Ca2+ pools. The results of the present study shown in Figure 6.1 are consistent with previous results showing that 2μM MeHg increased DA release from undifferentiated PC12 cells (Chapter 3) and the removal of extracellular Ca2+ did not appreciably alter this effect (Chapter 4). In the presence of extracellular Ca2+, pretreatment with BAPTA/AM did not alter basal DA release, but did attenuate MeHg-induced release (Figure 6.1; p<0.05). In the absence of extracellular Ca2+, a similar profile was observed. However, more stringent examination of the percentage change induced by MeHg in each treatment group demonstrated that only chelation of both extracellular and intracellular Ca2+ significantly diminished MeHg-induced DA release in undifferentiated PC12 cells (p<0.05). To confirm that BAPTA/AM attenuated the elevation in intracellular Ca2+ induced by MeHg, imaging studies were performed using single cell microfluorometry with the ratiometric Ca2+ indicating dye fura-2. In response to increasing Ca2+ concentrations, fura2 shows a peak signal at 340nm and minimum signal at 380nm. The ratio of peak 340 signal/ peak 380 signal is an accurate assessment of the concentration of intracellular Ca2+ 161 (Marty and Atchison, 1997). In the absence of extracellular Ca2+, the 340/380 fura-2 ratio increased gradually following application of 2μM MeHg (Figure 6.2), consistent with the first Ca2+ phase of MeHg exposure (Marty and Atchison, 1997). Preloading cells with 50μM BAPTA/AM for 30 min abolished this effect. 162 Figure 6.1. Intracellular Ca2+ partially contributes to MeHg-induced DA release. Undifferentiated PC12 cells were treated with 50µM BAPTA/AM in HBS for 30min prior to co-treatment with 0µM (white bars) or 2 µM (black bars) MeHg in the absence (Ca2+-free HBS) or presence (HBS) of extracellular Ca2+ for 60 min. Panel A) Concentrations of extracellular DA. Panel B) The percentage change of extracellular DA after treatment with MeHg in cells treated with either HBS or Ca2+-free HBS in the absence or presence of 50µM BAPTA/AM. The asterisk (*) indicates a value significantly different from HBS-Control (p ≤ 0.05). The hash (#) indicates a value significantly difference from both HBS-Control and HBS-MeHg (p ≤ 0.05). Values are means + SEM (n = 4, 3 replicates per n). 163 Figure 6.1 cont’d 164 Figure 6.2. Representative traces showing changes in the ratio of fura-2 intensity. Undifferentiated PC12 cells were pre-incubated with 2μM fura-2/AM in absence (orange) or presence (red) of 50μM BAPTA/AM for 30 min. Following pre-incubation, cells were perfused with 0-Ca2+ HBS. 2μM MeHg was added where indicated. Pre-treatment with BAPTA/AM prevented the rise in fura-2 fluorescence intensity induced by MeHg. No second phase of MeHg-induced increase in fura-2 intensity was observed because all cells were perfused in the absence of extracellular Ca2+. 165 6.3.2. Intracellular Ca2+ is essential for the maintenance of mitochondria bioenergetics To evaluate the contribution of different Ca2+ sources to mitochondrial dysfunction induced by MeHg, mitochondrial respiration was measured in undifferentiated PC12 cells treated with 2μM MeHg for 60 min in the absence or presence of intracellular and/or extracellular Ca2+. Following toxicant treatment, OCR was measured with the XF24 respirometer under basal conditions and following treatment with 5mM pyruvate, 1μM oligomycin A, 2μM FCCP, and 1μM antimycin A. Figure 6.3 depicts changes in OCR following treatment with these compounds. Quantification of these changes (Figure 6.4) revealed an essential role for intracellular Ca2+ source in maintaining mitochondrial respiratory function. MeHg attenuated basal respiration, spare respiratory capacity, and ATP generation, as demonstrated previously (Chapter 4; p<0.05). Chelation of extracellular Ca2+ alone was insufficient to reverse these effects (Chapter 5). However pre-treatment with BAPTA/AM caused marked decreases in basal respiration and ATP generation, and complete abolition of spare respiratory capacity (p<0.05). 166 Figure 6.3. Changes in mitochondrial bioenergetics induced by MeHg in absence or presence of either intracellular or extracellular Ca2+. Undifferentiated PC12 cells were pretreated for 30 min in the absence or presence of 50μM BAPTA/AM in HBS followed by a 60 min with HBS (red circle), 2μM MeHg in HBS (orange circle), 50μM BAPTA/AM in HBS (yellow triangle), 2μM MeHg + 50μM BAPTA/AM in HBS (green triangle), 0-Ca2+ HBS (green square), 2μM MeHg in 0-Ca2+ HBS (blue square), 50μM BATPA/AM in 0-Ca2+ HBS (purple diamond) or 2μM MeHg + 50μM BAPTA/AM in 0-Ca2+ HBS. Following pre-treatment, OCR (pMoles/min) was measured under basal conditions and following injections of 5mM pyruvate (A), 1μM oligomycin (B), 2μM FCCP (C), and 1μM antimycin A (D). Values are means ± S.E.M. (n = 3, 4-5 replicates per n). 167 Figure 6.4. Quantification of changes in mitochondrial bioenergetics induced by MeHg in absence or presence of either intracellular or extracellular Ca2+. Panel A) Basal respiration (OCR) was measured in the presence of 25mM glucose and 5mM pyruvate. Panel B) Spare respiration was calculated as the difference between maximum respiration (following treatment with 2μM FCCP) and basal respiration. Panel C) ATP generation was calculated as the difference between basal respiration and respiration following treatment with 1μM oligomycin A. The asterisk (*) indicates a value significantly different than vehicle control (p<0.05). The hash (#) indicates a value significantly different than HBS-Control (p ≤ 0.05). Values are means ± S.E.M. (n = 3, 4-5 replicates per n). 