Sirtuin 3 Expression and Acetylation of Three Downstream Targets in Painted Turtle Liver and Brain During Anoxia

Home , NDUFA9, Sirtuin 3, Sirtuin 4

Sirtuin 3 Expression and Acetylation of Three Downstream

Targets in Painted Turtle Liver and Brain During Anoxia

By:

Gabriele Nandal, B.Sc (Hons)

A thesis submitted in conformity with the requirements for the degree of Master of Science

Graduate Department of Cell and Systems Biology

University of Toronto

Sirtuin 3 Expression and Acetylation of Three Downstream Targets in Painted Turtle Liver and Brain During Anoxia

Gabriele Nandal Master of Science Department of Cell and Systems Biology University of Toronto 2019

ABSTRACT

The western painted turtle is a champion anaerobe; its anoxia tolerance is heavily enabled by metabolic depression. While cellular metabolism is highly regulated by reversible-acetylation of mitochondrial proteins; this phenomenon is relatively unexamined in the anoxia-tolerant turtle. Sirtuin 3 (SIRT3) is a mitochondrial global deacetylase involved in regulating many metabolic and stress-resistance processes, suggesting a potential role in the control of hypometabolism in the turtle. During early anoxia in the brain and liver, mitochondrial acetylation and SIRT3 protein levels increased. Cyclophilin D levels, a direct target of

SIRT3, were also elevated during anoxia. Another target of SIRT3, NF-kB p65 exhibited increased deacetylation during anoxia and reoxygenation in the brain; while there was no difference in prevalence or activity of manganese superoxide dismutase, an antioxidant target. This thesis provides preliminary evidence that SIRT3 and mitochondrial protein acetylation may play a role in the regulation of anoxia in the painted turtle.

ACKNOWLEDGMENTS

The thesis cumulates all my work from the past two and a half years and although the entirety of all my work and efforts barely appear to evident, this report is what I have to show for it. Much of scientific research appears to involve extensive troubleshooting and I was not immune to this. I can confidently say Western blots work based on the gods, the weather, or some other mystical force, and not because you are performing the protocol perfectly or because you want them to. While I often said throughout my Masters, “It’s not science, if you’re not crying”, there are several essential people I would like to thank for helping to dissuade my tears.

I would first like to thank my supervisor Dr. Les Buck for accepting me into his lab and letting me not only forge my own project but giving me the free reign to design it. His guidance helped me through the different avenues of my project as it evolved by connecting me with valuable resources. Also, as importantly, I would to thank Dr. Jim Eubanks for welcoming me into his lab and making me feel like own of his own. Dr. Eubanks was more than just a committee member, he provided me with essential advice, support, and wisdom that enabled the completion of my project and my thesis. Without Dr. Eubanks, I surely would still be struggling to obtain any data. I would also like to thank my other committee member Dr. Sergey Plotnikov for the guidance and critiques that pushed my project forward.

I also would like to thank my work-wife, Elena Sidorova-Darmos, for not only her friendship but her unwavering assistance. From showing me how to perform Western blots, analyze data, and everything in between, I would absolutely not have completed my thesis without her. Elena’s constant support to answer late my night calls about what was wrong with my blots, where the multi-channel pipette was, or how to perform some statistical test, was invaluable. Next, I would like to thank my lab mate Farah Al-Dajani, for always being available to help me with dissections and whose company made it so much more pleasant. I couldn’t have asked for a better partner-in-turtle-crime. I also need to thank my last-minute thesis editor, Dene Ringuette, whose late-night conversations at Krembil made working at night a pleasure and whose presence brightened many evenings.

Lastly, I could not have managed without the love and support from Mark Lutley who accompanied and encouraged me every weekend and every evening to Krembil. Without the motivation and emotional support he provided, there would have been many more tears. Finally, to my parents, thank you for your love and the foundation you provided me with to achieve great things in life.

iii

TABLE OF CONTENTS ABSTRACT ...... ii

ACKNOWLEDGMENTS ...... iii

LIST OF FIGURES AND TABLES ...... vii

LIST OF APPENDICES ...... viii

LIST OF ABBREVIATIONS ...... ix

CHAPTER 1: INTRODUCTION ...... 1

1.01 General introduction ...... 1

1.02 Post-translational protein modification ...... 1

1.03 Sirtuins ...... 2

1.04 Introduction to the mitochondrion ...... 4

1.05 Mitochondrial ROS production and antioxidant defenses ...... 5

1.06 Mitochondrial signaling ...... 6

1.07 Introduction to SIRT3 ...... 8

1.08 SIRT3 regulation of antioxidants and ROS ...... 9

1.09 SIRT3 regulation of mitochondrial metabolism ...... 10

1.10 SIRT3 influences cell survival and the mitochondrial permeability transition pore ...... 14

1.11 SIRT3 and hypoxia ...... 16

1.12 Expression and regulation of SIRT3 ...... 16

1.13 SIRTs and hibernation ...... 18

1.14 The vitality of oxygen ...... 20

1.15 Surviving anoxia ...... 21

1.16 Anoxic ATP production ...... 22 iv

1.17 Avoiding acidosis ...... 24

1.18 Metabolic depression ...... 26

1.19 Molecular changes during anoxia ...... 29

1.20 Anoxia intolerance ...... 31

1.21 Reoxygenation ...... 32

1.22 SIRT3 and the painted turtle ...... 34

1.23 Objectives and hypotheses ...... 34

CHAPTER 2: MATERIALS AND METHODS ...... 38

2.1 Animals ...... 38

2.2 Experimental protocol ...... 38

2.3 Tissue homogenization ...... 38

2.4 Mitochondrial isolation ...... 39

2.5 SIRT3 activity assay ...... 40

2.6 SOD activity assay ...... 41

2.7 Antibodies and western blotting ...... 42

2.8 Statistical Analysis ...... 43

CHAPTER 3: RESULTS ...... 44

3.1 SIRT3 expression ...... 44

3.2 NF-kB p65 expression and acetylated expression ...... 46

3.3 Cyclophilin D expression ...... 48

3.4 MnSOD expression ...... 50

3.5 Mitochondrial lysine acetylation ...... 52

CHAPTER 4: DISCUSSION ...... 53

4.1 SIRT3 regulation during anoxia ...... 53 v

4.2 Deacetylation of NF-kB p65 during anoxia ...... 55

4.3 CypD expression increases and could open the MPTP ...... 57

4.4 MnSOD during anoxia and reoxygenation ...... 60

4.5 Mitochondrial acetylation ...... 61

CHAPTER 5: CONCLUSION AND FUTURE REMARKS ...... 62

REFERENCES ...... 65

APPENDIX ...... 88

LIST OF FIGURES AND TABLES

Figure 1.1 Deacetylase activity by sirtuins ...... 3

Figure 1.2 Targets of Sirtuin 3 ...... 10

Figure 1.3 Alternative energy utilization regulated by SIRT3...... 13

Figure 1.4 SIRT3 influences the mitochondrial permeability transition pore...... 15

Figure 1.5 Anoxic survival is temperature-dependent...... 22

Figure 1.6 Blood pH and plasma lactate concentrations during anoxic submergence and

recovery at 3, 10, 15, and 20°C...... 25

Figure 1.7 Allocation of energy during normoxia vs. anoxia in turtle hepatocytes...... 27

Figure 1.8 Na+/K+ ATPase activity decreases during anoxia...... 28

Table 2.1: List of primary antibodies ...... 43

Figure 3.1 SIRT3 protein expression compared to control ...... 45

Figure 3.2 NF-kB p65 and NF-kB p65 K310 protein expression compared to control ...... 47

Figure 3.3 CypD protein expression compared to control ...... 49

Figure 3.4 MnSOD protein expression compared to control ...... 51

Figure 3.5 Mitochondrial acetylated lysine residues after 2hrs of anoxia...... 52

Figure 4.1 The proposed model of SIRT3 in the brain of anoxic freshwater turtle ...... 59

vii

LIST OF APPENDICES

APPENDIX A: Dose Response Blot ...... 88

APPENDIX B: Immunoblotting with heart and pectoralis tissues ...... 89

APPENDIX C: SIRT3 Activity ...... 90

APPENDIX D: SOD activity assay ...... 92

APPENDIX E: Coomassie Normalization ...... 93

APPENDIX F: Bioinformatics for Immunoblotting ...... 95

viii

LIST OF ABBREVIATIONS acetyl-CoA acetyl coenzyme A

ACS2 acetyl Co-A synthase 2

ADP adenosine diphosphate

AMP adenosine monophosphate

ANT adenine nucleotide translocator

ATP adenosine triphosphate cAMP cyclic adenosine monophosphate

CREB cAMP response element-binding protein

CypD cyclophilin D

DNA deoxyribonucleic acid

ETC electron transport chain

FAD flavin adenine dinucleotide

FADH2 flavin adenine dinucleotide

GABA gamma-aminobutyric acid

GDH glutamate dehydrogenase

GSH-Px glutathione peroxidase

HIF1α hypoxia-inducible factor 1α

HSP heat shock protein

IDH2 isocitrate dehydrogenase 2

LCAD long-chain acyl CoA dehydrogenase

LDH lactate dehydrogenase

MnSOD manganese superoxide dismutase ix

MPTP mitochondrial permeability pore

NAD+ nicotinamide adenine dinucleotide

NADH nicotinamide adenine dinucleotide

NAM nicotinamide

NF-kB nuclear factor kappa-light-chain-enhancer of activated B cells

NMDA N-methyl-D-aspartate

O2 oxygen

OPA1 optic atrophy 1

OTC ornithine transcarbamylase

PDH pyruvate dehydrogenase

PGC1-α peroxisome proliferator-activated receptor gamma coactivator 1-alpha

PKA protein kinase A

REDOX oxidation-reduction reaction

ROS reactive oxygen species

RT-PCR Reverse transcription polymerase chain reaction

SDH succinate dehydrogenase

SDS sodium dodecyl sulfate

SIRT silent information regulator (sirtuin)

SOD superoxide dismutase

TOM20 translocase of outer membrane 20

VDAC voltage-dependent anion channels

VLCAD very long-chain acyl-CoA dehydrogenase

CHAPTER 1: INTRODUCTION

1.01 General introduction

The western painted turtle is one of the most anoxia-tolerant vertebrates, meaning they have the ability to survive without oxygen for days to months. This adaptation likely arose from the seasonal ice cover it experiences in the northern part of its range. For the majority of vertebrates, including humans, an anoxic period of only a few minutes is a severe stress and can even be lethal. A family of proteins that play an important role in the response to stress, such as anoxic stress, are the sirtuins; they are deacetylases and/or ADP- ribosyltransferases. Members of this family reside in different cellular locales; the nucleus, cytoplasm, and mitochondria. Sirtuin 3 is the major deacetylase in the mitochondrial matrix and may play an important role in the anoxia-tolerance of the painted turtle. This is the focus of my thesis.

1.02 Post-translational protein modification

Post-translational modification is a form of covalent modification following protein synthesis that involves removing, adding, or altering a functional group on a protein. This changes its activity, protein-protein interactions, stability, and/or its subcellular localization.

Phosphorylation is the most prevalent form of post-translational modification, which is the addition of a phosphoryl group, however, there are many such as methylation, glycosylation, and ubiquitinylation (Khoury, Baliban, & Floudas, 2011). Another common post- translational modification is reversible acetylation, which transfers the acetyl group from acetyl coenzyme A (acetyl-CoA) to the amino (NH2-) group of a lysine residue. The addition of an acetyl group is regulated by histone acetyltransferases while the removal is regulated

by histone deacetylases. These enzymes often target histone proteins where acetylation or deacetylation alters the wrapping of deoxyribonucleic acid (DNA) around histone cores thereby affecting the binding ability of transcription factors to the DNA and thus transcription. Histone deacetylases tend to be associated with transcriptional silencing by increasing the tightness of DNA to histones; this condensed form limits access of transcription factors (Delcuve, Rastegar, & Davie, 2009). Histone acetylates and deacetylases control gene expression in this way but have other targets in addition to histones, such as transcription regulators, signaling factors, and metabolism (Freiman & Tjian, 2003; Lu,

Scott, Webster, & Sack, 2009).

1.03 Sirtuins

A focus of more recent research has been protein acetylation and deacetylation of non-histone proteins as a viable form of post-translation protein modification to regulate metabolic pathways. A family of protein deacetylases called silent information regulators, also referred to as sirtuins (SIRTs) are of particular interest due to their roles in regulating glucose metabolism, lipid metabolism, and oxidative stress responses in mammals (Schwer

& Verdin, 2008). Four genes involved in transcription silencing were first identified in budding yeast (Saccharomyces cerevisiae) (Ivy, Klar, & Hicks, 1986). The SIRT family consists of SIRT1-7 and are nicotinamide adenine dinucleotide (NAD+) -dependent deacetylases, meaning they couple the hydrolysis of acetyl-lysine residues with that of

NAD+, which is in turn an inhibitor of SIRTs (Fig. 1.1). Due to the dependence of SIRTs on

NAD+, their activity is directly related to the cell’s energy status via the amount of NADH and NAD+. SIRTs are distinct from other histone deacetylases and have been found to

possess other enzymatic activities (Mathias et al., 2014; Nishida et al., 2015; Tan et al.,

2014).

Structurally, each member of the SIRT family possesses a conserved catalytic core domain and NAD+ binding motif but have distinct N- and C-terminus regions that play a role in their unique localizations and functions (Landry et al., 2000; Marmorstein, 2004; Min,

Landry, Sternglanz, & Xu, 2001). SIRTs differ in their localization; SIRT1, SIRT6, and

SIRT7 are predominantly localized to the nucleus, while SIRT3-5 are primarily mitochondrial and SIRT2 is mostly cytoplasmic (Michishita, Park, Burneskis, & Barrett,

2005).

Figure 1.1 Deacetylase activity by sirtuins

Sirtuins act by deacetylating their target proteins. This reaction is coupled with the hydrolysis of NAD+ into nicotinamide (NAM) and ADP-ribose. The acetyl group removed from the target protein is then transferred onto the ADP-ribose, forming 2’-O-acetyl-ADP- ribose.

1.04 Introduction to the mitochondrion

Mitochondria play a role in a variety of cell functions like ATP production (Devin &

Rigoulet, 2007), reactive oxygen species (ROS) formation (Murphy, 2009), anti-oxidant defenses (Kalogeris, Bao, & Korthuis, 2014), energy metabolism (Vakifahmetoglu-Norberg,

Ouchida, & Norberg, 2017), apoptosis (Vakifahmetoglu-Norberg et al., 2017), fatty acid oxidation (Bartlett & Eaton, 2004), calcium homeostasis (Vandecasteele, Szabadkai, &

Rizzuto, 2002), and cell signaling (Brookes, Levonen, Shiva, Sarti, & Darley-Usmar, 2002;

Finkel, 2011; Tait & Green, 2012).

Mitochondria have an inner and outer membrane consisting of a phospholipid bilayer that separates the intermembrane space and mitochondrial matrix. The outer membrane is embedded with protein pores called porins allowing it to be permeable while the inner membrane is mostly impermeable. The matrix contains a vast number of enzymes involved with oxidative phosphorylation as well as mitochondrial DNA. However, more than 99% of mitochondrial proteins are encoded in the nuclear DNA which are translated in the cytoplasm and then translocated to the mitochondria (Kang, Fielden, & Stojanovski, 2018). The electron transport chain (ETC) is embedded in the inner membrane and in combination with oxidative phosphorylation, consists of 5 complexes.

ATP is the primary source of cellular energy and the brain relies mainly on glucose availability for ATP, but fatty acids can also be utilized (Freitas, Ferreira, Trevenzoli,

Oliveira, & De Melo Reis, 2017; Lenard & Berthoud, 2008). ATP can be produced through a variety of methods; the most efficient being glycolysis and oxidative phosphorylation.

Glycolysis involves the breakdown of glucose into the intermediate acetyl coenzyme A

(acetyl-CoA); this intermediate can also be derived through β-oxidation of fatty acids (L. Shi 4

& Tu, 2015). Acetyl-CoA enters the citric acid cycle where it is oxidized and the electrons

+ are used to reduce flavin adenine dinucleotide (FAD) and NAD to FADH2 and NADH, respectively. NADH and FADH2 pass their electrons through the ETC via a series of oxidation-reduction reactions (REDOX). Electrons from NADH or FADH2 enter either

Complex I or Complex II, respectively, to reduce the electron carrier CoQ to ubiquinol.

Finally, electrons are transferred to Complex III, cytochrome c, Complex IV and then to a terminal electron acceptor, oxygen (O2), resulting in water (H2O) as a byproduct. During the electron transfer, complexes I, II, and IV pump protons (H+) into the intermembrane space creating an electrochemical gradient with a high [H+] in the intermembrane space and a more negatively charged matrix. Complex V (also called F1Fo ATP synthase) utilizes the electrochemical gradient by allowing protons to flow back into the mitochondrial matrix while phosphorylating ADP into ATP.

1.05 Mitochondrial ROS production and antioxidant defenses

While mitochondria are the predominant source of ATP, they are also a source of

ROS production. Mitochondria generate ROS as a result of REDOX reactions, which are molecules derived from O2 that contain one or more unpaired electrons. The majority of ROS production occurs at complex I and III of the ETC, forming when electrons escape and react with O2. The incomplete reduction of O2 to superoxide or hydrogen peroxide (H2O2) produces highly reactive hydroxyl radicals that can cause oxidative stress by oxidizing lipids, proteins, and nucleic acids. Uncontrolled production of ROS can cause damage in the form of peroxidation of fatty acids in plasma membranes and organelles, oxidation of enzymes, depolymerization of polysaccharides, and single and double stranded DNA breaks (Blokhina,

Virolainen, & Fagerstedt, 2003; Tribble, Aw, & Jones, 1987).

Antioxidants are enzymes that scavenge ROS. MnSOD, GSH-Px, catalase and thioredoxins combat the oxidative damage that ROS can incur. The addition of one electron to O2 results in a superoxide anion which can be detoxified by manganese superoxide dismutase (MnSOD) to H2O2 and then converted to H2O by glutathione peroxidase (GSH-

Px) (Morris et al., 2017). In the presence of reduced transition metals, H2O2 can further be converted to hydroxyl radicals (Birben, Sahiner, Sackesen, Erzurum, & Kalayci, 2012).

While antioxidants convert ROS into less reactive species, oxidative stress or damage occurs when the rate of ROS production exceeds the capacity of antioxidants to detoxify ROS.

Another method of minimizing ROS levels and damage is to slow the rate of oxidative phosphorylation. Inducing a small proton leak through the inner mitochondrial membrane converts mitochondrial ROS-producing sites to a more oxidized state thus lowering the probability of ROS production and providing oxidative stress protection. In high concentrations, ROS can cause oxidative damage, however in low concentrations, it can act as a signaling molecule.

1.06 Mitochondrial signaling

One of the roles played by mitochondria is acting as a signaling organelle through the release of ROS and metabolites in order to regulate a variety of cellular processes (Finkel,

2011). For example, cysteine residues can be oxidized by ROS and activated. This residue in protein tyrosine phosphatases is highly susceptible to oxidation, where upon interaction with

H2O2 becomes activated (Finkel, 2011). Specific cysteine residue oxidation is thought to underlie most ROS-dependent signaling and is also the basis of ROS-mediated calcium

(Ca2+) signaling. Whereby oxidation of Ca2+ channels and pumps results in events like membrane excitability, neurotransmitter release, and gene expression (Clapham, 2007; Yan,

Wei, Zhang, Cheng, & Liu, 2006).

Mitochondria are also important in regulating cell death, where ROS toxicity is often involved in signaling apoptosis or necrosis. Apoptosis is programmed cell death initiated by signaling molecules or stressors. A multi-protein complex called the mitochondrial permeability transition pore (MPTP) is triggered by cellular disturbances like ROS and increased cytosolic Ca2+ (Du & Yan, 2010). Permeabilization of the outer mitochondrial membrane releases triggers for caspase-dependent and independent apoptotic pathways. In contrast to apoptosis, necrosis is unprogrammed cell death as a result of rapid ATP depletion and ion deregulation causing mitochondrial and cellular swelling and/or lysis (Golstein &

Kroemer, 2007).

The MPTP has been proposed to consist of several proteins: the adenine nucleotide translocase (ANT), the voltage-dependent anion channel (VDAC), a dimer of F0F1 ATP synthase, and cyclophilin D (CypD), where CypD plays a role in pore opening (Bernardi,

2013; Leung & Halestrap, 2008). Upon binding of the ANT to CypD, a tunnel-like structure is formed between the mitochondrial matrix and cytosol (Lemasters, Theruvath, Zhong, &

Nieminen, 2009). While there have been conflicting reports showing that pore formation can occur without ANT, CypD, or VDAC association (Baines, 2007; Basso et al., 2005;

Kokoszka et al., 2004), this is the most accepted model. CypD is also able to bind to ATP synthase, regulating its activity, and the ATP synthase has been further implicated in formation of the MPTP (Bernardi, 2013; Bonora et al., 2013; Chinopoulos et al., 2011;

Giorgio et al., 2009). Overall, mitochondria are essential in ATP and ROS production, as 7

well as regulation of cell death via the MPTP. Due to the important functions performed by the mitochondria, the dominant mitochondrial deacetylase, SIRT3, is of particular interest.

