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1 NITRIC OXIDE-DEPENDENT REGULATION OF AMP-ACTIVATED PROTEIN KINASE AND PEROXISOME-PROLIFERATOR-ACTIVATED RECEPTOR-γ CO-ACTIVATOR 1α IN SKELETAL MUSCLE By VITOR AGNEW LIRA A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2008

Transcript of NITRIC OXIDE-DEPENDENT REGULATION OF AMP...

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NITRIC OXIDE-DEPENDENT REGULATION OF AMP-ACTIVATED PROTEIN KINASE AND PEROXISOME-PROLIFERATOR-ACTIVATED RECEPTOR-γ CO-ACTIVATOR 1α

IN SKELETAL MUSCLE

By

VITOR AGNEW LIRA

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2008

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© 2008 Vitor Agnew Lira

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To my Parents Lucia and Edson, my Sister Cinthia, my Wife Carola, my Daughter Manuella, and my Friends who always supported me at both sides of the Equator.

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ACKNOWLEDGMENTS

First and foremost, I thank my mentor Dr. David Criswell for the friendship and the

unconditional support throughout my entire doctoral studies. I also would like to thank past

members of the Criswell’s lab. I am also grateful for the friendship and support from Tammy,

Jonathan and Joel Criswell.

I thank Dr. Scott Powers and Lou Powers for the friendship and support, which started

even before I came to the University of Florida.

I would like to name some very important friends that I came to know here in Gainesville

and that were very important for me during the last few years. Bruno Maciel, McNair Bolstick,

Carmen Valero Aracama and Luca Bolstick-Valero, Gisele, Jens and Gabriel Schoene, Marcio,

Belinda, André and Rebeca Pereira, Tânia Broisler, José Francisco, Tony and João Pedro

Figueiredo, Lucio, Valéria and Júlia Gordan, Abraão, Leandra, Luca and Enzo Dopazo, Flavio,

Catarina and Victoria Soares, Cláudio, Beatriz and João Pedro Varella, Guilhermo and Karina

Matias, Keith, Vera and Monique Blanchard, Thiago Resende, Luiz Augusto (Guto) and Camila

de Paula, Tristan Hromnick, Sikitti Punak, Marcos Ivanowski, Ryan Caserta, Alvaro Gurovich,

Carolina and Benjamin Valencia, Quinlyn Soltow, Zsolt Murlasits, Andreas Kavazis thank you

so much for the friendship and support.

I am also very thankful for the friendship and support from old time friends Fernando and

Sirlaine Filgueiras, Geraldo and Michele Maranhão Neto, José Ricardo Vianna, Walace

Monteiro, Sydnei Silva and family, André Siqueira Rodrigues and family, José Antonio Duarte

and family, Mario Pitaluga and family, Flávio Ferreira Pinto and family.

Finally, I am thankful for the experiences shared with my wife, daughter and family

throughout my life and career.

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TABLE OF CONTENTS page

ACKNOWLEDGMENTS ...............................................................................................................4 

LIST OF TABLES ...........................................................................................................................7 

LIST OF FIGURES .........................................................................................................................8 

ABSTRACT ...................................................................................................................................10

CHAPTER

1 INTRODUCTION ..................................................................................................................12 

2 LITERATURE REVIEW .......................................................................................................15 

Skeletal Muscle Plasticity and Metabolic Flexibility .............................................................16 Regulation of GLUT4 Expression in Skeletal Muscle ....................................................17 Mitochondrial Biogenesis in Skeletal Muscle .................................................................18 PGC-1α-dependent Regulation of GLUT4 Expression and Mitochondrial

Biogenesis ....................................................................................................................20 AMPK-dependent Regulation of GLUT4 Expression and Mitochondrial Function .......22 Nitric Oxide Production and Impact on Skeletal Muscle Phenotype ..............................23 Summary ..........................................................................................................................25 

3 MATERIALS AND METHODS ...........................................................................................27 

Experimental Designs .............................................................................................................27 Experiments 1 and 2 (Aim 1) Hypothesis 1a and 1b .......................................................27 Experiments 3, 4, 5, 6 and 7 (Aim 2) Hypotheses 2a, 2b, 2c ..........................................27 Experiment 8 (Aim 3) Hypothesis 3 ................................................................................28 Experiments 9 and 10 (Aim 4) Hypothesis 4 ..................................................................29 Experiments 11, 12 and 13 (Aim 5) Hypotheses 5a, 5b ..................................................29 Statistical Analysis ..........................................................................................................30 

Protocols .................................................................................................................................30 Experiments 1, 3, 4, 5 and 13 ..........................................................................................30 Experiments 2 and 8 ........................................................................................................31 Experiment 6 ...................................................................................................................32 Experiment 7 ...................................................................................................................32 Experiments 9 and 10 ......................................................................................................33 Experiments 11 and 12 ....................................................................................................34 

General Methods .....................................................................................................................34 Animal Measurements .....................................................................................................34 

Body weight and Tissue weight. ..............................................................................34 Western Blot .............................................................................................................34 Nuclear and Cytosolic Fractionation ........................................................................35 

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Cell Culture Measurements .............................................................................................36 Western Blot .............................................................................................................36 Quantitative Real-time PCR .....................................................................................37 Dual Luciferase Assay .............................................................................................38 Fluorescence-Based Real-Time Measurement of NO Production ...........................38 Citrate Synthase Assay .............................................................................................39 Cell Respiration ........................................................................................................39 

4 RESULTS ...............................................................................................................................49 