168 Figure 6.4 cont’d 169 6.3.3. Aberrant DA metabolism induced by MeHg is not dependent upon intracellular Ca2+ Chelating extracellular and/or intracellular Ca2+ had no effect on intracellular DOPAC formation (Figure 6.5). Concentrations of intracellular DOPET were markedly decreased by removal of intracellular Ca2+ (p<0.05). Treatment with 2μM MeHg for 60 min significantly diminished DOPAC and increased DOPET formation regardless of the absence or presence of intracellular or extracellular Ca2+ (p<0.05). 170 Figure 6.5. Effect of MeHg in the absence or presence of intracellular or extracellular Ca2+ on concentrations of intracellular metabolites. Undifferentiated PC12 cells were pre-treated with vehicle or 50μM BAPTA/AM in HBS for 30 min prior to co-treatment with 0µM (white bars) or 2µM (black bars) MeHg in the absence (Ca2+-free HBS) or presence (HBS) of extracellular Ca2+ for 60 min. Panel A) DOPAC and Panel B) DOPET. The asterisk (*) indicates a value significantly different from vehicle within treatment group (p ≤ 0.05). The hash (#) indicates a value significantly different than 0μM MeHg in HBS (p ≤ 0.05). Values are means + SEM (n = 4, 3 replicates per n). 171 6.4. Discussion MeHg targets both intracellular and extracellular pools of Ca2+ to disrupt cytosolic Ca2+ homeostasis (Chapter 3; Tiernan et al., 2013). Studies presented herein were designed to complement studies presented in Chapter 5, which demonstrated that extracellular Ca2+ does not mediate MeHg targeting of DA homeostasis or impaired mitochondrial bioenergetics. The present study sought to elucidate the role of intracellular Ca2+ as a primary mechanism in these effects. These data are consistent with the following conclusions: 1) intracellular Ca2+ at least in part mediates MeHg-induced DA release, 2) intracellular Ca2+ plays a crucial role in maintaining mitochondrial respiration and other parameters of bioenergetics including ATP generation and spare respiratory capacity, 3) the reductive pathway of DA metabolism, but not the oxidative pathway, is dependent upon the presence of intracellular Ca2+, and 4) aberrant DA metabolism induced by MeHg is not dependent upon the presence of intracellular Ca2+. 6.4.1. Role of intracellular Ca2+ in spontaneous and MeHg-mediated DA release Chelating intracellular Ca2+ causes a small, non-significant decline in basal DA release, and in the absence of both intracellular and extracellular Ca2+ it is comparable to basal DA release. These data together suggest that tonic DA release from undifferentiated PC12 cells is Ca2+ independent. This conclusion is somewhat perplexing considering tonic 172 DA release from undifferentiated PC12 cells is dependent upon functional synaptic vesicles (Katz and Miledi, 1967; Bajjalieh and Scheller, 1995; Sudhof, 2012), and Ca2+-dependent vesicular exocytosis plays a pivotal role in neurotransmitter release (Di Virgilio et al., 1984; Barrowman et al., 1986; Drapeau and Nachshen, 1988; Mochida et al., 1998; Parnas et al., 2000; Caldwell et al., 2013). However there is some evidence to support Ca2+-independent vesicular exocytosis (Drapeau and Nachshen, 1988). Cytosolic acidification can stimulate DA release from striatal synaptosomes in the absence of extracellular Ca2+ (Mochida et al., 1998). Additionally, ACh release from cultured superior cervical ganglion neurons is stimulated following focal membrane depolarization with a hyperosmotic sucrose solution in a Ca2+-free medium. This solution evoked release even when intracellular Ca2+ was chelated with BAPTA (Knight et al., 1989). In these studies, neurotransmitter release still required some stimulus to induced release, however secretion can be induced by endogenous factors independent of Ca2+ (Barrowman et al., 1986). For example, guanosine-5’-triphosphate analogs induce secretion from neutrophils in the absence of extracellular or intracellular Ca2+ (Atchison, 1986; Levesque and Atchison, 1987; 1988), suggesting that a G-protein-mediated secretory mechanism exists that is separate from Ca2+-mediated exocytosis. The present data provide intriguing evidence for a Ca2+independent mechanism of vesicular exocytosis in undifferentiated PC12 cells and warrant further investigation. 173 MeHg-induced DA release is attenuated in the absence of intracellular Ca2+, and this effect is most pronounced in the absence of both extracellular and intracellular Ca2+. These results corroborate studies at the neuromuscular junction (Hare and Atchison, 1995a; Chetty et al., 1996) and suggest that MeHg-induced DA release is dependent upon an increase in free cytosolic Ca2+. Furthermore, intracellular and extracellular Ca2+ pools work in concert to stimulate DA release following exposure to MeHg. The source of intracellular Ca2+ responsible for MeHg-induced DA release is unknown but both the SER and the mitochondria could contribute to the elevation of intracellular Ca2+ induced by MeHg. In general, MeHg first stimulates Ca2+ release from an IP3-sensitive pool in the SER (Limke et al., 2003). This release activates mitochondrial Ca2+ uptake, and Ca2+ is initially buffered into the mitochondrial matrix (Levesque and Atchison, 1991; Limke et al., 2003). Following excessive accumulation of mitochondrial Ca2+ and dissipation of the mitochondrial membrane potential, Ca2+ is extruded from the mitochondrial matrix into the cytosol (Levesque and Atchison, 1991; Levesque et al., 1992). Studies at the neuromuscular junction have identified mitochondria as the primary intracellular sequestration organelle responsible for MeHg-induced ACh release (Hare and Atchison, 1995a). However depending on the system, the contribution of each Ca2+ buffering organelle to MeHg-induced elevation in cytosolic Ca2+ is different. For example, in neuroblastoma cells an IP3-sensitive pool is the main contributor (Limke et al., 2003), 174 whereas in cerebellar granule cells it is the mitochondria (Denton, 2009; Glancy and Balaban, 2012; Pizzo et al., 2012). Therefore while both the SER and mitochondria likely participate to elevate intracellular Ca2+ following exposure to MeHg in undifferentiated PC12 cells, the magnitude of their contributions to MeHg-induced DA release has yet to be elucidated. 