1.07 Introduction to SIRT3

SIRT3 is the primary deacetylase in the mitochondria. While SIRT4 and SIRT5 are also localized to the mitochondria, they appear effect the mitochondrial acetylome significantly less than SIRT3. SIRT3-KO mice show globally elevated mitochondrial proteome acetylation in the liver, while SIRT4- or SIRT5-KO mice display no changes in mitochondrial acetylation (Lombard et al., 2007). SIRT3 is preferentially expressed in tissues with a high oxidative capacity like the brain, cardiac, hepatic, brown adipose tissue, and skeletal muscle, while lower levels are found in tissues with lower metabolic demands such as white adipose tissue, lungs, small intestine, and pancreas (Lombard et al., 2007; T. Shi,

Wang, Stieren, & Tong, 2005). SIRT3 is a nuclear-encoded gene, that upon translation, is targeted to the mitochondria (P. Onyango, Celic, McCaffery, Boeke, & Feinberg, 2002).

Once in the mitochondrial matrix, the amino-terminus is cleaved by mitochondrial processing peptidase and mitochondrial intermediate peptidase (Kobayashi et al., 2017), upon which, becomes enzymatically active.

SIRT3 has been linked with longevity and as a neuroprotective factor (Bellizzi et al.,

2005; Guarente, 2011; Outeiro, Marques, & Kazantsev, 2008; Rose et al., 2003; Srivastava &

Haigis, 2011). SIRT 3 also influences metabolic processes like the citric acid cycle, ETC, b- oxidation, ketogenesis, acetate metabolism, brown adipose tissue thermogenesis (Giralt &

Villarroya, 2012; T. Shi et al., 2005).

1.08 SIRT3 regulation of antioxidants and ROS

SIRT3 can protect against oxidative damage, as exhibited by SIRT3-deficient cells and tissues that possess high oxidative stress (Bause & Haigis, 2013; Eric L. Bell &

Guarente, 2011). This deacetylase possesses many known targets (Fig. 1.2), many of which

− are involved in ROS scavenging. One target of SIRT3 is MnSOD that converts O2 to H2O2.

Deacetylation by SIRT3 increases its catalytic activity, thereby decreasing mitochondrial superoxide (Tao et al., 2010). SIRT3 also regulates MnSOD expression via deacetylation of

FOXO3 which promotes this transcription factor’s nuclear localization and binding to the

MnSOD promoter (Jacobs et al., 2008; Wei et al., 2015). SIRT3-mediated deacetylation of

FOXO3 promotes the upregulation of other antioxidant enzymes like peroxiredoxin 3,5, and thioredoxin 2 (Tseng, Wu, Shieh, & Wang, 2014).

SIRT3 also influences mitochondrial membrane potential and ROS genesis, in addition to antioxidants. Overexpression has been shown to lower mitochondrial membrane potential and ROS levels, while increasing cellular respiration (Bell & Guarente, 2011; T.

Shi et al., 2005). It has been suggested that SIRT3 increases expression and activates the transcriptional regulator peroxisome proliferator-activated receptor gamma coactivator 1- alpha (PGC1-α) expression, which regulates genes involved in energy metabolism and mitochondrial biogenesis expression (Q. Wang et al., 2015). PGC1-α enables an increase in mitochondrial uncoupling protein 1, which decreases mitochondrial membrane potential by allowing proton flux back through the inner mitochondrial membrane (Azzu & Brand, 2010).

This dissipation of membrane potential lessens proton leak during oxidative phosphorylation and thereby mitigates ROS production as a result (Fink, Reszka, Herlein, Mathahs, & Sivitz,

2005). 9

Figure 1.2 Targets of Sirtuin 3

SIRT3 directly deacetylates and inactivates targets in red, all of which are involved in cell survival and stress responses. While deacetylation of targets in green result in activation, some are involved in energy production pathways and others are involved in stress responses. These are just some of the known targets of SIRT3, more exist and are under investigation.

1.09 SIRT3 regulation of mitochondrial metabolism

SIRT 3 regulates ATP production in the mitochondria through the deacetylation of several factors involved in its production. SIRT3-/- mice exhibit abnormally reduced ATP levels in tissues that normally exhibit high levels of ATP and tend to express elevated levels of SIRT3, including the heart, liver, kidney, and skeletal muscle (Ahn et al., 2008). SIRT3 likely regulates ATP production, in part, through complex I of the ETC considering that several components of complex I are hyperacetylated in SIRT3-/-, SIRT3 can physically interact with NDUFA9, (a subunit of complex I), and incubation of exogenous SIRT3 with mitochondria increases complex I activity (Ahn et al., 2008). Furthermore, SIRT3 has been directly shown to interact with two subunits of complex II, succinate dehydrogenase (SDH) 10

A and SDHB (Cimen et al., 2009; Finley, Haas, et al., 2011); SDH participates in both oxidative phosphorylation as well as the citric acid cycle. SIRT3-/- resulted in SDHA hyperacetylation and decreased SDH activity (Finley, Haas, et al., 2011). Subunits a and β of

ATP synthase are also deacetylated and activated by SIRT3 (Rahman et al., 2014).

SIRT3 not only regulates components of the ETC but also several factors involved in energy utilization, such as glutamate dehydrogenase (GDH), isocitrate dehydrogenase 2

(IDH2) and acetyl Co-A synthase 2 (ACS2). GDH is deacetylated and activated by SIRT 3; this metabolic enzyme is involved with producing a-ketoglutarate that is used to produce

ATP via the citric acid cycle and NADH (Lombard et al., 2007). Another enzyme of the citric acid cycle, isocitrate dehydrogenase 2 (IDH2), is also activated by SIRT 3; when deacetylated it promotes the regeneration of antioxidants and increases citric acid cycle activity (Someya et al., 2010). Acetyl Co-A synthase 2 (ACS2), a mitochondrial enzyme that converts acetate into acetyl-CoA, and was the first substrate of SIRT 3 identified (Schwer &

Verdin, 2008). SIRT 3 deacetylates and activates ACS2, providing more acetyl-CoA to feed into the citric acid cycle. ACS2 is particularly important during periods of limited glucose availability where the liver can release stored acetate for use in other tissues (Yamashita,

Kaneyuki, & Tagawa, 2001).

The metabolically-demanding brain must maintain a constant energy source for neuron function and viability (Mergenthaler, Lindauer, Dienel, & Meisel, 2013). Its main energy source is glucose. When glucose becomes limited, the brain must utilize energy stores like glycogen, however this is only viable for a short period of time (Waitt, Reed, Ransom, &

Brown, 2017). An alternative energy source when glycogen is depleted, are fatty acid reserves that generate acetyl-CoA via hepatic β-oxidation which are then converted into 11

ketone bodies (White & Venkatesh, 2011). Ketones are transported to the metabolically- demanding tissue where they are converted back into acetyl-CoA. In the liver it has been demonstrated that SIRT3 deacetylates and activates two key enzymes involved in fatty acid

β-oxidation pathway initiation: long-chain acyl CoA dehydrogenase (LCAD) (Bharathi et al.,

2013) and very long-chain acyl-CoA dehydrogenase (VLCAD) (Fig. 1.3) (Hallows et al.,

2011). Furthermore, SIRT3 has been shown to regulate 3-hydroxy-3-methylglutaryl-CoA synthase (HMGCS2), which mediates ketone body production in the liver (Hirschey et al.,

2010; Shimazu et al., 2010). During a 24 hour fasted phase, extra-hepatic tissues rely on fatty acids and ketone bodies produced by β-oxidation in the liver (Bauer et al., 2004). Thus

SIRT3 mediates crosstalk between tissues to enable nutrient shifts such as ketone production in the liver and usage in the brain (Fig. 1.3) (Dittenhafer-Reed et al., 2015).

Ketone bodies are used as an alternative fuel source for the brain when glucose is limited, as is lactate. Lactate can be converted into pyruvate by lactate dehydrogenase (LDH) which is then converted into acetyl-CoA by pyruvate dehydrogenase (PDH). PDH is the link between glycolysis and the citric acid cycle, transforming pyruvate into acetyl-CoA to generate ATP. SIRT3 has been shown to deacetylate LDH and PDH thereby increasing their activity (Fig. 1.3) (Cui et al., 2015; Ozden et al., 2014).

Figure 1.3 Alternative energy utilization regulated by SIRT3.

The brain is metabolically demanding organ. During low glucose availability, SIRT3 adipose tissue provides fatty acids to the liver where SIRT3 can deacetylate and activate long-chain acyl CoA dehydrogenase (LCAD) and very long-chain acyl-CoA dehydrogenase (VLCAD) as part of β-oxidation generating acetyl-CoA. SIRT3 also deacetylates and activates 3-hydroxy-3-methylglutaryl-CoA synthase (HMGCS2), which is involved in the production of ketone bodies from acetyl-CoA. Ketone bodies then travel to the brain where they are converted back into acetyl-CoA. ACS2 is also a target of SIRT3 which generates acetyl-CoA from free acetate. Lactate from peripheral tissues is another energy source which can be converted into pyruvate and acetyl-CoA by SIRT3- mediated activation of lactate dehydrogenase (LDH) and pyruvate dehydrogenase (PDH), respectively.

1.10 SIRT3 influences cell survival and the mitochondrial permeability transition pore

Mitochondria play an important role in apoptosis, however little is known about the role of mitochondrial SIRTs in apoptotic pathways. The formation and opening of the MPTP is key for the release of pro-apoptosis factors that activate cell death, and SIRT3 plays a role in its regulation (Halestrap & Pasdois, 2009). SIRT3 appears to control the MPTP; a loss of

SIRT3 results in activation of the MPTP in response to Ca2+ increases and SIRT3 knockouts are susceptible to Ca2+-induced mitochondrial swelling but could be rescued with an MPTP inhibitor (Hafner et al., 2010). SIRT3 directly deacetylates CypD at K166 and upon deacetylation, inhibits MPTP opening by decreasing the binding of CypD to ANT (Fig. 1.4)

(Hafner et al., 2010). Furthermore, deacetylation of CypD by SIRT3 initiates the dissociation of hexokinase II from the mitochondria, which is induces oxidative phosphorylation stimulation (Fig. 1.4) (Shulga, Wilson-Smith, & Pastorino, 2010).

SIRT 3 further controls cell survival through the deacetylation of Ku70; this promotes the sequestration of the pro-apoptotic factor protein Bax to Ku70 thereby preventing Bax- mediated cell damage (Sundaresan, Samant, Pillai, Rajamohan, & Gupta, 2008). When Bax is not sequestered it is able to translocate to the mitochondria and induce opening of the

MPTP as well as apoptosis (Green & Reed, 1998).

SIRT3 also protects cells from necrosis by reducing the expression of high mobility group box 1 (Pellegrini et al., 2012), which is exclusively released by necrotic cells making it a reliable measure of necrosis (Krysko, Berghe, Parthoens, D’Herde, & Vandenabeele,

2008). Another target of SIRT3 is optic atrophy 1 (OPA1) where its deacetylation activates this protein by elevating its GTPase activity and blocks mitochondrial-mediated apoptosis

(Samant et al., 2014). OPA1 is involved in maintaining the crista structures, promoting 14

mitochondrial fusion, cell protection from death stimuli (Olichon et al., 2003; Samant et al.,

2014). Similarly, SIRT3 is able to deacetylate mitochondrial p53 and promote survival and prevent cell arrest (Li et al., 2010).

Figure 1.4 SIRT3 influences the mitochondrial permeability transition pore.

The mitochondrial permeability transition pore (MPTP) is formed by the voltage- dependent calcium channel (VDAC), adenine nucleotide translocase (ANT) and cyclophilin D (CypD). SIRT3 knockouts are susceptible to opening of the MPTP and releases of Ca2+ into the cytosol, that induces apoptosis. Deacetylation of CypD by sirtuin 3 (SIRT3) inhibits opening of the MPTP and promotes the dissociation of hexokinase II (HKII) from VDAC, which in turn, promotes oxidative phosphorylation.

1.11 SIRT3 and hypoxia

SIRT3 has been shown to mediate resistance to various forms of cellular stress, including hypoxia (Pellegrini et al., 2012; Tseng et al., 2014; Q. Wang et al., 2015).

Increased expression of SIRT3 protects cells from hypoxia by inhibiting induction of the

MPTP, loss of membrane potential, and ROS accumulation (Pellegrini et al., 2012). In part, this is achieved by inhibiting HKII release, reducing the probability of the MPTP from opening, preventing mitochondrial Bax accumulation, and therefore apoptosis (Pellegrini et al., 2012).

Studies have implicated SIRT3 in the stabilization of hypoxia-inducible factor 1α

(HIF1α) which controls the transcription of genes involved in hypoxia responses. During normoxia, proline hydroxylation and lysine acetylation of the oxygen-dependent domain trigger its degradation (Jeong et al., 2002; Lee, Bae, Jeong, Kim, & Kim, 2004). However, reduced ROS production under hypoxic conditions inhibit HIF1α degradation allowing it to become stabilized and active (Giralt & Villarroya, 2012). One study demonstrated that

SIRT3 overexpression diminishes the activation of HIF1α and thereby decreases its transcriptional activity, while an absence of SIRT3 results in an increase in HIF1α stability

(Bell, Emerling, Ricoult, & Guarente, 2011). Another study, demonstrated similar findings where SIRT3-/- exhibited hyperactivation of HIF1α target genes which may be due to elevated levels of ROS allowing HIF1α stabilization (Finley, Carracedo, et al., 2011).

1.12 Expression and regulation of SIRT3

Hypoxia, is known to upregulate SIRT3 mRNA and protein levels , as are other forms of cellular stress (Tseng et al., 2014), including during caloric restriction and exercise

(Brandauer et al., 2015; Cheng et al., 2016; Palacios et al., 2009; T. Shi et al., 2005;

Tauriainen et al., 2011). Some data shows an increase in AMP-kinase activity upon exercise and/or caloric restriction due to elevated AMP:ATP ratios (Brandauer et al., 2015; Cantó &

Auwerx, 2011). One of AMP-kinase’s many targets is the transcriptional regulator cAMP response element-binding protein (CREB) which is known to activate PGC1-α (Palacios et al., 2009; Thomson et al., 2008). PGC1-α activates many mitochondrial and anti-oxidative defense factors as well as mitochondrial biogenesis (Onyango et al., 2010). Exercise and caloric withdrawal induce PGC1-α, hence this protein might play a plausible role in the upregulation of SIRT3 during these conditions (Buler, Andersson, & Hakkola, 2016). PGC1-

α does not directly bind to the SIRT3 promoter, instead it activates estrogen-related receptor-

α which does so (Giralt et al., 2011; Kong et al., 2010).

Energetic stress conditions involve pathways like the cAMP-PKA-CREB pathway, where cAMP can upregulate PGC1-α and in turn, mediate SIRT3 expression (Buler et al.,

2016). In addition cAMP can also regulate SIRT3 post-translationally as it has been shown to bind to the NAD+ -binding pocket of SIRT3 increasing its stabilization and catalytic activity (Z. Wang et al., 2015). Furthermore, SIRT3 activity is regulated by the metabolic state of the cell (Cheng et al., 2016; Palacios et al., 2009; L. Shi & Tu, 2015; Tauriainen et al., 2011). SIRTs are classified as NAD+-dependent deacetylases and NAD+ /NADH ratios tend to increase during caloric restriction and exercise thereby increasing the activity of

SIRT3 (Hipkiss, 2008). In addition, increases in ROS has been shown to promote SIRT3 expression (Y. Chen et al., 2011; Weir et al., 2012).

1.13 SIRTs and hibernation

Previous studies have implicated SIRTs in mammalian hibernation, regulating metabolic changes in terms of lipid catabolism, oxidative stress responses, and transcriptional suppression (Rouble & Storey, 2015). Starvation, cold temperatures, and low metabolic rates characterize mammalian hibernation (or torpor). Levels of SIRTs protein and activity were elevated in a tissue-specific manner in 13-lined ground squirrels; most notably,

SIRT3 increased during early and late torpor in liver and white adipose tissue (Rouble &

Storey, 2015). The authors implicate SIRT3 in regulating metabolic and oxidative stress reduction in these tissues (Rouble & Storey, 2015). Antioxidative responses are known to be active in torpid tissues in order to protect against cellular damage when ROS production surges upon arousal (Allan & Storey, 2012; Morin & Storey, 2007; Rouble & Storey, 2015).

Correspondingly, the ground squirrel exhibited decreased amounts of acetylated SIRT3 target, MnSOD, in liver and white adipose tissue during late torpor suggesting increased activity during this time (Rouble & Storey, 2015). Authors also found deacetylation of the p65 subunit of nuclear factor kappa-light-chain-enhancer of activated B cells (NF-kB) increased in the liver during late torpor, corresponding to suppression of NF-kB transcriptional activity (Rouble & Storey, 2015).

NF-kB is a transcription factor that is involved in the regulation of cellular stress responses. NF-kB target genes are involved in oxidative stress responses, apoptosis, transcription factors, and other essential cellular functions (Kaltschmidt et al., 2000;

Rothgiesser, Erener, Waibel, Lüscher, & Hottiger, 2010; Yeung et al., 2004).NF-kB is formed by homo- or hetero-dimers composed of the subunits p50, p52, p65 (also known as

RelA) , RelB, and c-Rel (Kumar, Takada, Boriek, & Aggarwal, 2004). It exists commonly as p50 and p65, sequestered in the cytoplasm in its inactive form associated with an inhibitory protein (IkB). A variety of stimuli, such as hypoxia (Koong, Chen, & Giaccia, 1994), can induce the phosphorylation and subsequent ubiquitinylation and degradation of IkB, thereby allowing NF-kB to translocate into the nucleus and induce expression of target genes (Kumar et al., 2004). However, NF-kB subunits, p65 and p50, and IkBa are also known to be found within the mitochondria, where they regulate mtDNA (Bottero et al., 2001; Cogswell et al.,

2003; Guseva, Taghiyev, Sturm, Rokhlin, & Cohen, 2004; Johnson, Witzel, & Perkins,

2011). However, in the absence of p53, p65 can be transported into the mitochondria, where it represses mitochondrial gene expression, oxygen consumption, and cellular ATP levels

(Johnson et al., 2011). In the case of the hypometabolic turtle, repressing genes involved in oxygen consumption and ATP production would likely be involved in metabolic suppression, conferring an adaption to anoxic survival; thus p65 may be important in regulating mitochondrial gene expression, like cytochrome c oxidase III (Cogswell et al., 2003).

Furthermore, SIRTs appear to deacetylate and inactivate NF-kB at lysine 310 (C.J. Chen,

Yu-Cai Fu, & Wang, 2013; Yeung et al., 2004). In fact, SIRT3 overexpression protects cells from Bax-mediated apoptosis by suppressing NF-kB-dependent transcription of downstream targets like Bcl-2 (C.J. Chen et al., 2013; Xu et al., 1999). In the ground squirrel, Rouble and

Storey reported an increase in K310 acetylation of p65 NF-kB during arousal in liver and brown adipose tissue suggesting a stimulation of NF-kB downstream targets, like MnSOD, that could enhance antioxidant response (Morgan & Liu, 2011; Rouble & Storey, 2015).

SIRT3 is involved in a medley of processes. Induced by cellular stresses like caloric

restriction and hibernation, SIRT3 can activate and deactivate a host of targets to combat these stresses. Being the major mitochondrial deacetylase, SIRT3 can post-translationally modify a great number of proteins, many of which are still unknown. The focus of this thesis is to assess the influence of SIRT3 and acetylation, during the anoxic stress (the complete lack of oxygen) in one of the most anoxia-tolerant vertebrates.

1.14 The vitality of oxygen

Oxygen’s essential role in sustaining life is through enabling aerobic ATP production.

Energy used by the cell is produced as ATP and hydrolyzed into ADP or AMP releasing usable energy in the process. Due to oxygen’s high REDOX potential and ability to accept electrons from reduced metabolic intermediations, like NADH and FADH2, oxygen acts as the final electron acceptor in the ETC. While anaerobic metabolism is able to produce some

ATP through the catabolism of glucose, termed glycolysis, this is limited and cannot sustain animal life for an extended period of time. However if oxygen is present, the amount of ATP that can be produced is increased from 2mol of ATP through the breakdown of 1 mol of glucose, to 30 mol of ATP in aerobic conditions (Brand, 2003).

Oxygen-independent pathways, like anaerobic glycolysis, are relatively inefficient forms of energy production. When oxygen is low (hypoxia) or absent (anoxia), oxygen- dependent pathways, like oxidative phosphorylation, are disrupted. Most vertebrates are highly sensitive to hypoxia and prolonged exposure to oxygen deprivation. If oxygen is lacking for more than a few minutes the ETC is halted, which is the main provider of ATP, and there is an increased reliance on anaerobic glycolysis. Some species have adapted to diminished or changes in oxygen availability due to oxygen constraints in their environment,

for example intertidal zones, high altitudes, or anoxic ice-locked lakes. These hypoxia- and anoxia-tolerant organisms are able to survive limited oxygen availability for small or long periods of time. During this time, these organisms experience decreased oxidative phosphorylation and increased cytotoxicity due to a build-up of toxic end products from anaerobic metabolism. However, hypoxia- and anoxia-tolerant organisms have developed physiological and biochemical coping mechanisms that allow them to survive, while anoxia- intolerant organisms suffer irreparable tissue damage and eventually death if oxygen deprivation continues.

1.15 Surviving anoxia

Majority of vertebrates suffer irreparable cell damage following minutes in anoxia.

