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1 EFFECTS OF PPAR GAMMA AGONIST ROSIGLITAZONE ON CARDIAC TISSUE AND CULTURED CARDIOMYOCYTES By DANA JOY DAVIS SCHREFFLER A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2013
2 2013 Dana Joy Davis Schreffler
3 To my family, and especial ly to my amazing husband, Jacob
4 ACKNOWLEDGMENTS Through this long journey, many people have helped me, taught me and supported me. I would like to thank all of you from the bottom of my heart. Special thanks go out to Dr. Wohlgemuth, who recognized my h ard work and ambition and made it p ossible for me to pursue a graduate degree. Dr. Wohlgemuth has been a mentor and a friend and has always exhibited endless patience and support. Thanks go to my committee members and undergraduate assistants who have helped me through my graduate endeavor s: Dr. John Driver and Dr. Sally Johnson Maria Guzman, Darah Kelley, Claire Kent and Yenny Ramirez Lee. Each of my undergraduate assistants showed immense dedication through their long hours and hard work. T hanks also go out to my family including my parents and amazing husband, Jacob, who has always provided me with unconditional love and everlasting support. None of this would have been possible wi thout him My mother and father have always been people I can count on, and without their love and supp ort and help, I would not have successfully made it to the point I am in my life.
5 TABLE OF CONTENTS page ACKNOWLEDGMENTS .................................................................................................. 4 LIST OF FIGURES .......................................................................................................... 8 ABSTRACT ..................................................................................................................... 9 1 INTRODUCTION .................................................................................................... 11 2 MATERIALS AN D METHODS ................................................................................ 14 Experimental Subjects and Design ......................................................................... 14 In vivo Experiments ................................................................................................ 14 Animals ............................................................................................................. 14 Experimental Treatme nt ................................................................................... 14 In vitro Experiments ................................................................................................ 15 Cells ................................................................................................................. 15 Experimental Treatment ................................................................................... 16 Processing of Cardiac Tissue ................................................................................. 18 Heart Size ......................................................................................................... 18 Preparation of Cardiac Tissue for Different Analyses ....................................... 18 Processing of AC16 Cells ....................................................................................... 19 Homo genization of AC16 Cells for Spectrophotometric Enzyme Activities ...... 19 Homogenization of AC16 Cells for Immunoblot ................................................ 20 Mitochondrial Function Measurements ................................................................... 21 Cytochrome c Oxidase Activity ......................................................................... 21 Citrate Synthase Activity ................................................................................... 21 High Resolution Respirometry .......................................................................... 22 Western blot analysis .............................................................................................. 23 Statistical Analysis .................................................................................................. 24 3 LITERATURE REVIEW .......................................................................................... 26 Peroxisome Proliferator Activated Receptors (PPARs) and Their Physiological Effects .................................................................................................................. 26 Peroxisome Proliferator Activated Receptor Gamma ( ........................ 28 Dependent Effects of TZDs ................................................................. 29 Receptor Independent Effects of TZDs ............................................................ 30 Mitochondrial Function ............................................................................................ 31 Autophagy ............................................................................................................... 33 Adverse Effects of Rosiglitazone Therapy .............................................................. 35 4 RESULTS ............................................................................................................... 36 Effect of Rosiglit azone on Mitochondrial Function in vivo and in vitro ..................... 37
6 Effect of Rosiglitazone on Hearts from Fischer 344 Rats ................................. 37 Summary of Analyses of Heart Size and Mitochondrial Function in Fischer 344 Rats ........................................................................................................ 40 Effect of Rosiglitazone on Mitochondrial Function in AC16 Cardiomyocytes .......... 40 Effects of Rosiglitazone on Mitochondrial Enzymatic Activities in vitro ............. 41 Effects of Rosiglitazone on Mitochondrial Respiration in vitro .......................... 43 Summary of Analyses of Mitochondrial Function in AC16 Cells ............................. 47 Effect of Rosiglitazone on Autophagy in vivo and in vitro ....................................... 48 5 DISCUSSION ......................................................................................................... 72 Heart Size is No t Affected by Rosiglitazone Administration .................................... 72 Mitochondrial Enzymes are Not Affected by Rosiglitazone Administration in vivo .. 73 Mitochondrial Enzymes are Not Affected by Rosiglitazone Application in vitro ....... 74 High Doses of Rosiglitazone Decreases Mitochondrial Respiration in vitro ............ 75 Effect of Rosiglitazone on Autophagy in Rat Heart and Cultured Cardiomyocytes ................................................................................................... 78 6 CONCLUSIONS AND IMPLICATIONS ................................................................... 80 APPENDIX A CYTOCHROME c OXIDASE ASSAY FOR MICROPLATE READER ..................... 83 B CITRATE SYNTHASE ACTIVITY PROTOCOL ...................................................... 92 LIST OF REFERENCES ............................................................................................... 97 BIOGRAPHICAL SKETCH .......................................................................................... 102
7 LIST OF TABLES Table page 4 1 Heart and body weight and heart weight normalized to body weight of Fischer 344 rats .................................................................................................. 52 4 2 Oxygen flux of AC16 cells under control conditions (CTRL) or after exposure to rosiglitazone (10 or 200 uM) for 4 or 24 ho urs. ............................................... 58 4 3 Statistical analysis of differences between routine respiration rates of AC16 cells under control conditions or after exposure to rosiglitazone (10 or 200 .......................................................................................... 60 4 4 Statistical analysis of differences between LEAK respiration rat es of AC16 cells under control conditions or after exposure to rosiglitazone (10 or 200 .......................................................................................... 62 4 5 Statistical analysis of differences between electron transport capacity (ETS) of AC16 cells under control conditions or after exposure to rosiglitazone (10 .............................................................................. 64 4 6 Statistical analysis of differences between LEAK Control Ratio (L/E) of AC16 cells under control conditions or after exposure to rosiglitazone (10 or 200 .......................................................................................... 66
8 LIST OF FIGURES Figure page 4 1 Heart weight n ormalized to body weight of Fischer 344 rats. ............................. 51 4 2 Enzymatic activity of citrate synthase in tissue extracts of hearts from Fischer 344 rats .............................................................................................................. 52 4 3 Enzymatic activity of cytochrome c oxidase in tissue extracts of hearts from Fischer 344 rats .................................................................................................. 53 4 4 C ytochrome c oxidase activity normalized to citrate synthase activity in tissue extracts of hearts from Fischer 344 rats. ............................................................ 54 4 5 Enzymatic activity of citrate synthase in AC16 cell s. .......................................... 55 4 6 C ytochrome c oxidase activity in AC16 cells. ..................................................... 56 4 7 Cytochrome c oxidase activity normalized to citrate synthase activity in AC16 cells. ................................................................................................................... 57 4 8 Routine respiration of AC16 cells. ...................................................................... 59 4 9 LEAK respiration of AC16 cells ........................................................................... 61 4 10 E TS respiration of AC16 cells ............................................................................. 63 4 11 Leak control ratio of AC16 cells .......................................................................... 65 4 12 Routine control ratio of AC16 cells. .................................................................... 67 4 13 Protein expression of LC3 I and II in rat heart tissue extracts ............................ 69 4 14 Protein expression of LC3 I and II in AC16 cell lysates. ..................................... 71
9 Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science EFFECTS OF THE PPAR GAMMA AGONIST ROSIGLITAZONE ON CARDIAC TISSUE AND CULTURED CELLS By Dana J oy D avis Schreffler August 2013 Chair: Stephanie E. Wohlgemuth Major: Animal Sciences The peroxisome proliferator ) is a ligand dependent transcription factor, responding to endogenous (i.e. fatty acids) and synthetic ligands. Thiazolidinediones are a family of synthetic ligands, including the anti diabetic drug rosiglitazone. Through activation of PPARy, Thiazolidinediones increase storage of fatty acids, thereby increasing insulin sensitivity of hepatic tissue and skeletal muscle. This has led to treatment of Type II Diabetes mellitus with rosiglitazone (marketed as Avandia by GlaxoSmithKline). However, adverse car diovascular events in patients administered rosiglitazone have prompted the FDA to place the drug in a restricted access program. We hypothesized that in rat heart and cultured adult human ventricular cardiomyocytes (AC16 cells) administration of rosiglitazone 1.) decreases mitochondrial function, and 2.) increases autophagy. To test our hypotheses, Fischer 344 rats (5, 12 and 18 months old) received 10mg/kg/day rosiglitazone for 19 days; and AC16 cells were exposed to varying doses of rosiglitazone for 4 a nd 24 hours. We then determined mitochondrial enzyme activities
10 and mitochondrial respiration as measures of mitochondrial function, and expression of the autophagy protein LC3 to assess autophagy. We found that rosiglitazone had no effect on mitochondrial enzyme activities, specifically cytochrome c oxidase and citrate synthase, in rat cardiac tissue and AC16 cells, and that the activity of these enzymes in rat heart was independent of age. Mitochondrial routine respiration and maximum electron transport capacity in AC16 cells were not affected by 10uM, but decreased by 200M rosiglitazone. Autophagy was not affected by rosiglitazone, both in vivo and in vitro.
11 CHAPTER 1 INTRODUCTION The peroxisome proliferator activated receptors (PPARs) are a superfamily of nuclear hormone receptors consisting of three isoforms: the heart. PPAR s are ligand dependent transcription factor s with two types of ligands: endogenous ligands such as fatty acids, and exogenous, synthetic ligands, such as drugs in the Thiazolidinediones family (Spiegelman 1998). The ligand activates PPAR by binding to it The activated PPAR associates with t he Retinoid X Receptor (RXR) forming the PPAR response element (Berger et al. 2005), which drives the transcription of specific genes that will induce mechanisms such as differentiation of adipocyte, lipid storage, fatty acid oxidation, and insulin sensit ization (Spiegelman 1998). The Thiazolidinedione (TZD) family is a group of synthetic PPAR ligands (with greatest affinity for ) which includes the compounds pioglitazone, troglitazone, and rosiglitazone. The TZDs insulin sensitizing effects led to its marketing as an anti diabetic drug under the name Avandia by the pharmaceutical company GlaxoSmithKline. Insulin resistance, a major contributor to Ty pe 2 Diabetes mellitus (T2DM), has been linked to decreased mitochondrial oxidative activity and ATP synthesis, as well as high levels of circulating triglycerides in muscle and liver (Lowell and Shulman 2005). TZDs such as rosiglitazone increase lipid storage and fatty acid utilization, thus combating the insulin resistant condition of the Type 2 Diabet es mellitus patient. Despite its positive effects on insulin resistance, administration of Rosiglitazone
12 has been associated with a number of adverse effects, such as increased risk of myocardial infarction and cardiovascular mortality ( Nissen & Wolski 20 1 0 ), and cardiac hypertrophy ( Festuccia et al. 2009). As a result, the FDA has restricted its use, and prescription of Rosiglitazone has become controversial in the United States (Pouwels & van Grootheest 2012). It has therefore been of great interest to identify the underlying causes of rosiglitazones negative effects on the heart. Recently, Rabol and colleagues (2010) evaluated the effects of Rosiglitazone on mitochondrial function in the skeletal muscle of patients with T2DM. The authors found that m itochondrial respiration was decreased in patients with T2DM compared to control subjects an effect that was exacerbated by administration of rosiglitazone. It is, however, not known whether rosiglitazone has a negative effect on mitochondrial respiration in the heart as well. In this study we hypothesized that administration of rosiglitazone decreases mitochondrial function in the heart. To test out hypothesis we investigated the effect of rosiglitazone on mitochondrial function in heart tissue from rats that had been administered rosiglitazone, and in a cultured cardiomyocyte cell line. In addition to its effects on mitochondrial metabolism, few in vitro studies have investigated the effect of activation on the cellular quality control mechanism aut ophagy. Autophagy is a cellular housekeeping mechanism by which cells typically sequester portions of cytoplasm which can include organelles protein aggregates and cellular waste, into autophagosomes and deliver these to lysosomes for degradation Besides its role as housekeeping mechanism, this process is activated in conditions of energy imbalance and serves to supply substrates for energy production.
13 The reports on the effect of activation on autophagy have been conflicting. While activat ion has been shown to induce autophagy in breast cancer cells (Zhou et al. 2009) and adrenocortical cancer cells (Cerquetti et al. 2011), Mahmood et al. human monocytederived macrophages, but there have been no reports on the effect of Both too much and too little autophagy can be detrimental to an organ system, making this a necessary niche to explore (Troncoso et al. 2013). In this study, we hypothesize that administration of rosiglitazone increases autophagy in the heart, leading to a reduction in heart remodeling and contributing to the likelihood of cardiovascular mortality. To test this hypothesis, we examined the effect of rosiglitazone on autophagy and on autophagic flux in heart tissue from rosiglitazone treated rats and in cultured cardiomyocytes. The objectives of the current study were (1) to assess the effect of rosiglitazone on mitochondrial function and content in the heart, (2) to determine if this effect is age dependent, and (3) to determine if rosiglitazone has an effect on autophagy in the heart.