6.4.2. Role of intracellular Ca2+ in mitochondrial bioenergetics Maintenance of intracellular Ca2+ homeostasis is essential for sustaining mitochondrial function. Chelating intracellular Ca2+ markedly diminishes respiration and ATP generation. These observations are aligned with a wealth of literature demonstrating the regulatory role of Ca2+ in ATP synthesis (Denton et al., 1972; Farah and Sossin, 2012). Indeed, Ca2+ activates at least three key enzymes that feed electrons in the ETC. Pyruvate dehydrogenase is stimulated when Ca2+ activates a phosphatase associated with the enzyme complex (Rutter and Denton, 1988; Mori et al., 2007; Jinsmaa et al., 2009), whereas isocitrate dehydrogenase, and α-ketoglutarate dehydrogenase are directly bound by Ca2+, which increases affinity for their substrates (Yoshino et al., 1966; Hamdy and Noyes, 1977; Yamada and Huzel, 1985; Hubbard and McHugh, 1996; Fox et al., 2012). Furthermore the mitochondrial ATP synthase is responsive to changes in cytosolic Ca2+, and Ca2+ readily binds the catalytic subunit of this enzyme (Schnaitman et al., 1967; Marchitti et al., 2008). Presently, buffering cytosolic Ca2+ would most likely cause the mitochondria to extrude 175 Ca2+ in an effort to maintain cytosolic Ca2+ levels. Loss of mitochondrial matrix Ca2+ would decrease activation of enzymes necessary for respiratory chain activity and ATP synthesis. Effects of buffering cytosolic Ca2+ and the resulting diminution of mitochondrial function have implications for the present work. No conclusions can be made concerning the role of intracellular Ca2+ in MeHg-impaired mitochondrial dysfunction. Furthermore, it is surprising that cellular functions, including DA release and metabolism, were not more severely affected by the loss of mitochondrial respiration and ATP generation. Perhaps the short time-course of the present experimental paradigm precluded the run-down of energy reserves and initiation of cell death. However no conclusions can be made regarding these events, as no measurement were taken to investigate effects of BATPA/AM on mitochondrial membrane potential, opening of the mitochondrial permeability transition pore, or markers of cell death. 6.4.3. Role of intracellular Ca2+ in basal and MeHg-impaired DA metabolism Intracellular DOPAC formation was not affected by buffering intracellular Ca2+ in the absence or presence of extracellular Ca2+. These data corroborate and extend results in Chapter 5 and suggest that canonical DA metabolism is independent of cytosolic Ca2+. Neither MAO nor ALDH are reported to require Ca2+ for enzymatic function. However because both enzymes are intimately associated with mitochondria (Petrash, 2004), the observation that DA metabolism proceeds in the face of a marked decrease in 176 mitochondrial respiration is confounding. One possible explanation is methodology. While neurochemistry can be measured from cell pellets harvested immediately after treatment, XF24 respirometry measurements are made during a time period of one to three hr following treatment. It is possible that mitochondrial dysfunction, as assessed using the XF24 respirometer, was not initiated until a time-point after neurochemical measurements were taken. The reductive pathway of DA metabolism is dependent upon the presence of intracellular Ca2+. DOPET formation was markedly reduced following treatment with BAPTA/AM. These data are novel in that a role for Ca2+ in AR function has yet to be established. There is some evidence to suggest that AR is a substrate for protein kinase C (Varma et al., 2003). Treatment with the protein kinase C stimulator bryostatin-1 directly increases phosphorylation of AR (Farah and Sossin, 2012), and because protein kinase C can be activated by Ca2+ (Mori et al., 2007; Jinsmaa et al., 2009), this represents one mechanism by which Ca2+ could modulate AR activity. Results presented herein are the first to demonstrate a direct effect of Ca2+ on DOPET formation. Exposure to MeHg was still able to elicit an aberrant DA metabolomic profile, even in the absence of both extracellular and intracellular Ca2+. These data suggest that MeHginduced perturbation of intracellular Ca2+ homeostasis does not contribute to changes in DA metabolism mediated by the toxicant. The observation that intracellular Ca2+ is required for mitochondrial bioenergetics, but not DA metabolism, suggests that the DA 177 metabolomic profile induced by MeHg is not in response to mitochondrial dysfunction. Therefore ALDH inhibition and decreased availability of its cofactor NAD could be due to oxidative stress induced by MeHg (Yoshino et al., 1966; Hamdy and Noyes, 1977; Fox et al., 2012) or inhibition of enzymes in the citric acid cycle that ultimately down-regulate the amount of NAD(H) (Haycock, 1990). 6.4.4. Summary The present work establishes a role for intracellular Ca2+ in MeHg-induced DA release, but suggests that aberrant DA metabolism induced by exposure to the toxicant is independent of altered intracellular Ca2+ homeostasis. Therefore, MeHg targets DA synthesizing cell by multiple mechanisms, including perturbation of intracellular Ca2+ homeostasis, impairment of mitochondrial function, and perhaps a yet to be elucidated mechanism responsible for inhibition of enzymes necessary for oxidative DA metabolism. 