Nitric Oxide, AMPK, PGC-1α and GLUT4 Expression in Skeletal Muscle .........................49 Nitric Oxide Increases αAMPK Phosphorylation and Upregulates PGC-1α and

GLUT4 Gene Expression in Myotubes........................................................................49 Both Nitric Oxide and cGMP Upregulate PGC-1α mRNA and Mitochondrial

Enzyme Activity in Myotubes. ....................................................................................50 Nitric Oxide-Dependent Upregulation of PGC-1α Gene Expression and

Mitochondrial Function Requires AMPK Activation. .................................................50 Pharmacological Evidence .......................................................................................50 Genetic Evidence ......................................................................................................51 

Presence of Either the CRE or the MEF2 site is Sufficient for the Nitric Oxide-Dependent Upregulation of PGC-1α Promoter Activity .............................................53 

Observations in eNOS(-/-) and nNOS(-/-) Mice ..................................................................53 AMPK Activation Increases NO Production in Both L6 and C2C12 Myotubes ............55 AMPK-Dependent Induction of PGC-1α and Mitochondrial Genes Requires

Endogenous NO Production in L6 Myotubes. .............................................................55 

5 DISCUSSION .........................................................................................................................76 

LIST OF REFERENCES ...............................................................................................................85 

BIOGRAPHICAL SKETCH .......................................................................................................100 

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LIST OF TABLES

Table page 4-1 Body weight and anatomical characteristics of eNOS(-/-) and nNOS(-/-) mice, as well

as of their respective wildtypes (eWT and nWT)... ...........................................................57

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LIST OF FIGURES

Figure page 3-1 Experimental designs to address whether nitric oxide (NO) transcriptionally

regulates PGC-1α expression.. ..........................................................................................41

3-2 Experimental designs to address whether nitric oxide (NO) and cGMP upregulate PGC-1α mRNA through AMPK activation.. .....................................................................42

3-3 Experimental designs to address whether nitric oxide (NO) upregulates mitochondrial function through AMPK activation.. ..........................................................43

3-4 Experimental design to address which isoform of the α catalytic subunit of AMPK is required for the NO-induced regulation of PGC-1α mRNA, ACC phosphorylation and HDAC5 phosphorylation.. ..........................................................................................44

3-5 Experimental design to test whether both MEF2- and CRE-binding sites in the PGC-1α promoter are required for the NO-induced upregulation of PGC-1α.. .........................45

3-6 Experimental designs to test whether basal levels of activation of AMPK, as well as phosphorylation, expression and localization of some of its downstream targets are altered in muscles from eNOS and nNOS knockout mice in comparison to their respective wildtypes. ..........................................................................................................46

3-7 Experimental designs to test whether AMPK activation causes an increase in endogenous NO production.. .............................................................................................47

3-8 Experimental design to test whether endogenous NO production is required for AMPK-dependent upregulation of PGC-1α mRNA and the mitochondrial genes F1ATP Synthase and Citrate Synthase.. .............................................................................48

4-1 αAMPK phosphorylation in response to NO treatments of 1 hour in L6 and C2C12 myotubes.. ..........................................................................................................................58

4-2 PGC-1α mRNA expression in L6 myotubes treated with NO donors for 3 hours, 8.5 hours and 16 hours.. ...........................................................................................................59

4-3 PGC-1α mRNA expression in C2C12 myotubes treated with NO donors for 3 hours, 8.5 hours and 16 hours.. ..........................................................................................59

4-4 GLUT4 mRNA expression in C2C12 myotubes treated with NO donors for 3 hours, 8.5 hours and 16 hours.. ..........................................................................................60

4-5 NO induces PGC-1α promoter activity in C2C12 myotubes. ...........................................60

4-6 PGC-1α mRNA expression in L6 myotubes treated for 3 hours.. .....................................61

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4-7 Chronic NO and cGMP treatments increase maximal Citrate Synthase activity in L6 myotubes.. ..........................................................................................................................61

4-8 AMPK inhibition prevents chronic NO-dependent upregulation of basal and maximal mitochondrial respiration in L6 myotubes.. .......................................................................62

4-9 Effect of α1AMPK siRNA and α2AMPK siRNA on αAMPK and PGC-1α gene expression.. ........................................................................................................................63

4-10 Knockdown of α1AMPK prevents NO-dependent increase in PGC-1α mRNA. .............65

4-11 NO induction of PGC-1α promoter activity does not require intact CRE and MEF2 sites in C2C12 myotubes.. .................................................................................................66

4-12 eNOS and nNOS expression in EDL and Soleus muscles from eNOS(-/-) and nNOS(-

/-) mice and their respective wildtypes.. .............................................................................67

4-13 Representative blots of proteins assessed in EDL and Soleus muscles of eNOS (-/-) and nNOS(-/-) mice and their respective wildtypes.. ...........................................................68

4-14 Protein expression of proteins related to AMPK signaling in the EDL muscle from eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes. .............................................69

4-15 Protein expression of proteins related to AMPK signaling in the Soleus muscle from eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes.. ............................................70

4-16 Representative blots of proteins assessed in nuclear and cytosolic fractions of the Plantaris muscle of eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes.. ............71

4-17 Protein expression of proteins related to AMPK signaling in nuclear and cytosolic fractions of the Plantaris muscle from eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes.. .........................................................................................................72

4-18 AMPK activation stimulates NO production in L6 myotubes.. .........................................73

4-19 AMPK activation stimulates NO production in C2C12 myotubes.. ..................................74

4-20 NOS inhibition blunts AMPK-dependent upregulation of PGC-1α, F1ATP Synthase and Citrate Synthase mRNAs in L6 myotubes.. ................................................................75