The western painted turtle (Chrysemys picta belli), however, can survive several months of anoxia, termed anaerobiosis, without any apparent cell damage. Other North American freshwater turtles are also well-established vertebrate models for anoxia tolerance, such as red-eared sliders (Trachemys scripta elegans) and midland painted turtles (C. picta marginata). Freshwater turtles avoid freezing temperatures on land during the winter by overwintering at the bottom of ice-locked ponds and lakes without access to the surface to breathe. These turtles are champion anaerobes, overwintering for 3-4 months at 3°C completely submerged with little to no access to oxygen (Ultsch & Jackson, 1982). However, the anoxic survival of freshwater turtles is highly dependent on temperature; the painted turtle can fully recover from 0.5 days of anoxia at 20°C, three days at 15°C, two weeks at

16°C, and up to 12 weeks at 3°C; conditions which most closely mimics their overwintering

(Fig. 1.5) (Ultsch & Jackson, 1982).

Figure 1.5 Anoxic survival is temperature-dependent. Duration of anoxic dives of Chrysemys picta bellii at various temperatures with full

recovery. Figure adapted from Jackson, 2000.

These turtles are facultative anaerobes, meaning they make ATP via aerobic pathways when oxygen is present, then make ATP via anaerobic pathways when it is absent.

The freshwater turtle has developed extensive mechanisms that enable long-term anaerobiosis. Some of the well-known biochemical adaptations that enable their survival without oxygen are (1) anaerobic pathways for ATP production, (2) large glycogen stores for prolonged anaerobiosis, (3) mechanisms to limit acidosis, and (4) metabolic depression

(Storey & Storey, 1990). Moreover, anoxia-tolerant organisms must be capable of surviving the early transition into anoxia, maintenance of a prolonged hypometabolic state, and reoxygenation during which ROS poses a threat.

1.16 Anoxic ATP production

The freshwater turtle possesses a low metabolic rate, which decreases further during

anoxic conditions. This low metabolic rate is pivotal to its long-term survival during anoxia.

Being an ectothermic reptile, the turtle’s metabolic rate is only 10-20% that of a similar-sized mammal at the same temperature (Bennett & Ruben, 1979). During anoxia, the freshwater turtle’s metabolic rate is decreased by roughly 90% compared to its normoxic rate (L T

Buck, Hochachka, Schön, & Gnaiger, 1993). The low rate of metabolism is functionally important in delaying the depletion of glycogen and slowing the build-up of acidic end products.

Low oxygen decreases ATP production; specifically, once blood oxygen levels fall below an arterial pO2 of about 20 Torr in freshwater turtles, oxidative phosphorylation is halted

(Greenway & Storey, 2000). Anaerobic glycolysis becomes the sole source of ATP during anoxia and therefore it is logical to assume glycolytic flux must be increased. This is achieved via three main anoxia-induced changes to glycolytic enzymes: dephosphorylation and phosphorylation, reversible binding of these enzymes to macromolecules or organelles, and allosteric regulation of enzymes by metabolites (Jackson, 2002).

In order to maintain anaerobic glycolysis during overwintering, freshwater turtles possess high glycogen stores principally in the liver, as well as skeletal muscle. In fact, glycogen comprises 15% of the turtle liver (Clark & Miller Jr, 1973). This large store of glycogen is sufficient to maintain stable plasma glucose and ATP levels during overwintering (Ultsch & Jackson, 1982). While ATP concentration drops somewhat after 1hr of anoxia at 20°C in red-eared sliders, levels return after 5hrs (Kelly & Storey, 1988).

Constant ATP levels have also been found in the brain (P. L. Lutz, Rosenthal, & Sick, 1985), heart (Wasser, Freund, Gonzalez, & Jackson, 1990), and liver (L T Buck et al., 1993).

1.17 Avoiding acidosis

An essential strategy for surviving anoxia is balancing ATP supply with ATP demand. While supply is limited due to reduced oxidative phosphorylation during anoxia, some ATP is produced through anaerobic glycolysis. However, lactate and protons are generated in equal proportions via anaerobic glycolysis, making lactic acid. Acidosis can be detrimental and is exacerbated in the hibernating turtle due to CO2 retention from reduced respiration, referred to as hypercapnia. In fact, the blood pH of the painted turtle can reach values of 6.7 during anoxic overwintering, in contrast with its normoxic blood pH at 3°C of

8.0 (Ultsch & Jackson, 1982). During early anoxia, hypercapnia is primarily responsible for acidosis, whereas during later submergence, the contribution of lactate increases (Herbert &

Jackson, 1985). pH and lactate accumulation during anoxia is also highly dependent on temperature (Fig. 1.6). At lower temperatures, there is a smaller increase in arterial CO2 but a larger increase in plasma lactate, compared to higher temperatures where the arterial CO2 increase is large and lactate increase is modest (Herbert & Jackson, 1985). At higher temperatures, like 15 and 20°C, the rapid rise in arterial CO2 accelerates the decrease in pH and contributes to a shorter anoxia survival (Herbert & Jackson, 1985). Compared to anoxic dives at 3°C, tolerance is enhanced due to reduced respiratory acidosis (Herbert & Jackson,

1985).

Figure 1.6 Blood pH and plasma lactate concentrations during anoxic submergence and recovery at 3, 10, 15, and 20°C.

pH is restored more rapidly than plasma lactate at each temperature. Lactate is also cleared faster at 3°C than at 20°C. Vertical lines mark anoxic and recovery periods. Figure adapted from Herbert & Jackson, 1985.

High blood acidity from respiratory and metabolic acidosis has negative consequences in most vertebrates, however the painted turtle has an exceptional buffering capacity enabling resistance to the negative effects of acidosis. The shell and skeleton bone of the turtle plays an essential role in buffering acidity, and extracellular buffering is

- enhanced through high concentrations of HCO3 that act as a sink for protons (Jackson,

2002). As anoxia continues, additional buffering is required because the buildup of lactic 25

acid will exceed the extracellular buffering capacity. Lactate uptake by the shell occurs during anoxia and is sequestered until reoxygenation or recovery, when lactate levels decrease (Jackson, 2002). Both Ca2+ and Mg2+ are released from the shell and skeleton as extracellular buffers in response to acidity (Jackson, Goldberger, Visuri, & Armstrong,

1999). Phosphate does not appear to be mobilized from the main molecule of the shell and bone, calcium phosphate, as plasma phosphate levels remained unchanged during anoxia

(Jackson et al., 1999). Rather calcium and magnesium carbonates are broken down during anoxic acidosis to act as extracellular buffers (Jackson, 2002).

1.18 Metabolic depression

During anoxia, there is a reduced supply of ATP; in order to match the reduced ATP supply with ATP demand, the painted turtle must reduce the demand of cells. This is achieved via metabolic depression and is common in other anoxia-tolerant organisms such as the crucian carp (Nilsson, 1992) and red-eared sliders (Milton & Prentice, 2007). Metabolic depression encompasses downregulating processes with high ATP utilization. Two of the most ATP-demanding cellular processes are ion pumping and protein synthesis which account for upwards of 80% of the cell’s energy budget (Fig. 1.7) (Boutilier, 2001). The early response of the turtle brain to anoxia (1-2hrs) is to decrease energy consumption by 70-

80% (Lutz, McMahon, Rosenthal, & Sick, 1984). Ion pumping consumes more than 50% of a normoxic neuron’s energy and therefore must be reduced in anoxia tolerant organisms in order to preserve energy loss (Jackson, 2002). This is achieved via (1) decreasing ion leakage or ion permeability, termed “channel arrest”, and (2) decreasing electrical activity, termed

‘spike arrest” (Hochachka, Buck, Dollt, & Landt, 1996).

Figure 1.7 Allocation of energy during normoxia vs. anoxia in turtle hepatocytes.

ATP-demanding cellular processes of isolated turtle hepatocytes decrease when cells become anoxic, with 74% of the total anoxic ATP demand being attributed to Na+ pumping. Figure adapted from Jackson, 2000.

The Na+/K+ ATPase is the major ATP-consuming ion pump and experiences a significant decrease in activity during anoxia, constituting 74% of total anoxic ATP turnover

(L T Buck et al., 1993). Channel arrest contributes to a reduction in ion pumping activity through the suppression of ion channel densities and ion channel leak. This decreases membrane permeability thereby lowering the ATP demands of maintaining transmembrane ion gradients (Fig. 1.8) (Hochachka et al., 1996). Adenosine plays a key role by decreasing

K+ efflux and down-regulating Ca2+ permeability (Buck & Bickler, 1998). Essential to maintaining transmembrane potential is maintaining extracellular glutamate levels and arresting N-methyl-D-aspartate (NMDA) receptor activity, which is a glutamate receptor and ion channel that is permeable to Na+, small amounts of Ca2+ into the cell, and K+ efflux. After 27

one hour of anoxia, this receptor was found to have a reduced probability of opening by 65% in the turtle cortex and that its downregulation was mediated by adenosine (Buck & Bickler,

1998). In fact, the MPTP silences NMDA receptors via Ca2+ release that occurs during the early onset anoxia in response to mitochondrial membrane depolarization, and this is mediated by ATP-sensitive potassium channels (Hawrysh & Buck, 2013).

Figure 1.8 Na+/K+ ATPase activity decreases during anoxia.

During anoxia, Na+ and K+ channel density and activity is reduced, leading to reduced Na+/K+ ATPase activity. This lowers the ATP demand of maintaining ion gradients across membranes. In the mitochondria, oxidative phosphorylation produces ATP driven by

proton flux through the F1Fo-ATPase during normoxia. Some protons also leak back to the mitochondrial matrix without producing ATP through a process called mitochondrial uncoupling. However during anoxia, glycolysis becomes the primary producer of ATP and

the F1Fo-ATPase maintains a mildly depolarized mitochondrial membrane potential by using ATP to translocate protons into the intermembrane space. Adapted from Boutilier, 2001.

Reduced channel activity not only contributes to maintaining membrane potential and decreased ion pumping, but also plays a role in spike arrest. Spike arrest is a reduction in brain electrical activity in order to preserve energy (Sick, Perez-Pinzon, Lutz, & Rosenthal,

1993). This can be achieved via decreasing of voltage-gated Na+ channel density and their probability of opening, as well as inhibiting ion channels involved in firing action potentials.

The anoxic turtle cerebellum exhibits a 42% decrease in the density of voltage-gated Na+ channels, likely elevating the action potential threshold and decreasing neuronal activity

(Perez-Pinzon, Chan, Rosenthal, & Sick, 1992; Sick et al., 1993). Spike arrest is also achieved through the release of inhibitory neurotransmitters, such as gamma-aminobutyric acid (GABA). After 4 hours of anoxia, extracellular GABA is elevated to 90 times its normoxic value and GABAA-receptor density increasing to 29% after 24 hours (Lutz &

Leone-Kabler, 1995; Nilsson & Lutz, 1991). In addition, NMDA receptors are inhibited by dephosphorylation during early anoxia and later by receptor renewal and internalization

(Bickler, Donohoe, & Buck, 2000). While Ca2+ influx is a ubiquitous signal for neurotransmitter release, decreased NMDA receptor activity protects against uncontrolled glutamate-activated Ca2+ influx (Bickler et al., 2000).

1.19 Molecular changes during anoxia

Mechanisms like channel arrest and spike arrest contribute to metabolic depression that the freshwater turtle employs to minimize ATP demand. While protein synthesis is also downregulated during anoxia, a select few proteins are upregulated. Protein synthesis in normoxic hepatocytes consumes 50% of the freshwater turtle’s ATP, however during anoxia, both protein synthesis and degradation are reduced by around 90% (Hochachka et al., 1996).

Heat shock proteins (HSPs) are molecular chaperones that protect against protein denaturation; their expression increases in tissues that experience high stress. During early anoxia Hsp72 is induced in the turtle brain for up to 8hrs before returning to normoxic levels around 12hrs of anoxia, while Hsc73 increases progressively over 12hrs of anoxia (Prentice,

Milton, Scheurle, & Lutz, 2004). The differential expression of these heat shock proteins suggests different roles, with Hsp72 playing a protective role during the initial transition to the anoxic, hypometabolic state and Hsc73 being more important for long term anoxia survival (Milton & Prentice, 2007). Hsc73 is not the only molecular change over long term anoxia. NF-kB exhibits maximal DNA binding at 6hrs of anoxia in the turtle brain (Lutz &

Prentice, 2002). Late translocation suggests it may be involved in long-term maintenance or a preemptive defense strategy against ROS damage upon re-oxygenation (Milton & Prentice,

2007).

Protein synthesis consumes vast amounts of ATP. In freshwater turtles it consumes

50% of the total ATP used during normoxia and to 6% during anoxia (Hochachka et al.,

1996). While synthesis and degradation are decreased during anoxia, regulatory changes in protein activity must be made in order to maintain the hypometabolic state as well as coordinate the entry to reoxygenation. Post-translational modifications of transcription factors, histones, and metabolic enzymes provide an efficient means to reduce or increase gene expression and protein activity during anoxia. For example, reversible protein phosphorylation is an effective mechanism to alter enzyme activity and is less ATP-costly than regulation of transcription or translation. In freshwater turtles, reversible protein phosphorylation controls several glycolytic enzymes; such as glycogen phosphorylase, phosphofructokinase, and pyruvate kinase (Mehrani & Storey, 1995; Storey, 1996). 30

Phosphorylation appears to be heavily prominent during anoxia in several tissues. Using 32P, global phosphoprotein levels increase by 1.6, 2.4, and 1.3 fold during anoxia in the freshwater turtle brain, heart, and liver, respectively (Brooks & Storey, 1993). Reversible protein phosphorylation is the most extensively studied but it is not the sole type of post- translation modification; others include acetylation, methylation, and ubiquitinylation. In red- eared sliders, fructose-1,6-bisphosphate aldolase and lactate dehydrogenase are regulated via acetylation during prolonged anoxia (Dawson, Bell, & Storey, 2013; Xiong & Storey, 2012).

Post-translational modification appears to be utilized as a low energy and rapid mechanism to regulate metabolic activity during anoxia. It requires far less energy than modulating protein levels via differential expression and thus is consistent with the energy-preserving state of the anoxic turtle.

1.20 Anoxia intolerance

In contrast to anoxia-tolerant organisms, anoxia-intolerant species experience no decrease in ATP demand during O2 deprivation. During hypoxia, intolerant species exhibit

“channel leak”: persistent Na+ current that increases the demand of ion balancing ATPases and triggers deleterious Ca2+ overload. Upon the onset of anoxia and a drop in cellular ATP levels, intolerant organisms, like the mammalian brain, experience hyperpolarization caused by the opening of K+ channels and increased K+ conductance (Krnjevic, 1993). This serves as a short-term defense to reduce neuronal activity by reducing the duration of action potentials as well as decreasing the release of excitatory neurotransmitters (Tanaka, Yoshida, Yokoo,

Mizoguchi, & Tanaka, 1995). However, if anoxia persists for several minutes, long term K+ channel activation is associated with a build-up of extracellular K+, gradual depolarization,

and activation of Na+ and Ca2+ channels. Excitotoxic cell death occurs within minutes of anoxia exposure through the release of excitatory neurotransmitters, such as glutamate and dopamine, causing an accelerated energy use and Ca2+ influx (Milton & Prentice, 2007).

Ca2+ influx in the cytoplasm initiates exocytotic cell death though the activation of

Ca2+-dependent proteases and apoptotic factors (Bickler & Buck, 1998). The opening of the

MPTP is triggered by cellular stress like Ca2+ and ROS, conditions likely during hypoxia and reperfusion (Halestrap, Kerr, Javadov, & Woodfield, 1998; Halestrap, Woodfield, &

Connern, 1997). Upon opening of the pore, the mitochondrial membrane potential collapses and cytochrome c is released (Borutaite, Morkuniene, & Brown, 1999).

1.21 Reoxygenation

Recovery from oxygen deprivation occurs upon reoxygenation, and during this time animals must avoid toxicity due to the overproduction of ROS. In mammals, anoxia or ischemia followed by reoxygenation is accompanied by a rapid burst of ROS production

(Halliwell and Gutteridge, 1997; Hashimoto et al. 2003; Schild and Reiser, 2005). During oxygen deprivation, electron carriers of the ETC become reduced and upon reintroduction to oxygen, these carriers are immediately oxidized. Another source of ROS is complex I.

During mammalian ischemia, succinate accumulates due to the reversal of SDH (Chouchani et al., 2016). Upon reperfusion, the accumulated succinate is re-oxidized, becoming a source of electrons by driving reverse electron transfer (Chouchani et al., 2016). Electrons are passed backwards form complex II to complex I, generating ROS using the proton motive force (Chouchani et al., 2016).

It is unknown if ROS generation is suppressed or effectively dealt with in the turtle as

part of their adaptive strategy to avoid damage upon reoxygenation. While ROS are present during normoxia and upon recovery, levels disappear after 4hrs of anoxia (Milton &

Prentice, 2007). Freshwater turtles escape ROS damage upon reoxygenation, in part, due to a diminished succinate accumulation during anoxia, where ischemic mice generate greater concentrations of this metabolic intermediate (Bundgaard, James, Joyce, Murphy, & Fago,

2018). However, ROS suppression may occur in turtle neurons (Milton, Nayak, Kesaraju,

Kara, & Prentice, 2007), and in the heart even with the accumulation of succinate

(Bundgaard et al., 2018). Another factor that contributes to the turtle’s reoxygenation tolerance is the low mitochondrial content in ectotherms compared to endotherms (Else &

Hulbert, 1985; Willmore & Storey, 1997).

Another factor that contributes to the turtle’s avoidance of reoxygenation damage is increased expression of antioxidants, such as catalase, superoxide dismutase (SOD), alkyl hydroperoxide, glutathione –S-transferase, and GSH-Px (W. Willmore & Storey, 1997).

Anoxic Midland painted turtle (C. picta marginata) hatchlings upregulated antioxidant genes in the heart and liver, including isozymes of SOD, GSH-Px, glutathione-S-transferase, and peroxiredoxin (Storey, 2006). While turtle liver GSH-Px and SOD activity levels are comparable to mammalian enzyme activities (Willmore & Storey, 1997), anoxia-tolerant reptiles do innately possess significantly higher levels of the antioxidant, ascorbic acid, than anoxia-intolerant species (Rice, Lee, & Choy, 1995). Overall, the turtle employs multiple strategies to minimize ROS damage upon reoxygenation. Although they possess similar antioxidant activity levels to mammals, their reoxygenation protection appears to come from reduced accumulation of succinate, upregulation of antioxidants, and fewer ROS-generating mitochondria. 33

1.22 SIRT3 and the painted turtle

Mitochondria have an essential role in both ATP metabolism and ROS production and mitigation. Therefore, this is a pivotal and particularly relevant organelle to the anoxia- tolerant freshwater turtle. The turtle must regulate decreased energy production during anoxic exposure and the threat of ROS upon reoxygenation, both of which involve regulating mitochondrial proteins. SIRT3 is the major deacetylase in the mitochondria and is activated under low energy conditions, such as elevated NAD+. The downstream effects of SIRT3 activation aim to limit further energy depletion, as exemplified when its abundance increased in fasting mice liver (Hebert et al., 2013) as well as the liver of 13-lined ground squirrels during torpor. SIRTs, particularly SIRT3, play a protective role in regulating apoptosis and

ATP production. However, their presence and role has yet to be characterized in a champion facultative anaerobe, like the freshwater turtle.

1.23 Objectives and hypotheses

The purpose of this study is to investigate the role of SIRT3 and its downstream partners in liver, brain, heart and pectoralis muscle samples of the western painted turtle, specifically with regard to the MPTP and antioxidant responses during anoxia.

While protein synthesis is downregulated during anoxia, the upregulation of SIRT 3 expression would be warranted because of its broad range of mitochondrial molecular targets that regulate ATP production and antioxidant defenses, among others. The liver and brain tissues were examined due to their role in energy metabolism: the brain, because it is the most energy-demanding organ and the most sensitive to oxygen deprivation, and the liver because it is the largest store of glycogen, hence would be heavily involved in glucose

mobilization and anaerobic glycolysis. The heart was examined because of its central role in transporting glucose from the liver to the brain and removing brain waste products.

Pectoralis muscle was selected to determine if responses were similar in a relatively quiescent tissue. Past studies have demonstrated post-translational protein modifications in these tissues, as well as changes in the SIRT 3 activity and targets during mammalian hibernation (Brooks & Storey, 1993; Rouble & Storey, 2015). To address this overall hypothesis, SIRT 3 and the levels of three of its targets were examined at the following time points of anoxia: 0hrs (control), 2hrs, 6hrs, 24hrs, and reoxygenation (6hrs of anoxia followed by 2hrs oxygenated recovery, henceforth referred to as 6+2hrs). The anoxic dive durations were chosen based on dive durations in similar studies. Kesaraju et al. (2009) examined HSPs during 1hr, 4hrs, and 24hrs, with changes in expression as early as 1hr following an anoxic exposure in the red eared slider. Prentice et al. (2004) also found changes in HSPs during after 6hrs and 12hrs of anoxia. Neuroglobin protein expression and mRNA levels exhibited significant increases after 4hrs of anoxia and 4hrs of reoxygenation

(Nayak, Prentice, & Milton, 2009). My hypothesis was guided by the following objectives:

Hypothesis:

The presence and upregulation of SIRT3 during anoxia is essential in the

western painted turtle in order to regulate NF-kB, antioxidant responses, and the

MPTP, which is fundamental to its survival during anoxia and the stress of

reoxygenation.