178 Chapter 7. General Discussion and Concluding Remarks Methylmercury is a potent bioaccumulative neurotoxicant. Despite general cellular and molecular targets, certain neuronal populations are selectively susceptible to toxicity. The present dissertation characterizes MeHg-induced toxicity in DA-synthesizing cells. Results describe how perturbation of intracellular Ca2+ and mitochondrial dysfunction induced by exposure to MeHg alter DA neurochemistry, and identify potential disturbances that could contribute to heightened susceptible of DA-synthesizing cells. A better understanding of DA neuronal targeting by MeHg elucidates how exposure to the environmental toxicant contributes to the etiology of PD. 7.1. Role of Ca2+ in MeHg-induced activation of DA synthesis Results from pharmacological studies in this dissertation demonstrate that MeHg stimulates DA synthesis and vesicular exocytosis, but does not interact with the membrane DA transporter, to induce DA release. Furthermore, the primary event initiating these effects is an increase in intracellular Ca2+. Ca2+ contributes to DA synthesis by potentiating phosphorylation of TH (Daubner et al., 2011). TH enzymatic activity is modulated by phosphorylation and dephosphorylation events at critical Ser residues, including Ser19, -31, and -40 (Lehmann et al., 2006). Phosphorylation at Ser40 directly increases enzymatic activity, whereas phosphorylation at Ser19 and -31 indirectly activate TH via a hierarchical mechanism, which increases the rate of phosphorylation at Ser40 (Albert et al., 1984; Vulliet et al., 1985). A multitude of protein kinases have been linked to TH phosphorylation, including PKC (Yamauchi and 179 Fujisawa, 1981; Vulliet et al., 1984; Atluri and Regehr, 1998; Lu and Trussell, 2000) and CaM-MPK (Levesque and Atchison, 1987; 1988; Haavik et al., 1989), both of which phosphorylate TH as Ser19 and -40 via a Ca2+ dependent mechanism (Haavik et al., 1989). Ca2+ has also been linked to dephosphorylation of TH. Two phosphatases, PP2A and PP2C account for 90% and 10%, respectively, of TH phosphatase activity (Wlodarchak et al., 2013). The effect of Ca2+ on PP2A activity in TH de-phosphorylation has not been investigated, however there is evidence to suggest its activity is regulated by Ca2+ (Bevilaqua et al., 2003). Specific investigation of PP2C has demonstrated that its activity is stimulated by certain divalent cations, including Mn2+ and Mg2+, however Ca2+ inhibits its activity (Olivera et al., 1994; Dunlap et al., 1995).. The ability of Ca2+ to facilitate phosphorylation and hinder de-phosphorylation indicates one potential mechanism by which MeHg stimulates DA synthesis. Western blot analysis of TH and pTH Ser40 in this dissertation demonstrate that MeHg increases phosphorylation of TH, without altering total protein. Phosphorylation is an acute regulatory mechanism, whereas protein synthesis provides chronic control. These data suggest that MeHg can regulate TH activity in an acute fashion, immediately following exposure. The precise role of Ca2+ has yet to be elucidated, however data presented herein support the hypothesis that MeHg stimulates TH phosphorylation via a Ca2+-dependent mechanism. First, MeHg-mediated DA release is dependent upon the presence of intracellular Ca2+. Secondly, de novo DA synthesis is necessary for MeHg-induced DA 180 release. Lastly, MeHg stimulates TH activity, as assessed by DOPA accumulation and Western blot. To investigate whether MeHg activates TH via a Ca2+-dependent mechanism, future analysis could include quantification of pTH Ser40 (and possibly pTH Ser19) by Western blot, as well as changes in DOPA accumulation following exposure to MeHg under Ca2+-free conditions. 7.2. Role of Ca2+ in MeHg-induced vesicular exocytosis The present studies illustrate a role for vesicular exocytosis in MeHg-induced DA release. Moreover, MeHg mobilizes vesicular DA stores in a Ca2+-dependent manner. Because MeHg was able to induce release in either the absence of extracellular Ca2+ or following inhibition of VGCC, extracellular Ca2+ does not appear to play an integral role. However, MeHg-induced DA release is attenuated in the absence of intracellular Ca2+, and this effect is enhanced in the absence of extracellular Ca2+. Taken together, these results suggest that release of Ca2+ from intracellular stores is the primary event mediating MeHginduced vesicular DA release, and influx of extracellular Ca2+ plays a secondary role promoting release. Ca2+ plays a ubiquitous role in vesicular exocytosis. Although neurotransmitter release is generally mediated by extracellular Ca2+ influx through VGCC (Atluri and Regehr, 1998; Lu and Trussell, 2000), intracellular Ca2+ can initiate exocytosis (Levesque and 181 Atchison, 1987; 1988). It follows that the elevation in intracellular Ca2+ induced by MeHg triggers release of DA-containing vesicles. However, this hypothesis is currently based on corollary data. Additional work is needed to demonstrate a direct effect of MeHg-induced intracellular Ca2+ elevation on vesicular DA release. Confocal imaging studies using the fluorescent membrane probe FM1-43 in combination with a Ca2+ indicating dye such as Fluo-4 would be able to visually and quantitatively illustrate a proximal and functional relationship between Ca2+ elevation and vesicular exocytosis. 7.3. Collaborative relationship between extracellular and intracellular Ca 2+ pools Studies investigating the source of Ca2+ that mediates MeHg-induced DA release suggest that release of Ca2+ from intracellular stores is the key event. At the neuromuscular junction, disruption of Ca2+ transport mechanisms into the mitochondria precluded MeHg from inducing Ca2+ release and subsequent ACh release (Levesque and Atchison, 1988). Agents that disrupted Ca2+ buffering by the SER were ineffective (Limke et al., 2004). These data suggest that mitochondria, rather than SER, play the critical role in MeHg-induced ACh release. However, studies investigating the contributions of different intracellular pools to the elevation in cytosolic Ca2+ induced by MeHg suggest that alterations in Ca2+ handling are complex and differ depending on cell type. 182 MeHg initiates a cascade of events that culminates in elevated cytosolic Ca2+ (Bearss et al., 2001). In cerebellar granule cells (Hare and Atchison, 1995a) and neuroblastoma cells (Hare and Atchison, 1995a) at least part of this Ca2+ originates from the SER, however the magnitude of contribution differs. In some cell types, such as neuroblastoma cells, the SER supplies the majority of released intracellular Ca2+. Treatment with the SER Ca2+ ATPase inhibitor thapsigargin depletes the SER Ca2+ content and reduces the amplitude of intracellular Ca2+ elevation by almost 70% (Bearss et al., 2001). In other cell types, such as cerebellar granule cells, the SER has a more limited storage capacity and treatment with thapsigargin, only reduces the amplitude of intracellular Ca2+ elevation by 30% (Limke et al., 2003). Mitochondria contribute the bulk of intracellular Ca2+ in these cell types. Prior depletion of mitochondrial Ca2+ with mitochondrial membrane depolarization reduces the amplitude of Ca2+ elevation by ~65% (Kass and Orrenius, 1999). Neurons are constantly buffering large fluctuations of intracellular Ca2+ (Marty and Atchison, 1997), and as such have similar Ca2+ buffering systems in place. Because DAsynthesizing cells more closely resemble cerebellar granule cells, than neuroblastoma cells, it follows that Ca2+ handling would also be more similar in these cell types. Consequently, the intracellular Ca2+ pool responsible for MeHg-induced DA release is most likely the mitochondria. However, there are differences between DA-synthesizing cells and cerebellar granule cells. Experiments elucidating the source of intracellular Ca2+ that initiates MeHg183 induced DA release are necessary to confirm this hypothesis. Pharmacological manipulation of SER and mitochondrial Ca2+ pools in combination with Ca2+ imaging (i.e. fura-2 fluorescence changes) and neurochemistry could be used to evaluate sources of intracellular Ca2+ that trigger MeHg-induced DA release. Buffering extracellular Ca2+ did not significantly attenuate MeHg-induced DA release, whereas buffering intracellular Ca2+ did. However the ability of intracellular Ca2+ to stimulate DA release was augmented by the presence of extracellular Ca2+, because MeHg-induced DA release in the absence of both sources was the only condition that significantly attenuated the percentage change in extracellular DA induced by MeHg. These data suggest that different Ca2+ sources collaborate to stimulate neurotransmitter release. Because VGCCs did not contribute to release in the present studies, other membrane channels permeable to Ca2+ or membrane depolarization could mediate extracellular Ca2+ influx. Certain subtypes of the glutamate N-methyl-D-aspartate receptor are permeable to Ca2+, however evidence suggests that inhibition of these channels does not attenuate the elevation in intracellular Ca2+ induced by MeHg (Saito et al., 2003). Furthermore, PC12 cells do not express functional N-methyl-D-aspartate receptors (Patrick and Barchas, 1976), making it unlikely that these channels would contribute to extracellular Ca2+ influx in the present model. PC12 cells do, however, express nicotinic ACh receptors (Bondy and McKee, 1991). Although MeHg interacts with these channels to increase intracellular Ca2+ 184 in PC12 cells, evidence suggests that cellular effects mediated by this channel are related to an intracellular mechanism rather than Ca2+ entry (personal communication, Dr. William Atchison). There may be a role for membrane depolarization in MeHg-induced DA release. MeHg depolarizes the plasma membrane (Limke et al., 2004). Furthermore, there is a temporal relationship between membrane depolarization and MeHg-induced mitochondrial dysfunction (Bemis and Seegal, 1999; Dreiem et al., 2009). Excessive accumulation of mitochondrial Ca2+ depletes ATP and inhibits its synthesis. Loss of ATP would inhibit plasma membrane transporters, including the Na+/K+ ATPase. Because this exchanger maintains the transmembrane potential, its inhibition would depolarize the plasma membrane, facilitating entry of extracellular Ca2+. Thus release of Ca2+ from mitochondrial stores coincides with membrane depolarization and influx of extracellular Ca2+. Together these events could initiate vesicular DA exocytosis. 7.4. Aberrant DA metabolism induced by MeHg Previous studies have established that MeHg inhibits oxidative DA metabolism (Taylor and DiStefano, 1976; Chakrabarti et al., 1998; Beyrouty et al., 2006; Castoldi et al., 2006), however this is the first study to indicate that this inhibitory profile culminates in the up-regulation of the reductive DA metabolic pathway to form DOPET, and accumulation of the toxic intermediate DOPAL. The present results demonstrate that, in an intact system, MeHg inhibits DA metabolism at the level of ALDH, and not MAO as previously asserted 185 (Goedde and Agarwal, 1990). However there are other possible explanations. For example, the aberrant DA metabolomic profile induced by MeHg in undifferentiated PC12 cells could be a cell-type specific response, such that in other immortalized cell lines or primary cultures, MAO and not ALDH may be impaired. Enzyme assays demonstrate that MeHg does not directly inhibit ALDH. PC12 cells only express the ALDH2 isoform, which is localized to the mitochondrial matrix (Marchitti et al., 2007). While ALDH2 plays an integral role in DA metabolism (Galter et al., 2003; Anderson et al., 2011b), another isoform ALDH1A1, localized to the cytosol, has been reported to catalyze DOPAL oxidation in NSDA neurons (Yoshino et al., 1966). Therefore it would be of interest to assess the potential contribution of ALDH1A1 to buffering DOPAL in NSDA neurons, and determine how MeHg interacts with this isoform in vivo. Perhaps different subcellular localizations or subtle differences in protein structure impart a contrasting interaction with MeHg that could influence NSDA neuronal