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

NITRIC OXIDE-DEPENDENT REGULATION OF AMP-ACTIVATED PROTEIN KINASE AND PEROXISOME-PROLIFERATOR-ACTIVATED RECEPTOR-γ CO-ACTIVATOR 1α

IN SKELETAL MUSCLE

By

Vitor Agnew Lira

December 2008 Chair: David S. Criswell Major: Health and Human Performance

Over nutrition and the associated mitochondrial dysfunction in skeletal muscle are central

players in the development of insulin resistance. PGC-1α, AMPK and NO are involved in

mitochondrial biogenesis, and tend to be downregulated in insulin resistant states. Our lab has

recently provided evidence for a positive feedback loop between NO and AMPK that upregulates

GLUT4 expression in skeletal muscle. Our central hypothesis was that NO and AMPK act

synergistically to upregulate PGC-1α mRNA expression, GLUT4 expression and stimulate

mitochondrial biogenesis. We demonstrated for the first time that NO regulates PGC-1α

transcriptionally and that normal αAMPK function is required for the NO-dependent increase in

mitochondrial respiration in skeletal muscle cells. More specifically, we provide evidence for an

important role of α1AMPK expression in the NO-mediated regulation of PGC-1α in L6

myotubes. Further, our observations suggest that NO does not require intact CRE and MEF2

sites at the PGC-1α promoter to induce its activity. In mouse skeletal muscle, ablation of the

nNOS gene results in lower expression of GLUT4 in the glycolytic EDL only, paralleled by a

trend towards reduced markers of AMPK-dependent signaling. Finally, we demonstrated that

endogenous NO production is required for AMPK-dependent upregulation of PGC-1α, F1ATP

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Synthase and Citrate Synthase in L6 myotubes. Altogether these results corroborate our

previously proposed model that NO and AMPK interact through a positive feedback loop that

impacts metabolic adaptations in skeletal muscle.

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CHAPTER 1 INTRODUCTION

A striking worldwide increase in the number of people with the metabolic syndrome, also

known as insulin resistance syndrome, has taken place due to over nutrition and physical

inactivity. This condition is associated with mitochondrial dysfunction and dysregulation of the

metabolic gene transcriptional co-factor, peroxisome-proliferator-activated receptor-γ co-

activator 1α (PGC-1α) in skeletal muscle (81, 99, 116, 131).

Nitric Oxide has emerged as a critical regulator for mitochondrial biogenesis in different

tissues. Further, endothelial Nitric Oxide Synthase (eNOS) and the neuronal isoform of NOS

(nNOS) are both reduced in muscle of rodents and humans presenting peripheral insulin

resistance (13, 128, 149, 165). AMP-activated protein kinase (AMPK) also induces

mitochondrial biogenesis in skeletal muscle, and its activity seems to be reduced in obesity and

diabetes (11, 136, 139). Although the AMPK-dependent regulation of PGC-1α, the glucose

transporter GLUT4 and mitochondrial genes is well accepted and remains a major focus of

research, the mechanisms by which NO regulates mitochondrial biogenesis remain to be defined.

It is currently accepted that the NO-dependent signal involves increased production of cyclic-

GMP (cGMP) and upregulation of the transcription cofactor PGC-1α. However, the mechanism

by which NO affects PGC-1α expression is still elusive.

Recently we have shown that both NO and cyclic GMP (cGMP), a downstream effector of

the NO-dependent signal, upregulate GLUT4 expression in skeletal muscle cells through AMPK

activation. In addition, we also observed that endogenous NO is required for the upregulation of

AMPK activity by the AMP mimetic, AICAR (89). These findings suggest that NO-induced

mitochondrial biogenesis and PGC-1α regulation could be AMPK-dependent as well. Our

central hypothesis is that NO modulates AMPK activity, upregulating PGC-1α activity and

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expression. Such signaling positively impacts GLUT4 expression and mitochondrial function.

Our experiments were designed to address the following specific aims.

Specific Aim 1: To determine whether NO transcriptionally regulates PGC-1α.

Hypothesis 1a: NO will upregulate PGC-1α mRNA in both L6 and C2C12 myotubes.

Hypothesis 1b: NO treatments will increase PGC-1α-luciferase promoter activity in C2C12.

Specific Aim 2: To test whether NO upregulates PGC-1α mRNA, and mitochondrial function through AMPK activation.

Rationale: We have shown that pharmacological inhibition of AMPK prevents NO-induced

upregulation of GLUT4. Muscle Enhancer Factor 2 (MEF2) serves as a transcription factor for

both GLUT4 and PGC-1α. Therefore, AMPK-mediated phophorylation of Histone Deacetylase

5 (HDAC5), causing its dissociation from MEF2 and exclusion from the nucleus, could be

essential for NO-induced upregulation of PGC-1α and mitochondrial genes. In addition, it has

been shown that α2 is the most abundant catalytic isoform of AMPK in adult skeletal muscle (2,

31, 78, 114), but some of the AMPK-dependent adaptations have been attributed to α1AMPK as

well (114). We will address these issues with both pharmacological inhibition and genetic

knockdown of the catalytic subunits of AMPK in L6 myotubes.

Hypothesis 2a: Specific pharmacological inhibition of AMPK will prevent the NO-dependent increase in PGC-1α mRNA.

Hypothesis 2b: Chronic NO treatment will increase Citrate Synthase Activity and improve mitochondrial function. Pharmacological inhibition of AMPK will prevent this adaptation.