Objective 1: The presence of SIRT 3 has not been demonstrated in an anoxia-tolerant animal. The presence and expression of this protein will allow for greater insight its role in regulating metabolism and essential cell functions during anoxia. Overall mitochondrial acetylation levels are expected to decrease indicating increased deacetylase activity and/or post-translational modification during anoxia. In addition, the anoxic and reoxygenating turtle will possess greater SIRT3 protein levels and activity compared to normoxic conditions. Lysine acetylation and SIRT3 expression will be quantified via immunoreactive protein levels. SIRT3 activity will be measured via an activity assay.

Objective 2: NF-kB is a transcription factor that regulates apoptosis and oxidative stress responses. In fact, NF-kB p65 has been shown to increase in the liver of another anoxia- tolerant freshwater turtle, red-eared sliders; DNA-binding activity of NF-κB increased during anoxia and promoted the transcription of pro-survival genes Bcl-xL and Bcl-2 (Krivoruchko

& Storey, 2010b). In contrast, the hibernating squirrel liver exhibited increased deacetylation of NF-kB p65, which correlates to decreased transcriptional activity (Rouble & Storey,

2015). Thus, it is predicted than the painted turtle will exhibit similar results to the red-eared slider, in which transcriptional activity is activated during anoxia. It is expected that during anoxia deacetylation of NF-kB p65 will decrease and will be measured by measuring the ratio of acetylated NF-kB p65 (K310) to total NF-kB immunoreactivity, corresponding to enhanced transcriptional activity.

Objective 3: The MPTP is known to open in early anoxia and allow small amounts of controlled Ca2+ efflux (Hawrysh & Buck, 2013), however, during long-term anoxia the pore should remain closed in order to maintain mitochondrial membrane potential and prevent excessive Ca2+ release that can trigger cell death (Bickler & Buck, 1998). Given that inhibition of the pore is essential for long-term anoxia survival, and CypD expression is directly correlated to opening of the pore (Matas et al., 2009), I expect CypD (K166) levels will decrease during anoxia and reoxygenation to maintain closure of the MPTP so membrane potential can be preserved and Ca2+ efflux can be minimized. This will be quantified by measuring immunoreactive levels of CypD prevalence.

Objective 4: Antioxidant defenses are crucial to avoid the damage of oxidative stress during reoxygenation. Following an anoxic or ischemic bout, mammals experience a burst of ROS production upon reoxygenation (McCord, 1985; Li & Jackson, 2002). Turtles, however, are able to potentially ameliorate this damage through high antioxidant levels and suppression of

ROS formation. SIRT 3 deacetylates MnSOD thereby activating it, therefore because ROS formation is predicted to be suppressed in the turtle, it is predicted that the amount of acetylated MnSOD will remain high during anoxia and reoxygenation. This will be quantified by measuring the ratio of acetylated MnSOD (K68) to the total amount of MnSOD immunoreactive protein levels. It is also expected that the activity levels will not change throughout anoxia or reoxygenation.

CHAPTER 2: MATERIALS AND METHODS

2.1 Animals

This study was approved by the University of Toronto Animal Care Committee and conforms to the relevant guidelines issued by the Canadian Council on Animal Care regarding the care and use of experimental animals. Adult females Chrysemys picta turtles

(carapace length ~15cm, weight ~300g) were used; purchased from Niles Biological Inc.

(Sacramento, California, USA). The animals are housed in large circular indoor ponds (3m in diameter × 1.5 m deep) with a flow-through dechlorinated fresh water system maintained at

~ 18°C. Turtles are kept on a 12hrs light:12hrs dark photoperiod and given continuous access to food.

2.2 Experimental protocol

Turtles were placed in rat cages and dived at tank temperature of ~ 18°C for 2, 6, and

24 hours, as well as 6 hours of anoxia followed by two hours of reoxygenation at room temperature. Underwater, the turtles’ necks were clamped and then quickly decapitated to avoid aeration. This method has previously been demonstrated to be proficient in making the turtles anoxic (Ramaglia & Buck, 2004). Subsequently a bone saw and scalpel were used to separate the carapace and plastron. Liver, brain, heart and pectoralis muscle tissues were dissected and stored at -80°C until homogenization.

2.3 Tissue homogenization

Tissue was ground in liquid nitrogen using a mortar and pestle. Tissue used for western blots and SIRT3 activity assay was prepared as follows: 60 µL per 1 mg of tissue was homogenized in RIPA buffer [150mM NaCl, 50mM Tris-HCl, 1mM PMSF, 0.1% Triton-X, 38

0.1% SDS, 1 phosphatase inhibitor cocktail tablets (PhosSTOP, Roche Diagnostics,

Mississauga, ON, Canada)]. Tissue homogenization was done with sample on ice for approximately 10s (OmniTip Homogenizer, Omni International, Marietta, GA, USA) with homogenizer outfitted with flat, plastic tip (OmniTip plastic homogenizer probe, Omni

International, Marietta, GA, USA). Homogenates were then centrifuged at 12000 x g for 20 min at 4°C. Supernatant was removed and diluted 10x with RIPA buffer. Tissue used in

SOD activity assay was prepared according to manufacturer’s instructions. Homogenization

(OmniTip Homogenizer, Omni International, Marietta, GA, USA) was performed in HEPES buffer (20mM HEPES, 1mM EGTA, 210mM of mannitol, and 70mM sucrose, pH=7.2) using 7000µL/gram of tissue and centrifuged at 1500 x g for 5 min at 4°C. The supernatant was removed at frozen at -80°C until assayed.

Tissue samples were analyzed for protein content with a bicinchoninic acid (BCA) assay (Thermo Scientific, Oakville, ON, Canada). Using a plate spectrometer (Spectra max

384 Plus, Molecular Devices, Sunnyvale, CA, USA), the protein concentration of each sample was determined using a standard curve of bovine albumin serum (BSA) 2mg/ml serial dilutions. Samples were then stored at -80°C for future use.

2.4 Mitochondrial isolation

Control (0hrs) and dived turtles for 2hrs were decapitated according to methods previously described, then whole brain and partial liver tissues were excised and placed on ice. Immediately after dissection isolation of mitochondrial fraction was performed according to Chinopoulos et al., except whole brains were used (2011). Samples were individually homogenized in MSEGTA Buffer (225 mM mannitol, 75 mM sucrose, 5 mM

HEPES (pH 7.4), 1 mM EGTA, dissolved in water) that was supplemented with 0.2 mg/ml bovine serum albumin (BSA) and centrifuged at ~500 x g x 5 min. The supernatant was then centrifuged at 14,000 x g x 10 min. The resulting pellet was resuspended in 200 ul of 12%

Percoll-MSEGTA solution (100% Percoll-MSEGTA buffer: 225 mM mannitol, 75 mM sucrose, 5 mM HEPES (pH 7.4), 1 mM EGTA, dissolved in 100% Percoll), this suspension was then layered over 1ml of 24% Percoll-MSEGTA solution. The prepared density gradient was then centrifuged at 18,000 x g x 15 min. Following centrifugation, 700 µl was aspirated of the top portion of the sample and 1.2 ml of MSEGTA was added and mixed by inversion and centrifuged at 18,000 x g x 5min. After centrifugation 1.5ml of supernatant was aspirated and 1.5ml of MSEGTA was added and mixed by inversion. The resulting sample was centrifuged at 14,000 x g x 5 min. The pellet was then lysed in lysis buffer consisting of 50 mM Tris—pH 8.0, 1% NP40, 150 mM NaCl, 1mM EDTA, and 1mM PMSF. The samples were then centrifuged at 21,000 × g for 10 min. The supernatant was then collected, and total protein concentration determined using the BCA assay. The resulting protein samples were then stored at -80°C.

2.5 SIRT3 activity assay

Total SIRT3 activity was measured in brain tissue using SIRT3 Activity Assay Kit

(Abcam, ab156067) according to the manufacturer’s instructions. 20µg of protein prepared in

RIPA buffer according to previously described methodology, 5µL of SIRT3 Assay Buffer

(provided with kit), 5µL of Developer (provided in kit), 5µL of Enzyme Sample (provided in kit) and 5µL of Fluoro-Substrate Peptide or Fluoro-Deacetylated Peptide (provided in kit) was loaded into a black 96-well microplate in duplicate. Fluorescence was measured for

20min at an excitation of 340-360 nm and emission at 440-460 nm using a microtiter plate fluorometer (ClarioSTAR Plus, BMG Laboratories, Ortenburg, BW, Germany). The rate of reaction was calculated based on the change in fluorescence over time. Quality and efficacy of this kit was uncertain, and results were not included in this thesis (see Appendix C).

2.6 SOD activity assay

Total SOD activity was measured using Superoxide Dismutase Activity Assay Kit

(Cayman Chemicals, 706002) according to manufacturer’s instructions. Tissue homogenates from liver, brain, and heart were prepared in homogenization buffer (20mM HEPES, 1mM

EGTA, 210 mM mannitol, and 70mM sucrose, pH=7.2) using 10 µL per 1 mg wet weight of tissue. Samples were centrifuged at 1500 x g for 5 min at 4°C. Supernatant was removed and protein content was determined using BCA assay. A standard curve was prepared using 0,

20, 40, 80, 120, 160, and 200 µL of SOD stock solution (SOD standard diluted with 1.99 mL of Sample Buffer, both provided with the kit) along with Sample Buffer to make a total well volume of 1000 µL. Next, 10 µL of each prepared standard and 200 µL of Radical Detector

(undiluted form provided in kit) was loaded in duplicate into 96-well plate. Sample wells contained 200 µL of Radical Detector and 10 µL of sample. To initiate reaction, 20 µL of

Xanthine Oxidase (provided in kit) was added to all wells. Plate was covered and incubated on shaker for 30 min at room temperature. Absorbance was measured at 450 nm using plate spectrometer (Spectra max 384 Plus, Molecular Devices, Sunnyvale, CA, USA). The absorbance of 0 U/mL SOD standard was divided by itself and by all other standards to yield the linearized rate and then plotted as a function of SOD activity (U/mL), providing the standard curve. Activity of samples was obtained using the linear regression from the

standard curve substituting the linearized rate for each sample. Total SOD activity was calculated using the following formula, then normalized to protein content determined from

BCA assay to yield U/mg:

� �� − � �� 0.23�� = ∗ ∗ �� 0.01��

Data obtained from assay were not similar to those in the literature and thus are not included in this thesis (see Appendix D).

2.7 Antibodies and western blotting

Quantification of protein concentration of the samples obtained was determined using the BCA assay as previously described and 10-20µg of total protein samples were resolved using western blotting as outlined in Sidorova-Darmos (2014). A dose response blot was performed to assess amount of necessary protein loading required (see Appendix A).

Primary antibodies used and dilutions as are listed in Table 2.1. Horseradish peroxidase- linked secondary antibodies used were anti-rabbit raised in sheep (1:10000; GE Healthcare, cat # NA934V) or anti-mouse raised in donkey (1:10000; GE Healthcare, cat # NA931V). To allow normalization for minor variances in protein loading between sample lanes, gels were stained with Coomassie Blue (0.1 % Coomassie blue R-250, 10 % acetic acid, 40 % methanol) for 2hrs and destained with 10% acetic acid overnight (see Appendix E). Loading controls such as GAPDH have not been proven reliable when comparing across different tissues and/or specific cell types from the same animal (Eaton et al., 2013). In other words, total protein loading was used for normalization as an alternative to a single housekeeping gene as a loading control. Target protein bands were identified by running a standard protein 42

ladder (Cell Signaling, #14208) and 7µL of a mammalian positive control (Mouse, Mus musculus) of liver alongside. Similarity between SIRT3 found in mouse and the western painted turtle was compared to determine efficacy of antibodies (see Appendix F).

X-Ray Films were scanned, and the optical density of bands was quantified using Image J version 1.816 software (https://imagej.nih.gov/ij/), with film background subtracted to give the final densitometric values.

Table 2.1: List of primary antibodies Antibody Species Dilution Used Company, Catalogue #

Acetylated-Lysine Rabbit, polyclonal 1:1000 Cell Signaling, #9441

Cyclophilin F Mouse, polyclonal 1:1000 Abcam, ab110324

MnSOD Rabbit, polyclonal 1:1000 Abcam, ab13533

MnSOD acetyl K68 Rabbit, polyclonal 1:1000 Abcam, ab137037

NF-kB p65 Rabbit, polyclonal 1:500 Abcam, ab16502

NF-kB p65 K310 Rabbit, polyclonal 1:1000 Genetex, GTX86963

SIRT2 Rabbit, polyclonal 1:1000 Sigma-Aldrich, S8447

SIRT3 Rabbit, polyclonal 1:1000 Cell Signaling, #5490

Tom 20 Rabbit, monoclonal 1:1000 Cell Signaling, #42406

2.8 Statistical Analysis

Statistical analysis was performed using one-way or two-way ANOVA, and

Bonferroni post-hoc test, as well as ROUT outlier test, in GraphPad Prism version 5.00

(GraphPad Software, San Diego CA). All results are expressed as mean ± standard error of means (SEM) and p < 0.05 was set as the level of statistical significance.

CHAPTER 3: RESULTS

This thesis aimed to characterize the level of reversible acetylation in the mitochondrial proteome, primarily, the role of the major mitochondrial deacetylase, SIRT3.

Measurement of lysine acetylation levels revealed that mitochondrial acetylation increased during early anoxia. Contrary to the prediction that SIRT3 would be upregulated and remain elevated during anoxia, protein levels remained close to control values with two exceptions; late anoxia in the liver and potentially early anoxia in the brain. Furthermore, the SIRT3 target, NF-kB p65, only increased in expression in terms of its deacetylated state in the brain during anoxia and reoxygenation. MnSOD exhibited minimal changes in terms of activity, prevalence, and acetylation status. Lastly, CypD expression increased during the middle of the painted turtle’s anoxic bout only to return to control values.

3.1 SIRT3 expression

Protein synthesis during anoxia can be ATP-costly, thus it is heavily downregulated during anoxia, however, select proteins are upregulated. Rouble & Storey (2015) found increased SIRT3 expression during late torpor in the liver of 13-lined ground squirrels, I predicted that the painted turtle would exhibit similar findings, thus suggesting a role for

SIRT3 during hypometabolism. In the brain, expression of SIRT3 increased 3.11-fold after

2hrs of anoxia and there was a 1.79-fold increase after 24hrs of anoxia, compared to control

(Fig. 3.2A). In the liver, there was a significant increase in SIRT3 levels by 2.34-fold at

24hrs of anoxia compared to control; this increase was significantly different compared to

2hrs and 6hrs (Fig. 3.2B). There was also a 1.69-fold increase in SIRT3 expression upon reoxygenation (6+2hrs) in the liver (Fig. 3.2B). Therefore, SIRT3 expression increases

during late anoxia in the liver of the painted turtle, similarly to the ground squirrel, and thus may be important in regulating late stages of hypometabolism.

A B

Brain Liver 500 500

400 400

300 300 a a,b 200 200 b

(% of 0hrs) (% of 0hrs) b b

Protein Prevalence 100 Protein Prevalence 100

0 0

0hrs 2hrs 6hrs 0hrs 2hrs 6hrs 24hrs 24hrs 6+2hrs 6+2hrs

0 2 6 6+2 24 0 2 6 6+2 24

Figure 3.1 SIRT3 protein expression compared to control

Representative Western blots of showing the prevalence of SIRT3 in the (A) brain and (B) liver. The histograms in each panel show the densitometric mean and SEM normalized to Coomassie blue staining control. The expression level for each is presented relative to the expression level in control (0hrs) cohort. (Error bars represent SEM, n=5 for all except, ROUT outlier test performed, and one data point removed from brain at 24hrs resulting in n=4 for that treatment group). “a” indicates significant difference between 24hrs and 0hrs (p= 0.008), 2hrs (p=0.002), and 6hrs (p=0.046). One-way ANOVA with Bonferroni post- hoc test.

3.2 NF-kB p65 expression and acetylated expression

NF-kB DNA binding is enhanced in the anoxia-tolerant red-eared slider

(Krivoruchko & Storey, 2010b), however in the 13-lined ground squirrel acetylation of NF- kB p65 K310 decreases corresponding to less transcriptional activity (Rouble & Storey,

2015). In the anoxic painted turtle, Total NF-kB p65 protein levels in the brain, exhibited increases of 1.56-fold and 1.57-fold after 2hrs and 6hrs of anoxia, respectively, compared to control (Fig. 3.3A). The brain also experienced increases in NF-kB p65 protein levels of

1.77- and 1.43-fold after 6 + 2hrs of anoxia and 24hrs of anoxia, respectively (Fig. 3.3A).

However, in the liver, NF-kB p65 increased more dramatically by 2.40-fold after the 6+2hrs treatment compared to anoxia, and 2.92-fold after 24hrs of anoxia compared to control (Fig.

3.3B). In contrast, NF-kB p65 K310 experienced a significant 0.40- and 0.43-fold decrease after 2hrs and 6hrs of anoxia, respectively in the brain (Fig. 3.5A). During 6+2hrs and 24hrs of anoxia in the brain, NF-kB p65 K310 also exhibited a significant 0.41- and 0.53-fold decrease compared to 0hrs (Fig. 3.3C). While in the liver, no significant changes in NF-kB p65 K310 occurred compared to control, there was a 0.51-fold decrease after 6+2hrs of anoxia (Fig. 3.3D). In contrast to the red-eared slider but similarly to the 13-line ground squirrel, acetylated status of NF-kB p65 decreases during anoxia; this would mean transcriptional activity appears is downregulated in brain of anoxic painted turtles.

A B Brain Liver

400 400

300 300

200 200 (% of 0hrs) (% of 0hrs) 100 100 Protein Prevalence Protein Prevalence

0 0

0hrs 2hrs 6hrs 0hrs 2hrs 6hrs 24hrs 24hrs 6+2hrs 6+2hrs

0 2 6 6+2 mouse 0 2 6 6+2 24

C D Brain Liver 2.0 2.0 a 1.5 1.5

1.0 1.0

b b b b 0.5 0.5 NFkB K310:Total NFkB NFkB K310:Total NFkB 0.0 0.0

0hrs 2hrs 6hrs 0hrs 2hrs 6hrs 24hrs 24hrs 6+2hrs 6+2hrs

mouse 0 2 6 6+2 24 mouse 0 2 6 6+2 24

Figure 3.2 NF-kB p65 and NF-kB p65 K310 protein expression compared to control Representative Western blots of NF-kB p65 in the (A) brain and (B) liver, and NF-kB p65 K310 in (C) brain and (D) liver. Histograms (A), (B), (C), and (D) show the densitometric mean normalized to Coomassie blue staining control. Expression levels are presented relative to 0hrs. (A) and (B) show total NF-kB p65, (C) and (D) show ratio of NF-kB p65 K310 to total NF-kB p65 expression levels. (Error bars represent SEM, n=5 for all except n=4 for 0hrs group of (C) using ROUT outlier test). “a” indicates significant difference between 0hrs and 2hrs (p= 0.0095), 6hrs (p= 0.0120), 6+2hrs (p= 0.010), and 24hrs (p= 0.0360). One-way ANOVA with Bonferroni post-hoc test.

3.3 Cyclophilin D expression

CypD is a major regulator of the MPTP and its formation. I predicted that CypD expression would remain low during anoxia, however there was one finding contrary to my prediction. After 6hrs of anoxia in the brain, turtles experienced a significant 3.11-fold increase compared to control, with all other treatments unchanged (Fig. 3.5A). The liver also experienced a 4.07-fold increase after 6hrs of anoxia compared to control (Fig. 3.5B). At

6hrs of anoxia, CypD prevalence possesses a large SEM in the liver; this resulted from some individuals exhibiting greater band density than others. The large variances could suggest some turtles exhibit an increase in CypD expression while others do not. CypD expression increased in the brain and there was an apparent similar trend in the liver, indicating an open conformation of the MPTP.

A B

Brain Liver 600 600

a 400 400

a,b

(% of 0hrs) 200

(% of 0hrs) 200 b b b Protein Prevalence Protein Prevalence

0 0

0hrs 2hrs 6hrs 0hrs 2hrs 6hrs 24hrs 24hrs 6+2hrs 6+2hrs

0 2 6 6+2 24 0 2 6 6+2 24

Figure 3.3 CypD protein expression compared to control

Representative Western blots of showing the prevalence of CypD in the (A) brain and (B) liver. The histograms in each panel show the densitometric mean and SEM normalized to Coomassie blue staining control. The expression level for each are presented relative to the expression level in control (0hrs) cohort. (Error bars represent SEM, n=5). “a” denotes significantly difference between 6hrs and 0hrs (p= 0.0107), 6+2hrs (p= 0.0092) and 24hrs (p= 0.0137). One-way ANOVA with Bonferroni post-hoc test.

3.4 MnSOD expression

Antioxidant prevalence or activity was not predicted to be high during anoxia in the painted turtle due to the lack of O2 and ROS production at this time. During reoxygenation mammals experience a burst of ROS production. An animal adapted to periods of oxygen deprivation and reoxygenation should possess enhanced oxidative stress protection. Thus, it was predicted that the anoxia-tolerant painted turtle would increase =the expression of antioxidants. To assess the antioxidant capacity and its potential regulation by SIRT3,

MnSOD expression and acetylated MnSOD was measured. Total MnSOD expression in the brain increased by 1.81 after 6+2hrs treatment compared to control (Fig. 3.4A). While in the liver, the was a significant change in expression of MnSOD in the 24hrs of anoxia group compared to the 6+2hrs reoxygenation group, with a 1.23-fold increase compared to control

(Fig. 3.4B). In contrast, proportion of acetylated MnSOD remained unchanged through anoxia and reoxygenation in both the brain and liver (Fig. 3.4C).