susceptibility. Based on neurochemical results, it was hypothesized that MeHg inhibits ALDH activity indirectly by decreasing availability of the ALDH cofactor NAD, and that inhibition of the respiratory chain by MeHg mediates decreased synthesis of NAD. While there is some evidence to support this hypothesis, there is also indication that it is incorrect. MeHg markedly inhibits both basal respiration and spare respiratory capacity. However chelating intracellular Ca2+ severely impairs mitochondrial respiration, but DA metabolism and impairment by MeHg proceed without hindrance. Thus the original hypothesis must be rejected; mitochondrial dysfunction does not decrease the availability of NAD or contribute to aberrant DA metabolism induced by MeHg. 186 Decreased availability of NAD could result from many deleterious circumstances. MeHg inhibits the activity of certain enzymes in the citric acid cycle, including succinate dehydrogenase (Hamdy and Noyes, 1977) and malate dehydrogenase (LeBel et al., 1990; Sarafian et al., 1994; Gatti et al., 2004). Inhibition of malate dehydrogenase, in particular, would decrease oxidation of NADH to NAD. Additionally, inhibition of certain citric acid cycle enzymes would down-regulate overall activity of the cycle and further decrease NADH oxidation. Oxidative stress induced by MeHg (Singh et al., 2008) may also contribute to decreased NAD synthesis and impaired DA metabolism. There is evidence to suggest that under conditions of oxidative stress, NADH is converted to NADPH by a compensatory metabolic network involving NADH kinase and malic enzyme (Yee and Choi, 1996; Gatti et al., 2004; Cuello et al., 2010). Up-regulation of this network could account for the observed decrease in both NAD and NADH, however, it is unlikely that this pathway would provide a substantial defense against oxidative stress. NADPH is a cofactor for many endogenous antioxidant systems, including glutathione peroxidase and catalase, both of which are inhibited by MeHg (LeBel et al., 1990; Sarafian et al., 1994; Gatti et al., 2004). Oxidative stress and the production of free radicals induced by MeHg (Jinsmaa et al., 2009) may also contribute to aberrant DA metabolism. Products of oxidative stress, 4hydroxynonenal and malondialdehyde, significantly enhance DOPAL production in PC6-3 cells, a subline of the PC12 cell line (Franco et al., 2007; Mori et al., 2007; Grotto et al., 2011; Deng et al., 2012; Liu et al., 2012). Both compounds were found to inhibit DA metabolism and increase DOPAL production at low micromolar concentrations targeting the aldehyde oxidative and/or reductive pathways. Accumulation of these and other lipid peroxidation 187 products following MeHg exposure (Pearl et al., 2000) could contribute to DOPAL accumulation. Additional analysis is needed to confirm the inhibitory effect of decreased NAD availability on ALDH activity, and to identify potential roles for citric acid cycle inhibition and/or oxidative stress in these processes. Attempts to reverse aberrant DA metabolism with NAD add-back proved futile because NAD(H) can stimulate TH activity and DA release (Minnema et al., 1989; Gassó et al., 2000; Yuan and Atchison, 2007), confounding an analysis of the metabolomic profile. Perhaps treatment with the NAD precursor nicotinamide would overcome these confounding variables. Studies investigating the roles of oxidative stress and citric acid cycle activity could include antioxidant treatment, citric acid cycle substrate addition, and isolated enzyme activity assays. 7.5. Pathways by which MeHg targets DA neurons The present dissertation provides evidence for at least two pathways by which MeHg targets DA-synthesizing cells. A model depicting these mechanisms is illustrated in Figure 7.1. MeHg induces the release of Ca2+ from intracellular stores, including the mitochondria and SER, which elevates cytosolic Ca2+. This event has two consequences: 1) activation of Ca2+-dependent protein kinases, such as PKC and CaM-MPK, increases phosphorylation of TH at Ser19 and -40, which stimulates DA synthesis and increases the amount of cytosolic DA, and 2) Ca2+-dependent vesicular exocytosis triggers release of stored DA. Newly synthesized DA may be packaged into vesicles and released, however, stimulation of DA synthesis may overwhelm the capacity of VMAT to package DA. 188 Remaining cytosolic DA would be partitioned for metabolism. A second primary event mediated by MeHg, that may involve oxidative stress, inhibits oxidative DA metabolism. This event shunts DOPAL along the reductive metabolic pathway, however because AR has a lower affinity for DOPAL, metabolism proceeds more slowly and the toxic intermediate accumulates. 189 Figure 7.1. Proposed model identifying molecular targets of MeHg toxicity in DAsynthesizing cells. This dissertation has established at least two mechanisms by which MeHg alters DA homeostasis. 1) MeHg increases cytosolic Ca2+ primarily by causing release from intracellular stores. Elevated cytosolic Ca2+ activates protein kinases that phosphorylate TH and stimulate DA synthesis. Ca2+ in turn triggers vesicular exocytosis. 2) MeHg inhibits DA metabolism at the level of ALDH. The exact mechanism by which MeHg impairs ALDH activity remains unclear, however decreased availability of NAD is likely involved. Oxidative stress may also contribute. As a result of aberrant DA metabolism, a reductive DA metabolic pathway becomes active in response to elevated concentrations of the toxic intermediate DOPAL. MeHg, methylmercury; Ca2+, calcium; PKC, Ca2+/phospholipid dependent protein kinase; CaM-MPK, Ca2+/calmodulindependent multiprotein kinase; TH, tyrosine hydroxylase; DA, dopamine; MAO, monoamine oxidase; DOPAL, dihydroxyphenylacetaldehyde; ALDH, aldehyde dehydrogenase; DOPAC, dihydroxyphenylacetic acid; AR, aldehyde reductase; DOPET, dihydroxyphenylethanol; SER, smooth endoplasmic reticulum. 