14 CHAPTER 2 MATERIALS AND METHODS Experimental Subjects and Design In vivo E xperiments Animals Fischer 344 rats were purchased from the National Institute of Aging (NIA) colony at Harlan Laboratories (Indianapolis, IN). The experimental groups consisted of rats of three ages: 5 months (n=19), reflecting a young age; 12 months (n=21), reflecting adul thood; and 18 months (n=21), reflecting onset of senescence. Rats were received and housed for 1 week prior onset of the experimental treatment. Rats were individually housed and maintained on a 12h light/dark cycle under controlled conditions. Health sta tus, body weight and food intake were monitored daily. The rats had free access to regular rat chow, and to tap water. Principles of laboratory animal care (NIH publication No. 8623, revised 1985) were followed and all procedures were approved by the Univ ersity of Floridas animal care and use committee. Experimental T reatment Seven days after arrival, the experimental treatments and assessments were started. All rats were given plain frozen strawberry milk treats starting two days prior to water maze trai ning, and vehicle treats (2.5% ethanol/straw b erry milk, v/v) throughout training and testing to familiarize the rats with the treat and administration paradigm. Based on water maze performance (used for a concurrent study), rats of each age were randomly a ssigned to the experimental groups: the treatment group, which received rosiglitazone (5 mo nths, n = 11 ; 12 mo nth s, n = 10 ; and 18 mo nth s, n = 10) or the control group, which received vehicle (5 mo nth s, n =9 ; 12 mo nth s, n = 11; and 18
15 mo nth s, n = 11). After completing the initial water maze training and testing, the experimental drug exposure was started in which rats were either given 500 L frozen strawberry milk treats containing rosiglitazone (5 mg/kg; Cayman Chemical Company, Ann Arbor, MI, USA) o r containing vehicle ( 2.5% ethanol v/v) twice daily for 19 days. Rats were all euthanized in the morning. Before euthanization, rats were weighed, subsequently sedated using isofluorane, and then euthanized with a guillotine. Following decapitation, the c hest was opened and the heart excised. The heart was immediately flushed with saline, blotted dry, weighed, and then flashfrozen in liquid nitrogen. The tissues were stored in liquid nitrogen until time of use All rats in this study were treated in accor dance with the policies set forth by the University of Florida Institutional Animal Care and Use Committee (IACUC) and the National Institutes of Health (NIH) regarding the ethical use of animals for experimentation. In vitro E xperiments Cells The in vivo study was complemented with in vitro experiments using an immortalized cardiac cell line reflecting as closely as possible the heart tissue examined in the in vivo experiments Using cultured cells had the advantage that the effects of different concentrations of rosiglitazone and durations of rosiglitazone exposure could be evaluated. AC 16 human derived ventricular cardiomyocytes (Garbern et al. 2013) were generously donated by Dr. Leeuwenburgh, U F and used to evaluate the effect of rosiglitazone on 1) mitochondrial function using High Resolution Respirometry (HRR), 2) mitochondrial enzyme activity via spectrophotometric enzyme activity assays and 3) expression of autophagy regulatory proteins us ing Western
16 Blotting. The cells were maintained in Dulbeccos Modified Earles Medium F 12 50/50 containing 10% FBS, 29mM sodium bicarbonate, 2.5mM glutamine, 17.5mM glucose, 1.25mM sodium pyruvate, and 50IU peni cillin mixed with 50uM streptomycin at pH 7. 2 in an incubator at 37 C with 5%CO2 Experimental T reatment Rosiglitazone treatment: For a rosiglitazone exposure experiment, cells were seeded into 6well plates (Corning) in normal growth medium and grown to 90% confluency, which amounted to approximately 0.5 x 106 cells. Prior to the experimental treatment, each well was washed once with PBS. Cells were then exposed to growth in DMSO (purchased from Cayman Chemicals) for 4 or 24 hours. In order to limit the cells exposure to DMSO, a 100 mM rosiglitazone stock solution was prepared, aliquoted and stored at 20 C, and heated to 72 C prior to use. At the beginning of the experiment, an appropriate amount of rosiglitazone s tock solution was added to the exposure the cells were either lysed and the cell lysates frozen at 80 C until performance of biochemical analyses (enzyme activity assays; Western Blot; see below), or trypsinized, washed in normal culture media and injected into the respirometer chamber for determination of cellular respiration by High Resolution Respirometry (see below). Experimental replicates were performed on different d ays using a different cell passage. Each experiment included control cells, which were maintained in normal growth medium and exposed for the same amount of time than the simultaneously rosiglitazonetreated cells.
17 Measurement of autophagy and autophagic f lux Control cells were maintained in normal growth medium without rosiglitazone, with vehicle (0.1% DMSO). The autophagic flux was quantified by adding Bafilomycin A1 (50nM final concentration) to a duplicate well of each treatment. Bafilomycin A1, a proton pump inhibitor, which prevents the fusion of the autophagosome with the lysosome by neutralizing the lysosomal pH, causes the accumulation of autophagosomes (Rubinsztein et al. 2009). Because the final degradation of the autophagosomes and their contents is prevented, the autophagosomal membranebound form of the autophagy marker protein LC3II accumulates. LC3II accumulation can then be utilized to determine the autophagic flux by calculating the difference of LC3 II protein level between the cells treated without and the cells treated with Bafilomycin A1 within the same treatment (Rubinsztein et al. 2009). Two independent experiments were performed to determine the effect of rosiglitazone on autophagic flux in AC16 cells. Cells for measurement of cellular respiration Cells used for determination of cellular respiration in the absence and presence of rosiglitazone were maintained in T 25 flasks. Per experiment two flasks were prepared and cells grown to 90% confluency. For the experiment, t he cells in one flask were maintained in normal growth medium (control), and the cells in the other flask final concentration; see above). At the beginning of the experim ental exposure, the normal growth medium was exchanged for either control medium or treatment medium. At the end of the experimental exposure, the medium was removed, cells were washed once with PBS, subsequently trypsinized and a cell count performed. Cells were added
18 to each of the two respirometer chambers in a final concentration of 0.3 0.5 x 106 mL1 cells. Each respirometer assessment was performed on control and rosiglitazonetreated cells side by side. Cellular respiration was measured in five treatments groups: (n=4). (for more details see High Resolution Respirometry below) Processing of Cardiac Tissue Heart S ize Heart weight was determined and then normalized to body weight. Normalized heart weight was analyzed across treatments and ages. Preparation of C ardiac Tissue for Different A nalyses Homogenization of cardiac tissu e for spectrophotometric enzyme activities The rat hearts, collected and stored in liquid nitrogen as described above, were homogenized using a liquid nitrogencooled Biopulverizer (BioSpec products, Bartlesville, OK). The powdered tissue (1520mg) was tra nsferred to a liquid nitrogen2PO4 buffer with 2mM EDTA; pH 7.2; Adhihetty et al. ) added. The tissue suspension was mixed by tapping and then further diluted 20fold (w/v) with extraction buffer. Following incubation and shaking on a thermomixer (Eppendorf, Hamburg, Germany) at room temperature for 15 minutes at 1400 rpm, the samples were sonicated on ice using a sonic dismembrator (Fisher) at a setting of 7 (on a scale of 1 to 20) for 3 x 3 seconds. The samples were then again diluted with extraction buffer to a final 80fold dilution. The final tissue extracts were centrifuged at 14,000 x g for 2 minutes at room temperature.
19 The supernatant was transferred to a new tube and the protein concentration determined using the Bradford assay (Fisher). Homogenization of cardiac tissue for immunoblot For immunobloting, the cryopulverized heart tissue (see above) was suspended ma), 1mM EDTA (Fisher), 2% SDS (Sigma), 10mM TRIS base (Fisher Scientific), and 1% Protease Inhibitor Cocktail (ThermoScientific); pH 8. The samples were vortexed for 15 seconds, and heated at 95 C 3 x 3 minutes with 15 seconds of vortexing in between. Th e samples were then centrifuged for 10 minutes at 12,000 x g at room temperature, transferred to a new tube and the protein concentration determined using the Bradford assay (Fisher). Processing of AC 16 C ells Homogenization of AC1 6 Cells for Spectrophotome tric Enzyme A ctivities AC16 cells were seeded to confluency in a 6well plate and treated for 4 hours with 0 (vehicle only), 10uM, 100uM or 200uM rosiglitazone as described above. After the four hour exposure to rosiglitazone or vehicle, the experimental m edia was removed, and the cells quickly washed with icecold PBS (Phosphate Buffered Saline, ( 0.25 M sucrose, 40mM potassium chloride, 2 mM EGTA, 1 mg/ml bovine serum albumin, and 20 mM Tris HCl; pH 7.2; Campian et al. 2007) was added to each well followed by rotating the plate at 4 C for 5 minutes. The cells were scraped off and transferred into a microcentrifuge tube. The cell suspensions were then rotated at 4 C for 30 minutes. A fterwards, the cells were lysed by sonicating on ice using a sonic dismembrator (Fisher Scientific) at setting 3 (on a scale of 1 to 20) for 10 one second
20 bursts. The cell lysates were then centrifuged at 2000 x g for 10 minutes at 4 C, and the supernatant transferred to a new tube. Prior to enzymatic activity assays, the protein concentration in the lysates was determined using the Bradford assay (Fisher Scientific) according to the manufacturers instructions. Cell lysates were stored at 80C until enzy me assays (see below). Ho mogenization of AC16 Cells for I mmunoblot AC16 cells were seeded to confluency in a 6well plate and treated for four hours with 0 (vehicle only), 10uM, 100uM or 200uM rosiglitazone, with or without bafilomycin A1 (25 nM final conc entration). At the end of the exposure, the experimental media was removed, and each well rinsed with icecold PBS buffer, as described above. Then, 100uL of icecold RIPA lysis buffer (ThermoScientific, Rockford, IL), containing 1mM PMSF (Sigma, St. Louis, MO), and 1% Protease inhibitor cocktail (ThermoScientific, Rockford, IL) was added to each well. The plates were then rotated at 4 C for 5 minutes. The cell lysates were scraped off and transferred into individual microcentrifuge tubes and rotated at 4 C for 30 minutes. Cell lysates were the sonicated on ice with a sonic dismembrator for 10 one second bursts at a setting of 7 (on a scale of 1 to 20). After sonication, the samples were centrifuged at 14000 x g at 4C for 10 minutes, and the supernat ant transferred to new tubes. Protein concentration was determined using the BCA assay (ThermoScientific) according to the manufacturers instructions. Samples were aliquoted and stored at 80 C until further analysis.
21 Mitochondrial Function Measurements Cytochrome c Oxidase A ctivity Cytochrome c Oxidase (COX) activity in AC16 cell lysates was determined with a spectrophotometric enzyme activity assay modified after Smith (1955). Briefly, a solution of reduced horse heart cytochrome c (cyt c; 2 mg/mL; Sigm a, St. Lou s, MO), serving as substrate for COX, was prepared in 100 mM KPO4 buffer (pH 7.0) using 57 mM sodium dithionite (in 10 mM KPO4) as a reducing agent. Enzyme activity was determined from the rate of oxidation of reduced cyt c at 30 C, measured as reduction in absorbance at 550 nm using a microplate reader (Synergy HT, Biotek Instruments, protein. See Appendix I for detailed protocol. Citrate S yntha se A ctivity The activi ty of Citrate Synthase (CS) in AC16 cell lysates was determined with a spectrophotometric enzyme activity assay as described in Kuznetsov et al. (2010). In this assay, the production of Coenzyme A by CS from Oxaloacetate and Acetyl CoA is coupled to the i rreversible reaction of Coenzyme A with DTNB (5,5dithiobis 2 nitrobezoate), producing TNB (thionitrobenzoic acid). The absorbance of TNB is measured at a wavelength of 412 nm using a microplate reader. The reaction mix contained a final concentration of 0.25% Triton X 100, 0.31 mM Acetyl CoA (30 mM stock in water), 0.1 mM DTNB (1.01 mM stock in 1.0 M Tris HCl buffer, pH 8.1), and 8 addition of 0.5 mM Oxaloactetate (10 mM stock in 0.1 M triethanolamine, pH 8.0). See Appendix II for detailed protocol.