Hypothesis 2c: Genetic knockdown of the α2 subunit of AMPK will be sufficient to blunt the effect of NO on AMPK activity, assessed by means of Acetyl CoA Carboxylase (ACC) phosphorylation, and will prevent the NO-mediated increase in HDAC5 phosphorylation and PGC-1α mRNA.

Specific Aim 3: To ascertain whether NO-induced upregulation of PGC-1α promoter activity requires the MEF2-binding site and/or the CRE-binding site in the promoter.

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Rationale: Both the MEF2- and the cAMP response element (CRE)-consensus sequences

play a fundamental role in the activation of the PGC-1α promoter in response to exercise and

other stimuli (4, 50, 51). We will use plasmids containing the mouse PGC-1α promoter with site-

directed mutagenesis of these sequences (i.e. ΔMEF2 and ΔCRE) to test their requirement for the

NO-induced increase in PGC-1α mRNA in C2C12.

Hypothesis 3: Only the ΔMEF2-PGC-1α promoter will be unresponsive to NO and AICAR treatments. This will provide genetic evidence for the MEF2 role in the NO/AMPK regulation of PGC-1α and consequent mitochondrial biogenesis.

Specific Aim 4: To test whether genetic ablation of either the nNOS, or the eNOS gene will affect GLUT4 and PGC-1α expression as well as basal levels of AMPK activation in skeletal muscle in vivo.

Hypothesis 4: Basal levels of AMPK activation, phosphorylation of some of its downstream targets (e.g. ACC and HDAC5), as well as PGC-1α nuclear localization and GLUT4 expression will be reduced in muscles of both eNOS and nNOS knockout animals in comparison to their respective wildtypes.

Specific Aim 5: To determine whether endogenous NO production is required for the AMP analog 5-aminoimidazole-4-carboxamide-1-β-D-ribofuranoside (AICAR)-dependent upregulation of PGC-1α and mitochondrial genes.

Rationale: AICAR, which serves as an AMP analog, has been the drug of choice to activate

AMPK in adult skeletal muscle and in muscle cell lines due to its high specificity (75, 78, 94,

132). We will test whether NOS inhibition affects AMPK-dependent signaling in L6 myotubes.

Hypothesis 5a: AMPK activation will induce increased NO production in both L6 and C2C12.

Hypothesis 5b: NOS inhibition will blunt AICAR-induced upregulation of PGC-1α, F1ATP Synthase, and Citrate Synthase mRNAs in L6 myotubes.

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CHAPTER 2 LITERATURE REVIEW

The number of people with the metabolic syndrome, also known as insulin resistance

syndrome, has been increasing dramatically over the last two decades (33, 116, 166, 169).

Approximately 85% of people diagnosed with type 2 diabetes in the United States are

overweight, and 55% are obese (97). In fact, type 2 diabetes and obesity are just two examples of

diseases comprising the metabolic syndrome, and therefore associated with insulin resistance

(116). The economic burden of these metabolic disorders is enormous. For example, the current

estimate of 150 million people with diabetes is set to rise to 220 million by 2010 (166), and the

direct medical costs in the United States alone are expected to increase from $92 billion in 2002

to $138 billion by 2020 (62).

Skeletal muscle is extremely plastic and accounts for 70-90% of glucose disposal under

insulin-stimulated conditions (27, 28, 80). Insulin resistant states in humans are generally not

associated with reductions in GLUT4 expression, which is the main glucose transporter in

skeletal muscle (5, 39, 111). Instead, the compromised capacity to transport glucose seems to

derive from a loss of sensitivity of the receptor and the associated downstream kinases to insulin

(93, 102, 131). However, it has been shown that skeletal muscle-specific overexpression of

GLUT4 improves glucose disposal in animal models of insulin resistance (86, 119, 148).

Therefore, the study of the mechanisms regulating GLUT4 expression could provide useful

insights into therapeutic targets for the treatment of insulin resistant states.

Over nutrition and the associated mitochondrial dysfunction in skeletal muscle are central

players in the development of insulin resistance (102, 109, 112, 116). Mitochondrial dysfunction

is also a common feature of both type 2 diabetes and obesity. Although it is still debatable

whether mitochondrial dysfunction is the main cause for the development of insulin resistance,

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recent studies suggest that improving mitochondrial function has a direct and positive impact on

insulin sensitivity in peripheral tissues, including skeletal muscle (84, 102). Consequently, it is

critical to understand the cellular mechanisms involved in the regulation of mitochondrial

volume and function. The focus of this dissertation is to investigate a signaling pathway

potentially linked to the regulation of both GLUT4 and mitochondrial genes in skeletal muscle.

In this chapter we will review skeletal muscle metabolic plasticity and how it affects the

whole body metabolic status under insulin stimulated conditions. In this context we will discuss

mechanisms involved in mitochondrial biogenesis and GLUT4 regulation, with special emphasis

to the roles of peroxisome proliferator-activated receptor γ coactivator 1α (PGC-1α), AMP-

activated protein kinase (AMPK), and nitric oxide (NO).

Skeletal Muscle Plasticity and Metabolic Flexibility

Skeletal muscle accounts for approximately 45% of body mass, and is the main site for

glucose disposal under insulin stimulated conditions (28, 32). During exercise glucose transport

into muscle occurs through mechanisms that are independent of the action of insulin (154, 164).