A B Brain Liver 250 250

200 200

150 b 150 a,b a,b a,b a 100 100 (% of 0hrs) (% of 0hrs)

Protein Prevalence 50

0 0

0hrs 2hrs 6hrs 24hrs 0hrs 2hrs 6hrs 6+2hrs 24hrs 6+2hrs

mouse 0 2 6 6+2 24 mouse 0 2 6 6+2 24 C D Brain Liver 1.5 1.5

1.0 1.0

0.5 0.5 SODK68:Total SOD SODK68:Total SOD

0.0 0.0

0hrs 2hrs 6hrs 0hrs 2hrs 6hrs 24hrs 24hrs 6+2hrs 6+2hrs

mouse 0 2 6 6+2 24 mouse 0 2 6 6+2 24

Figure 3.4 MnSOD protein expression compared to control

Representative Western blots of showing the prevalence of MnSOD in the (A) brain and (B) liver and MnSODK68 in the (C) brain and (D) liver. Histograms (A), (B), (C), and (D) show the densitometric mean normalized to Coomassie blue staining control where expression levels are presented relative to 0hrs. (A) and (B) show total MnSOD, while (C) and (D) show ratio of acetylated MnSOD to total MnSOD expression levels. (Error bars represent SEM, n=5). “a” and “b” denotes significant difference between 6+2hrs and 24hrs, p= 0.0406. One-way ANOVA with Bonferroni post-hoc test

3.5 Mitochondrial lysine acetylation

In order to measure mitochondrial acetylation levels, staining for TOM20 was used to assess purity of isolated mitochondria and used for normalization. Expression of acetylated lysine residues in the mitochondria decreased significantly in the brain after 2hrs of anoxia resulting in a 0.75-fold decrease compared to control (Fig. 3.1). A similar trend was observed in the liver, however results were insignificant (Fig. 3.1). This provides evidence of post- translational modification occurring during anoxia, suggesting deacetylation is occurring during early anoxia in the brain. This could be a potential mechanism for hypometabolic regulation during this time.

Acetylated Lysine Brain 200 0hrs 2hrs Acetyl-Lys 150 TOM 20 100 * 0 2 (% of 0hrs) 50 Liver Protein Prevalence

0 Acetyl-Lys

Brain Liver TOM 20

0 2

Figure 3.5 Mitochondrial acetylated lysine residues after 2hrs of anoxia.

Relative protein levels of acetylated lysine residues in isolated mitochondria after 2hrs of anoxia represented as a percent (%) of control (0hrs) in brain and liver. The histograms in each panel show the densitometric mean and SEM normalized to Coomassie blue staining control and TOM20. (Error bars represent SEM, n=3) “*” indicates significant difference from its respective control (0hrs), p= 0.012. Student t-test performed.

CHAPTER 4: DISCUSSION

Oxygen-based metabolism is critical for sufficient generation of ATP for all vertebrates; thus oxygen deprivation is lethal to these species. There are a select few vertebrates, however that are able to withstand this deprivation for periods as long as several months. The Western painted turtle is capable of surviving anoxia for as much as three months at 3°C and recover with no apparent cell damage (Jackson, 2002; Ultsch & Jackson,

1982). This animals is enabled to do so by employing a multitude of strategies: (1) reliance on glycolysis and large hepatic glycogen stores for ATP production, (2) acid buffering by releasing bicarbonate from their shell and lactate storage into the shell, and (3) reducing metabolic rate and ATP demands via suppression of gene expression, ion pumping, protein translation, metabolic flux, and neuronal activity (Buck & Pamenter, 2018; Jackson, 2000).

4.1 SIRT3 regulation during anoxia

SIRT3 upregulation has been demonstrated following cellular stresses such as: caloric deprivation, exercise (Brandauer et al., 2015; Cheng et al., 2016; Palacios et al., 2009; T. Shi et al., 2005; Tauriainen et al., 2011) and is protective during ischemic stress (He, Zeng, &

Chen, 2016; Morris et al., 2017; Porter, Urciuoli, Brookes, & Nadtochiy, 2014). Ischemia- induced SIRT3 expression positively regulates autophagy and protects cortical neurons from damage (Dai et al., 2017). I predicted SIRT3 would provide protection from anoxic stress and oxidative stress upon reoxygenation to the painted turtle and thus would be upregulated.

I found SIRT3 protein levels increased maximally at 24hrs in the liver, which could be a preparatory mechanism to maximize ATP production upon reoxygenation. Turtles exposed to anoxia at 20°C fully recover after 12hrs (Ultsch & Jackson, 1982), and while turtles in this

experiment survived a 24hr dive at 18°C, this may have been the extent of their survival.

Thus, the increase in SIRT3 during the turtle’s final hours of survival might be to poise the liver for imminent and reoxygenation. Where upon reoxygenation, SIRT3 is ready to activate targets LCAD and VLCAD to oxidize fatty acids as well as HMGCS2 and mobilize ketone bodies to brain. As well as activate PDH and LDH to convert the accumulated lactate into acetyl-CoA, and/or activate ACS2 to convert acetate into acetyl-CoA, and when oxygen is present acetyl-CoA will be used to generate ATP. A similar trend may occur in the brain where SIRT3 mildly increases in the brain at 2hrs, which may serve a similar function as in the liver: to prepare for imminent reoxygenation (Fig. 4.1), by activating similar targets such as ACS2, PDH, and LDH. Apparent upregulation of SIRT3 in the brain may also be attributed decreased protein degradation and not necessarily increased protein translation due to the short time of anoxic exposure and the amount of time needed to induce transcription and translation.

SIRT3 is classified as an NAD+ dependent deacetylase, meaning NAD+ is a necessary cofactor for the enzymatic action of SIRT3 to be performed however, it experiences a substantially smaller increase than the ischemic mouse (Bundgaard et al., 2019). The elevated NADH/NAD+ ratio that occurs during anoxia drives the conversion of a- ketoglutarate to oxaloacetate and then malate in the cytosol producing NAD+ in the process

(Bundgaard et al., 2019). Mitochondrial malate import leads to succinate accumulation, which has been previously demonstrated to accumulate in the anoxic painted turtle (Buck,

2000). While the NADH/NAD+ ratio does increase during anoxia, there might be sufficient levels of NAD+ to maintain SIRT3 function particularly in the mitochondria. Mitochondrial levels of NAD+ are shown to remain at physiological levels following genotoxic stress and 54

maintain cell viability even when cytoplasmic pools are depleted (Yang et al., 2007).

Furthermore, the ischemic brain may be able to maintain NAD+ levels as levels have been demonstrated to remain unchanged in terms of its NAD+ concentrations (Paschen, Oláh, &

Mies, 2000). This may be mediated by neuroglobin, an O2 storage protein in the brain, that possesses many roles including regenerating NAD+; its mRNA expression has been demonstrated to be strongly upregulated during hypoxia and reoxygenation, with a lesser increase during anoxia in the brain of red-eared sliders (Milton, Nayak, Lutz, & Prentice,

2006). Furthermore, cardiomyocytes exhibited increases in a stress-responsive gene,

NAMPT, which boosts mitochondrial NAD+ levels and interestingly, enhances SIRT3 activity (Yang et al., 2007). Similarly, hypoxia induced HIF-2a, upregulates NAMPT, which stimulates NAD+ synthesis and SIRT activation (Oh et al., 2015). Based on these studies, it is possible mitochondrial NAD+ levels are maintained during anoxia in the turtle, mediated by neuroglobin or NAMPT, and are able to sustain SIRT3 activity.

4.2 Deacetylation of NF-kB p65 during anoxia

Many transcription factors are known to be involved in stress responses, however

NF-kB might play an essential role due to its rapidity of activation and the numerous pathways it controls; this transcription factor induces the expression of several target genes involved in immune response, antioxidant defenses, cell growth, and apoptosis upon activation (Kaltschmidt et al., 2000; Rothgiesser et al., 2010; Yeung et al., 2004). Due to NF- kB mediating of a plethora of stress responses, its activation could enable anoxic survival by the painted turtle.

Phosphorylation and dissociation of IkB from NF-kB is induced by various stimuli.

This allows NF-kB to translocate to the nucleus, and possibly the mitochondria, to promote gene transcription (Johnson et al., 2011; Kumar et al., 2004). Elevated levels of phosphorylated IkB have been demonstrated in the liver of anoxic red-eared sliders, suggesting activation of NF-kB, in addition to increased p65 protein levels (Krivoruchko &

Storey, 2010b). Not only does increased activation of NF-kB take place during anoxia, target genes of this transcription factor were also assessed in the turtle. Transcript levels of NF-kB antioxidant target genes: MnSOD and CuSOD, were upregulated in the anoxic red-eared slider liver, which suggests a role in cell protection and/or preparation for oxidative stress upon reoxygenation (Krivoruchko & Storey, 2010b). Anti-apoptotic genes, Bcl-2, and Bcl- xL, are also controlled by NF-kB, and experienced upregulation of transcript levels

(Krivoruchko & Storey, 2010b). Both anti-apoptotic factors reside in the outer mitochondrial membrane and maintain membrane integrity (Janumyan et al., 2003), thus these genes may be preventing apoptosis in the turtle during anoxia.

In this thesis, the turtle brain experienced a decrease in acetylated NF-kB p65 throughout anoxia and reoxygenation, which suggests suppressed transcriptional activity.

Deacetylation of NF-kB p65 corresponds with decreased transcriptional activity of its target genes (Rothgiesser et al., 2010; Yeung et al., 2004). These findings are in contrast with those of the red-eared slider but similar to that of the hibernating ground squirrel. Late torpor decreased K310 acetylation of NF-kB p65, suggesting that transcriptional activity is suppressed (Rouble & Storey, 2015). In the case of the painted turtle, anoxia generally suppresses gene expression and protein translation in order to minimize ATP expenditure.

The brain, in particular, must be stringent with ATP usage because it must maintain ion

gradients to preserve neuron viability. The liver, in contrast to the brain, maintained the proportion of acetylated NF-kB p65 during anoxia and reoxygenation suggesting suppression of NF-kB p65 is unnecessary or some transcriptional activity is necessary. Lastly, while it is possible that deacetylation could be mediated by another deacetylase, deacetylation of mitochondrial NF-kB p65 is most likely performed by SIRT3 given that it is the dominant mitochondrial deacetylase (Fig. 4.1).

4.3 CypD expression increases and could open the MPTP

Mitochondria mediate apoptotic and necrotic cell death and the MPTP plays a critical role in regulating this as it controls mitochondrial swelling and the release of apoptotic mediators. Vulnerability to MPTP opening has been associated with oxidative stress (Chien,

1999; Giordano, 2005), accumulation of mitochondrial matrix Ca2+ (Balke & Shorofsky,

1998; Bernardi, 2013; Yano, Ikeda, & Matsuzaki, 2005), hypoxia (Giordano, 2005), and high matrix pH (Zoratti & Szabò, 1995). Recent studies have found that CypD plays an important role in regulation of the MPTP and thus susceptibility to cell death. Gene inactivation of

CypD in mice resulted in inhibition of MPTP opening and protection from cell death induced by ischemia-reperfusion oxidative stress, while over expression lead to activation of MPTP opening and susceptibility to cell death (Baines et al., 2005). Increased abundance of CypD and translocation to the mitochondria has also been found to increase the mitochondria’s vulnerability to stress (Matas et al., 2009).

In this thesis, CypD protein levels were found to be low throughout anoxia and reoxygenation, suggesting a closed conformation of the MPTP, except for a large increase in expression at 6hrs of anoxia in the brain, with a similar trend occurring in the liver. While I

predicted the MPTP would remain closed throughout experimentation in order to prevent cell death; low expression of CypD would suggest that for the most part, this occurred. However, the sudden increase in expression at 6hrs suggests an opening of the MPTP at this time.

While it is possible that increased expression of CypD may not be the sole promoter of an open MPTP, alkaline conditions would be the most probable secondary cause due to the turtle’s extraordinary buffering capacity, where these animals can maintain an extracellular pH above 7.0 for at least 3 weeks of anoxia at 3°C (Ultsch & Jackson, 1982). However, this is not likely because while extracellular pH is high during anoxia in the painted turtle, mitochondrial matrix pH actually acidifies during anoxia (Hawrysh & Buck, 2019).

Therefore, if the MPTP is opened by the increase in CypD protein levels, it is likely the only inducer of an open MPTP.

An open MPTP would normally confer activation of necrosis or apoptosis as formation of the MPTP is associated with cell death in anoxia-intolerant species. However, moderate opening of the MPTP is also associated with anoxic survival in the painted turtle.

Activation of the MPTP by CypD mediates moderate Ca2+ release during early anoxia, which silences NMDA receptors and ultimately aims to preserve ATP during anoxia (Hawrysh &

Buck, 2013). An open MPTP during the middle of an anoxic bout in the turtle would likely not increase its vulnerability to stress like in mammals, instead it likely confers prolonged survival. This could be a transition from short term to longer-term survival where silencing

NMDA receptors again like in early anoxia is necessary to preserve unnecessary ATP usage

(Hawrysh & Buck, 2013). SIRT3 also has the possibility to counteract the actions of increased CypD expression, by deacetylation and inhibition of MPTP formation to some degree. However, increase in CypD prevalence during the middle of the turtle’s anoxic bout 58

in unmatched by any corresponding increase in SIRT3 during this time to inactivate CypD.

Thus, it is likely that CypD promotes formation of the MPTP.

Figure 4.1 The proposed model of SIRT3 in the brain of anoxic freshwater turtle

Brain SIRT3 increases during early anoxia, concomitantly with increased deacetylated NF- kB p65 which is maintained throughout anoxia and reoxygenation. This corresponds with less transcriptional activity like to preserve ATP. While deacetylation of NF-kB p65 has not been confirmed to be performed by SIRT3, it is represented here given that SIRT3 is the dominant mitochondrial deacetylase. Increased CypD expression also occurs during the middle of an anoxic bout which may cause opening of the MPTP and Ca2+ release. However, SIRT3 has the ability to deacetylate and inactivate CypD, which would result in inhibition of the MPTP. While no increase in MnSOD expression or activation was seen, the turtle may possess adequate levels of this antioxidant to scavenge ROS produced upon reoxygenation. Horizontal arrows represent activation and inhibition arrows are flat- topped, while increases in expression are represented by thicker vertical arrows.

4.4 MnSOD during anoxia and reoxygenation

The 13-lined ground squirrel experiences increases in ROS production during arousals from torpor due to shivering thermogenesis, but does not upregulate expression or activity or MnSOD or CuSOD in heart, brain, or liver (Page, Peters, Staples, & Stuart, 2009).

The squirrel’s enhanced oxidative stress resistance does not appear to be due to upregulation of antioxidants in the major oxidative tissues. Anoxia-tolerant freshwater turtles are known to possess high innate concentrations of antioxidants (Rice et al., 1995), however, there remains some debate about whether they experience a ROS burst upon reoxygenation like anoxia-intolerant animals. Turtles could have adequate antioxidant enzyme levels to deal with ROS efficiently, however, they appear to employ multiple strategies to suppress reoxygenation-induced damage. In the Midland painted turtle hatchlings, several antioxidant enzymes experience upregulation during anoxia: Cu/ZnSOD, glutathione peroxidase, glutathione-S-transferase, and peroxiredoxin (Storey, 2007). In adult red eared slider, liver glutathione-S-transferase experienced a reduction in activity during anoxia that rose again upon reoxygenation; the activity in the liver and red muscle during anoxia increased by 52% and 80% respectively (Willmore & Storey, 1997). In this thesis, MnSOD levels remains relatively unchanged; acetylation of MnSOD was also static during anoxia and reoxygenation. This means that MnSOD does not appear to be involved in anoxia tolerance or reoxygenation. The freshwater turtle does possess other antioxidants that may be sufficient to combat ROS damage. However, it is also possible that the freshwater turtle suppresses

ROS production upon reoxygenation and thus does not experience significant upregulation of antioxidants. The anoxia-tolerant turtle actually suppresses succinate production during anoxia which becomes a source of ROS upon reoxygenation (Bundgaard et al., 2019, 2018). 60

These animals also inherently possess fewer mitochondria than endotherms (Else & Hulbert,

1985; Willmore & Storey, 1997), thus would innately be capable of producing less ROS.

These factors contribute to the possibility of turtles inhibiting ROS production upon reoxygenation, unlike mammals who experience reperfusion damage.

4.5 Mitochondrial acetylation

Regulation of protein activity through post-translational modification during anoxia in the hypometabolic turtle has been examined in several studies. Altered expression of transcription factors (NF-kB, FoxO, and HIF-1) and their genes, as well as changes in histone acetylation and histone deacetylation have been reported (Krivoruchko & Storey,

2010b, 2010a, 2013). Phosphorylation also occurs during anoxia in the turtle brain and liver

(Brooks & Storey, 1993); as does acetylation of two essential glycolytic enzymes: fructose-

1,6-bisphosphate aldolase and lactate dehydrogenase (Dawson et al., 2013; Xiong & Storey,

2012). During early anoxia, I found that the painted turtle exhibited a decrease in acetylated mitochondrial lysine residues in the brain, meaning decreased mitochondrial acetylation or increased deacetylation. A similar trend may also be occurring in the liver. However, the definitive presence of a mitochondrial acetyltransferase has not yet been identified (Baeza,

Smallegan, & Denu, 2016). Deacetylation may be occurring in turtle mitochondria, likely by the dominant mitochondrial deacetylase, SIRT3.

CHAPTER 5: CONCLUSION AND FUTURE REMARKS

The painted turtle utilizes biochemical and physiological mechanisms to enable its long-term anoxic survival. Suppression of protein synthesis and metabolism during anoxia aims to minimize energy expenditure and balance ATP supply with ATP demand. Post- translational modification is an ATP-inexpensive and rapid method of altering protein activity. While phosphorylation is the most prevalent form of post-translational modification, acetylation (and deacetylation) is the second most prevalent, and is relatively unexamined in the anoxia-tolerant turtle (Khoury, Baliban, & Floudas, 2011). This type of protein modification was examined within the scope of SIRT3 in this thesis.

The anoxia tolerance of the freshwater turtle has been explored with regards to their antioxidant response (Willmore & Storey, 1997), heat shock proteins (Prentice et al., 2004), transcription factors such as NF-kB and FoxO (Krivoruchko & Storey, 2010b, 2013), and post-translational modification of glycolytic enzymes (Brooks & Storey, 1993; Dawson et al., 2013; Mehrani & Storey, 1995; K. B. Storey, 2007; Xiong & Storey, 2012). While this thesis helped elucidate the role of post-translational modification by SIRT3, further investigation is needed into this deacetylase and its activity. During anoxia, protein synthesis is, for the most part suppressed, however, some proteins do warrant the energy expenditure

(ATP) it requires. This is exhibited in terms of hepatic SIRT3 levels during late anoxia, which could be involved in poising the tissue for reoxygenation. Results in this thesis suggest limited regulation of brain and liver SIRT3. Future work could examine mRNA levels of

SIRT3 as well as CypD, MnSOD, and NF-kB in these tissues utilizing real-time reverse transcription polymerase chain reaction (RT-PCR) as well as successfully measuring SIRT3 enzymatic activity in order to determine if patterns seen in protein expression are reflected in 62

mRNA expression. This would also help tease apart the role of decreased protein degradation and protein synthesis. Furthermore, due to the use of mouse and human specific antibodies, future work should employ positive and negative controls: SIRT3 upregulation through plasmid transfection and knockdown with siRNA in isolated turtle cells, respectively.

Deacetylation of the mitochondrial proteome occurred during early anoxia likely via

SIRT3, the major mitochondrial deacetylase. While this thesis only examined early anoxia, a longer-term analysis of mitochondrial acetylation would further illuminate the role of SIRT3 deacetylation and this type of post-translational modification as a means of metabolic suppression. This thesis also found increased deacetylation of NF-kB p65 during anoxia and reoxygenation in the brain. ChIP-sequencing can be integrated with other genomic assays used to determine gene expression mediated by this transcription factor during anoxia.

CypD expression increased in brain and liver tissues to potentially open the MPTP and silence NMDA receptors during a transition from short to long-term anoxic survival.

Investigation into deacetylation of CypD using immunoprecipitation followed by western blotting could determine levels of acetylated CypD during anoxia. Further research into confirmation of an open pore is also essential to fully elucidate the role of the MPTP during anoxia. Another target of SIRT3, MnSOD, exhibited no changes in deacetylation or expression levels during anoxia or reoxygenation, suggesting that the turtle does not require further activation of this antioxidant to combat ROS damage upon reoxygenation. However, the mechanisms by which the turtle suppresses ROS synthesis remain in need of further investigation especially with regards to succinate suppression being a key factor in preventing ROS synthesis.

Overall, this thesis has characterized mitochondrial acetylation as well as 63

deacetylation of known SIRT3 targets, providing preliminary results into the role of post- translational modification in the mitochondria as a survival mechanism during anoxia and reoxygenation or the painted turtle. Anoxia survival is multi-faceted. This study has furthered our understanding of this animal’s adaptive strategies to anoxia, however, future studies are needed to illuminate the role of the MPTP, SIRT3, and ROS suppression.