22 High Resolution R espirometry Oxygen consumption of intact AC 16 cells was measured as an indicator of mitochondrial function using a high resolution respirometer (Oroboros O2K Oxygraph, Innsbruck, Austria). In brief, a polarographic oxygen sensor detects the oxygen concentration inside a closed respirometer chamber, which contains the cells. The data output is reported as oxygen concentration over the course of time and as its first derivative, which is the oxygen consumption rate, or oxygen flux. After an experimental treatment, cells were trypsinized and counted, and 300,000500,000 cells/mL added to each of the two 2mL chambers of the oxygraph. Oxygen consumption was measured in the two chambers simultaneously, such that oxygen flux of control and Rosiglitazonetreated cells were assessed side by side. The inhibitors oligomycin (2ug/ml), rotenone (final concentration 0.5 uncoupler FCCP ( carbonyl cyanide 4(trifluoromethoxy) phenylhydrazone; respirometer chamber in a specifi c sequence: 1. Oxygen flux of intact cells in culture media (Routine respiration; physiologically coupled respiration in the intact cell); 2. Addition of the ATP synthase inhibitor oligomycin (LEAK state; respiration compensating for the proton leak proton slip, and cation cycling); 3. Addition of FCCP (uncoupled respiration, or excess capacity; respiration in the presence of a protonophore (i.e. FCCP) which cycles across the inner mitochondrial membrane with transport of protons and dissipation of the electrochemical proton gradient); 4. Addition of the Complex I inhibitor rotenone (respiration with Complex II substrates only); and 5. Addition of the Complex III inhibitor antimyc in A (residual, nonmitochondrial, oxygen consumption). The residual oxygen consumption is subtracted as extramitochondrial
23 from the mitochondrial oxygen consumption. Oxygen consumption rate, or oxygen flux, is reported in pmol O2 s1 (106 cells. Based o n these recorded oxygen flux measures the following respiratory control ratios were calculated: Routine Control Ratio (Routine/ETS), a ratio expressing how close routine respiration operates to ETS capacity, and LEAK Control Ratio (LEAK/ETS), which is the ratio between respiration in the presence of the ATP synthase inhibitor oligomycin, and maximal electron transport system capacity (ETS). Western blot a nalysis Expression of LC3 B, an integral autophagy regulating protein, and GAPDH, glyceraldehyde 3phosphate dehydrogenase a housekeeping protein, in cardiac tissue and AC16 cells was determined using western blot analysis of tissue extracts and cell lysates, respectively. Cardiac tissue homogenates and AC16 cells were homogenized and lysed respectively, as described above, and the tissue extracts and cell lysates prepared for SDS PAGE electrophoresis followed by Western Blot. Prior to loading on a denaturing SDS polyacrylamide gel, extracts and lysates were mixed 1:2 with Laemmli buffer (62.5 mM Tris HCl, 2 % SDS, 25% Glycerol, 0.01% Bromophenol Blue, pH 6.8; mercaptoethanol, and boiled at lysate) were applied to precast Tris HCl gels (Criterion system, BioRad) formulated as any kDa acrylamide (proprietary formulation). Gels were run at 200V for approximately 50 minutes using Tris Glycine SDS running buffer (BioRad). After electrophoretic separation, the proteins wer e transferred to polyvi polyvinylidene difluoride (PVDF) membrane (Immobilon P, 0.45 lm, Millipore, Billerica, MA) using a semidry blotter (BioRad) at a constant 7V for 2.5 hours. The transfer buffer (Tris Glycine; BioRad)
24 contained 15% methanol. After the transfer, the membranes were rinsed in dH2O and methanol, and let dry for at least 15 min to optimize protein binding to the membrane. Transfer efficiency and quality was subsequently verified by staining the membrane with Ponceau S (SigmaAldrich). Membranes were then blocked in Tris buffered saline (137mM NaCL, 2.7mM KCl, and 24.76mM Tris Base) with 0,05% Tween20 (Fisher) (TBS t) containing 5 % (w/v) nonfat dry milk (Blotting Grade Blocker, BioRad, Hercules, CA) for one hour at room temperature; subseq uently washed in TBS (without Tween), and incubated overnight in the corresponding primary antibody (see below) at 4 C. Subsequently, membranes were washed in TBS t and incubated with horseradish perioxidase (HRP) conjugated secondary antibody (SigmaAldr ich) at room temperature for one hour. Membranes were then washed in TBS t, followed by TBS. Finally, the DuoLux chemiluminescent/fluorescent substrate for HRP (Vector Laboratories, Burlingame, CA) was applied for 5 min, followed by a quick rinse with Tris HCl (100 mM, pH 9.5), and the chemiluminescent signal captured with a Syngene G BOX imager (Synoptics Group, Frederick, MD).The digital images were analyzed using SynGene software (Synoptics Group, Frederick, MD). Density of the target band was normalized to GAPDH ( Glyceraldehyde 3phosphate dehydrogenase), and expressed in arbitrary optical density units. Antibodies used were LC3B (1:1000, Cell Signaling, Danvers, MA), GAPDH (1:1000, Cell Signalling), and HRP linked secondary antibodies (1:50,000, Sigma). Add clone names Statistical Analysis For both heart tissue and cultured cells, the experiments were fully crossed, twofactor designs with two or three levels for each factor: a) heart: 1) age (5, 12 and 18
25 months) and rosiglitazone (vehicle without rosiglitazone control, and vehicle with rosiglitazone treatment); and b) cultured cells: exposure time (4 and 24 hours) and T wo W ay analysis of variance (ANOVA) will allow us to distinguish between the effects of age ( in vivo study) or exposure time ( in vitro study) respectively, and the presence of the drug rosiglitazone, and a possible interaction effect between age/exposure time and drug. Post hoc multiple comparisons for all ANOVA tests were performed with the Tukey HSD procedure. A Kruskal Wallis ANOVA on Ranks test was performed when data were not normally distributed. Enzyme activities in AC16 cells were compared between experimental groups in a One W ay ANOVA, with dose of rosiglitazone as the independent factor (only one exposure time was tested) Heart size normalized to body weight and enzyme activities in heart tissue extracts were analyzed using a TwoWay ANOVA with age and dose of rosiglitazone as independent factors. Protein expression data were analyzed using a One Way ANOVA (AC16 cells) or a Two Way ANOVA (heart), respectively. All statistical analyses were conducted using SigmaPlot v s. 12 (Systat Software, Inc.,
26 CHAPTER 3 LITERATURE REVI EW A constant supply of energy is essential for maintenance of heart activity. The oxidation of fatty acids, which support 6090% of cardiac energy supply, and subsequent oxidation of reduction equivalents in the mitochondrial oxidative phosphorylation (OXPHOS) system. The heart relies heavily on oxidative phosphorylation for energy production, which accounts for over 90% of the ATP produced, and there is a complex interplay of receptors and co factors that must be engaged to activate energy production pathways in order meet these demands (Kodde et al. 2006). The nuclear hormone receptor family of peroxisome proliferator activated receptors (PPARs) is an intr insic part of the regulatory system to ensure energy balance in the heart. Peroxisome Proliferator A ctivated Receptors (PPARs) and Their P hysiological E ffects The members of the PPAR family are transcription factors that are major regulators of energy hom eostasis and metabolic functions (Tyagi et al. 2011). The expressed in the liver, where it plays a major role in fatty acid oxidation and lipid ch is most widely found in skeletal muscle tissue, where it in white adipose tissue, the liver, and at moderate levels in the heart, regulates adipocyte differentiation, fatty acid storage and glucose metabolism (Edvardsson et al. 1999; Ehrenborg & Krook 2009). All three PPAR subtypes function as lipid sensors, which, when activated, will associate with a retinoid X receptor (RXR). The heterodimer will translocate into the nucleus and bind to a specific DNA sequence, the peroxisome
27 proliferator response element (PPRE). The association of the heterodimer with the PPRE activates transcription of specific genes involved in the metabolic functions outlined above. The PPARs are activated by small lipophilic ligands such as fatty acids, which occur endogenously, and synthetic drugs such as compounds in the insulin sensitizing Thiazolidinedione (TZD) family (Tyagi et al. 2011). Their role in regulation of metabolic functions and t he well defined therapeutic actions of their synthetic ligands make PPARs ideal targets for metabolic disease intervention. agonists are employed as anti atherosclerotic and hypolipidemia agents, to combat dyslipidemia associated with metabolic syndrome (Berger et al. 2005 ; Tyagi et al. 2011). Obesity and inflammation have also been treated with PPAR agonists. Activation of enuate obesity in rodents by increasing hepatic fatty acid oxidation and decreasing the levels of triglycerides demonstrated to inhibit the expression of inflammatory cytokines (Tyagi et al. 2011). resistance. Their role in regulation of metabolic functions and the well defined therapeutic actions of their synthetic ligands make PPARs ideal targets for metabolic disease intervention. agonists are employed as anti atherosclerotic and hypolipidemia agents, to combat dyslipidemia associated with metabolic syndrome (Berger et al. 2005 ; Tyagi et al. 2011). Obesity and inflammation have also been treated with PPAR agonists. Activation of
28 oxidation and decreasing the levels of triglycerides responsi ble for adipose hypertrophy, inflammatory cytokines (Tyagi et al. 2011). as a therapeutic approach to insulin resistance. Peroxisome Proli ferator to 30fold higher than in other tissues, such as the liver and cardiac tissue (Spiegelman 1998). gulator for adipocyte differentiation as well as regulating fatty acid storage and glucose metabolism (Tyagi et al. 2011). In adipose tissue, by altering the expression of genes that are involved in lipid uptake, lipid metabolism and insulin action, thereby enhancing adipocyte insulin signaling, lipid uptake and anabolic lipid metabolism and attenuating free fatty acid release. In hepatic and su levels of circulating free fatty acids. cell population is sustained, thus allowing for more efficient gl ucose clearance (Berger et al. 2005). In addition to reducing been demonstrated to cause an accumulation of small adipocytes while promoting apoptosis in large, mature adipocytes. This shift in adipocyte population favors insulin sensitivity, as the small adipocytes are intrinsically more insulin sensitive than large adipocytes (Zieleniak et al. agonists ideal targets for the treatment of Type II diabetes mellitus (T2DM).
29 T2DM is a metabolic syndrome primarily associated with decreased insulin sensitivity of tissues, generally stemming from defects in the secretion of insulin by s (Lowell & Shulman, 2005) Lifestyle choices, such as poor diet and lack of exercise, combined with genetic predisposition for diabetes are major risk factors for the onset of T2DM (Riserus et al. 2009). Thiazolidinediones (TZDs) are synthetic ligands that TZDs, derivatives of the thiazolidinedione, include the compounds troglitazone, pioglitazone, and rosiglitazone. Pharmacological administration of TZDs were approved for treatment of T2DM by the FDA starting with troglitazone in 1996, manufactured under the names Rezulin or Romozin. Troglitazone has since been removed from the market in the US due to its hepatotoxic side effects (Koffarnus et al 2013; Yau et al. 2013). Pioglitazone is currently marketed under the name Actos by Takeda and approved for prescription to treat T2DM, and rosiglitazone is currently marketed under the name Avandia by GlaxoSmithKline. Rosiglitazone has been removed from market in Europe and New Zealand, and the FDA has restricted its use in the US due to the increased risk of adverse cardiovascular effects such as myocardial infarction and cardiac death (Koffarnus et al. 2013 ; Nissen & Wolski 2007). Dependent E ffe cts of TZDs Insulin sensitizing effect. Insulin resistance has been linked to decreased mitochondrial oxidative activity and ATP synthesis, as well as high levels of circulating triglycerides in muscle and liver (Lowell & Shulman, 2005). Oxidative phosphor ylation of fatty acids is normally responsible for about 6090% of the ATP synthesis in the human heart ( Stanley et al. 1997 ) In the diabetic patient, both glucose and lactate uptake is
30 impaired, leading to an even greater reliance on oxidative phosphory lation of free fatty acids (Stanley et al. 1997). At the same time, oxidation of fatty acids is impaired, leading to higher levels of circulating free fatty acids. It is thought that these high levels of circulating free fatty acids compete with two enzymes required for glucose utilization, pyruvate dehydrogenase and phosphofructokinase, causing inhibition of the glycolytic pathway, and thus increasing insulin resistance (Lowell & Shulman 2005). Due to their insulin sensitizing effects, TZDs are widely pr escribed for the treatment of T2DM. acid storage and utilization and glucose homeostasis (Koffarnus et al. 2013). In experiments studying the efficacy of TZDs, it was determined that these compounds improve glucose tolerance and insulin resistance by increasing insulin sensitivity of et al. 2013; Fryer et al. 2002). Receptor Independent E ffects of TZDs In addition to its receptor dependent actions, such as fatty acid storage and use, insulin sensitizing effects and anti inflammatory effects, TZDs have been shown to exert receptor independent actions (Feinstein et al. 2005). These effects, which are independent of direct effects on gene transcription, may be the underlying cause for some of the effects that are not metabolism related. For example, TZDs have been shown to activate the adenosine monophosph ate activated protein kinase ( AMPK ) pathway, inhibit inflammatory cytokines, suppress cell proliferation and disrupt mitochondrial function (Feinstein et al. 2005; Bolten et al. 2007). Bolten and colleagues (2007) and Brunmair and colleagues (2001) suggested that TZDs activate the AMPK pathway. AMPK is a master regulator of energy metabolism. It
31 is activated by an increase in the intracellular AMP/ATP ratio, and considered a sensor for cellular energy imbalance (Feinstein et al. 2005; Dyck & Lopaschuk 2006). In a 2002 study by Fryer and colleagues, it was demonstrated that TZDs, specifically rosiglitazone, directly activate the AMPK pathway in skeletal muscle, which lead to the inhibition of mitochondrial OXPHOS. In the same journal article, it was stated that long term activation of AMPK has been shown to increase the expression of a number of proteins that lead to an increase in insulin sensitivity. These overlapping effects, both metabolic effects and insulin sensitizing effects, of AMPK and TZDs makes it difficult to determine if TZDs are causing mitochondrial dysfunction and thus stimulating AMPK due to the subsequent energy imbalance or if it is a direct effect of TZDs on AMPK. Feinstein and colleagues (2005) speculate that activation of AMPK may be dependent upon TZD induced changes in metabolic state and mitochondrial impairment. Mitochondrial Function Mitochondria are intracellular doublemembrane enclosed organelles that generate most of the cells ATP. Under physiological conditions, a series of enzyme complexes and electron carriers (collectively termed electron transport chain or system) located at the inner mitochondrial membrane transfer electrons from the reduction equivalents NADH and FADH2, produced in glycolysis and the citric acid cycle, to the ultimate electron acceptor oxygen, thereby producing H2O. Enzyme complex I, II and IV of the electron transport system (ETS) utilize the energy released by this electron transfer to pump protons from the mitochondrial matrix into the intermem brane space (IMS), thereby creating an electrochemical gradient across the inner mitochondrial membrane. This electrochemical gradient is utilized to produce ATP from ADP and inorganic phosphate, and is catalyzed by the inner mitochondrial membrane enzyme
32 ATP synthase, which allows the exergonic transport of protons back into the matrix. The process of electron transport coupled with ADP phosphorylation is commonly referred to as oxidative phosphorylation (Spinazzi et al. 2012; Gnaiger 2012). There are sever al methods to assess mitochondrial function. The most common approach is to determine mitochondrial oxygen consumption or respiration. This approach takes advantage of the fact that oxygen is the terminal electron acceptor in the mitochondrial ETS, The rat e of oxygen consumption in a closed (air tight) chamber containing living cells (or biological material such as tissue or isolated mitochondria) can therefore be used as an indicator of mitochondrial electron transport function (Spinazzi et al. 2012). Depe nding on the biological material assessed (tissue, intact cells, permeabilized cells, isolated mitochondria), a series of different energy substrates, inhibitors of mitochondrial respiratory enzyme complexes, and mobile ion carrier (also called uncoupler o f oxidative phosphorylation) are sequentially titrated into the measuring chamber and subsequent oxygen consumption recorded There are four major states of mitochondrial respiration used to assess their overall function. Routine respiration measures respi ration of the cell sample in an unaltered state, prior to offering substrates or ADP. LEAK respiration is measured after inhibition of the ATP synthase and accounts for any mitochondrial respiration not linked to the phosphorylation system, such as proton leaks or slips, cation cycling, and ROS production (Gnaiger 2012). ETS is defined as excess, or maximum, capacity of the electron transport system, and is measured in the presence of an uncoupler, such as FCCP ( Carbonyl cyanide4 (trifluoromethoxy)phenylhy drazone) By uncoupling proton flow from electron transfer, true maximum capacity can be assessed demonstrating full
33 electron transfer efficiency. Finally, residual oxygen consumption (ROX) is assessed in the presence of antimycin A, a complex III inhibitor. When antimycin A is introduced, it effectually shuts down the electron transport system allowing the contribution to respiration by the cell to be accounted for. These respiratory states can then be compared in ratio form to assess other important param eters of mitochondrial function such as coupling efficiency (LEAK/ETS), and the routine control ratio (Routine/ETS). Assessing LEAK/ETS determines how coupled oxygen consumption is to the phosphorylation system. Routine/ETS demonstrates to what extent routine is contributing to the overall capacity of the electron transport system. Another method to evaluate mitochondrial function is to determine enzymatic activity of mitochondrial enzymes using a spectrophotometric enzyme assay (see materials and methods section). Most commonly, complex IV of the ETS, or cytochrome c oxidase, and citrate synthase, an enzyme of the citric acid cycle located in the mitochondrial matrix, are used as markers of mitochondrial function. Moreover, activity of citrate synthase has been shown to correlate well with mitochondrial content and has therefore been used as a marker for this parameter (Campian et al. 2007; ChrzanowskaLightowlers et al. 1993). Autophagy AMPK will employ autophagy when the AMP:ATP ratio is high. This will lead to a signaling cascade that will culminate in the degradation of aggregate proteins, thereby restoring the supply of amino acids for energy production (Vacek et al. 2011) Autophagy is cellular housekeeping mechanism by which cellular constituents, s uch as proteins, protein aggregates, and cell organelles, are sequestered within a double-
34 membrane bound organelle, the autophagosome, which subsequently fuses with a lysosome, which delivers the enzymes for the degradation of the sequestered cellular mate rial (Cuervo 2008; Rubinsztein et al. 2009). Autophagy is thereby a catabolic process that serves the degradation and recycling of cellular constituents. It is a highly regulated process, and a variety of stimuli can initiate this process, such as the disr uption of mitochondrial membrane potential, nutrient deficiency, and pharmacological agents such as rapamycin (Cuervo 2008 ; Cerquetti et al. 2011). One of the major regulatory pathway regulators at which some of the inducers and inhibitors of autophagy merge for autophagy is mTOR, the mammalian target of rapamycin (Vacek et al. 2011) AMPK is an example of a kinase that merges at the mTOR complex to induce autophagy. Both, too much autophagy and too little autophagy can be detrimental to an organ system. Deregulation of autophagy has been implicated in a number of pathologies such as cardiovascular disease, diabetes and cancer (Troncoso et al. 2013). or otherwise, on autophagy, with the findings being cell specific results. For example, et al. 20 09) and adrenocortical cancer cells (Cerquetti et al. 2011), while Mahmood et al. (2011) derived macrophages, but there are no reports on the effects in heart cells. Because both AMPK and mitochondrial dysfunction can induce autophagy, it is a niche that must be explored.