However, one bout of exercise is sufficient to transiently improve insulin sensitivity in muscle

(16, 120), which seems to persist for as long as glycogen levels remain low (16). Exercise

training on the other hand induces more pronounced phenotypic changes, which are mostly due

to upregulation of several proteins involved in both glycolytic and oxidative metabolisms. Such

improved phenotype involves, but is not exclusive to, a shift in expression from fast-to-slow

twitch-specific proteins (e.g. increased levels of type 1 myosin heavy chain and myoglobin),

increased GLUT4 expression, and improved mitochondrial function and fuel oxidation capacity

(45, 63-65, 84, 125, 144). Interestingly, physical inactivity causes a fast reversal of these

adaptations, thereby reducing insulin sensitivity and limiting metabolic flexibility in the tissue

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(108, 117). Therefore, regular physical activity has been used not only in the prevention of

metabolic disorders, such as obesity and type 2 diabetes, but also as an adjunct therapy for

insulin resistant states in general (45, 51, 56, 57, 59, 129, 145). Taken together these evidences

represent an important motivation for scientists to understand the mechanisms involved in

skeletal muscle adaptation to metabolic challenging events, such as caloric restriction and

exercise or increased contractile activity.

Regulation of GLUT4 Expression in Skeletal Muscle

Glucose transport in eukaryotic cells is mediated by a family of transporters (GLUT), and

13 isoforms have been identified and cloned (77, 79, 92). GLUT4 is the major glucose

transporter in skeletal muscle, but is also expressed in other insulin-responsive tissues, such as

white and brown adipose, and heart (15, 79, 143, 150, 155, 172). The translocation hypothesis

suggests that intracellular vesicles containing GLUT4, in response to insulin and other stimuli

translocate and fuse with the plasma membrane, thereby allowing the contact of the transporter

with the glucose molecule in the interstitial fluid (14, 24). Glucose, and other 6-carbon

carbohydrates, is transported in an ATP-independent manner, via a facilitative diffusion

mechanism (71).

The expression of GLUT4 is highly regulated in muscle, being stimulated during muscle

regeneration and in response to increased contractile activity (101, 103, 171). Exercise induces a

very rapid upregulation of GLUT4 expression in both rodent and human muscle. In rats most of

the adaptive response occurs within 18h after a single bout of exercise, with reports showing that

the entire response to chronic exercise (~twofold increase) can be achieved with two days of

exercise (66-68, 121). In humans GLUT4 expression may increase by ~100% with exercise

regimens ranging from 7 days to 14 weeks (49, 69, 70).

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Various transcription factors and cofactors are involved in GLUT4 transcription regulation,

including myocyte enhancer factor 2 (MEF2) (90, 94, 100), GLUT4 enhancer factor (GEF)

(107), PGC1-α (75, 95), MyoD (123), thyroid hormone receptor (TRalpha1) (123) and Kruppel-

like factor 15 (47). However, MEF2 seems to be the most important factor in GLUT4 regulation

since it physically interacts with many of the other factors at the GLUT4 promoter (94, 95, 123).

Several lines of evidence support the notion that MEF2 is involved in the exercise/activity-

dependent regulation of GLUT4. First, MEF2 is regulated by calcium-related pathways and is

involved in fiber-type determination in skeletal muscle (160). Second, MEF2 activity is higher in

the oxidative soleus in comparison to the glycolytic EDL muscle, which correlates with the

pattern of contractile activity of these muscles and the level of GLUT4 expression across fiber

types (101, 171). Third, mutation of the MEF2 binding site at the GLUT4 promoter drastically

reduces the activity of the promoter in skeletal muscle (101). Fourth, MEF2 activity is repressed

by its physical association with class 2 histone deacetylases (HDAC4-9). Recently HDAC5 has

been identified as a critical regulator of GLUT4 in response to AMPK activation (94). AMPK is

activated in response to energy depriving events, such as increased contractile activity. We

hypothesize that MEF2 is involved in the NO-dependent regulation of GLUT4, and that NO

regulates MEF2 binding activity by activating AMPK and inducing the associated

phosphorylation of HDAC5.

Mitochondrial Biogenesis in Skeletal Muscle

Mitochondria are unique organelles in the sense that they possess part of their own genetic

material in a circular double-stranded DNA molecule (mtDNA) (124). Mitochondrial DNA

encodes 13 protein subunits of the respiratory chain complexes involved in the electron transport

system. In addition, mtDNA encodes 22 tRNAs and 2 ribosomal RNAs. However, the vast

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majority of proteins involved in oxidative metabolism located in mitochondria are nuclear

encoded and therefore synthesized outside of the organelle and subsequently transported to the

mitochondria (21, 22, 153). As pointed out by Scarpulla in a recent review, mitochondria are

genetically semiautonomous (124), and mitochondrial nuclear dependence can be further

supported by the fact that mitochondrial transcription, translation and DNA replication are all

controlled by factors encoded in the nuclear genome and regulated at that level (9, 21, 22, 124).

Mitochondrial dysfunction in muscle has been related to neurodegenerative disease (126),

diabetes (91, 127), obesity (1) and aging (30). Conversely, the upregulation of mitochondrial

mass and function, also referred as mitochondrial biogenesis, is induced by a series of

environmental conditions involving metabolic challenges, of which exercise, cold exposure and

caloric restriction are the most used as research models (3, 4, 104, 106, 159).

Mitochondrial biogenesis is regulated by several transcription factors and cofactors, such

as mitochondrial transcription factor A (Tfam) (37, 38), mitochondrial transcription factor B

(TFB1M and TFB2M) (35, 44), nuclear respiratory factors (NRF-1 and NRF-2) (34, 124, 151),

specificity protein 1 (Sp1) (23), early growth response gene-1 (Egr-1) (74, 96), estrogen-related

receptor (ERRα) (72,73), peroxisome proliferator activated receptors (PPARα/δ) (8, 43, 46, 73),

PGC-1α (161) and others (124).