REFERENCES

Ahn, B. H., Kim, H. S., Song, S., L., H., I., Liu, J., Vassilopoulos, A., … Finkel, T. (2008). A role for the mitochondrial deacetylase Sirt3 in regulating energy homeostasis. Proceedings of the National Academy of Sciences, 105(38), 14447–14452. Allan, M., & Storey, K. (2012). Expression of NF-jB and downstream antioxidant genes in skeletal muscle of hibernating ground squirrels, Spermophilus tridecemlineatus. Cellular Biochemistry, 30(2), 166–174. Azzu, V., & Brand, M. D. (2010). The on-off switches of the mitochondrial uncoupling proteins. Trends in Biochemical Sciences, 35(5), 298–307. https://doi.org/10.1016/j.tibs.2009.11.001 Baeza, J., Smallegan, M. J., & Denu, J. M. (2016). Mechanisms and Dynamics of Protein Acetylation in Mitochondria. Trends in Biochemical Sciences, 41(3), 231–244. https://doi.org/10.1016/j.tibs.2015.12.006 Baines, C. P. (2007). The mitochondrial permeability transition pore as a target of cardioprotective signaling. American Journal of Physiology-Heart and Circulatory Physiology, 293(2), H903–H904. https://doi.org/10.1152/ajpheart.00575.2007 Baines, C. P., Kaiser, R. A., Purcell, N. H., Blair, N. S., Osinska, H., Hambleton, M. A., … Molkentin, J. D. (2005). Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death. Nature, 434(7033), 658–662. https://doi.org/10.1038/nature03434 Balke, C. W., & Shorofsky, S. R. (1998). Alterations in calcium handling in cardiac hypertrophy and heart failure. Cardiovascular Research, 37(2), 290–299. https://doi.org/10.1016/S0008-6363(97)00272-1 Bartlett, K., & Eaton, S. (2004). Mitochondrial β-oxidation. European Journal of Biochemistry, 271(3), 462–469. https://doi.org/10.1046/j.1432-1033.2003.03947.x Basso, E., Fante, L., Fowlkes, J., Petronilli, V., Forte, M. A., & Bernardi, P. (2005). Properties of the Permeability Transition Pore in Mitochondria Devoid of Cyclophilin D. Journal of Biological Chemistry , 280(19), 18558–18561. https://doi.org/10.1074/jbc.C500089200 Bauer, M., Hamm, A. C., Bonaus, M., Jacob, A., Jaekel, J., Schorle, H., … Katzenberger, J. 65

D. (2004). Starvation response in mouse liver shows strong correlation with life-span- prolonging processes. Physiological Genomics, Vol. 17, pp. 230–244. https://doi.org/10.1152/physiolgenomics.00203.2003 Bause, A. S., & Haigis, M. C. (2013). SIRT3 regulation of mitochondrial oxidative stress. Experimental Gerontology, 48(7), 634–639. https://doi.org/https://doi.org/10.1016/j.exger.2012.08.007 Bell, E L, Emerling, B. M., Ricoult, S. J. H., & Guarente, L. (2011). SirT3 suppresses hypoxia inducible factor 1α and tumor growth by inhibiting mitochondrial ROS production. Oncogene, 30, 2986. Retrieved from https://doi.org/10.1038/onc.2011.37 Bell, Eric L., & Guarente, L. (2011). The SirT3 Divining Rod Points to Oxidative Stress. Molecular Cell, 42(5), 561–568. https://doi.org/https://doi.org/10.1016/j.molcel.2011.05.008 Bellizzi, D., Rose, G., Cavalcante, P., Covello, G., Dato, S., De Rango, F., … Franceschi, C. (2005). A novel VNTR enhancer within the SIRT3 gene, a human homologue of SIR2, is associated with survival at oldest ages. Genomics, 85(2), 258–263. Bennett, A. F., & Ruben, J. A. (1979). Endothermy and Activity in. Science, 206(4419), 649–654. https://doi.org/10.1126/science.493968 Bernardi, P. (2013). The mitochondrial permeability transition pore: a mystery solved? . Frontiers in Physiology , Vol. 4, p. 95. Bharathi, S. S., Zhang, Y., Mohsen, A.-W., Uppala, R., Balasubramani, M., Schreiber, E., … Goetzman, E. S. (2013). Sirtuin 3 (SIRT3) Protein Regulates Long-chain Acyl-CoA Dehydrogenase by Deacetylating Conserved Lysines Near the Active Site. Journal of Biological Chemistry , 288(47), 33837–33847. https://doi.org/10.1074/jbc.M113.510354 Bickler, P. E., & Buck, L. T. (1998). Adaptations of vertebrate neurons to hypoxia and anoxia: maintaining critical Ca2+ concentrations. The Journal of Experimental Biology, 201(Pt 8), 1141–1152. Bickler, P. E., Donohoe, P. H., & Buck, L. T. (2000). Hypoxia-Induced Silencing of NMDA Receptors in Turtle Neurons. 20(10), 3522–3528. Birben, E., Sahiner, U. M., Sackesen, C., Erzurum, S., & Kalayci, O. (2012). Oxidative

Stress and Antioxidant Defense. World Allergy Organization Journal, 5(1), 9–19. https://doi.org/10.1097/WOX.0b013e3182439613 Blokhina, O., Virolainen, E., & Fagerstedt, K. (2003). Antioxidants, oxidative damage and oxygen deprivation stress: a review. Annals of Botany, 2(91), 179–194. Bonora, M., Bononi, A., De Marchi, E., Giorgi, C., Lebiedzinska, M., Marchi, S., … Wieckowski, M. R. (2013). Role of the c subunit of the FO ATP synthase in mitochondrial permeability transition. Cell Cycle, 12(4), 674–683. Borutaite, V., Morkuniene, R., & Brown, G. C. (1999). Release of cytochrome c from heart mitochondria is induced by high Ca2+ and peroxynitrite and is responsible for Ca2+- induced inhibition of substrate oxidation. Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease, 1453(1), 41–48. https://doi.org/https://doi.org/10.1016/S0925-4439(98)00082-9 Bottero, V., Rossi, F., Samson, M., Mari, M., Hofman, P., & Peyron, J.-F. (2001). IκB-α, the NF-κB Inhibitory Subunit, Interacts with ANT, the Mitochondrial ATP/ADP Translocator. Journal of Biological Chemistry , 276(24), 21317–21324. https://doi.org/10.1074/jbc.M005850200 Boutilier, R. G. (2001). Mechanisms of cell survival in hypoxia and hypothermia. The Journal of Experimental Biology, 204(Pt 18), 3171. https://doi.org/10.1074/jbc.273.6.3320 Brand, M. (2003). Approximate yield of ATP from glucose, designed by Donald Nicholson. Biochemistry and Molecular Biology Education, 31(1), 2–4. https://doi.org/10.1002/bmb.2003.494031010178 Brandauer, J., Andersen, M. A., Kellezi, H., Risis, S., Frøsig, C., Vienberg, S. G., & Treebak, J. T. (2015). AMP-activated protein kinase controls exercise training- and AICAR- induced increases in SIRT3 and MnSOD . Frontiers in Physiology , Vol. 6, p. 85. Retrieved from https://www.frontiersin.org/article/10.3389/fphys.2015.00085 Brookes, P. S., Levonen, A.-L., Shiva, S., Sarti, P., & Darley-Usmar, V. M. (2002). Mitochondria: regulators of signal transduction by reactive oxygen and nitrogen species1, 21Guest Editor: Harry Ischiropoulos2This article is part of a series of reviews on “Reactive Nitrogen Species, Tyrosine Nitration and Cell Signaling.” The full list .

Free Radical Biology and Medicine, 33(6), 755–764. https://doi.org/https://doi.org/10.1016/S0891-5849(02)00901-2 Brooks, P., & Storey, B. (1993). De novo protein synthesis and protein phosphorylation during anoxia and recovery in the red-eared turtle. American Journal of Physiology- Regulatory, Integrative and Comparative Physiology, 265(6), R1380–R1386. Buck, & Bickler. (1998). Adenosine and anoxia reduce N-methyl-D-aspartate receptor open probability in turtle cerebrocortex. Journal of Experimental Biology, 2(201), 289–297. Buck, L T. (2000). Succinate and alanine as anaerobic end-products in the diving turtle (Chrysemys picta bellii). Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 126(3), 409–413. https://doi.org/https://doi.org/10.1016/S0305-0491(00)00215-7 Buck, L T, Hochachka, P. W., Schön, A., & Gnaiger, E. (1993). Microcalorimetric measurement of reversible metabolic suppression induced by anoxia in isolated hepatocytes. Am J Physiol, 265(5 Pt 2), R1014-9. https://doi.org/10.1152/ajpregu.1993.265.5.R1014 Buck, Leslie T, & Pamenter, M. E. (2018). The hypoxia-tolerant vertebrate brain: Arresting synaptic activity. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 224, 61–70. https://doi.org/https://doi.org/10.1016/j.cbpb.2017.11.015 Buler, M., Andersson, U., & Hakkola, J. (2016). Who watches the watchmen? Regulation of the expression and activity of sirtuins. The FASEB Journal, 30(12), 3942–3960. https://doi.org/10.1096/fj.201600410RR Bundgaard, A., James, A. M., Gruszczyk, A. V, Martin, J., Murphy, M. P., & Fago, A. (2019). Metabolic adaptations during extreme anoxia in the turtle heart and their implications for ischemia-reperfusion injury. Scientific Reports, 9(1), 2850. https://doi.org/10.1038/s41598-019-39836-5 Bundgaard, A., James, A. M., Joyce, W., Murphy, M. P., & Fago, A. (2018). Suppression of reactive oxygen species generation in heart mitochondria from anoxic turtles: the role of complex I <em>S</em>-nitrosation. The Journal of Experimental Biology, 221(8), jeb174391. https://doi.org/10.1242/jeb.174391

Cantó, C., & Auwerx, J. (2011). Calorie Restriction: Is AMPK a Key Sensor and Effector? Physiology, 26(4), 214–224. https://doi.org/10.1152/physiol.00010.2011 Chen, C.-J., Yu-Cai Fu, W. Y., & Wang, W. (2013). SIRT3 protects cardiomyocytes from oxidative stress-mediated cell death by activating NF-κB. Biochemical and Biophysical Research Communications, 430(2), 798–803. Chen, Y., Zhang, J., Lin, Y., Lei, Q., Guan, K., Zhao, S., & Xiong, Y. (2011). Tumour suppressor SIRT3 deacetylates and activates manganese superoxide dismutase to scavenge ROS. EMBO Reports, 12(6), 534 LP – 541. https://doi.org/10.1038/embor.2011.65 Cheng, A., Yang, Y., Zhou, Y., Maharana, C., Lu, D., Peng, W., … Mattson, M. P. (2016). Mitochondrial SIRT3 Mediates Adaptive Responses of Neurons to Exercise and Metabolic and Excitatory Challenges. Cell Metabolism, 23(1), 128–142. https://doi.org/https://doi.org/10.1016/j.cmet.2015.10.013 Chien, K. R. (1999). Stress Pathways and Heart Failure. Cell, 98(5), 555–558. https://doi.org/https://doi.org/10.1016/S0092-8674(00)80043-4 Chinopoulos, C., Konràd, C., Kiss, G., Metelkin, E., Töröcsik, B., Zhang, S. F., & Starkov, A. A. (2011). Modulation of F0F1-ATP synthase activity by cyclophilin D regulates matrix adenine nucleotide levels. The FEBS Journal, 278(7), 1112–1125. Chouchani, E. T., Pell, V. R., James, A. M., Work, L. M., Saeb-Parsy, K., Frezza, C., … Murphy, M. P. (2016). A Unifying Mechanism for Mitochondrial Superoxide Production during Ischemia-Reperfusion Injury. Cell Metabolism, 23(2), 254–263. https://doi.org/10.1016/j.cmet.2015.12.009 Cimen, H., Han, M., Yang, Y., Tong, Q., Koc, H., & Koc, E. C. (2009). Regulation of succinate dehydrogenase activity by SIRT3 in mammalian mitochondria. Biochemistry, 49(2), 304–311. Clapham, D. E. (2007). Calcium Signaling. Cell, 131(6), 1047–1058. https://doi.org/https://doi.org/10.1016/j.cell.2007.11.028 Clark, V. M., & Miller Jr, A. T. (1973). Studies on anaerobic metabolism in the fresh-water turtle (Pseudemys scripta elegans). Comparative Biochemistry and Physiology Part A: Physiology, 44(1), 55–62.

Cogswell, P. C., Kashatus, D. F., Keifer, J. A., Guttridge, D. C., Reuther, J. Y., Bristow, C., … Baldwin, A. S. (2003). NF-κB and IκBα Are Found in the Mitochondria: EVIDENCE FOR REGULATION OF MITOCHONDRIAL GENE EXPRESSION BY NF-κB . Journal of Biological Chemistry , 278(5), 2963–2968. https://doi.org/10.1074/jbc.M209995200 Cui, Y., Qin, L., Wu, J., Qu, X., Hou, C., Sun, W., … Liu, J. (2015). SIRT3 Enhances Glycolysis and Proliferation in SIRT3-Expressing Gastric Cancer Cells. PLOS ONE, 10(6), e0129834. Retrieved from https://doi.org/10.1371/journal.pone.0129834 Dai, S.-H., Chen, T., Li, X., Yue, K.-Y., Luo, P., Yang, L.-K., … Jiang, X.-F. (2017). Sirt3 confers protection against neuronal ischemia by inducing autophagy: Involvement of the AMPK-mTOR pathway. Free Radical Biology and Medicine, 108, 345–353. https://doi.org/https://doi.org/10.1016/j.freeradbiomed.2017.04.005 Dawson, N., Bell, R., & Storey, K. (2013). Purification and properties of white muscle lactate dehydrogenase from the anoxia-tolerant turtle, the red-eared slider, Trachemys scripta elegans. Enzyme Research, 1–8. Delcuve, G. P., Rastegar, M., & Davie, J. R. (2009). Epigenetic control. Journal of Cellular Physiology, 219(2), 243–250. https://doi.org/10.1002/jcp.21678 Devin, A., & Rigoulet, M. (2007). Mechanisms of mitochondrial response to variations in energy demand in eukaryotic cells. American Journal of Physiology-Cell Physiology, 292(1), C52–C58. https://doi.org/10.1152/ajpcell.00208.2006 Dittenhafer-Reed, K. E., Richards, A. L., Fan, J., Smallegan, M. J., Fotuhi Siahpirani, A., Kemmerer, Z. A., … Denu, J. M. (2015). SIRT3 Mediates Multi-Tissue Coupling for Metabolic Fuel Switching. Cell Metabolism, 21(4), 637–646. https://doi.org/https://doi.org/10.1016/j.cmet.2015.03.007 Du, H., & Yan, S. (2010). Mitochondrial permeability transition pore in Alzheimer’s disease: cyclophilin D and amyloid beta. Molecular Basis of Disease, 1802(1), 198–204. Eaton, S. L., Roche, S. L., Llavero Hurtado, M., Oldknow, K. J., Farquharson, C., Gillingwater, T. H., & Wishart, T. M. (2013). Total protein analysis as a reliable loading control for quantitative fluorescent Western blotting. PLoS One, 8(8), e72457. Else, P. L., & Hulbert, A. J. (1985). An allometric comparison of the mitochondria of

mammalian and reptilian tissues: the implications for the evolution of endothermy. Journal of Comparative Physiology. B, Biochemical, Systemic, and Environmental Physiology, 156(1), 3–11. Fink, B. D., Reszka, K. J., Herlein, J. A., Mathahs, M. M., & Sivitz, W. I. (2005). Respiratory uncoupling by UCP1 and UCP2 and superoxide generation in endothelial cell mitochondria. American Journal of Physiology-Endocrinology and Metabolism, 288(1), E71–E79. https://doi.org/10.1152/ajpendo.00332.2004 Finkel, T. (2011). Signal transduction by reactive oxygen species. The Journal of Cell Biology, 194(1), 7 LP – 15. https://doi.org/10.1083/jcb.201102095 Finley, L. W. S., Carracedo, A., Lee, J., Souza, A., Egia, A., Zhang, J., … Haigis, M. C. (2011). SIRT3 Opposes Reprogramming of Cancer Cell Metabolism through HIF1α Destabilization. Cancer Cell, 19(3), 416–428. https://doi.org/https://doi.org/10.1016/j.ccr.2011.02.014 Finley, L. W. S., Haas, W., Desquiret-Dumas, V., Wallace, D. C., Procaccio, V., Gygi, S. P., & Haigis, M. C. (2011). Succinate Dehydrogenase Is a Direct Target of Sirtuin 3 Deacetylase Activity. PLOS ONE, 6(8), e23295. Retrieved from https://doi.org/10.1371/journal.pone.0023295 Freiman, R. N., & Tjian, R. (2003). Regulating the Regulators: Lysine Modifications Make Their Mark. Cell, 112(1), 11–17. https://doi.org/10.1016/S0092-8674(02)01278-3 Freitas, R. H., Ferreira, D. G., Trevenzoli, H. I., Oliveira, D. K., & De Melo Reis, A. R. (2017). Fatty Acids, Antioxidants and Physical Activity in Brain Aging. Nutrients , Vol. 9. https://doi.org/10.3390/nu9111263 Giordano, F. J. (2005). Oxygen, oxidative stress, hypoxia, and heart failure. The Journal of Clinical Investigation, 115(3), 500–508. https://doi.org/10.1172/JCI24408 Giorgio, V., Bisetto, E., Soriano, M. E., Dabbeni-Sala, F., Basso, E., Petronilli, V., … Lippe, G. (2009). Cyclophilin D modulates mitochondrial FOF1 ATP synthase by interacting with the lateral stalk of the complex. Journal of Biological Chemistry, 98(3), 382. Giralt, A., Hondares, E., Villena, J. A., Ribas, F., Díaz-Delfín, J., Giralt, M., … Villarroya, F. (2011). Peroxisome Proliferator-activated Receptor-γ Coactivator-1α Controls Transcription of the Sirt3 Gene, an Essential Component of the Thermogenic Brown

Adipocyte Phenotype. Journal of Biological Chemistry , 286(19), 16958–16966. https://doi.org/10.1074/jbc.M110.202390 Giralt, A., & Villarroya, F. (2012). SIRT3, a pivotal actor in mitochondrial functions: metabolism, cell death and aging. Biochemical Journal, 444(1), 1–10. https://doi.org/10.1042/bj20120030 Golstein, P., & Kroemer, G. (2007). Cell death by necrosis: towards a molecular definition. Trends in Biochemical Sciences, 32(1), 37–43. https://doi.org/https://doi.org/10.1016/j.tibs.2006.11.001 Green, D. R., & Reed, J. C. (1998). Mitochondria and Apoptosis. Science, 281(5381), 1309 LP – 1312. https://doi.org/10.1126/science.281.5381.1309 Greenway, S. C., & Storey, K. B. (2000). Mitogen‐activated protein kinases and anoxia tolerance in turtles. Journal of Experimental Zoology, 7(287), 477–484. Guarente, L. (2011). Sirtuins, aging, and metabolism. Cold Spring Harb Symp Quant Biol, 76, 81–90. Guseva, N. V, Taghiyev, A. F., Sturm, M. T., Rokhlin, O. W., & Cohen, M. B. (2004). Tumor Necrosis Factor–Related Apoptosis-Inducing Ligand–Mediated Activation of Mitochondria-Associated Nuclear Factor-κB in Prostatic Carcinoma Cell Lines<a id="xref-fn-1-1" class="xref-" href="#fn- 1"><sup>1</s. Molecular Cancer Research, 2(10), 574 LP – 584. Retrieved from http://mcr.aacrjournals.org/content/2/10/574.abstract Hafner, A. V., Dai, J., Gomes, A. P., Xiao, C.-Y., Palmeira, C. M., Rosenzweig, A., & Sinclair, D. A. (2010). Regulation of the mPTP by SIRT3-mediated deacetylation of CypD at lysine 166 suppresses age-related cardiac hypertrophy. Aging, 2(12), 914. Halestrap, A. P., Kerr, P. M., Javadov, S., & Woodfield, K.-Y. (1998). Elucidating the molecular mechanism of the permeability transition pore and its role in reperfusion injury of the heart. Biochimica et Biophysica Acta (BBA) - Bioenergetics, 1366(1), 79– 94. https://doi.org/https://doi.org/10.1016/S0005-2728(98)00122-4 Halestrap, A. P., & Pasdois, P. (2009). The role of the mitochondrial permeability transition pore in heart disease. Biochimica et Biophysica Acta (BBA) - Bioenergetics, 1787(11), 1402–1415. https://doi.org/https://doi.org/10.1016/j.bbabio.2008.12.017