35 Adverse Effects of Rosiglitazone Therapy identified and applied as anti diabetic agent s, they have also been associated with elevated risk of myocardial infarction and adverse cardiovascular events, such as congestive heart failure, and cardiovascular ischemia, morbidity and mortality (Nissen & Wolski 2007, 2011). In a meta analysis by Nis sen and Wolski in 2007, it was determined that treatment with rosiglitazone significantly increased the risk of myocardial infarctions. These findings are of great importance because there are more than 23 million people with diabetes in the United States and 300 million worldwide. Cardiovascular disease is the leading cause of death in patients with T2DM, which represents 68% of all causes of mortality in these patients (Nissen & Wolski 2007). In a follow up metaanalysis in 2011, Nissen and Wolski reported a 2839% increase in the risk of myocardial infarctions after treatment with rosiglitazone. The reason for these adverse side effects has yet to be fully elucidated, but it is currently thought to be due to higher levels of circulating low density lip oprotein cholesterol, which is increased by 23% after treatment with rosiglitazone (Nissen & Wolski 2011). In comparative efficacy trials, rosiglitazone raised triglyceride levels and increased low density lipoprotein cholesterol levels over pioglitazone. induces cardiac hypertrophy (Festuccia et al. 2009). These effects can lead to death, especially in patients already predisposed to congestive heart failure, as both diabetes and congestive heart failure induces alterations in energy metabolism and mitochondrial function (Garnier et al. 2003; Spinazzi et al. 2012)
36 CHAPTER 4 RESULTS Rosiglitazone is a PPARy agonist, which is prescribed to treat Type II Diabetes mellitus due to its insulin sensitizing effects. With more and more diabetic patients taking rosiglitazone, cardiovascular side effects became apparent. This prompted investigations aiming to underst and the underlying biological mechanisms by which rosiglitazone affects cardiac function. In this study, we hypothesized that rosiglitazone impairs cellular bioenergetics and cellular quality control mechanisms, thereby ultimately leading to cardiac dysfunction. Specifically, we hypothesized 1) that rosiglitazone impairs mitochondrial function in the heart, thus compromising energy balance in heart cells; and 2) that rosiglitazone suppresses autophagic activity in heart cells, which could lead to disruption of cellular homeostasis and heart remodeling. To test the first hypothesis (section 4.1), we measured the weight of hearts from Fischer 344 rats that had been administered 10mg/kg/day rosiglitazone for 19 days, and determined cardiac mitochondrial respiratory function (indicated by cytochrome c oxidase and citrate synthase activity). To complement these analyses, we exposed cultured cardiomyocytes (AC16 cells, a human derived ventricular cardiac cell line) to rosiglitazone and assessed mitochondrial functi on using enzyme activity assays and high resolution respirometry. To test the second hypothesis (section 4.2), we analyzed the autophagic response in hearts from rosiglitazone treated Fischer 344 rats and AC16 cells by determining the expression of an autophagy marker protein. In summary, we found that 1) routine respiration and maximal electron transport capacity of the mitochondrial electron transport system were decreased by high concentrations of rosiglitazone in vitro but 2) the activity of the mitoch ondrial enzyme
37 cytochrome c oxidase was decreased by low concentrations of rosiglitazone in vitro or was not affected at all in vivo Furthermore, we report that 3) autophagic flux was not affected by rosiglitazone in vivo and in vitro Effect of Rosigl itazone on Mitochondrial Function in vivo and in vitro Effect of Rosiglitazone on H e arts from Fischer 344 R ats Effect of r osiglitazone administration on heart size in Fischer 344 rats Fischer 344 rats of three different ages were administered 10mg Rosiglitazone/kg/day for 19 days. At the end of this period the rats were weighed to determine their body weight and subsequently euthanized (see Methods section). In order to determine if rosiglitazone administration had an effect on heart size the heart was col lected from each rat after euthanization, flushed with PBS, blotted dry and weighed. For further analysis, heart weight was normalized to body weight (HW/BW, mg g1, Figure 41). Heart weight and body weight increased with age, independent of treatment (Table 4 1; age 5 and 12 months, and 5 and 18 months, but not between ages 12 and 18 months (Table 4 1). However, heart weight normalized to body weight (HW/BW) decreased with age, independent of tr eatment (ageeffect, p<0.001; Figure 41). Rosiglitazone treatment had no effect on HW/BW compared to controls of the same age (treatment effect, p=0.502, 5 mo; 0.615, 12 mo; 0.263, 18 mo). Effect of r osiglitazone administration on mitochondrial e nzyme a ct ivities in heart from Fischer 344 r ats In order to determine if mitochondrial function in heart tissue of Fischer 344 rats was affected by rosiglitazone administration, we analyzed the enzymatic activity of two mitochondrial enzymes, cytochrome c oxidase and citrate synthase, in heart tissue
38 extracts (Figure 42, 4 3 and 44). Cytochrome c oxidase (COX) is one of the electron and proton transporting enzyme complexes (complex IV) of the electron transport system, and its activity has been used as a marker of mitochondrial function (Campian et al. 2007). Citrate synthase (CS) is an enzyme of the Krebs cycle (citric acid cycle), located in the mitochondrial matrix. It has been shown to correlate with mitochondrial content, and has therefore used as a marker enzyme of this parameter (Spinazzi et al. 2012). In order to evaluate the specific activity of COX, its enzymatic activity i s commonly normalized to CS activity. Normalizing COX activity, the indicator of mitochondrial function, to CS activity, measure of mitochondrial content, allows us to assess mitochondrial function per mitochondrial unit. Citrate synthase activity CS activity in tissue extracts from rat heart was not affected by age or rosiglitazone treatment (p=0.248 and p=0.535, respectively, Figure 42), and there was no interaction effect of age and treatment (p=0.543). Specifically, tissue extracts of hearts from 5 months old rats that had received vehicle or rosiglitazone showed an average CS activity of 0.935 0.125 and 0.936 0.0731 mol/min/mg protein, respectively, and this difference was not statistically significant (p=0.994). CS activity did not significa ntly change in 12 months old rats within the respective treatment groups and was 0.883 0.082 mol/min/mg protein in vehicle treated rats (p=0.752), and 0.894 0.10 mol/min/mg protein in rosiglitazone treated rats (p=0.470), respectively, or in 18 month s old rats, which averaged 0.902 0.0.89 in vehicle treated rats (p=0.827), and 0.828 0.177 mol/min/mg protein in rosiglitazone treated rats (p=0.219). No significant change in CS activity was determined in both vehicle and rosiglitazone treated rats at the age of 18 months compared to 12 months old rats (p=0.741, vehicle; p=0.458
39 rosiglitazone). Finally, there was no significant difference in CS activity between vehicle and Rosiglitazone treated rats at any age (p=0.994, age 5 months; p=0.852, age 12 m onths; p=0.221, age 18 months). Cytochrome c oxidase activity COX activity in tissue extracts from rat heart was not affected by age or rosiglitazone treatment (p=0.634 and p=0.459, respectively, Figure 43), and there was no interaction effect of age and treatment (p=0.190). Specifically, tissue extracts of hearts from 5 months old rats that received vehicle or rosiglitazone had an average COX activity of 0.582 0.176 and 0.607 0.208 mol/min/mg protein, respectively, and this difference was not statis tically significant (p=0.784). COX activity did not significantly change in 12 months old rats within the respective treatment groups, and was 0.456 0.167 mol/min/mg protein in vehicle treated rats (p=0.290), and 0.615 0.087 mol/min/mg protein in ros iglitazone treated rats (p=0.928), respectively, or in 18 months old rats, which averaged 0.593 0.137 mol/min/mg protein in vehicle treated rats (p=0.907), and 0.523 0.218 mol/min/mg protein in rosiglitazone treated rats (p=0.591). No significant change in COX activity was determined in both vehicle and rosiglitazone treated rats at the age of 18 months compared to 12 months old rats (p=0.332, vehicle; p=0.659, rosiglitazone). Finally, there was no significant difference in COX activity between vehicl e and rosiglitazone treated rats at any age (p=0.784, age 5 months; p=0.067, age 12 months; p=0.447, age 18 months). C ytochrome c oxidase activity normalized to citrate synthase activity Normalized COX activity, hereafter referred to as normalized COX ac tivity, in tissue extracts from rat heart was not affected by age or rosiglitazone treatment (p=0.860 and p=0.345, respectively, Figure 44), and there was no interaction effect of
40 age and treatment (p=0.386). Specifically, tissue extracts of hearts from 5 months old rats that received vehicle or rosiglitazone had an average normalized COX activity of 0.636 0.215 and 0.655 0.245 mol/min/mg protein, respectively, and this difference was not statistically significant (p=0.869). Normalized COX activity di d not significantly change in 12 months old rats within the respective treatment groups and was 0.522 0.202 mol/min/mg protein in vehicle treated rats (p=0.496), and 0.700 0.146 mol/min/mg protein in rosiglitazone treated rats (p=0.893), respectively or in 18 months old rats, which averaged 0.657 0.133 mol/min/mg protein in vehicle treated rats (p=0.856), and 0.639 0.267 mol/min/mg protein in rosiglitazone treated rats (p=0.884). No significant change in normalized COX activity was determined i n both vehicle and rosiglitazone treated rats at the age of 18 months compared to 12 months old rats (p=0.516, vehicle; p=0.919, rosiglitazone). There was no significant difference in normalized COX activity between vehicle and rosiglitazone treated rats at any age (p=0.869, age 5 months; p=0.092, age 12 months; p=0.871, age 18 months). Su mmary of A nalyses of Heart Size and Mitochondrial Function in Fischer 344 R ats In summary, we found that rosiglitazone administration to Fischer 344 rats had no effect on heart weight and heart weight normalized to body weight. However, heart weight normalized to body weight decreased from 5 to 12 and 18 months of age. Furthermore, neither rosiglitazone administration nor age had an effect on activities of the mitochondrial enzymes cytochrome c oxidase and citrate synthase in rat heart. Effect of R osiglita zone on Mitochondrial Function in AC16 C ardiomyocytes In addition to the experiments and analyses on rat heart tissue, we conducted experiments on the effects of rosiglit azone in vitro which allowed manipulation of
41 rosiglitazone concentrations and exposure times. We used AC 16 cells, which are immortalized human adult ventricular cardiomyocytes, expressing adult cardiomyocyte myosin heavy chain (Garbern et al. 2013). This made them a good in vitro complement to the in vivo study. We measured mitochondrial respiration and enzymatic activity of two mitochondrial enzymes, cytochrome c oxidase and citrate synthase, using high resolution r espirometry (HRR), and spectrophotometric enzyme activity assays, respectively. Effects of Rosiglitazone on Mitochondrial Enzymatic Activities in vitro In order to determine if the dose of rosiglitazone application has an effect on mitochondrial function i n cardiomyocytes, we exposed AC16 cells to four doses (0, 10M, 100M, or 200M) of rosiglitazone for four hours and assessed the activities of two mitochondrial enzymes, cytochrome c oxidase and citrate synthase (Figures 45, 4 6 and 47). Citrate synthas e activity CS activity in AC16 cell lysates was not affected by rosiglitazone dose (p=0.249, Figure 45). Specifically, cells exposed to vehicle (0.01% DMSO) showed an average CS activity of 0.568 0.0156 mol/min/mg protein, cells exposed to 10M rosiglitazone showed an average CS activity of 0.0318 0.0059 mol/min/mg protein, cells exposed to 100M rosiglitazone showed an average CS activity of 0.0468 0.0207 mol/min/mg protein, and cells exposed to 200M rosiglitazone showed an average CS act ivity of 0.0527 0.0125 mol/min/mg protein. Activities in the rosiglitazone groups were not statistically different from control (0 vs. 10M: p=0.196; 0 vs. 100M: p=0.674; 0 vs. 200M: p=0.740).
42 Cytochrome c oxidase activity COX activity in AC16 cell lysates was not affected by rosiglitazone dose (p=0.151, Figure 46). Specifically, cells exposed to vehicle (0.01% DMSO) showed an average COX activity of 0.0324 0.0212 mol/min/mg protein, cells exposed to 10M rosiglitazone showed an average COX activi ty of 0.0097 0.00045 mol/min/mg protein, cells exposed to 100M rosiglitazone showed an average COX activity of 0.0165 0.00205 mol/min/mg protein, and cells exposed to 200M rosiglitazone showed an average COX activity of 0.0173 0.00451 mol/min/mg protein. When activities of the rosiglitazone groups were compared to the control group, 10M rosiglitazone treated cells had a significantly lower COX activity (p<0.05), but the other treatment groups were not significantly different (0 vs. 100M: p=0.211; 0 vs. 200M: p=0.127). C ytochrome c oxidase activity normalized to citrate synthase activity Normalized COX activity of AC16 cells exposed to rosiglitazone was not affected by the dose of rosiglitazone treatment (p=0.262 Figure 47). Specifically, cells exposed to vehicle (0.01% DMSO) showed an average normalized COX activity of 0.540 0.212 mol/min/mg protein, cells exposed to 10M rosiglitazone showed an average normalized COX activity of 0.310 0.0426 mol/min/mg protein, cells exposed to 100M ros iglitazone showed an average normalized COX activity of 0.390 0.126 mol/min/mg protein, and cells exposed to 200M rosiglitazone showed an average normalized COX activity of 0.342 0.123 mol/min/mg protein. Activities in the rosiglitazone groups were not statistically different from control (0 vs. 10M: p=0.215; 0 vs. 100M: p=0.151; 0 vs. 200M: p=0.199).