190 7.6. Factors influencing sensitivity of DA-synthesizing cells to MeHg One of the goals of this dissertation was to identify mechanisms specific to DA neurons that would predispose them to MeHg toxicity. While MeHg-induced DA release, perturbation of intracellular Ca2+ homeostasis, and mitochondrial dysfunction certainly have deleterious consequences, these mechanisms are not specific to DA neurons. MeHg induces the release of many neurotransmitters, including NE, ACh, γ-aminobutyric acid, and glutamate (Roos et al., 2012). Similarly, Ca2+ dysregulation and mitochondrial dysfunction are ubiquitous mechanisms of action (Weiss et al., 2002; Weiss, 2011). Accumulation of cytosolic DA and DOPAL would selectively and discretely heighten the sensitivity of DA neurons to MeHg toxicity. Non-vesicular DA has been described as an endogenous neurotoxin (Graham, 1978; Donaldson et al., 1982; Halliwell and Gutteridge, 1984; Stokes et al., 1999). It undergoes oxidation to form cytotoxic radicals and quinones either spontaneously or in the presence of transition metals, such as Mn2+ and Fe2+ (Akagawa et al., 2006). These oxidation events produce ROS and lipid peroxidation products (Kristal et al., 2001; Burke et al., 2003) that can severely damage cellular macromolecules. DOPAL is 100-1000-fold more toxic than DA (500ng versus 20μg; Burke et al., 2003; 2003). While injections of exogenous DA or DOPAL each cause focal, non-specific lesions in the substantia nigra, DOPAL toxicity occurs at much lower concentrations (Mattammal et al., 1995). In striatal synaptosomes and differentiated PC12 cells, DOPAL neurotoxicity is attributed to uptake and accumulation in terminals by high-affinity DA transport (Anderson et al., 2011a), and in mescencephalic cultures, DOPAL causes specific, selective 191 damage to TH-immunoreactive neurons. Like DA, DOPAL oxidizes to produce quinones, which increases its likelihood to cross-link proteins (Li et al., 2001) and generate free radicals (Marchitti et al., 2007). Accumulation of DA and particularly DOPAL following MeHg may be a critical endogenous event that triggers cell death in DA-synthesizing cells. Concentrations of DOPAL as low as 6μM can exert significant toxicity in vitro (Donaldson et al., 1982; Halliwell and Gutteridge, 1984; Li et al., 2001), well within the range observed presently. Future investigation concerning the contribution of DA and DOPAL oxidation products to MeHg-induced toxicity is of interest. Furthermore, because transition metals can increase the likelihood of both DA and DOPAL oxidation (Grace and Bunney, 1983), a better understanding of their interaction with MeHg would also be beneficial. The L-type VGCC is another potential mechanism by which MeHg may exert selective susceptibility in DA neurons in vivo. While results in this dissertation suggest that MeHg does not interact with the L-type VGCC to induce DA release in PC12 cells, studies were only conducted while the channel was at rest; changes in MeHg-induced DA release were not assessed following VGCC activation. The L-type VGCC is essential for NSDA neuronal function and targeting by MeHg may be a critical event in discrete DA neurotoxicity. Adult NSDA neurons maintain an autonomous rhythm, generating action potentials with a frequency of 2-4 Hz in the absence of synaptic input (Nedergaard et al., 1993; Bonci et al., 1998). Unlike most neurons that rely on monovalent cation channels to drive pacemaker activity, NSDA neurons engage somatodendritic L-type VGCCs (Chan et al., 2007) that contain the CaV1.3 pore-forming subunit (Koschak et al., 2001; Chan et al., 192 2007). L-type VGCC with this subunit open at relatively hyperpolarized potentials (50mV), and the influx of Ca2+ depolarized the cell to spike threshold (Sinnegger-Brauns et al., 2009). The CaV1.3 VGCC is rare, accounting for only ~10% of all L-type VGCCs in the brain (Surmeier, 2007), and is normally positioned near the synapse where it is only activated episodically. Sustained engagement of the CaV1.3 VGCC during pacemaking in NSDA neurons places a metabolic burden on these cells which is hypothesized to render them susceptible to degeneration in PD (Sirois and Atchison, 1996). MeHg interacts with the L-type VGCC (Shafer and Atchison, 1991b; Shafer, 1998). Immediately following application, MeHg blocks whole cell currents carried by Ba2+or Ca2+ through L-type VGCCs in PC12 cells (Denny et al., 1993; Marty and Atchison, 1997). However, L-type VGCC may also allow influx of extracellular Ca2+ during the second Ca2+ phase induced by MeHg (Marty and Atchison, 1997), as treatment with a L-type VGCC antagonist delays the time-to-onset of second phase influx (Tanner et al., 1999). Given these effects, a thorough analysis of the interaction between MeHg and CaV1.3 VGCCs in NSDA neurons may provide evidence of another mechanism by which MeHg selectively targets DA neurons. 7.7. Biological and clinical relevance to PD Twin studies have demonstrated that environment contributes to the idiopathic form of PD. Concordance for monozygotic twins, although higher before age 51, is the same as that for dizygotic twins after the age of 51 (Monnet-Tschudi et al., 2006). The authors 193 discuss the possibility that heredity is not a major contributor in most cases of PD, and call for investigation of non-genetic risk factors. The data presented in this dissertation provide evidence that the environmental toxicant MeHg targets DA homeostasis, which alters neurochemistry and may contribute to DA neuronal degeneration. However work is still needed to establish the relationship between MeHg-induced changes in DA homeostasis and NSDA toxicity in PD. Environmental exposure to MeHg could contribute to PD either through slow accumulation during long-term exposure to low concentrations, or