Halestrap, A. P., Woodfield, K.-Y., & Connern, C. P. (1997). Oxidative Stress, Thiol Reagents, and Membrane Potential Modulate the Mitochondrial Permeability Transition by Affecting Nucleotide Binding to the Adenine Nucleotide Translocase. Journal of Biological Chemistry , 272(6), 3346–3354. https://doi.org/10.1074/jbc.272.6.3346 Hallows, W. C., Yu, W., Smith, B. C., Devires, M. K., Ellinger, J. J., Someya, S., … Zhao, S. (2011). Sirt3 promotes the urea cycle and fatty acid oxidation during dietary restriction. Molecular Cell, 41(2), 139–149. Hawrysh, P. J., & Buck, L. T. (2013). Anoxia-mediated calcium release through the mitochondrial permeability transition pore silences NMDA receptor currents in turtle neurons. Journal of Experimental Biology, 216(23), 4375–4387. https://doi.org/10.1242/jeb.092650 Hawrysh, P. J., & Buck, L. T. (2019). Mitochondrial matrix pH acidifies during anoxia and is maintained by the F(1)F(o)-ATPase in anoxia-tolerant painted turtle cortical neurons. FEBS Open Bio, 9(4), 571–581. https://doi.org/10.1002/2211-5463.12612 He, X., Zeng, H., & Chen, J.-X. (2016). Ablation of SIRT3 causes coronary microvascular dysfunction and impairs cardiac recovery post myocardial ischemia. International Journal of Cardiology, 215, 349–357. https://doi.org/https://doi.org/10.1016/j.ijcard.2016.04.092 Hebert, A., Dittenhafer-Reed, K., Yu, W., Bailey, D., Selen, E., Boersma, M., … Westphall, M. (2013). Calorie restriction and SIRT3 trigger global reprogramming of the mitochondrial protein acetylome. Molecular Cell, 49(1), 186–199. Herbert, C. V, & Jackson, D. C. (1985). Temperature Effects on the Responses to Prolonged Submergence in the Turtle Chrysemys picta bellii. I. Blood Acid-Base and Ionic Changes during and following Anoxic Submergence. Physiological Zoology, 58(6), 655–669. https://doi.org/10.1086/physzool.58.6.30156070 Hipkiss, A. R. (2008). Energy metabolism, altered proteins, sirtuins and ageing: converging mechanisms? Biogerontology (Dordrecht), 9(1), 49–55. https://doi.org/10.1007/s10522- 007-9110-x Hirschey, M. D., Shimazu, T., Goetzman, E., Jing, E., Schwer, B., Lombard, D. B., … Verdin, E. (2010). SIRT3 regulates mitochondrial fatty-acid oxidation by reversible

enzyme deacetylation. Nature, 464, 121. Retrieved from https://doi.org/10.1038/nature08778 Hochachka, P., Buck, L., Dollt, C., & Landt, S. (1996). Unifying theory of hypoxia tolerance: Molecular/metabolic defense and rescue mechanisms for surviving oxygen lack (oxygen sensing/hypoxia defense/turtle hepatocytes/turtle brain). Biochemistry, 93(September), 9493–9498. https://doi.org/10.1073/pnas.93.18.9493 Ivy, J. M., Klar, A. J., & Hicks, J. B. (1986). Cloning and characterization of four SIR genes of Saccharomyces cerevisiae. Molecular and Cellular Biology, 6(2), 688 LP – 702. https://doi.org/10.1128/MCB.6.2.688 Jackson, D. C. (2000). Living without oxygen : lessons from the freshwater turtle. Comparative Biochemistry and Physiology Part A: Physiology, 125, 299–315. Jackson, D. C. (2002). Hibernating without oxygen: Physiological adaptations of the painted turtle. Journal of Physiology, 543(3), 731–737. https://doi.org/10.1113/jphysiol.2002.024729 Jackson, D. C., Goldberger, Z., Visuri, S., & Armstrong, R. (1999). Ionic exchanges of turtle shell in vitro and their relevance to shell function in the anoxic turtle. The Journal of Experimental Biology, 202(5), 513–520. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/9929454 Jacobs, K. M., Pennington, J. D., Bisht, K. S., Aykin-Burns, N., Kim, H.-S., Mishra, M., … Gius, D. (2008). SIRT3 interacts with the daf-16 homolog FOXO3a in the mitochondria, as well as increases FOXO3a dependent gene expression. International Journal of Biological Sciences, 4(5), 291–299. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/18781224 Janumyan, Y. M., Sansam, C. G., Chattopadhyay, A., Cheng, N., Soucie, E. L., Penn, L. Z., … Yang, E. (2003). Bcl-xL/Bcl-2 coordinately regulates apoptosis, cell cycle arrest and cell cycle entry. The EMBO Journal, 22(20), 5459–5470. https://doi.org/10.1093/emboj/cdg533 Jeong, J.-W., Bae, M.-K., Ahn, M.-Y., Kim, S.-H., Sohn, T.-K., Bae, M.-H., … Kim, K.-W. (2002). Regulation and Destabilization of HIF-1α by ARD1-Mediated Acetylation. Cell, 111(5), 709–720. https://doi.org/https://doi.org/10.1016/S0092-8674(02)01085-1

Johnson, R. F., Witzel, I.-I., & Perkins, N. D. (2011). p53-Dependent Regulation of Mitochondrial Energy Production by the RelA Subunit of NF-κB. Cancer Research, 71(16), 5588 LP – 5597. https://doi.org/10.1158/0008-5472.CAN-10-4252 Kalogeris, T., Bao, Y., & Korthuis, R. J. (2014). Mitochondrial reactive oxygen species: A double edged sword in ischemia/reperfusion vs preconditioning. Redox Biology, 2, 702– 714. https://doi.org/https://doi.org/10.1016/j.redox.2014.05.006 Kaltschmidt, B., Kaltschmidt, C., Hofmann, T., Hehner, S., Dröge, W., & Schmitz, L. (2000). The pro- or anti-apoptotic function of NF-kappaB is determined by the nature of the apoptotic stimulus. Journal of Biochemistry, 267(12), 3828– 3835. Kang, Y., Fielden, L. F., & Stojanovski, D. (2018). Mitochondrial protein transport in health and disease. Seminars in Cell & Developmental Biology, 76, 142–153. https://doi.org/https://doi.org/10.1016/j.semcdb.2017.07.028 Kelly, D. A., & Storey, K. B. (1988). Organ-specific control of glycolysis in anoxic turtles. American Journal of Physiology: Regulatory, Integrative and Comparative Physiology, 5(255), 774–779. Khoury, G. A., Baliban, R. C., & Floudas, C. A. (2011). Proteome-wide post-translational modification statistics: frequency analysis and curation of the swiss-prot database. Scientific Reports, 1, 90. https://doi.org/10.1038/srep00090 Kobayashi, M., Takeda, K., Narita, T., Nagai, K., Okita, N., Sudo, Y., … Higami, Y. (2017). Mitochondrial intermediate peptidase is a novel regulator of sirtuin-3 activation by caloric restriction. FEBS Letters, 591(24), 4067–4073. https://doi.org/10.1002/1873- 3468.12914 Kokoszka, J. E., Waymire, K. G., Levy, S. E., Sligh, J. E., Cai, J., Jones, D. P., … Wallace, D. C. (2004). The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore. Nature, 427(6973), 461–465. https://doi.org/10.1038/nature02229 Kong, X., Wang, R., Xue, Y., Liu, X., Zhang, H., Chen, Y., … Chang, Y. (2010). Sirtuin 3, a New Target of PGC-1α, Plays an Important Role in the Suppression of ROS and Mitochondrial Biogenesis. PLOS ONE, 5(7), e11707. Retrieved from https://doi.org/10.1371/journal.pone.0011707

Koong, A. C., Chen, E. Y., & Giaccia, A. J. (1994). Hypoxia Causes the Activation of Nuclear Factor κB through the Phosphorylation of IκBα on Tyrosine Residues. Cancer Research, 54(6), 1425 LP – 1430. Retrieved from http://cancerres.aacrjournals.org/content/54/6/1425.abstract Krivoruchko, A., & Storey, K. B. (2010a). Epigenetics in anoxia tolerance: a role for histone deacetylases. Molecular and Cellular Biochemistry, 342(1), 151–161. https://doi.org/10.1007/s11010-010-0479-5 Krivoruchko, A., & Storey, K. B. (2010b). Molecular mechanisms of turtle anoxia tolerance: A role for NF-κB. Gene, 450(1), 63–69. https://doi.org/https://doi.org/10.1016/j.gene.2009.10.005 Krivoruchko, A., & Storey, K. B. (2013). Anoxia-responsive regulation of the FoxO transcription factors in freshwater turtles, Trachemys scripta elegans. Biochimica et Biophysica Acta (BBA) - General Subjects, 1830(11), 4990–4998. https://doi.org/https://doi.org/10.1016/j.bbagen.2013.06.034 Krivoruchko, A., & Storey, K. B. (2015). Turtle anoxia tolerance: Biochemistry and gene regulation. Biochimica et Biophysica Acta (BBA) - General Subjects, 1850(6), 1188– 1196. https://doi.org/https://doi.org/10.1016/j.bbagen.2015.02.001 Krnjevic, K. (1993). Membrane current activation during hypoxia in hippocampal neurons. In P. Hochachka, P. Lutz, T. Sick, M. Rosenthal, & G. van den Thilart (Eds.), Surviving Hypoxia: Mechanisms of Control and Adaptation (pp. 365–388). Boca Raton: CRC Press. Krysko, D. V, Berghe, T. Vanden, Parthoens, E., D’Herde, K., & Vandenabeele, P. B. T.-M. in E. (2008). Chapter 16 Methods for Distinguishing Apoptotic from Necrotic Cells and Measuring Their Clearance. In Programmed Cell Death,General Principles forStudying Cell Death, Part A (Vol. 442, pp. 307–341). https://doi.org/https://doi.org/10.1016/S0076-6879(08)01416-X Kumar, A., Takada, Y., Boriek, A. M., & Aggarwal, B. B. (2004). Nuclear factor-κB: its role in health and disease. Journal of Molecular Medicine, 82(7), 434–448. https://doi.org/10.1007/s00109-004-0555-y Landry, J., Sutton, A., Tafrov, S. T., Heller, R. C., Stebbins, J., Pillus, L., & Sternglanz, R.

(2000). The silencing protein SIR2 and its homologs are NAD-dependent protein deacetylases. Proceedings of the National Academy of Sciences, 97(11), 5807 LP – 5811. https://doi.org/10.1073/pnas.110148297 Lee, J.-W., Bae, S.-H., Jeong, J.-W., Kim, S.-H., & Kim, K.-W. (2004). Hypoxia-inducible factor (HIF-1)α: its protein stability and biological functions. Experimental &Amp; Molecular Medicine, 36, 1. Retrieved from https://doi.org/10.1038/emm.2004.1 Lemasters, J. J., Theruvath, T. P., Zhong, Z., & Nieminen, A. L. (2009). Mitochondrial calcium and the permeability transition in cell death. Bioenergetics, 1787(11), 1395– 1401. Lenard, N. R., & Berthoud, H.-R. (2008). Central and Peripheral Regulation of Food Intake and Physical Activity: Pathways and Genes. Obesity, 16(S3), S11–S22. https://doi.org/10.1038/oby.2008.511 Leung, A. W. C., & Halestrap, A. P. (2008). Recent progress in elucidating the molecular mechanism of the mitochondrial permeability transition pore. Biochimica et Biophysica Acta (BBA) - Bioenergetics, 1777(7), 946–952. https://doi.org/https://doi.org/10.1016/j.bbabio.2008.03.009 Li, S., Banck, M., Mujtaba, S., Zhou, M.-M., Sugrue, M. M., & Walsh, M. J. (2010). p53- Induced Growth Arrest Is Regulated by the Mitochondrial SirT3 Deacetylase. PLOS ONE, 5(5), e10486. Retrieved from https://doi.org/10.1371/journal.pone.0010486 Lombard, D., Alt, F., Cheng, H., Bunkenborg, J., Streeper, R., Mostoslavsky, R., … Yang, Y. (2007). Mammalian Sir2 homolog SIRT3 regulates global mitochondrial lysine acetylation. Molecular and Cellular Biology, 27(24), 8807–8814. https://doi.org/10.1128/MCB.01636-07 Lu, Z., Scott, I., Webster, B. R., & Sack, M. N. (2009). The Emerging Characterization of Lysine Residue Deacetylation on the Modulation of Mitochondrial Function and Cardiovascular Biology. Circulation Research, 105(9), 830–841. https://doi.org/10.1161/CIRCRESAHA.109.204974 Lutz, P. L., & Leone-Kabler, S. L. (1995). Upregulation of the GABAA/benzodiazepine receptor during anoxia in the freshwater turtle brain. American Journal of Physiology- Regulatory, Integrative and Comparative Physiology, 268(5), R1332–R1335.

Lutz, P. L., McMahon, P., Rosenthal, M., & Sick, T. J. (1984). Relationships between aerobic and anaerobic energy production in turtle brain in situ. American Journal of Physiology-Regulatory, Integrative and Comparative Physiology, 247(4), R740–R744. https://doi.org/10.1152/ajpregu.1984.247.4.R740 Lutz, P. L., Rosenthal, M., & Sick, T. J. (1985). Living without oxygen: turtle brain as a model of anaerobic metabolism. Molecular Physiology, 8, 411–425. Lutz, P., & Prentice, H. M. (2002). Sensing and Responding to Hypoxia, Molecular and Physiological Mechanisms1. Integrative and Comparative Biology, 42(3), 463–468. https://doi.org/10.1093/icb/42.3.463 Marmorstein, R. (2004). Structure and chemistry of the Sir2 family of NAD + -dependent histone/protein deactylases . Biochemical Society Transactions, 32(6), 904–909. https://doi.org/10.1042/bst0320904 Matas, J., Tien Sing Young, N., Bourcier-Lucas, C., Ascah, A., Marcil, M., Deschepper, C. F., & Burelle, Y. (2009). Increased expression and intramitochondrial translocation of cyclophilin-D associates with increased vulnerability of the permeability transition pore to stress-induced opening during compensated ventricular hypertrophy. Journal of Molecular and Cellular Cardiology, 46(3), 420–430. https://doi.org/https://doi.org/10.1016/j.yjmcc.2008.10.020 Mathias, R. A., Greco, T. M., Oberstein, A., Budayeva, H. G., Chakrabarti, R., Rowland, E. A., … Cristea, I. M. (2014). Article Sirtuin 4 Is a Lipoamidase Regulating Pyruvate Dehydrogenase Complex Activity. Cell, 159(7), 1615–1625. https://doi.org/10.1016/j.cell.2014.11.046 Mehrani, H., & Storey, K. (1995). Enzymatic control of glycogenolysis during anoxic 56 submergence in the freshwater turtle Trachemys scripta. The International Journal of Biochemistry & Cell Biology, 27(8), 821–830. Mergenthaler, P., Lindauer, U., Dienel, G. A., & Meisel, A. (2013). Sugar for the brain: the role of glucose in physiological and pathological brain function. Trends in Neurosciences, 36(10), 587–597. https://doi.org/https://doi.org/10.1016/j.tins.2013.07.001 Michishita, E., Park, J. Y., Burneskis, J. M., & Barrett, J. C. (2005). Evolutionarily

Conserved and Nonconserved Cellular Localizations and Functions of Human SIRT Proteins. Molecular Biology of the Cell, 16(10), 4623–4635. https://doi.org/10.1091/mbc.E05 Milton, S. L., Nayak, G., Kesaraju, S., Kara, L., & Prentice, H. M. (2007). Suppression of reactive oxygen species production enhances neuronal survival in vitro and in vivo in the anoxia-tolerant turtle Trachemys scripta. Journal of Neurochemistry, 101(4), 993– 1001. https://doi.org/10.1111/j.1471-4159.2007.04466.x Milton, S. L., Nayak, G., Lutz, P. L., & Prentice, H. M. (2006). Gene transcription of neuroglobin is upregulated by hypoxia and anoxia in the brain of the anoxia-tolerant turtle Trachemys scripta. Journal of Biomedical Science, 13(4), 509—514. https://doi.org/10.1007/s11373-006-9084-8 Milton, S. L., & Prentice, H. M. (2007). BEYOND ANOXIA: THE PHYSIOLOGY OF METABOLIC DOWNREGULATION AND RECOVERY IN THE ANOXIA- TOLERANT TURTLE. Comp Biochem Physiol A Mol Integr Physiol, 147(2), 277–290. https://doi.org/10.1038/s41598-017-13872-5 Min, J., Landry, J., Sternglanz, R., & Xu, R.-M. (2001). Crystal Structure of a SIR2 Homolog–NAD Complex. Cell, 105(2), 269–279. https://doi.org/10.1016/S0092- 8674(01)00317-8 Morgan, M., & Liu, Z. (2011). No crosstalk of reactive oxygen species and NF-κB signaling. Cell Research, 21(1), 103–115. Morin, P., & Storey, K. (2007). Antioxidant defense in hibernation: cloning and expression of peroxiredoxins from hibernating ground squirrels, Spermophilus tridecemlineatus. Archives of Biochemistry and Biophysics, 461(1), 59–65. Morris, G., Anderson, G., Dean, O., Berk, M., Galecki, P., Martin-Subero, M., & Maes, M. (2017). The glutathione system: a new drug target in neuroimmune disorders. Molecular Neurobiology, 50(3), 1059–1084. https://doi.org/10.2307/302397 Murphy, M. P. (2009). How mitochondria produce reactive oxygen species. Biochemical Journal, 417(1), 1 LP – 13. https://doi.org/10.1042/BJ20081386 Nayak, G., Prentice, H. M., & Milton, S. L. (2009). Role of neuroglobin in regulating reactive oxygen species in the brain of the anoxia-tolerant turtle Trachemys scripta.

Journal of Neurochemistry, 110(2), 603–612. https://doi.org/10.1111/j.1471- 4159.2009.06157.x Nilsson, G. (1992). Evidence for a Role of Gaba in Metabolic. Journal of Experimental Bilogy, 259, 243–259. Nilsson, G. E., & Lutz, P. L. (1991). Release of inhibitory neurotransmitters in response to anoxia in turtle brain. American Journal of Physiology - Regulatory, Integrative and Comparative Physiology, 261(1), R32–R37. Nishida, Y., Rardin, M. J., Carrico, C., Hellerstein, M., Gibson, B. W., Verdin, E., … Najjar, R. (2015). SIRT5 Regulates both Cytosolic and Mitochondrial Protein Malonylation with Glycolysis as a Major Resource SIRT5 Regulates both Cytosolic and Mitochondrial Protein Malonylation with Glycolysis as a Major Target. Molecular Cell, 59(2), 321–332. https://doi.org/10.1016/j.molcel.2015.05.022 Oh, H., Kwak, J. S., Yang, S., Gong, M. K., Kim, J. H., Rhee, J., … Chun, J.-. (2015). Reciprocal regulation by hypoxia-inducible factor-2α and the NAMPT-NAD+-SIRT axis in articular chondrocytes is involved in osteoarthritis. Osteoarthritis and Cartilage, 23(12), 2288–2296. https://doi.org/https://doi.org/10.1016/j.joca.2015.07.009 Olichon, A., Baricault, L., Gas, N., Guillou, E., Valette, A., Belenguer, P., & Lenaers, G. (2003). Loss of OPA1 Perturbates the Mitochondrial Inner Membrane Structure and Integrity, Leading to Cytochrome c Release and Apoptosis . Journal of Biological Chemistry , 278(10), 7743–7746. https://doi.org/10.1074/jbc.C200677200 Onyango, I., Lu, J., Rodova, M., Lezi, E., Crafter, A. B., & Swerdlow, R. H. (2010). Regulation of neuron mitochondrial biogenesis and relevance to brain health. Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease, 1802(1), 228–234. https://doi.org/https://doi.org/10.1016/j.bbadis.2009.07.014 Onyango, P., Celic, I., McCaffery, J. M., Boeke, J. D., & Feinberg, A. P. (2002). SIRT3, a human SIR2 homologue, is an NAD- dependent deacetylase localized to mitochondria. Proceedings of the National Academy of Sciences, 99(21), 13653 LP – 13658. https://doi.org/10.1073/pnas.222538099 Outeiro, T. F., Marques, O., & Kazantsev, A. (2008). Therapeutic role of sirtuins in neurodegenerative disease. Biochim Biophys Acta, 1782(6), 363–369.