43 Effects of Rosiglitazone on Mitochondrial Respiration in vitro In addition to determination of mitochondrial enzyme activity, mitochondrial respirat ion was measured, using a High Resolution Respirometer. AC16 cells were exposed to rosiglitazone and oxygen consumption of intact cells in culture media was assessed following the exposure. A low (10M) or a high (200M) dose of rosiglitazone was applied f or a duration of 4 hours or 24 hours (for number of replicates for each treatment see Table 42). Each rosiglitazone exposure experiment was accompanied by a control exposure (cell culture media). Based on the titration protocol outlined in the Methods sec tion, the following respiratory states, and flux and coupling control ratios were analyzed and calculated (for definitions refer to Methods section): Routine respiration (R; oxygen consumption rate, or oxygen flux, of intact cells in culture media in the absence of inhibitors or uncouplers), LEAK respiration (L; oxygen flux in the presence of an inhibitor of the phosphorylation system, e.g. oligomycin), ETS capacity (E; oxygen flux in the presence of an uncoupler, e.g. FCCP, which dissipates the mitochondri al membrane potential), Leak control ratio (L/E), and Uncoupling control ratio (E/R). Routine respiration: Routine respiration rate under control conditions was 46.1 18.5 and 42.6 15.6 pmol s1 106 cells (4 h and 24 hour exposure, respectively; Table 4 2, Figure 48). When cells were exposed to 10 M r osiglitazone for 4 or 24 hours, routine respiration amounted to 50.9 13.1 and 42.5 11.9 pmol s1 106 cells, respectively. Cells exposed to 200 M r osiglitazone for 4 or 24 hours had a routine respir ation rate of 24.5 13. 7 and 27.0 3.2 pmol s1 106 cells, respectively. There was no overall effect of duration of exposure on routine respiration (p=0.557), and duration of exposure had no effect on routine respiration within each dose group
44 (p=0.607 0.408, and 0.798 for 0, 10, and 200 M rosiglitazone, respectively). However, there was an overall significant effect of rosiglitazone dose on routine respiration (p=0.007). That is, the overall mean of routine respiration in cells exposed to 200 M rosi glitazone (4h and 24 h combined) was lower than the overall mean of routine respiration of control cells (4h and 24h combined; p=0.012), and of cells exposed to 10 M rosiglitazone (4h and 24h combined; p=0.012), independent of duration of the exposure. Ro utine respiration of control cells (4h and 24h combined) and cells exposed to 10 M rosiglitazone (4h and 24h combined) were not statistically different (p=0.704). When differences between doses within the same exposure duration were analyzed, routine respiration was decreased in cells treated for 4 hours with 200 M compared to 10 M rosiglitazone (p=0.025) and to control cells (p=0.037), but not when cells were treated for 24 hours (p=0.248, 200 M 10 M; p=0.239, 200 M control). Routine respiration of control cells and cells treated with 10 M rosiglitazone did not differ from each other, independent of duration of exposure (4h: p=0.592, 24h: p=0.985). Finally, there was no interaction effect of dose x duration (p=0.741). LEAK respiration: LEAK respi ration (LEAK; L) was measured after the inhibition of the phosphorylation system through the ATP synthase inhibitor oligomycin. This inhibition increases the electrochemical proton gradient across the inner mitochondrial membrane. Oxygen flux is depressed to a level determined by proton leak. LEAK respiration in AC16 cells was not affected by rosiglitazone (dose effect: p=0.246) or duration of exposure to vehicle or rosiglitazone (duration effect: p=0.973), and there was no interaction affect between these factors (p=0.796) (Figure 49). Specifically, the overall means of LEAK respiration in cells exposed to 0, 10 and 200 M rosiglitazone
45 (4h and 24 h combined) did not statistically differ from each other, independent of duration of the exposure (p=0.983, 010 M; p=0.305, 0200 M; p=0.314, 10200 M). When differences between doses within the same exposure duration was analyzed, LEAK respiration was unchanged in cells treated for 4 hours with 10 or 200 M compared to control cells (p=0.802, 010 M; p=0.282, 0 200 M; p=0.456, 10200 M, Table 42). LEAK respiration was also unaffected in cells treated for 24 hours with 10 or 200 M rosiglitazone compared to control cells (p=0.763, 010 M; p=0.760, 0200 M; p=0.794, 10200 M). ETS capacity: The maximal e lectron transport capacity (ETS capacity; E) is experimentally induced by titration of an uncoupler (FCCP), and is characterized by the collapse of the mitochondrial membrane potential across the inner mitochondrial membrane. ETS capacity in AC16 cells was not affected by duration of exposure to vehicle or rosiglitazone (p=0.558), but was significantly decreased by rosiglitazone (dose effect: p=0.001; Figure 410). Specifically, the overall mean of ETS capacity in cells exposed to 200 M rosiglitazone (4h and 24 h combined) was lower than the overall mean of ETS capacity of control cells (4h and 24h combined; p=0.004), and of cells exposed to 10 M rosiglitazone (4h and 24h combined; p=0.002), respectively, independent of duration of the exposure (Table 42) ETS capacity of control cells (4h and 24h combined) and cells exposed to 10 M rosiglitazone (4h and 24h combined) were not statistically different (p=0.261). When differences between doses within the same exposure duration was analyzed, ETS capacity was decreased in cells treated for 4 hours with 200 M compared to 10 M rosiglitazone (p=0.018) and to control cells (p=0.042). ETS capacity in cells treated for 24 hours with 200 M rosiglitazone was
46 significantly or tended to be lower compared to control ( p=0.048) and cells exposed to 10 M rosiglitazone (p=0.054), respectively. ETS capacity of control cells and cells treated with 10 M rosiglitazone did not differ from each other, independent of duration of exposure (4h: p=0.279, 24h: p=0.623). Finally, there was no interaction effect between duration of exposure and rosiglitazone dose (p=0.894). LEAK control ratio: The Leak control ratio (LCR; L/E) is an index of uncoupling at constant ETS capacity, and indicates how much LEAK respiration is contributing to the overall capacity of the electron transport system. LCR in AC16 cells was not affected by duration of exposure to vehicle or rosiglitazone (p=0.379), but was significantly increased by rosiglitazone (dose effect: p=0.004; Figure 411). Specifically, independent of duration of the exposure, the overall mean of LCR in cells exposed to 200 M rosiglitazone (4h and 24 h combined) was higher than the overall mean of LCR in control cells (4h and 24h combined; p=0.016), and of cells exposed to 10 M rosiglit azone (4h and 24h combined; p=0.005), respectively (Table 4 2). LCR of control cells (4h and 24h combined) and cells exposed to 10 M rosiglitazone (4h and 24h combined) were not statistically different (p= 0.218). When differences between doses within the same exposure duration was analyzed, LCR was not significantly different in cells treated for 4 hours independent of rosiglitazone dose (p=0.258, 010 M; p=0.266, 0 200M; p=0.093, 10 M 200 M). LCR in cells treated for 24 hours with 200 M rosiglitazon e was significantly elevated compared to control (p=0.039) and cells exposed to10 M rosiglitazone (p=0.036), respectively. LCR of control cells and cells treated with 10 M rosiglitazone did not differ from each other after 24 h of exposure
47 (p=0.549). Finally, there was no interaction effect between duration of exposure and rosiglitazone dose (p=0.724). Routine control ratio: The routine control ratio (R/E) is used to evaluate how close routine respiration operates to ETS capacity. R/E in AC16 cells was not affected by duration of exposure to vehicle or rosiglitazone (p=0.609), but was significantly decreased by rosiglitazone (dose effect: p=0.029; Figure 412). Specifically, independent of duration of the exposure, the overall mean of R/E in cells exposed to 200 M rosiglitazone (4h and 24 h combined) was higher than the overall mean of R/E of cells exposed to 10 M rosiglitazone (4h and 24h combined; p=0.025). R/E of control cells (4h and 24h combined) was not statistically different from cells exposed to 10 M rosiglitazone (4h and 24h combined; p=0.150), and 200 M rosiglitazone (4h and 24h combined; p=0.148). When differences between doses within the same exposure duration was analyzed, R/E was increased in cells treated for 24 hours with 200 M compared to 10 M rosiglitazone (p=0.019), while there were no statistical differences between the other treatments (4h exposure: p=0.459, 010 M; p=0.914, 0200 M; p=0.508, 10200 M; 24h exposure: p=0.082, 0200 M, p=0.211, 010 M) (Table 42). Finally, the re was no interaction effect between duration of exposure and rosiglitazone dose (p=0.255). Summary of A nalyses of Mitochondrial Function in AC16 C ells In summary, we found that, independent of duration of exposure, routine respiration and ETS capacity of AC16 cells were decreased when cells were exposed to 200 M rosiglitazone, while LEAK respiration was unaffected. This was reflected in an increase in LEAK control ratio (LCR; L/E) and routine control ratio (RCR; R/E). Analysis of mitochondrial enzymes showed no effect of rosiglitazone on mitochondrial function as
48 determined by activities of COX and CS. COX activity appeared to be depressed by exposure to 10M rosiglitazone, but the effect was removed when COX activity was normalized to CS activity. Effect of Rosiglitazone on Autophagy in vivo and in vitro In this part of the study, we investigated the effect of rosiglitazone administration on autophagy in vivo in rat heart, and in vitro in AC16 cardiomyocytes. We measured in heart tissue extracts and cell l ysates, respectively, protein expression of the autophagy marker MAP LC3 (microtubuleassociated protein light chain 3). LC3 is essential for the expansion of the early autophagosome, and becomes posttranslationally modified. PreLC3 is processed by the au tophagy protein Atg4 to its cytosolic form, LC3I. LC3 I is then activated by Atg7, transferred to Atg3 and lipidated to its membrane bound form, LC3 II, localized to the preautophagosome structure and the autophagosomes (Abeliovich et al. 2000). Its posttranslational modification from cytosolic LC3I to membrane bound LC3 II makes it a useful marker of autophagy (Rubinsztein et al. 2009). In this study, protein expression of LC3I and LC3II was analyzed in rat heart tissue extracts, and calculated the rat io of LC3 II/I as a measure of autophagic activity in the heart. However, these measures represent only a snapshot of autophagic activity in the biological material fixed at a given time. T herefore the investigation of autophagy in rat cardiac tissue was complemented with in vitro experiments. Here, AC16 cells were exposed to rosiglitazone in the presence and absence of bafilomycin A1 (BafA1), a lysosomal inhibitor (Rubinsztein et al. 2009). BafA1 inhibits the fusion of the autophagosome with the lysosome, which leads to an accumulation of the autophagosomebound form LC3II. The difference of accumulated LC3II between
49 BafA1 treated and BafA1untreated cells can then be utilized to determine the autophagic flux. We found that autophagy, using the protein ex pression of LC3II and LC3 I as an indicator, was not affected by age (p=0.901) or treatment (p=0.755) in the hearts of Fischer 344 rats supplemented with 10mg/kg/day rosiglitazone for 19 days (Figure 1413). Specifically 5 months of age did not significantly differ from 12 months or 18 months (p=0.923 and p=0.967, respectively) and 12 months did not significantly differ from 18 months (p=0.953). Within the vehicle treatment 5 months did not differ from 12 months or 18 months (p=0.791 and p=0.854, respectiv ely) and 12 months did not significantly differ from 18 months (p=0.876). Similar results were obtained in the rosiglitazone treated group, where 5 months did not differ from 12 months or 18 months (p=0.964 and p=0.891, respectively) and 12 months did not significantly differ from 18 months (p=0.975). We next analyzed autophagic flux in the presence of rosiglitazone in AC16 cells, by exposing cells to fed conditions (control), starvation (used to induce autophagy through nutrient deprivation) and varying doses of rosiglitazone, and then treating a duplicate sample of cells with BafA1. BafA1 is a proton pump inhibitor, which prevents the fusion of the autophagosome with the lysosome by neutralizing the lysosomal pH, causes the accumulation of autophagosomes ( Rubinsztein et al. 2009). Because the final degradation of the autophagosomes and their contents is prevented, the autophagosomal membranebound form of the autophagy marker protein LC3II accumulates. LC3II accumulation can then be utilized to determine the autophagic flux
50 by calculating the difference of LC3II protein level between the cells treated without and the cells treated with BafA1 within the same treatment. We first assessed whether autophagy is inducible in the AC16 cell via starvation. Altho ugh there is an increase in autophagosome formation in the starved cells compared to over control cells with mean LC3II/LC3I ratios of 0.568 0.310 and 1.582 0.237, respectively, and LC3II/LC3I ratios of 3.086 1.614 and 6.115 3.89 in BafA1treated c ontrol and starved cells, respectively, the overall effect of 4 hours of starvation was not statistically significant (p=0.150). We next assessed autophagic flux in AC16 cells exposed to rosiglitazone by calculating the foldincrease caused by addition of BafA (LC3 II/I ratio of cells in the presence of BafA1divided by the value in the absence of BafA1). Treatment with rosiglitazone did not have an effect on autophagic flux (p=0.219; Figure 1414). Specifically, LC3 II/I ratio increased 4.71 2.12 fold in control cells, 10.05 7.40 in cells treated with 10M rosiglitazone, 19.16 12.92 in cells treated with 100M rosiglitazone, and 7.66 4.15 in cells treated with 200M rosiglitazone. Comparisons between the autophagic flux in cells treated with different doses of rosiglitazone were not statistically significant (0 10M: p=0.860; 0100M: p=0.305; 0200M: p= 0.884; 10100M: p=0.674; 10200M: p=0.749; 100200M: p=0.460; Figure 1414 D). In summary, autophagy was not affected by treatment with rosiglit azone in vivo or in vitro TZDs or otherwise, on autophagy, to corroborate or contradict our findings.
51 Age (months) 51218HW/BW (mg/g) 0.00.51.01.52.02.53.0 control Rosi. a,bbaccd,ede Figure 41 Heart weight (HW, mg) normalized to body weight (BW, g) of Fischer 344 rats age 5, 12 and 18 months that received rosiglitazone or vehicle (control) for 19 days prior to euthanization. (for number of animals per group refer to Methods section). Data are r epresented as mean SD. Same indices indicate significant difference between groups. a: p< 0.001, b: p=0.004, c: p=0.005, d: p=0.014, e: p=0.038.