exposure during a critical period during development (Choi et al., 1978). MeHg is a more potent neurotoxicant to the developing nervous system as compared to the adult nervous system (Amin-Zaki et al., 1978; Grandjean et al., 1997). Indeed, early life exposure to MeHg severely impairs neurobehavioral performance in several domains, including language, attention, memory, motor, and visuospatial function (McKeown-Eyssen and Ruedy, 1983; Davis et al., 1994; Weil et al., 2005). Adult exposure does not cause such pronounced deficits (Ngim and Devathasan, 1989; Wermuth et al., 2000; 2008; Petersen et al., 2008a; 2008b). Currently, a higher incidence of PD is only correlated with lifetime exposure to MeHg (Langston et al., 1999). However a mechanistic hypothesis suggests that early exposure to MeHg may reduce the number of neurons in vulnerable areas of the brain (e.g. the substantia nigra) below those needed to sustain function in the face of neuronal attrition associated with advanced aging (Klein and Westenberger, 2012). Few studies have investigated the effects of chronic, low-concentration MeHg exposure and its relevance to PD. A better understanding of how chronic, low-concentration exposure to MeHg induces oxidative stress and mitochondrial dysfunction, as well as dysregulation of DA homeostasis would 194 provide evidence to support current hypotheses about the impact of early life exposure to toxicants in neurodegenerative disease, and elucidate roles for exposure during a vulnerable period or exposure throughout one’s lifetime. Although genetic risk factors may not play the decisive role in the etiology of PD, their contribution is of significant importance. Moreover, understanding the interaction between genetic and environmental risk factors is essential in gaining a more complete understanding of how PD develops. To date, 28 distinct chromosomal regions have been associated with PD (Polymeropoulos et al., 1997), and MeHg exposure may be linked to at least three, parkin, DJ-1, and α-synuclein. Mutations in the gene α-synuclein have been linked to autosomal dominant PD (Spillantini et al., 1998), however the protein also plays a role in idiopathic PD being a major component of Lewy bodies, abnormal aggregates of protein found in the substantia nigra of PD patients at autopsy (Piao et al., 2000). α-Synuclein is widely distributed throughout the brain (Xu et al., 2002), and as such mutations alone cannot account for the selective disturbances in DA neurons. Indeed, transfection of α-synuclein is toxic in DAsynthesizing neurons but not non-DA containing neurons (Betarbet et al., 2000; Giasson and Lee, 2000). Therefore α-synuclein-mediated toxicity depends on the synthesis of DA. αSynuclein accumulation in Lewy bodies has been attributed to oxidative stress (Burke et al., 2003). It has been proposed that DOPAL and the generation of hydroxyl radicals associated with its oxidation trigger aggregation of α-synuclein in DA neurons (Li et al., 2001). Data presented in this dissertation provides evidences for a mechanism by which MeHg may interact selectively with DA neurons to induce α-synuclein accumulation and associated toxicity. In a hypothetical scenario, MeHg exposure would interfere with ALDH- 195 mediated DA metabolism, leading to accumulation of DOPAL. Being a highly unstable intermediate, DOPAL would readily oxidize, and in the process produce a hydroxyl radical (Hashimoto et al., 1999; Xu et al., 2002). The associated oxidative stress would accelerate aggregration of α-synuclein in its toxic form selectively in DA neurons (Bonifati et al., 2003). A similar situation could be envisioned for the PD-associated protein DJ-1 (Mitsumoto and Nakagawa, 2001; Mitsumoto et al., 2001). Although the exact function of DJ-1 is unknown, it is thought to monitor or buffer effects of excessive oxidative stress (Di Monte, 2003). Mutations in DJ-1 would compromise antioxidant function and increase susceptibility of DA neurons to oxidative stress (Shimura et al., 2000; Zhang et al., 2000). While there is currently no evidence linking MeHg exposure to mutations in DJ-1, a potential connection warrants further investigation. The production of free radicals associated with MeHg intoxication and aberrant DA metabolism could potentially have deleterious consequences in combination with a mutation in DJ-1. Lastly, there is evidence to suggest that MeHg interacts with the protein parkin in DA neurons. Parkin is a component of the multiprotein E3 ubiquitin ligase complex (Sherman and Goldberg, 2001), which in turn is part of the ubiquitin-proteosome system that targets misfolded proteins for degradation (Kitada et al., 1998; Mizuno et al., 2001). Loss of function mutations in the gene that encodes parkin cause autosomal recessive juvenile PD (Martinez-Finley et al., 2013). Mutations would abolish its E3 ligase activity and result in the accumulation of misfolded parkin substrates. A recent study has reported that early-life exposure to MeHg exacerbates the loss of DA neurons associated with parkin knockout in C. elegans. Evidence suggests that mercury accumulates more readily in parkin 196 knockout worms, and cell death might be due to an inability to mount an antioxidant defense against MeHg toxicity as compared to wildtype counterparts. Future studies investigating the relationship between MeHg-impaired DA metabolism and oxidative stress in the face of parkin mutations would contribute greatly to the understanding of gene/ environment interactions. 7.8. Concluding Remarks MeHg is a bioaccumulative environmental neurotoxicant that preferentially targets discrete regions of the brain, including the NSDA system. Results presented in this dissertation describe the deleterious consequences of MeHg exposure in DA-synthesizing cells. 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