Mitochondrial function is greatly influenced by muscle contractile activity. First, at an

acute stage contraction induces a dramatic increase in substrate flux through β-oxidation and

Tricarboxylic Cycle (TCA) (84, 144). This increase in substrate flux is most likely the result of

an initial change in intracellular AMP-to-ATP ratio leading to AMPK activation. AMPK

phosphorylates and inhibits Acetyl Coa Carboxylase (ACC) causing the levels of its product

Malonyl CoA to drop sharply. As a result Carnitine Palmitoyl Transferase (CPT1) is released

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from the allosteric inhibition mediated by Malonyl CoA, and fatty-acid transport to mitochondria

is stimulated (55, 144). Secondly, chronic contraction, or exercise training, causes increases in

mitochondrial volume and improved oxidation capacity (63). More importantly, improvements in

mitochondrial number and function in skeletal muscle display a very strong correlation with

improvements in whole body insulin sensitivity in type 2 diabetics (146) and obese patients

(147).

Mitochondrial bioegenesis in muscle can be induced by calcium-dependent pathways,

AMPK activation, and NO. Although the extent of participation of each of these upstream

players in the adaptive events related to exercise, caloric restriction and cold exposure requires

more investigation, the upregulation of PGC-1α mRNA and protein levels, with subsequent

upregulation of other transcription factors such as Tfam, NRF-1 and NRF-2, seem to be a

common feature. Therefore understanding the regulation of PGC-1α by these pathways, as well

as determining the degree of redundancy among those pathways may bring about important

insights for therapeutic alternatives to improve mitochondrial function. In the following sections

we will examine the roles of PGC-1α, AMPK and NO in the regulation of mitochondrial

biogenesis and GLUT4 expression in skeletal muscle.

PGC-1α-dependent Regulation of GLUT4 Expression and Mitochondrial Biogenesis

The transcription coactivator PGC-1α stimulates transcription of nuclear- and

mitochondrial-encoded metabolic genes, as well as mitochondrial DNA replication. Such

regulation is due to the ability of PGC-1α to increase the expression and activity of Nuclear

Respiratory Factors 1 and 2 (NRF-1 and NRF-2, respectively), and Mitochondrial Transcription

Factor A (Tfam or mTFA) (160). Ectopic expression of PGC-1α in transgenic mice induces a

conversion from the fast-glycolytic type 2b/2x fibers to the mitochondria rich type 1/2a fibers

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(88). GLUT4 expression in muscle cells is also regulated by PGC-1α, which can bind to and

coactivate Muscle Enhancer Factor 2 (MEF2) in the GLUT4 promoter (95). PGC-1α is

upregulated by contraction and is involved in most of the metabolic adaptations that occur in

muscle thereafter (3, 4, 125, 162). On the other hand, insulin resistance, diabetes and moderate-

to-severe obesity are related to lower PGC-1α levels in muscle and white adipose tissue (84, 111,

134, 136).

In the last 8 to 9 years much attention has been dedicated to the study the PGC-1α

promoter and the elements that stimulate its activity. Interestingly, PGC-1α interacts with MEF2

in its own promoter and stimulates transcription, thereby controlling its own expression (50).

However, the PGC-1α promoter also has a cAMP responsive element (CRE), which together

with the MEF2 binding-site is responsible for the PGC-1α induction mediated by calcineurin

(CaN) and calcium/calmodulin dependent protein kinase IV (CAMK IV) (50). More recent

studies using site-directed mutagenesis in the PGC-1α promoter in exercising mice in vivo have

shown that actually both binding sites must be intact for the exercise induction of the gene to

take place (3, 4).

Although the upregulation of PGC-1α expression seems to be important for the adaptive

changes related to mitochondrial biogenesis and GLUT4 to occur, it does not seem to account for

the initial upregulation of some mitochondrial protein mRNAs of very short half-life induced by

increased contractile activity (159). In fact, recent studies have shown that PGC-1α is also post-

translationally regulated (75, 159). This seems to be the case of sirtuin-dependent regulation of

PGC-1α in response to caloric restriction and AMPK-dependent regulation of PGC-1α in

response to AICAR treatment (75). These evidences raise the exciting notion that PGC-1α

expression accounts for more delayed responses, as well as long-term maintenance of the level of

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expression of mitochondrial proteins and GLUT4, but that activation of existing PGC-1α

activation should also be considered in an experimental setting. Since most of PGC-1α is found

in the cytosol, but can translocate to the nucleus in response to certain stimuli (159), nuclear

localization can be used as an indirect measure of overall state of activation of the protein.

AMPK-dependent Regulation of GLUT4 Expression and Mitochondrial Function

The 5’-AMP-activated protein kinase (AMPK) is also downregulated in diseases

characterized by an unbalanced metabolic state, such as diabetes and obesity (11, 136). Due to

the adenylate kinase reaction in muscle, and also to its very low basal levels, AMP concentration

is much more dramatically affected than that of ADP or ATP in response to increased metabolic

rate. Therefore, AMP serves as a metabolic stress signal, to which AMPK is very sensitive to

(40, 52-54). AMPK is a heterotrimeric enzyme (α1, α2, β1, β2, γ1, γ2, γ3), which is activated by

both direct AMP allosteric regulation and phosphorylation of its α catalytic subunit, the latter

being mediated by an upstream AMPK kinase (157). The major AMPK kinase in skeletal muscle

is LKB-1 (82). AMPK is involved in acute and delayed responses that improve the cell capacity

to resynthesize ATP. Acutely, AMPK stimulates glucose transport and fatty acid oxidation in

skeletal muscle (6, 132). AMPK is also known to phosphorylate both the endothelial and

neuronal isoforms of nitric oxide synthase (eNOS and nNOS, respectively) (18-20, 26). The

associated delayed responses, or adaptations, to AMPK activation include the upregulation of

proteins involved in substrate availability (e.g. GLUT4) and oxidation capacity (e.g. PGC-1α

and mitochondrial genes) (158, 167). These changes represent important adaptive responses

triggered by metabolic challenges such as exercise, energy deprivation, and hypoxia (140, 158,

167, 170).