52 Table 41 Heart (HW) and body weight (BW) and heart weight normalized to body weight (HW/BW) of Fischer 344 rats aged 5, 12 and 18 months, rats were treated for 19 days with either vehicle (control) or 10 mg rosiglitazone/kg/day (rosi). Data are represented as mean SD. Superscripts indicate significant difference between groups with ak: p=0.014, l: Age (mo) Treatment n HW BW HW/BW 5 control 8 843 60 a,b 358 23 e,f 2.36 0.2 i 5 rosi 8 814 93 c,d 353 41 g.h 2.31 0.13 k,l 12 control 11 996 63 a 468 27 e 2.13 0.13 i 12 rosi 9 978 55 c 467 15 g 2.10 0.14 k 18 control 10 985 111 b 446 39 f 2.21 0.17 18 rosi 8 1004 67 d 472 31 h 2.13 0.1 l Age (months) 5 10 15 20mol min-1 mg-1 prot. 0.0 0.2 0.4 0.6 0.8 1.0 1.2 control Rosiglitazone Figure 42 1 mg1 protein) in tissue extracts of hearts from Fischer 344 rats age 5, 12 and 18 months that received rosiglitazone or vehicle (control) for 19 days prior to euthanization (for number of animals per group refer to Methods section) Data are represented as mean SD.
53 Age (months) 5 10 15 20mol min-1 mg-1 prot. 0.0 0.2 0.4 0.6 0.8 1.0 control Rosiglitazone Figure 43. 1 mg1 protein) in tissue extracts of hearts from Fischer 344 rats age 5, 12 and 18 months that received rosiglitazone or vehicle (control) for 19 days prior to euthanization. (for number of animals per group refer to Methods section). Data are represented as mean SD.
54 Age (months) 5101520COX activity / CS activity 0.00.20.40.60.81.0 control Rosiglitazone Figure 44. COX activity normalized to CS activity in tissue extracts of hearts from Fischer 344 rats age 5, 12 and 18 months that received rosiglitazone or vehicle (control) for 19 days prior to euthanizat ion. (for number of animals per group refer to Methods section). Data are represented as mean SD.
55 Figure 45. 1 mg1 protein) in AC16 cell that were exposed to rosiglitazone or vehicle (control) for 4 hours prior to cell lysis (for number of experiments per group refer to Methods section). Data are represented as mean SD. Same indices indicate significant difference b etween groups. Rosiglitazone Dose 0uM10uM100uM200uMCS Activity (umol/min/mg) 0.000.020.040.060.08
56 Figure 46 COX activity in AC16 cell that were exposed to rosiglitazone or vehicle (control) for 4 hours prior to cell lysis (for number of experiments per group refer to Methods section). Data are represented as mean SD. Sa me indices indicate significant difference between groups. Rosiglitazone Dose 0uM10uM100uM200uMCOX Activity (umol/min/mg) 0.000.010.020.030.040.050.06 a
57 Figure 47. COX activity normalized to CS activity in AC16 cell that were exposed to rosiglitazone or vehicle (control) for 4 hours prior to cell lysis (for number of experiments per group refer to Methods section). Data are represented as mean SD. Same indices indicate significant difference between groups. Rosiglitazone Dose 0uM10uM100uM200uMCOX Activity Normalized to CS Activity (umol/min/mg) 0.00.20.40.60.81.0
58 Table 42. Oxygen flux (pmol s1 106 cells) of AC16 cells under control conditions (CTRL) or after exposure to rosiglitazone (10 or 200 uM) for 4 or 24 hours. Oxygen flux measured as: Routine respiration (Routine), LEAK respiration (LEAK), electron transport capacity (ETS). LEAK Control Ratio (LEAK/ETS; L/E) and Routine Control Ratio (Routine/ETS; R/E) were calculated from indicated oxy gen flux. Data represent mean standard deviation. n: number of replicates indicates independent replication of experiment Condition n Routine LEAK ETS L/E R/E CTRL 4 h 10 46.1 18.5 27.2 14.4 74.1 33.3 0.381 0.118 0.646 0.111 CTRL 24 h 10 42.6 15.6 24.3 6.7 72.6 27.3 0.379 0.164 0.614 0.147 R osi. 4h 4 50.9 13.1 25.7 4.6 91.0 20.3 0.288 0.056 0.557 0.065 24h 5 42.5 11.9 26.0 5.8 79.7 18.2 0.334 0.078 0.532 0.070 4h 5 24.5 13.7 18.2 11.8 39.5 23.8 0.494 0.194 0.639 0.119 24h 4 27.0 3.2 20.5 3.8 36.2 7.3 0.579 0.132 0.760 0.112
59 Rosiglitazone 0 mM0.01 mM0.2 mMR (pmol O2 s-1 10-6 cells) 010203040506070 4h exposure 24h exposure a a,b bcdc.d Figure 48 Routine respiration of AC16 cells measured in pmol s1 106 cells, compared between two exposure durations (4 hours and 24 hours) and three doses of Data are represented as mean SD. Same indices indicate significant difference. a: p=0.012, b: p=0.012, c: p=0.037, d p=0.0 25.
60 Table 43 Statistical analysis of differences between routine respiration rates of AC16 cells under control conditions or after exposure to rosiglitazone (10 or 200 values were determined using Twoway ANOVA and post hoc anal 0.05. *: time of exposure (duration of exposure) Treatment groups compared p value Dose effect 0.007 Time effect* 0.557 Dose x Time* interaction 0.741 0 0.704 0 0.012 10 Rosiglitazone 0.012 4h Control / 24h Control 0.607 4h 0.408 4h 0.798 0.592 4h: Control 0.037 0.025 0.985 0.239 0.248
61 Figure 49 LEAK respiration of AC16 cells measured in measured in pmol s1 106 cells, compared between two exposure durations (4 hours and 24 hours) and three Data are represented as mean SD.
62 Table 4 4 Statistic al analysis of differences between LEAK respiration rates of AC16 cells under control conditions or after exposure to rosiglitazone (10 or 200 values were determined using Twoway ANOVA and post hoc analysis. Statistical difference 0.05. Treatment groups compared p value Dose effect 0.246 Time effect* 0.973 Dose x Time* interaction 0.796 0 0.983 0 0.305 10 0.314 0.802 4h: Control 0.282 0.456 0.763 0.760 Rosiglitazone 0.794 *: time of exposure (duration of exposure)
63 Figure 410. ETS respiration of AC16 cells measured in pmol s1 106 cells, compared between two exposure durations (4 hours and 24 hours) and three doses of Data are represented as mean SD. Same indices indicate significant difference. a: p=0.004, b: p=0.002, c: p=0.042, d: p=0.018, e: p=0.048.
64 Table 45. Statistical analysis of differences between electron transport capacity (ETS) of AC16 ce lls under control conditions or after exposure to rosiglitazone (10 values were determined using Twoway ANOVA and post hoc analysis. Statistical difference was considered Treatment groups compared p value Dose effect 0.246 Time effect* 0.973 Dose x Time* interaction 0.796 0 0.983 0 0.305 10 0.314 0.802 4h: Control 0.282 0.456 0.763 0.760 0.794 *: time of exposure (duration of exposure)
65 ()Rosiglitazone 0 mM 0.01 mM 0.2 mML/E 0.0 0.2 0.4 0.6 0.8 4h exposure 24h exposure a a,b b c c,d d Figure 411. Leak control ratio (L/E) of AC16 cells compared between two exposure Data are represented as mean SD. Same indices indicate significant difference. a: p=0.004, b: p=0.002, c: p=0.042, d: p=0.018, e: p=0.048.
66 Table 46 Statistical analysis of differences between LEAK Control Ratio (L/E) of AC16 cells under control conditions or after exposure to rosiglitazone (10 or 200 values were determined using Twoway ANOVA and 0.05. Treatment groups compared p value Dose effect 0.004 Time effect* 0.379 Dose x Time* interaction 0.724 0 0.218 0 0.016 10 0.005 0.258 4h: Control 0.266 Rosiglitazone 0.093 0.549 0.039 0.036 *: time of exposure (duration of exposure)
67 Rosiglitazone 0 mM0.01 mM0.2 mMR/E 0.00.20.40.60.81.0 4h exposure 24h exposure aabb Figure 412. Routine control ratio (Routine/ETS, R/E) of AC16 cells compared between two exposure durations (4 hours and 24 hours) and three doses of SD. Same indices indicate significant differen ce. a: p=0.025, b: p=0.019.
68 Table 47 Statistical analysis of differences between Respiratory Control Ratio (R/E) of AC16 cells under control conditions or after exposure to rosiglitazone (10 or values were determined using Twoway ANOVA and post hoc analysis. Statistical difference was considered significant with p Treatment groups compared p value Dose effect 0.029 Time effect* 0.609 Dose x Time* interaction 0.255 0 0.150 0 0.148 10 0.025 0.459 4h: Control 0.914 0.508 0.211 0.082 0.019 *: time of exposure (duration of exposure)
69 A. B. Figure 413. Protein expression of LC3 I and II in rat heart tissue extracts. A.) Western Blot of LC3 I and LC3II. Lanes annotated with months of age (5, 12, and 18) and treatment (R: rosiglitazone; V: vehicle). B.) Ratio of LC3II/LC3I in rat cardiac tissue (n=3 per treatment group). Data are represented as mean SD. 0 0.2 0.4 0.6 0.8 1 1.2 1.4 1.6 1.8 2 5 Months 12 Months 18 MonthsLC3II/LC3I Vehicle Rosiglitazone
70 A. B. Figure 414. Protein expression of LC3 I and II in AC16 cell lysates. A.) Western Blot of LC3 I and II in AC16 cells that had been exposed to varying doses of rosiglitazone for 4 hours (0, 10, 100, 200 uM ) or that had been starved for the same duration, in the absence or presence of 50 nM bafilomycin A1. B.) Ratio of LC3II/LC3I in AC16 cell lysate (n=3 per treatment group). Data are represented as mean SD. Autophagy is inducible in AC16 cells by starvati on. C.
71 D. Figure 414. Continued C.) Measurement of autophagy as a ratio of LC3II/LC3I in AC16 cells that have been exposed to varying doses of rosiglitazone (n=3). D.) Fold increase after treatment with bafilomycin a1 (Baf/no Baf). Bafilomycin a1 (50nM) is used to assess autophagic flux by inhibiting the formation of the autophagosome to the lysosome, thus preventing degradation.
72 CHAPTER 5 DISCUSSION Rosiglitazone was approved by the FDA for administration in the treatment of Type 2 Diabetes mellitus (T2DM) in 1999. With the increase in prescriptions of this medication for T2DM, a concurrent increase in adverse cardiovascular events, such as myocardial infarctions, cardiovascular mortality and cardiac hypertrophy, became apparent. Diabetic p atients are already predisposed to heart complications (Vinokur et al. 2013), and rosiglitazone clearly exacerbated these symptoms (Starner et al. 2008). to determine wh We hypothesized that 1.) Administration of rosiglitazone will decrease mitochondrial function in the heart, and 2.) Administration of rosiglitazone will increase autophagy in the heart. We tested our hypotheses on cardiac tissue of Fischer 344 rats that had been administered 10mg/kg/day rosiglitazone for 19 days, and cultured cardiomyocytes exposed to varying doses of rosiglitazone. In brief, we found that i.) high doses of rosiglitazone decreased mitochondrial routine respiration and excess capacity in AC16 cells, ii.) mitochondrial enzyme activities (cytochrome c oxidase and citrate synthase) were not affected by rosiglitazone in vivo or in vitro iii.) autophagic flux was not affected by rosiglitazone in vivo or in vitro Heart Size is Not A ffected by Rosiglitazone Administration (F estuccia et al. 2009). Rosiglitazone administration has been shown to induce cardiac hypertrophy (Lee et al. 2010). In this particular study, 8 week old mice were
73 administered 10mg/kg/day rosiglitazone for 4 weeks, and an increase in heart weight to body weight was observed. Lee and colleagues (2010) concluded that the increase in heart size was attributed to an increase in adipose deposition caused by rosiglitazone the energy sensing AMPK pathway, which has been shown to induce cardiac hypertrophy in patients with T2DM (Tian et al. 2001; Dyck & Lopaschuk, 2006; Festuccia et al. 2009). We hypothesized that administration of rosiglitazone would increase the size of the heart relative to body weight. In this study, Fischer 344 rats were administered 10mg/kg/day for 19 days. In order to determine if this dose and duration of treatment with rosiglitazone had an effect on heart weight, we assessed heart weight normalized to body weight of the Fischer 344 rats after the drug treatment. We determined that 19 days of 10 mg/kg/day rosiglitazone did not have an effect on heart size. We did determine, however, a drug independent agerelated increase in heart weight and normalized heart weight between the young and the middleaged and old rat, which was likely due to normal heart growth. It is likely that our results do not corroborate with aforementioned studies due to the duration of rosiglitazone administration in our study. Mito chondrial Enzymes are N ot A ffected by Rosiglitazone Administration in vivo The activities of the mitochondrial enzymes cytochrome c oxidase and citrate synthase are widely used markers of mitochondrial function and content, respectively. Cytochrome c oxidase (COX) is one of the electron and proton transporting enzyme complexes (also named Complex IV) of the electron transport system. Citrate synthase (CS) is a component of the Krebs cycle (citric acid cycle). A commonly applied method to assess COX activit y per mitochondrial unit is to normalize the activity of cytochrome c
74 oxidase to that of citrate synthase ( Campian et al. 2007; ChrzanowskaLightowlers et al. 1993). We analyzed the activity of these enzymes in cardiac tissue of Fischer 344 rats that had b een administered 10 mg/kg/day rosiglitazone or vehicle for a period of 19 days before euthanization. We found that neither age nor treatment, nor an interaction of age and treatment affected CS and COX activity, and COX activity per mitochondrial unit. The se findings are corroborated by a 2010 study by Rabol and colleagues on skeletal muscle from diabetic and healthy patients, in which the authors demonstrated that citrate synthase activity and the protein contents of Complexes I V were not affected by rosi glitazone treatment (4mg/day for 12 weeks). This dose is significantly less than the dose administered to the rats in our study, equating to approximately 0.04mg/kg/day compared to our 10mg/kg/day administration, but the metabolic profiles of the two study organisms also differ greatly in relation to body size. Julie and colleagues (2008) reported that treatment with troglitazone caused mitochondrial dysfunction in livers of patients with Type II Diabetes, although no specific assays of mitochondrial functi on were performed in this study. Mitochondrial Enzymes are Not A ffected by Rosiglitazone Application in vitro To supplement our in vivo work, the activities of cytochrome c oxidase and citrate synthase were analyzed in cultured cardiomyocytes (AC16 cells) that were exposed to period. We found that mitochondrial enzyme activities of COX and CS, representing mitochondrial function and content, respectively, in AC16 cells were not affected by exposure to rosiglitazone regardless of the dose, except in the case of the 10uM rosiglitazone, which decreased COX activity from control. Scatena and colleagues