Ozden, O., Park, S.-H., Wagner, B. A., Song, H. Y., Zhu, Y., Vassilopoulos, A., … Gius, D. (2014). SIRT3 deacetylates and increases pyruvate dehydrogenase activity in cancer cells. Free Radical Biology & Medicine, 76, 163–172. https://doi.org/10.1016/j.freeradbiomed.2014.08.001 Page, M. M., Peters, C. W., Staples, J. F., & Stuart, J. A. (2009). Intracellular antioxidant enzymes are not globally upregulated during hibernation in the major oxidative tissues of the 13-lined ground squirrel Spermophilus tridecemlineatus. Comparative Biochemistry and Physiology Part A: Molecular & Integrative Physiology, 152(1), 115– 122. https://doi.org/https://doi.org/10.1016/j.cbpa.2008.09.032 Palacios, O. M., Carmona, J. J., Michan, S., Chen, K. Y., Manabe, Y., Ward 3rd, J. L., … Tong, Q. (2009). Diet and exercise signals regulate SIRT3 and activate AMPK and PGC-1alpha in skeletal muscle. Aging, 1(9), 771–783. https://doi.org/10.18632/aging.100075 Paschen, W., Oláh, L., & Mies, G. (2000). Effect of Transient Focal Ischemia of Mouse Brain on Energy State and NAD Levels. Journal of Neurochemistry, 75(4), 1675–1680. https://doi.org/10.1046/j.1471-4159.2000.0751675.x Pellegrini, L., Pucci, B., Villanova, L., Marino, M. L., Marfe, G., Sansone, L., … Tafani, M. (2012). SIRT3 protects from hypoxia and staurosporine-mediated cell death by maintaining mitochondrial membrane potential and intracellular pH. Cell Death and Differentiation, 19(11), 1815–1825. https://doi.org/10.1038/cdd.2012.62 Perez-Pinzon, M. A., Chan, C. Y., Rosenthal, M., & Sick, T. J. (1992). Membrane and synaptic activity during anoxia in the isolated turtle cerebellum. American Journal of Physiology - Regulatory, Integrative and Comparative Physiology, (263), R1057– R1063. Porter, G. A., Urciuoli, W. R., Brookes, P. S., & Nadtochiy, S. M. (2014). SIRT3 deficiency exacerbates ischemia-reperfusion injury: implication for aged hearts. American Journal of Physiology. Heart and Circulatory Physiology, 306(12), H1602-9. https://doi.org/10.1152/ajpheart.00027.2014 Prentice, H., Milton, S., Scheurle, D., & Lutz, P. (2004). The Upregulation of Cognate and Inducible Heat Shock Proteins in the Anoxic Turtle Brain. Journal of Cerebral Blood

Flow & Metabolism, 27(4), 826–828. https://doi.org/10.1097/01.WCB.0000126565.27130.79 Rahman, M., Nirala, N. K., Singh, A., Zhu, L. J., Taguchi, K., Bamba, T., … Acharya, U. R. (2014). Drosophila Sirt2/mammalian SIRT3 deacetylates ATP synthase β and regulates complex V activity. The Journal of Cell Biology, 206(2), 289 LP – 305. https://doi.org/10.1083/jcb.201404118 Rice, M. E., Lee, E. J. K., & Choy, Y. (1995). High Levels of Ascorbic Acid, Not Glutathione, in the CNS of Anoxia-Tolerant Reptiles Contrasted with Levels in Anoxia- Intolerant Species. Journal of Neurochemistry, 64(4), 1790–1799. https://doi.org/10.1046/j.1471-4159.1995.64041790.x Rose, G., Dato, S., Altomare, K., Bellizzi, D., Garasto, S., Greco, V., … De Benedictis, G. (2003). Variability of the SIRT3 gene, human silent information regulator Sir2 homologue, and survivorship in the elderly. Experimental Gerontology, 38(10), 1065– 1070. https://doi.org/10.1016/S0531-5565(03)00209-2 Rothgiesser, K., Erener, S., Waibel, S., Lüscher, B., & Hottiger, M. (2010). SIRT2 regulates NF-jB dependent gene expression through deacetylation of p65 Lys310. Journal of Cellular Science, 123(4), 4251–4258. Rouble, A., & Storey, K. (2015). Characterization of the SIRT family of NAD+ -dependent protein deacetylases in the context of a mammalian model of hibernation, the thirteen- lined ground squirrel. Cryobiology, 71(2), 334–343. https://doi.org/10.1016/j.cryobiol.2015.08.009 Samant, S. A., Zhang, H. J., Hong, Z., Pillai, V. B., Sundaresan, N. R., Wolfgeher, D., … Gupta, M. P. (2014). SIRT3 Deacetylates and Activates OPA1 To Regulate Mitochondrial Dynamics during Stress. Molecular and Cellular Biology, 34(5), 807 LP – 819. https://doi.org/10.1128/MCB.01483-13 Schwer, B., & Verdin, E. (2008). Conserved metabolic regulatory functions of sirtuins. Cell Metabolism72, 72(2), 104–112. Shi, L., & Tu, B. P. (2015). Acetyl-CoA and the regulation of metabolism: mechanisms and consequences. Current Opinion in Cell Biology, 33, 125–131. https://doi.org/https://doi.org/10.1016/j.ceb.2015.02.003

Shi, T., Wang, F., Stieren, E., & Tong, Q. (2005). SIRT3, a Mitochondrial Sirtuin Deacetylase, Regulates Mitochondrial Function and Thermogenesis in Brown Adipocytes. Journal of Biological Chemistry , 280(14), 13560–13567. https://doi.org/10.1074/jbc.M414670200 Shimazu, T., Hirschey, M. D., Hua, L., Dittenhafer-Reed, K. E., Schwer, B., Lombard, D. B., … Verdin, E. (2010). SIRT3 Deacetylates Mitochondrial 3-Hydroxy-3-Methylglutaryl CoA Synthase 2 and Regulates Ketone Body Production. Cell Metabolism, 12(6), 654– 661. https://doi.org/https://doi.org/10.1016/j.cmet.2010.11.003 Shulga, N., Wilson-Smith, R., & Pastorino, J. G. (2010). Sirtuin-3 deacetylation of cyclophilin D induces dissociation of hexokinase II from the mitochondria. Journal of Cell Science, 123(894–902). Sick, T. J., Perez-Pinzon, M., Lutz, P. L., & Rosenthal, M. (1993). Maintaining coupled metabolism and membrane function in anoxic brain: a comparison between the turtle and rat. Surviving Hypoxia: Mechanisms of Control and Adaptation, 351–364. Someya, S., Yu, W., Hallows, W. C., Xu, J., Vann, J. M., Leeuwenburgh, C., … Prolla., T. A. (2010). Sirt3 mediates reduction of oxidative damage and prevention of agerelated hearing loss under caloric restriction. Cell, 143(5), 802–812. Srivastava, S., & Haigis, M. C. (2011). Role of sirtuins and calorie restriction in neuroprotection: implications in Alzheimer’s and Parkinson’s diseases. Curr Pharm Des, 17(31), 3418–3433. Storey, K. (1996). Metabolic adaptations supporting anoxia tolerance in reptiles: recent advances. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 113(1), 23–35. Storey, K. (2006). Reptile freeze tolerance: metabolism and gene expression. Cryobiology, 52(1), 1–16. Storey, K. B. (2007). Anoxia tolerance in turtles: Metabolic regulation and gene expression. Comparative Biochemistry and Physiology Part A: Molecular & Integrative Physiology, 147(2), 263–276. https://doi.org/https://doi.org/10.1016/j.cbpa.2006.03.019 Storey, K. B., & Storey, J. M. (1990). Metabolic Rate Depression and Biochemical Adaptation in Anaerobiosis, Hibernation and Estivation. The Quarterly Review of

Biology, 65(2), 145–174. Sundaresan, N., Samant, S., Pillai, V., Rajamohan, S., & Gupta, M. (2008). SIRT3 is a stressresponsive deacetylase in cardiomyocytes that protects cells from stress-mediated cell death by deacetylation of Ku70. Molecular Cell Biology, 28, 6384–6401. Tait, S. W. G., & Green, D. R. (2012). Mitochondria and cell signalling. Journal of Cell Science, 125(4), 807 LP – 815. https://doi.org/10.1242/jcs.099234 Tan, M., Peng, C., Anderson, K. A., Chhoy, P., Xie, Z., Dai, L., … Zhao, Y. (2014). Article Lysine Glutarylation Is a Protein Posttranslational Modification Regulated by SIRT5. Cell Metabolism, 19(4), 605–617. https://doi.org/10.1016/j.cmet.2014.03.014 Tanaka, T., Yoshida, M., Yokoo, H., Mizoguchi, K., & Tanaka, M. (1995). The role of ATP- sensitive potassium channels in striatal dopamine release: an in vivo microdialysis study. Pharmacology Biochemistry and Behavior, 52(4), 831–835. Tao, R., Coleman, M. ., Pennington, J. D., Ozden, O., Park, S. ., Jiang, H., … Olivier, A. K. (2010). Sirt3-mediated deacetylation of evolutionarily conserved lysine 122 regulates MnSOD activity in response to stress. Molecular Cell, 40(6), 893–904. Tauriainen, E., Luostarinen, M., Martonen, E., Finckenberg, P., Kovalainen, M., Huotari, A., … Mervaala, E. (2011). Distinct effects of calorie restriction and resveratrol on diet- induced obesity and fatty liver formation. Journal of Nutrition and Metabolism. Thomson, D. M., Herway, S. T., Fillmore, N., Kim, H., Brown, J. D., Barrow, J. R., & Winder, W. W. (2008). AMP-activated protein kinase phosphorylates transcription factors of the CREB family. Journal of Applied Physiology, 104(2), 429–438. https://doi.org/10.1152/japplphysiol.00900.2007 Tribble, D., Aw, T., & Jones, D. (1987). The pathophysiological significance of lipid peroxidation in oxidative cell injury. Hepatology, 2(7), 377–386. Tseng, A. H.-H., Wu, L.-H., Shieh, S.-S., & Wang, D. L. (2014). SIRT3 interactions with FOXO3 acetylation, phosphorylation and ubiquitinylation mediate endothelial cell responses to hypoxia. Biochemical Journal, 464(1), 157 LP – 168. https://doi.org/10.1042/BJ20140213 Ultsch, G. R., & Jackson, D. C. (1982). Long-term submergence at 3 degrees C of the turtle Chrysemys picta bellii in normoxic and severely hypoxic water. I. Survival, gas

exchange and acid-base status. The Journal of Experimental Biology, 97, 87–99. Vakifahmetoglu-Norberg, H., Ouchida, A. T., & Norberg, E. (2017). The role of mitochondria in metabolism and cell death. Biochemical and Biophysical Research Communications, 482(3), 426–431. https://doi.org/https://doi.org/10.1016/j.bbrc.2016.11.088 Vandecasteele, G., Szabadkai, G., & Rizzuto, R. (2002). Mitochondrial calcium homeostasis: Mechanisms and molecules. IUBMB Life, 52(3–5), 213–219. https://doi.org/10.1080/15216540152846028 Waitt, A. E., Reed, L., Ransom, B. R., & Brown, A. M. (2017). Emerging Roles for Glycogen in the CNS . Frontiers in Molecular Neuroscience , Vol. 10, p. 73. Retrieved from https://www.frontiersin.org/article/10.3389/fnmol.2017.00073 Wang, Q., Li, L., Li, C. Y., Pei, Z., Zhou, M., & Li, N. (2015). SIRT3 protects cells from hypoxia via PGC-1α- and MnSOD-dependent pathways. Neuroscience, 286, 109–121. https://doi.org/https://doi.org/10.1016/j.neuroscience.2014.11.045 Wang, Z., Zhang, L., Liang, Y., Zhang, C., Xu, Z., Zhang, L., … Wang, Z. (2015). Cyclic AMP Mimics the Anti-ageing Effects of Calorie Restriction by Up-Regulating Sirtuin. Scientific Reports, 5, 12012. Wasser, J. S., Freund, E. V., Gonzalez, L. A., & Jackson, D. C. (1990). Force and acid–base state of turtle cardiac tissue exposed to combined anoxia and acidosis. American Journal of Physiology, 259, R15–R20. Wei, L., Zhou, Y., Qiao, C., Ni, T., Li, Z., You, Q., … Lu, N. (2015). Oroxylin A inhibits glycolysis-dependent proliferation of human breast cancer via promoting SIRT3- mediated SOD2 transcription and HIF1α destabilization. Cell Death &Amp; Disease, 6, e1714. Retrieved from https://doi.org/10.1038/cddis.2015.86 Weir, H. J. M., Murray, T. K., Kehoe, P. G., Love, S., Verdin, E. M., O’Neill, M. J., … Balthasar, N. (2012). CNS SIRT3 Expression Is Altered by Reactive Oxygen Species and in Alzheimer’s Disease. PLOS ONE, 7(11), e48225. Retrieved from https://doi.org/10.1371/journal.pone.0048225 White, H., & Venkatesh, B. (2011). Clinical review: Ketones and brain injury. Critical Care, 15(2), 219. https://doi.org/10.1186/cc10020

Willmore, W. G., & Storey, K. (1997). Glutathione systems and anoxia tolerance in turtles. American Journal of Physiology-Regulatory, Integrative and Comparative Physiology, 273(1), R219–R225. https://doi.org/10.1152/ajpregu.1997.273.1.R219 Willmore, W., & Storey, K. (1997). Antioxidant systems and anoxia tolerance in a freshwater turtle Trachemys scripta elegans. Molecular and Cellular Biochemistry, 1– 2(170), 177–185. Xiong, Z., & Storey. (2012). Regulation of liver lactate dehydrogenase by reversible phosphorylation in response to anoxia in a freshwater turtle. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 163(2), 221–228. Xu, Y., Kiningham, K. K., Devalaraja, M. N., Yeh, C.-C., Majima, H., Kasarskis, E. J., & Daret K. St Clair. (1999). An Intronic NF-kB Element Is Essential for Induction of the Human Manganese Superoxide Dismutase Gene by Tumor Necrosis Factor-a and Interleukin-1b. DNA and Cell Biology, 18(9), 709–722. Yamashita, H., Kaneyuki, T., & Tagawa, K. (2001). Production of acetate in the liver and its utilization in peripheral tissues. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids, 1532(1), 79–87. https://doi.org/https://doi.org/10.1016/S1388- 1981(01)00117-2 Yan, Y., Wei, C., Zhang, W., Cheng, H., & Liu, J. (2006). Cross-talk between calcium and reactive oxygen species signaling. Acta Pharmacologica Sinica, 27(7), 821–826. https://doi.org/10.1111/j.1745-7254.2006.00390.x Yang, H., Yang, T., Baur, J. A., Perez, E., Matsui, T., Carmona, J. J., … Sinclair, D. A. (2007). Nutrient-Sensitive Mitochondrial NAD+ Levels Dictate Cell Survival. Cell, 130(6), 1095–1107. https://doi.org/https://doi.org/10.1016/j.cell.2007.07.035 Yano, M., Ikeda, Y., & Matsuzaki, M. (2005). Altered intracellular Ca2+ handling in heart failure. The Journal of Clinical Investigation, 115(3), 556–564. https://doi.org/10.1172/JCI24159 Yeung, F., Hoberg, J., Ramsey, C., Keller, M., Jones, D., Frye, R., & Mayo, M. (2004). Modulation of NF‐κB‐dependent transcription and cell survival by the SIRT1 deacetylase. The EMBO Journal, 23(12), 2369–2380. Zoratti, M., & Szabò, I. (1995). The mitochondrial permeability transition. Biochimica et

Biophysica Acta (BBA) - Reviews on Biomembranes, 1241(2), 139–176. https://doi.org/https://doi.org/10.1016/0304-4157(95)00003-A

APPENDIX

APPENDIX A: Dose Response Blot

mouse 2.5µg 5µg 10µg 20µg

Appendix Figure 1: Representative dose response blot

An immunoblot with brain from a 2hrs-anoxia exposed painted turtle. Lanes are as labelled, with double the protein content loaded from lanes left to right (2.5, 5, 10, 20 µg respectively). Total soluble protein was resolved on a 12% SDS-polyacrylamide gel at 100V and transferred onto a nitrocellulose membrane overnight at 30V. The blot was blocked with 5% Skim milk powder for 1hr at room temperature and incubated overnight at 4°C with primary antibody, MnSOD (Abcam, ab13533) using a concentration of 1:10000. Blot was then washed 3x15min with TBST, incubated with anti-rabbit secondary antibody for 2hrs, and washed 3x15min with TBST before enhanced chemiluminescence imaging.

APPENDIX B: Immunoblotting with heart and pectoralis tissues

A B

0hrs 2hrs 8hrs mouse 6hrs 24hrs

C D

0hrs 2hrs 8hrs mouse 6hrs 24hrs

Appendix Figure 2: CypD and MnSOD expression in heart

20µg of prepared heart samples from painted turtles were resolved on a 12% SDS- polyacrylamide gel at 100V and transferred onto a nitrocellulose membrane overnight at 30V. Blots were blocked with 5% Skim milk powder for 1hr at room temperature and incubated overnight at 4°C with primary antibody. Primary antibody used for blots (A) and (B) was CypD (Abcam, ab110324), and MnSOD (Abcam, ab13533) for (C) and (D) at a concentration of 1:1000. Blots were then washed 3x15min with TBST, incubated with anti-rabbit secondary antibody for 2hrs, and washed 3x15min with TBST before enhanced chemiluminescence imaging.

Western immunoblots with heart and pectoralis muscle were inconsistent often blank. This could be due to use of a flat tipped-homogenizer that is designed for soft tissue (Omni Tip, Omni International, F30750). Use of the saw-toothed tip (Omni Tip, Omni International, F30750H) designed for hard tissue or use of a sonicator would been more efficient at interrupting cell membranes. Although BCA assay indicated protein content in samples, without proper interruption of cell membranes, samples might have consisted of primarily actin. Furthermore, increasing protein loading for 50µg has been previously shown to generate better Western immunoblotting results.

APPENDIX C: SIRT3 Activity

SIRT3 Activity Assay (ab156067) was performed according to the manufacturer’s instructions with 20 µg cell lysate samples loaded per well in duplicate. No change in fluorescent activity was seen with Fluoro-Substrate Peptide (provided in kit). While change in fluorescence was seen with Fluoro-Deacetylated Peptide (provided in kit), there was only enough for ~10 wells.

Appendix Figure 3: SIRT3 activity levels in control (0hrs) brain

Raw fluorescence of 0hrs brain samples. Each letter in legend represents a different individual with samples run in duplicate, i.e. A01 and A02 correspond to samples from the same individual, B01 and B02 represent a different individual from A01 and A02, also run in duplicate.

Appendix Table 1: SIRT3 Activity for 0hrs and 2hrs brain

Slopes of 0hrs and 2hrs brain samples calculated based on raw fluorescent data, resulting in activity level. Data in Slope/Activity column represent activity levels per individual, averaged over 2 wells since samples were run in duplicate. Data in Average activity column represent mean activity among all individuals of the same treatment.

Treatments Slope/Activity Average activity

0hrs 3.21 3.152 4.96 2.415 2.885 2.29

2hrs 2.275 1.18125 -0.115 0.39 2.175

APPENDIX D: SOD activity assay

SOD activity

0.5 0hrs 0.4 2hrs 6hrs 0.3 6+2hrs 0.2 24hrs Activity U/mg 0.1

0.0 Brain Liver Muscle

Appendix Figure 4: Total SOD activity

Histograms exhibit mean SOD enzyme activity levels (units of activity/mg of protein) during anoxia and reoxygenation in brain, liver, and skeletal muscle tissues at assay temperature of 37°C. (Error bars represent SEM, n=3-4). Two-way ANOVA with Bonferroni post-hoc test.

Values obtained in the thesis do not coincide with those found in current literature. SOD activity levels in various organs from red eared sliders range from 15-50 U/mg (Willmore & Storey, 1997). There is a at least a 30-fold difference in activity levels between my values and those of Willmore and Storey (1997). Furthermore, after 20hrs of anoxia, authors found a decrease in liver and brain SOD activity by 30 and 15% respectively (Willmore & Storey, 1997), while my results indicated a 38 and 39% increase following 24hrs of anoxia (the most similar exposure), respectively. The significantly lower activity levels determined from my experiments is not likely due to interference of any reagents listed in the manufacturer’s instructions because homogenization was performed in buffer. It is possible repeated thaw/freezing cycles decreased enzyme activity or prolonged thawing promoted prot4ease degradation of samples.

APPENDIX E: Coomassie Normalization

All blots were normalized to Coomassie as a loading control. Normalization was performed by highlighting total lane loading in Image J version 1.816 software (https://imagej.nih.gov/ij/) measuring densitometry and normalizing to the gel’s background (Appendix Fig. 3). This yielded the Coomassie control value. Next, the target protein band densitometry was measured by selecting the corresponding band, measuring its densitometry in Image J, then normalizing this value to the film’s background (Appendix Fig. 4). This value was then divided by Coomassie control value to yield the final densitometric value.

0hrs 2hrs

Appendix Figure 5: Coomassie stained gel from the acetylated lysine immunoblots on brain in Fig. 3.1

A Coomassie stained gel loaded with four painted turtles exposed to 0hrs of anoxia and three turtles exposed to 2hrs of anoxia, as labelled. Normalization was performed by highlighting each lane (as indicated by the box in red) and measuring its densitometry and then normalizing this to the gel’s background densitometric value. Corresponding acetylated lysine stained membrane in Fig. 3.5.

80 57

22 17 mouse 0hrs 2hrs 6hrs 6+2hrs 24hrs

Appendix Figure 6: MnSOD expression in turtle liver

An ECL developed film of a membrane stained with MnSOD (Abcam, ab13533) loaded with mouse, two samples of 0hrs, 2hrs, 6hrs, 6+2hrs, and 24hrs-exposed anoxic turtle livers, as labelled. Densitometry was performed by selecting each lane’s band of interest (as indicated by the red box in the above image), measuring its densitometry and normalizing to the film’s background. Protein ladder (Cell Signaling, #14208) was loaded alongside samples and was visible on membranes. Marker was used to make ladder visible on film by aligning the film and membranes in the developing cassette.

APPENDIX F: Bioinformatics for Immunoblotting

The following bioinformatics tools were used to determine the molecular weight of Sirtuin 3 (SIRT3) in the Western painted turtle as well as the similarity to mouse SIRT3 given the primary antibody used is specific to mouse. Note that the Western painted turtle (Chrysemys picta bellii) has been fully sequenced.

The amino acid sequence was obtained for SIRT3 from NCBI (The National Center for Biotechnological Information) at https://www.ncbi.nlm.nih.gov/. An example is shown below illustrating the stepwise determinant of SIRT3 molecular weight using the Western painted turtle genome.

Appendix Figure 7: Western painted turtle SIRT3 on NCBI

Screenshot of how SIRT3 sequence and molecular weight was obtained on NCBI using the western painted turtle genome.

The coding sequence was obtained ExPASy Bioinformatics Resource Portal was used as described by Bjelleqvist et al., (1993) and the molecular weight was determined to be 43.7 kDa.

Appendix Figure 8: ExPASy

Screenshot of SIRT3 sequence inputted on ExPASy to determine the molecular weight.

The coding sequence of mouse SIRT3 was also compared to that of the Western painted turtle using NCBI Blast (https://blast.ncbi.nlm.nih.gov/Blast.cgi) by aligning the two sequences. A 73.44% identity match was found between the two animal sequences.

Appendix Figure 9: Comparison of SIRT3 amino acid sequence in Western painted turtle with mouse

Protein sequences for SIRT3 in the Western painted turtle was compared to that of the mouse (sequences obtained as in Appendix Figure 7) and inputted into the multiple alignment tool of NCBI Blast. Screenshot of output imaged above.