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Several studies have shown that the pattern of expression of the catalytic subunits of

AMPK (i.e. α1 and α2), as well as basal levels of AMPK activity differ between slow-twitch and

fast-twitch predominant muscles (2, 31, 114). Of note, AMPK expression is also responsive to

increased metabolic activity, with both α1 and α2 expression increasing in response to exercise

(114).

Due to its beneficial effects in muscle phenotype and metabolism, AMPK has been

considered as a pharmacological target for interventions directed to improve insulin sensitivity

(41, 55). The AMP analog 5-aminoimidazole-4carboxamide-1- -D-ribofuranoside (AICAR) is

one of several drugs that can induce phosphorylation and activation of AMPK and has been

widely used in research models investigating AMPK-dependent signaling in different tissues (75,

78, 94,132).

Nitric Oxide Production and Impact on Skeletal Muscle Phenotype

NO is a reactive nitrogen molecule that is formed enzymatically by nitric oxide synthases

(NOS), via the conversion of L-arginine to L-citrulline. Skeletal muscle expresses nNOS, eNOS,

and the inducible iNOS isoforms (118, 137). Interestingly, eNOS and nNOS expressions are

downregulated in insulin resistant states and iNOS is upregulated (42, 113, 128, 149, 165). Both

eNOS and nNOS synthesize NO at low levels, whereas iNOS expression results in much higher

NO production (76, 113, 137). Another interesting observation is the differential expression of

the constitutive NOS isoforms (i.e. eNOS and nNOS) across fiber types. eNOS is preferentially

expressed in slow-twitch fibers, whereas nNOS is more abundant in fast-twitch fibers (61, 137),

suggesting that: a) they may play different roles in NO-mediated signaling, and b) their

expression is related to the determination and/or maintenance of muscle phenotype and the

related differences in muscle physiology across fiber types. Our lab has provided some evidence

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for a role of NO in fiber type determination. NOS activity is required for functional overload-

induced increase in myosin heavy chain type 1 (MHC 1) in the plantaris muscle (133) and NO

donors induce MHC 1 expression and enhance calcium-induced nuclear accumulation of the

transcription factor Nuclear Factor of Activated T-Cells (NFAT) in skeletal muscle myotubes

(29).

NO synthesis increases during skeletal muscle contraction (7, 112, 130), and many of its

signaling effects are mediated by activation of soluble guanylate cyclase (sGC), leading to

increased production of cGMP (85, 118). However, cGMP levels seem to increase in response to

contraction mostly in fast-twitch fibers (85). Such increase correlates with nNOS expression and

is observed in eNOS(-/-), but not in nNOS(-/-) mice (85). Some evidence suggests that NO is

involved in some early responses in gene expression induced by exercise in humans (137).

Altogether these observations suggest that if NO were to play a role in exercise-related

adaptations in skeletal muscle, and the signals were conserved across species, nNOS expression

should be required. Interestingly, both NO and cGMP are involved in mitochondrial biogenesis

in different cell types (104, 105), but mitochondrial biogenesis is compromised only in eNOS(-/-)

mice in response to cold exposure and caloric restriction (104-106). More recently, it was

observed that neither nNOS nor eNOS is required for the exercise-induced increase in markers of

mitochondrial biogenesis, including PGC-1α expression (152).

We have recently shown that NO regulates GLUT4 expression via AMPK activation. Of

note is the observation that specific pharmacological activation of AMPK with AICAR, and the

subsequent induction of GLUT4, is blunted by inhibition of endogenous NO production.

Considering these findings together with reports showing that AMPK phosphorylates and

activates NOS (18-20, 26), we proposed the existence of a positive feedback mechanism between

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NO production and AMPK activity in skeletal muscle. According to this positive feedback loop,

basal levels of NO are required for normal AMPK activation and signal, and once AMPK is

activated NO production is increased, which increases AMPK activity even further (89).

However, several questions remain to be addressed. For example, it is unknown whether this

mechanism is present in adult skeletal muscle and if so, whether the involvement of nNOS and

eNOS would be related to their level of expression across different fiber types.

Skeletal muscle GLUT4 upregulation shares several similarities with mitochondrial

biogenesis, such as AMPK participation and transcription stimulation by MEF2, and PGC-1α

(95, 105, 167, 170). AMPK is known to phosphorylate Histone Deacetylase 5 (HDAC5), and

thereby mediate its dissociation from MEF2 and nuclear export. This pathway seems to be very

important for the regulation of GLUT4 expression mediated by AMPK (94). Also, it has been

shown that the HDAC5/MEF2-pathway is required for the PGC-1α regulation in cardiomyocytes

(25). At present, it remains to be tested whether NO-mediated upregulation of PGC-1α, GLUT4

and mitochondrial biogenesis are related and whether they require AMPK activation. Further, it

is still unknown whether NO transcriptionally regulates PGC-1α expression and if so, which cis-

elements at the PGC-1α promoter are required for the NO-dependent induction of the gene.