75 (2004) reported similar results. The authors found that the activity of cyctochrome c oxidase did not differ from control with treatment with ciglitazone, another TZD member (10 60 cells. Interesting to note is that COX activity of AC16 cells significantly decreased from con normalized to CS activity as indicator of mitochondrial content, the effect dissipated, because CS activity was also lower in those cells, albeit not significantly. These data sugge st a decrease in mitochondrial content in this treatment group and preservation of COX function. Removal of depolarized, damaged or dysfunctional mitochondria by autophagy, specifically termed mitophagy, could be the cause for the observed tendency of decr eased mitochondrial content after exposure to 10uM rosiglitazone. Further replications of these experiments will have to clarify this connection, because although we determined a numerical increase in autophagic flux in the presence of lower rosiglitazone doses, the difference to control conditions was not significant. et al. 2011). In support of our speculation, it has been hypothesized that the high doses of TZDs administered for their anti diabetic effects are acting through a receptor independent pathway (Festuccia et al. 2009). This could mean that the effects we have observed PG independent effect on the mitochondria (Bolten et al. 2007). High Doses of Rosiglitazone D ecreases Mitochondrial Respiration in vitro Normal mitochondrial function is essential for life. It is the driving force behind all biological systems requiring ATP. The heart, especially, relies on proper mitochondrial
76 function to supply the constant amount of energy needed to perform its function. In the diabetic patient, many subcellular organelles become dysfunctional, i ncluding the mitochondria, often leading to cardiomyopathy and heart disease (Xu et al. 2012). It has also been reported that TZDs, like rosiglitazone, have been implicated in mitochondrial dysfunction (Brunmair et al. 2011; Rabol et al. 2010; Julie et al 2008; Sanz et al. 2011). We first set out to determine if mitochondrial enzyme activities were affected by rosiglitazone, which we determined were not affected in vivo and in vitro We next exposed cultured cardiomyocytes in vitro to high (200uM) and low (10uM) doses of rosiglitazone for two different durations, acute (4 hours) and 24 hours and subsequently assessed mitochondrial respiration using High Resolution Respirometry (HRR) Specifically, we measured routine respiration (R), LEAK respiration (L), ETS (E; maximum electron transport capacity), and calculated LEAK control ratio (L/E) and routine control ratio (R/E). We determined that duration of exposure to rosiglitazone did not affect any of these parameters. However, the high dose of rosiglitazone, but not the low dose, caused a decrease of routine respiration and ETS capactity, and both doses had no effect on LEAK respiration. These results demonstrate that rosiglitazone can impair cellular respiration and maximum capacity of electron transport, but that the drug does not affect the leakiness of the inner mitochondrial membrane, suggesting that there was not an increase in ROS production. Interestingly, the significant increase in routine control ratio (R/E) at the higher rosiglitazone dose reveal s that the effect of the drug on maximum capacity is stronger than on routine respiration. The underlying cause for the decreased respiration and maximum capacity of the electron transport chain could lie in damage to one or more of the mitochondrial compl exes. We found, however,
77 that cytochrome c oxidase activity, which is Complex IV of the electron transport system, was unaffected by rosiglitazone, and therefore can be eliminated as the limiting factor of the electron transport system. Our respiration experiments were performed on intact cells, and we could not distinguish between activities of individual mitochondrial complexes. Further analysis of mitochondrial respiration in permeabilized cells will be required to determine if other complexes contribute to the observed effects of the drug on cellular respiration. In a 2010 study, Rabol and colleagues reported that rosiglitazone inhibited Complex I of the electron transport system. A study by Sanz and colleagues (2011) reported that specific activities of Complexes I and III were decreased by 50% and 35%, respectively, by TZD treatment of hepatocytes isolated from Wistar rats after a 30 minute incubation period in 25uM or 100uM rosiglitazone. Hoffman and colleagues (2012) identified an off target affinity of the glitazones for binding to dehydrogenases like those found in the electron transport system. These findings are corroborated by a 2004 study by Scatena and colleagues that identified a strong inhibitory effect of rosiglitazone on NADHcytochrome c r eductase (combined enzyme activity of complexes I III). In another study, Pan and colleagues (2006) reported alterations in the inner mitochondrial membrane by rosiglitazone in heart mitochondria of obese compared to nonobese rats. The authors detected changes in the composition of cardiolipin, a phospholipid localized in the inner mitochondrial membrane and essential for optimal function of Complexes I, III, and IV of the electron transport system, (Pan et al. 2006). Specifically, they measured an increase in lineolic acid and a decrease in DHA content. DHA is an omega 3 fatty acid, which is known to exert cardioprotective
78 effects, thus a decrease in DHA content could be pathologically significant (Pan et al. 2 006). However, it is not known whether altered cardiolipin composition has an effect on mitochondrial complex assembling and function, and maybe the formation of the respiratory supercomplexes (the physical association of all respiratory complexes into a supercomplex). Effect of Rosiglitazone on Autophagy in Rat Heart and Cultured Cardiomyocytes Autophagy is a cellular housekeeping mechanism by which cellular constituents are degraded. Autophagy is activated by several conditions including nutrient or amino acid deprivation, energy imbalance, or activation of the energy sensor AMPK. AMPK leads to an induction of autophagy, which in turn increases the degradation of proteins and thereby replenishes the pool of substrates for energy production. Autophagy has also been shown to be induced by dysfunctional mitochondria, thereby removing these potentially damaging organelles before they can cause further harm (for example by an increased release of free radicals). We hypothesized that treatment with rosiglitazone would induce an increase in autophagic activity, due to 1) its activation of AMPK (Fryer et al. 2002), and 2) its negative effect on mitochondrial function, which would induce mitophagy, the autophagic removal of (dysfunctional) mitochondria. We found that autophagy, as indicated by the ratio of LC3II to LC3 I protein expression, is not significantly affected in rat cardiac tissue after supplementation with 10mg/kg/day rosiglitazone for 19 days, or by age. In general, it has been shown in a variety of tissues and species that autophagy is decreased with age (Cuervo 2009; Yen and Klionsky 2008), but age was not a statistically significant factor in our study. This could be due to to the high variation of protein expression in the rats regardless of age, and the number of animals per group analyzed will have to be increased to draw a definite conclusion.
7 9 A reason for the lack of an ageeffect could be that the oldest age in our in vivo study was 18 months. Fischer 344 rats of that age are not yet considered old, but rather at the transition to senescence, and the decline in autophagic cellular housekeeping might not have commenced. Similar to rat heart tissue, exposure of AC16 cells to rosiglitazone had no significant effect on autophagic activity. Taken together, we cannot raw clear conclusions based on the results of both our in vivo and in vitro experiments. More experiments will have to be performed, both in vivo and in vitro to increase the power of the analysis. However, if we assume that the numeri cal increase in autophagic flux (fold increase after BafA1 Figure 414D) in AC16 cells in the presence of 10 and 100 uM rosiglitazone compared to controls is a true trend, at least in vitro one could speculate whether this (at this point speculative) incr ease in autophagy represents an increase in mitophagy, the autophagic removal of (dysfunctional) mitochondria. An increase in mitophagy could be interpreted as a cellular defense mechanism by which further cellular stress caused by damaged, dysfunctional m itochondria is prevented or at least attenuated. The decrease in CS activity (as an indicator of mitochondrial content), albeit nonsignificant, could suggest increased mitophagy. However, this is speculative and has to be tested with further experiments.
80 CHAPTER 6 CONCLUSIONS AND IMPLICATIONS The main finding of our study is that mitochondrial function is impacted by rosiglitazone in vitro In particular, maximum electron transport capacity and routine respiration of AC16 cells were decreased by exposur e to high doses of rosiglitazone (200M), but normalized COX activity was not affected regardless of dose. The precise reason for the observed hindrance of maximum electron transport capacity is not clear in our study, but other groups have speculated on t he reason for the negative impact of rosiglitazone on mitochondrial function. For example, Hoffman and colleagues (2012) assessed the off target binding properties of rosiglitazone, and determined that some of the off target binding partners of the TZDs are dehydrogenases involved in alteration of mitochondrial respiration, leading to changes in metabolism, energy production and insulin sensitivity. In addition, a 2004 study Scatena and colleagues determined that a 96 hour exposure to ciglitazone induced d erangement of the mitochondrial electron system, specifically by inhibiting NADHcytochrome c reductase activity, representing combined activity of Complexes I, II, and III of the electron transport system. And in 2010 Rabol and colleagues reported that rosiglitazone (4mg/day for 12 weeks) affects mitochondrial respiration by inhibiting Complex I of the electron transport system. Interestingly, Pan et al. (2006) found that treatment with rosiglitazone caused cardiolipin remodeling in the heart mitochondria of rats, which could impact optimal function of Complexes I, III, and IV of the electron transport system. In this study, we investigated the effects of rosiglitazone on heart cell bioenergetics by assessing mitochondrial function. The impaired cellular r espiration that we detected in the presence of high rosiglitazone concentrations in vitro could be an
81 underlying cause for the adverse cardiovascular effects reported in T2DM patients who received rosiglitazone. The heart depends on sufficient aerobic ener gy supply. This implies that the heart employs effective energy sensing signaling pathways and tight regulation of substrate supply. The AMP activated protein kinase ( AMPK ) is an important master regulator of metabolism (Zaha & Young 2012) and cellular e nergy sensor, and it is apparent that the AMPK regulatory pathway is particularly important in the heart. The heart has very little reserved energy and depends on AMPK and proper mitochondrial function to sense energy status and to keep the ATP balance intact (Dyck & Lopaschuk 2006). Furthermore, activity of AMPK as a master regulator of metabolic has far reaching implications in cellular processes like autophagy and free fatty acid oxidation (Guo et al. 2013; Feinstein et al. 2005). AMPK is activated during acute exposure to high concentrations of rosiglitazone (200M rosiglitazone for 30 minutes) in skeletal muscle cells (Fryer et al. 2002; Skrobuk et al. 2009;Rabol et al. 2010) but whether its stimulation in the diabetic heart is adaptive or maladaptive is unclear. Alterations in AMPK activation have been linked to cardiac hypertrophy, myocardial ischemia and glycogen storage cardiomyopathy (Dolinsky & Dyck, 2006). In hypertrophic cardiac tissues, the AMP:ATP ratio is increased more than 4 fold over control hearts, which would lead to stimulation of AMPK (Tian et al. 2001 ; Dyck & Lopaschuk 2006 ). There have been many contradictory reports on the role of AMPK in heart disease (Dolinsky & Dyck, 2006; Fryer et al. 2002; Festuccia et al. 2009; Dyck & Lo paschuk, 2006). Further clarification of its direct role in cardiac hypertrophy is required to determine if its activation is advantageous or detrimental to cardiovascular health.
82 AMPK activation has also been linked to upregulation of autophagy (Cerquett i et al. 2011). Interestingly, Festuccia and coworkers (2009) have demonstrated that rosiglitazone induced cardiac hypertrophy was associated with increased turnover of myofibrillar proteins in rats administered 15mg/kg/day for 21 days. The increased turnover was linked to increased protein synthesis, orchestrated by the activation of the mTOR signaling pathway Similarly to AMPK, the mammalian target of rapamycin, mTOR, is a master regulator of metabolism and anabolic cellular processes. Activation of mTOR stimulates protein synthesis and causes inhibition of catabolic processes, such as autophagy. However, whether the observed activation of mTOR in the hypertrophic hearts was associated with an inhibition of autophagy was not reported. But one could specul ate that an inhibition of autophagy would cause the AMP:ATP ratio to shift, causing activation of AMPK, and thereby substantiating the cardiac hypertrophy. Ultimately, we conclude that decreased mitochondrial routine respiration and decreased maximum capac ity of the electron transport system is not caused by a decrease in COX IV or CS activity. Further analysis of each individual mitochondrial complex will be required for elucidation into direct site of rosiglitazone action. Others have reported alterations in the composition of the inner mitochondrial membrane caused by TZD treatment, possibly causing a derangement of the supercomplexes. Lastly, our autophagy results were inconclusive. Further replications of the autophagy experiments are required to clarif y the role of rosiglitazone on autophagy.
83 APPENDIX A CYTOCHROME c OXIDASE ASSAY FOR MICROPLATE READER Theory Tissue extract containing cytochrome c oxidase (COX) is added to the test solution containing fully reduced cytochrome c. The rate of cytochrome c oxidation by COX is measured over time as a reduction in absorbance at 550 nm. The reaction is carried out at 30 C. Technical note 1. Experience has shown that it is important to use sample homogenates in the same freeze thaw cycle number. The repeated free zing thawing seems to enhance COX activity, probably due to further breaking of membranes but this is just our current working hypothesis. However, it is not clear how often this freezethaw cycle could be pursued for maximal activity. 2. Pay attention to f ootnotes! 3. Check molecular weights (F W ) of the chemical and doublecheck the amount to weigh in for a desired molarity! Reagents1 1. 20 mM KCN (6.512 mg/5 mL dH2O) 2. 100 mM K Phosphate Buffer make up 0.1 M KH2PO4 (680.45 mg/50 mL dH2O) make up 0.1 M K2HPO4.3H2O (use stock K2HPO4, 870.9 mg/50 mL dH2O) 1 Check molecular weights (Fw) of the chemical and doublecheck the amount to weigh in for a desired molarity!
84 mix in equal proportions, pH to 7.0 3. 10 mM K Phosphate Buffer dilute 0.1 M KPO4 Buffer prepared above 1:10 with ddH2O 4. Extraction Buffer (0.1 M KH2PO4 Buffer with 2 mM EDTA (29.22 mg/50 mL); pH 7.2)2 5. Test Solution Dissolve 50 mg (entire content of horse heart cytochrome c bottle3) in 2.5 mL of 10 mM KPO4 buffer Store at 20C in aliquots (cover tubes with foil)4 Test solution preparation: (Make daily and wrap in foil) o Make up a small volume (1mL) of 10 mg/ml sodium dithionite5 in 10 mM KPO4 stock solution Needs to be accurate, otherwise reduction wont work correctly; make in a 1.5 ml tube (use within twenty minutes); o Add the following reagents in order (see table below). This is important, otherwise reduction wont work correctly. o Make enough test solution for the appropriate number of wells (2 wells per sample = duplicates). 2 Adhihetty et al. 2009 3 Cyt c is the chemical that we run out of m ost quickly! Pay attention to what we have and order accordingly for your experiment! Sigma C775250mg ($94) 4 The volume of the aliquots depends on the planned runs! Ex.: 50 uL 5 Na2S2O4: Fw 174.11; 10mg/mL = 57 mM
85 o NOTE: Test solution color should change from dark red/brown to bright pink, indicating cytochrome c was successfully reduced. o The reducti on quality can be assessed, if necessary, by measuring absorbance at 550 and 565 nm. A550/A565 = 10 20. If <10, then not reduced enough. Table A 1. Preparation of test solution. Test Solution (reduced cytochrome c, 2 mg/ml) 23 ml (enough for 82 microplate wells) Solution of Horse Cytochrome c dissolved in 10 mM KPO4 2.3 ml 10 mg/ml Dithionite in 10 mM KPO4 (make fresh) 92 ul dithionite solution dH2O 18.4 ml 100mM KPO4 2.3 ml Procedure Sample preparation 1. Place powdered6 muscle samples in liquid N2 storage. 2. Extraction Buffer to a 1.5 ml Eppendorf tube per sample on ice. (One tube per sample). a. Place tube on scale Tare then add sample as described in step #3. Weigh ~ 10 20 mg. 6 The powder homogenization of the samples can be performed on another day and the samples stored in liq. N2 or 80 C.
86 3. Add about 710 mg tissue to each tube reco rding exact tissue mass. Mix by tapping. 4. Add the volume of Extraction Buffer required to obtain a 20 fold dilution. a. 5. Mix in thermomixer @ 1400 rmp for 15 min @ Room Temperature (RT). 6. Make up Test Solution during this time and wrap in foil. 7. Following mixing, sonicate each tube 3 x 3 seconds with tube immersed as far as possible in ice water. Clean the probe between samples with 70% ethanol. 7 8. Make 80fold dilution of sample (This is a 1:4 dilution of your 1:20 sample: 70 ul of 20 fold sample + 210 ul of EB = 280 ul total). Keep samples on ice. 9. Centrifuge 80 fold for 2 min at 14,000xg a. Collect 80 fold supernatant and keep samples in ice. 10. Collect the supernatant into a new tube and discard the pellet. 11. At this point, set aside 1520uL of your sample for protein determination using the Bradford Assay. This may also serve as a stopping place, in that case store samples at 80C. Make sure ALL samples have been through the same amount of freezethaw cycles. 7 The desired intensity of sonication for this step of sample prep has nowhere been defined, unfortunately.
87 Plate reader preparation 12. Open Gen5 plate reader program. Select New Experiment and select Cytochrome c Oxidase Protocol Click the PRE HEATING setting, enter 30C and select ON. (Do not run assay until t emperature has reached 30C.) Reading should be in Kinetic mode @ 550 nm. Reduced cytochrome c preparation 13. for duplicates) of 96well microplate and incubate in plate reader at 30C for 10 minutes to stabilize the temperature and absorbance.8 14. In Gen5, select Procedure icon, the parameters should be set to a Kinetic Read. a. Select Kinetic for Reading Type. b. Select Absorbance for Reader and 550 nm for wavelength (dropdown menu). P roceed with prep and samples 15. Get a second, clean 96 well plate, pipette samples into empty wells (run duplicates of each sample). fold extract of muscle tissue Start of assay 8 In the plate reader templat e the reading of the samples is set such that the duplicates are in the same column. Each column will be read separately (see below for more explanation).
88 16. Remove microplate with Test Solution from the te mperature incubator (as long as it has been incubating for 10 minutes @ 30C). Place this plate beside the plate with the sample extracts in it. 17. Make sure the volume is equal in al l the pipette tips, and that no significant air bubbles have entered any of the tips. 18. Be sure to double check plate reader settings prior to reading plate. 19. Pipette the Test Solution into the wells with the sample extracts (the second plate). As soon as all the Test Solution has been expelled from the tips (do not wait for the second push from the multipipette), place the plate onto the tray of the plate reader and with the other hand on the mouse, press the OK button. (Speed at this point is paramount, as there is an unavoidable latency period between the time of pressing the OK button and the time of the first reading.) a. If running more than 4 samples at a time (4 samples take up 8 wells in the 96well microplate for a duplicate measurement of each sample), plate must be read multiple times. It is critical to read the absorbance as soon as reaction has been started. In order to read absorbance as soon as reaction has been started, the reaction must be started (= add tes t solution to samples) and read (= follow absorbance change for a set amount of time) separately for each column 9 of the plate. That is, add test solution to a column of 8 wells (A though H), read the plate according to the protocol; 9 columns are numbered 1 through 8; rows are named a through H
89 at the end the plate c omes out and is ready to have next columns reaction started. So, start the reaction in the next column of 8 wells. 20. KCN (COX inhibitor) to one of the wells to measure any absorbance changes that is independent of COX. This would be your negative control: COX is inhibited and even though the reaction should have been started, the specific catalysis of cytochrome c oxidation wont proceed since the enzyme is inhibited. If there is a absorbance change occurring, it is likely independent of COX activity and therefore unspecific (unless inhibition was unsuccessful; never rule this possibility out!). This activity then needs to be subtracted from the specific activity in your sample. Preparation of assay program in KC4: this might have to be done above at #10 21. To obtain the rate of change of absorbance over different time periods, select Options and enter the amount of time f or which you would like a rate of change of absorbance to be calculated. The graph, along with one rate (at whichever time interval is selected) for each sample can be printed on a single sheet of paper, and the results can be saved. Calculation COX activ = dEsee footnote 10 /min x total volume (ml) x 80 (dilution) 10 dE/min: absorbance change over time (slope)
90 18.5 ( mol/ml11) x sample vol (ml) x pathlength (cm)see legend for equation below Equation development: Generic formula for enzyme activity used here: [ ] = With: dE: change in absorbance with time [dE/min]; this is the slope of your absorbancereading curve (linear part) total sample dilution: sample prep dilution; in this assay: 80 : molar extinction coefficient [SI unit is: mM-1 cm-1; this is the same as: mol L-1 cm-1 -1 cm-1], -1 cm-1 L: pathlength of the optical path in the spectrophotometer (here: 0.552 cm for plate read in BioTek spectrophotometer plate reader) accordingly: = 80 . = . 11 : extinction coefficient [mM-1 cm-1 = mol L-1 cm-1 -1 cm-1-1 cm-1)
91 dE is the slope of the linear part of the absorbance change at the given wavelength that you will obtain from your plate readers readings over the set period of time. Look at the absorbance change and make sure the linear part is taken. There is the possibi lity that the later part of the reaction time becomes substrate limiting and the curve levels off.
92 APPENDIX B C ITRATE SYNTHASE ACTIVITY PROTOCOL Table B 1. Reaction r eagents Compound n ame Final Concentrations Supplier Comments 10% Triton X 100 Final Concentration: 0.25% Promega 10% Triton X 100 stored on shelf Acetyl CoA AcCoA Stock: [30 mM] in water Final Concentration in assay: 0.31mM Sigma (A2181 25 mg) Aliquot in 30 or 25 store @ 80C DTNB DTNB Stock: [1.01 mM ] in 1 M Tris HCl buffer, pH 8.1; 2mg DTNB + 5mL Tris HCl Final Concentration in assay: 0.1 mM Stored in chemical cabinet at RT Citrate Synthase (positive control) 1:500 dilution in 0.1 M Tris HCl buffer, pH 7.0 Sigma (C3260) Positive Control Citrate Synthase (Sigma, C3260); stored @ 48 C Oxaloacetate (make fresh daily) OAA Stock: [10mM] in 0.1 M trie thanolamineHCL (TEA) buffer pH 8.0; 6.6 mg OAA + 5 mL of 0.1 M TEA buffer Final Concentration in assay: 0.5mM Sigma (O4126) OAA powder stored @ 20C Make 0.1 M TEA buffer dilution fresh before use 1 Kuznetsov A.V., Lassnig B. Gnaiger E. (2010). Laboratory protocol: Citrate synthase. Mitochondrial marker enzyme. Mitochondrial Physiology Network. 08.14(110). Procedure Sample preparation12: NOTE: If running COX activity assay on the same samples, follow the sample preparation instructions in COX protocol and use 80 fold dilution of samples for citrate synthase, skip ahead to protein determination step. 1. Homogenize tissue using BioPulverizer m ethod (see tissue homogenization protocol) 2. Record the weight of tissue added to Cell Lytic MT buffer in 0.6 mL tube (about 5 12 Sample prepared for COX activity assay (80 fold dilution) can be used as well
93 3. Vortex tubes until mixture is homogenous. 4. Centrifuge for 10 minutes @ 12,000 x g and @ 4C 5. Collect supernatant in a new prechilled Eppendorf tube; keep samples on ice. 6. Run a Bradford13 assay to determine protein concentration of samples. Protein Determination via Bradford2 Assay: 1. Prepare BSA protein standards in CelLytic14 buffer and store in the fridge (4C). 2. Aliquot about 20 mL of Bradford reagent and let warm up to room temperature. 3. wells in a 96well microplate, run triplicates for all standards and samples. 4. Using a 15 of Bradford reagent per well. 5. Remove any bubbles using a pin and incubate at room temperature for at least 5 min. Positive control preparation: 1. Dilute Citrate Synthase enzyme (CS) in a 1:500 dilution using 0.1 M Tris HCl buffer, pH 7.0; a. 2. Keep diluted enzyme on ice until activity assay is performed. Plate reader preparation: (Synergy HT from BioTek; Gen5 operating software) 1. Ensure that program is set to read the appropriate wells on the plat e (procedure > plate layout). Set Temp to 30C. 2. Run kinetic program at 412 nm using the following procedure (Sigma Citrate Synthase Protocol): a. Shake: medium intensity for 2 s b. READ: 2 min total, 20 s intervals c. Plate out/Plate in: Add 10ul of 10mM Oxaloacetate to all wells using multichannel pipet d. Shake e. READ: 2 min total, 20 second intervals Start of assay: 1. Based on protein concentration of samples, calculate the amount of sample o each well. 13 Protein assay: check compatibility of the cell lysis buffer used for sample prep with the assay chemicals. Change assay accordingly, if necessary. 14 Or in the lysis buffer that was used to prepare the sample! 15
94 Table B 2. Sample preparation per reaction mix 2. Prepare reaction mixture for each sample and positive control. Follow the scheme shown in the table below. Using the Excel template sheet, enter the amount of sample needed (orange cell); the E xcel template will then calculate the amount of each component required for the reaction mix. Table B 3. Preparation of reaction mix. The table above shows several different columns: a) Reaction Mix : every component for correct final concentration w ithout sample, without 5the starter OAA, with only the total amount of water to bring volume the react ion starter OAA will be added later; b) Reaction Mix + 5% : it usually is advisable to make a bit more of a bulk (with x=190sample vol) will be added to each designated well, and ul timately c) Horse Reaction Mix for duplicate measurements : for this everything in the previous column is multiplied by two to provide enough reaction mix for a duplicate run of the same sam ple. Except the volume of water; for the correct volume of water the sample volume*2 has been subtracted (in the example above: (2*171.0765) (2*2.1) = 337.9). Same calculations were applied to the cow example and the CS standard/pos control mix. From thi s sample specific Reaction Mix 190 minus the sample volume will be pipetted into each
95 of two wells. Together with sample and 10 uL OAA the volume will add up to 200 uL total reaction volume. 3. Consider the usefulness of preparing a bulk mix of TritonX + AcCoA + DTNB, duplicate sample with 5% safety excess. Multiply this with the number of samples you plan on running. For individual sample reaction mix preparation take bine with the desired volume of water [(2*171.0765) (2*sample vol)]171.0765*2 4. Once reaction mix is prepared as described, follow the guidelines on the table for pipetting the reaction mix and sample for a total of 190 ul per well. 5. Read plate as described previously. Meanwhile, prepare to add 10 ul of Oxaloacetate per well when instructed to do so (use multichannel pipet). Immediately read plate a second time. a. This will follow the following reader protocol @ 30C. 1. Run kinetic program at 412 nm using the Ci trate Synthase Protocol 2. Shake: medium intensity for 2 s 3. READ: 2 min total, 20 s intervals ii. The second part of the kinetic reading starts when plate comes out: 1. Plate out/Plate in: Add 10ul of 10mM Oxaloacetate to all wells using multichannel pipet 2. Shake 3. RE AD: 2 min total, 20 second intervals
96 Activity Calculation: 1. Use excel sheet to calculate citrate synthase activity using the formula shown in the table below. Table B 4. Activity calculation of citrate synthase activity Citrate Synthase Activity Sample ddE [min 1] dF final umol/mL/min = IU/mL umol/min/mg = IU/mg prot CS 250 0.2134 250 1421.302217 163.3680709 CS 500 0.1101 500 1466.592072 168.5738013 Horse 0.1396 1 3.320621727 0.461197462 Cow 0.1404 1 2.010972692 0.467668068 0 0 IU/mL ((ddE*Vtot)/(L*13.6*1*Vs))*dF IU/mg protein ((ddE*Vtot)/(L*13.6*1*Vs*mg/mL prot))*dF
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102 BIOGRAPHICAL SKETCH Dana Joy Davis Schreffler (born Dana Joy Davis) was born to Frank E. Davis and Christina J. Davis in Phoenixville, Pennsylvania. Dana has a younger brother, Frank E. Davis Jr. In January 1991, her family moved to Palm Coast, Florida where she attended elementary through high school. She graduated from Flagler Palm Coast High School in May 2005 and in August 2005 she began her undergraduate studies at the Pennsylvania State University Hazelton campus. In 2007, Dana transferred to main Penn State campus in State College, PA, where she participated in undergraduate research in the Langkilde lab. Dana graduated from Penn State in 2009 with a Bachelor of Science in General Biology. In August 2009, Dana moved to back to Palm Coast, FL where she began work as a laboratory technician in Dr. Dirk Buchers neuroscience lab at the Whitney Labs of UF working on DNA mapping of the dopamine receptor in Homarus americanus At the Whitne y Labs, Dana also performed in silico research for the director, Dr. Peter Anderson, collecting an inventory of marine species in the Atlantic Ocean off the east coast of Florida. In May 2011, Dana began working as a laboratory technician for Dr. Stephanie Wohlgemuth in the Department of Animal Sciences. After working for Dr. Wohlgemuth for a few months, Dana began earning her Master of Science degree as Dr. Wohlgemuths first graduate student. In March 2012, Dana married Jacob D. Schreffler and changed her name to Dana Joy Davis Schreffler. In June 2013, Dana successfully defended her thesis entitled, Effects of the Dana intends to pursue a career as a forensic scientist for the state of Pennsylvania.