Finally, it is important to note that in the last decade a clear progress has been made in

understanding the NO-dependent signal in skeletal muscle. However, the role of the different

NOS isoforms in muscle metabolism and adaptation requires further investigation. We attempted

to address some of these issues in the experiments that integrate this dissertation.

Summary

Skeletal muscle plays a crucial role in whole body glucose homeostasis due to its large

mass and rapid response to insulin. For this reason a defect in the insulin signal within skeletal

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muscle, as observed in obese individuals, is a major determinant of peripheral insulin resistance

and the development of type 2 diabetes. Improvement of mitochondrial function and increased

expression of the glucose transporter GLUT4 have been shown to improve muscle-dependent

glucose uptake in response to insulin. NO has been associated with GLUT4 expression and

mitochondrial biogenesis. These events share some common signaling molecules, such as

AMPK activation and upregulation of PGC-1α expression and activity. Therefore we

hypothesize that NO regulates GLUT4 and PGC-1α expression via activation of AMPK. Here

we address this central question with experiments involving several models including culture of

muscle cell lines, use of siRNAs to knockdown AMPK expression, analysis of PGC-1α promoter

activity, and analysis of the expression of key proteins related to AMPK activity in adult skeletal

muscle of mice carrying null mutations of either nNOS or eNOS genes.

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CHAPTER 3 MATERIALS AND METHODS

Experimental Designs

Experiments 1 and 2 (Aim 1) Hypothesis 1a and 1b

L6 and C2C12 myoblasts were seeded, grown and differentiated in 6-well plates and 24-

well plates, respectively. L6 and C2C12 myotube cultures were randomly assigned to one of the

following groups and treated for 3, 8½, or 16 hours (Experiment 1) (Figure 3-1):

Experiment 1: 1) Control, 2) SNAP (25μM), 3) DETA-NO (50μM).

C2C12 were transiently transfected with a 2kb PGC-1α promoter-luciferase plasmid

together with pRL-CMV plasmid to control for transfection efficiency, and then randomly

assigned to one of the following groups and treated for 9 hours (C2C12, Experiment 2). Another

set of cells was transfected with the control plasmid (pGL3) together with pRL-CMV. These

cells served as a control for possible non-specific treatment effects on the luciferase gene (Figure

3-1):

Experiment 2: 1) Control, 2) DETA-NO (25μM), 3) DETA-NO (50μM).

Experiments 3, 4, 5, 6 and 7 (Aim 2) Hypotheses 2a, 2b, 2c

L6 were seeded, grown and differentiated in 6-well plates. Cells were randomly assigned

to one of the following groups and treated for either 3h for subsequent mRNA quantification

(Experiment 3, Figure 3-2), or for 48h (Experiment 4, Figure 3-2). Another set of cells was

treated for 12 hours/day during 5 consecutive days (Experiment 5) (Figure 3-3):

Experiment 3: 1) Control, 2) Compound C (40μM), 3) DETA-NO (50μM), 4) 8-Br-cGMP

(2mM), 5) Compound C (40μM) + DETA-NO (50μM);

Experiment 4: 1) Control, 2) DETA-NO (50μM), 3) ODQ (1μM), 4) ODQ (1μM) +

DETA-NO (50μM);

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Experiment 5: 1) Control, 2) SNAP (10μM), 3) 8-Br-cGMP (2mM);

Experiment 6: 1) Control, 2) DETA-NO (50μM), 3) Compound C (20μM), 4) DETA-NO

(50μM) + Compound C (20μM).

For the siRNA experiment (Experiment 6), L6 myotubes were grown and differentiated in

6-well plates. After 2 days of differentiation cells were transfected with siRNAs for 48 hours,

and then treated. Preliminary experiments were conducted to test for transfection efficiency, and

to optimize the concentration of each siRNA used (i.e. α1AMPK and α2AMPK), as well as to

establish the duration of transfection. Once those optimization experiments were concluded the

actual experiments started with treatments conducted for either 1h (protein measurements) or 3h

(mRNA measurements) (Figure 3-4):

Experiment 7: 1) Control, 2) unrelated siRNA+DETA-NO (50μM), 3) α1 siRNA+ DETA-

NO (50μM), 4) α2 siRNA+ DETA-NO (50μM), 5) α1+α2 siRNAs + DETA-NO (50μM).

Experiment 8 (Aim 3) Hypothesis 3

C2C12 were seeded, grown and differentiated in 24-well plates. Cells were transiently

transfected with either an intact 2kb PGC-1α promoter-luciferase plasmid, or with mutated

PGC1α promoters (either ΔMEF2-, or ΔCRE-luciferase plasmid) and then assigned to one of the

following groups and treated for 9 hours. Another set of cells was transfected with the control

plasmid (pGL3) together with pRL-CMV. These cells served as a control for possible non-

specific treatment effects on the PGC-1α promoter (Figure 3-5):

Experiment 8: 1) Control, 2) AICAR (0.65mM), 3) DETA-NO (50μM).

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Experiments 9 and 10 (Aim 4) Hypothesis 4

Basal levels of expression and phosphorylation status of certain proteins were compared in

Soleus and EDL muscles from knockout animals (either eNOS-/-, or nNOS-/-) and their respective

wildtypes (WTe and WTn). In addition, nuclear and cytosolic fractions of the Plantaris muscles

from the same animals were isolated and compared to provide some insight on differential

localization of certain proteins between wild-types and knockout animals (Figure 3-6):

Experiment 9: 1) WTe, 2) eNOS-/-

Experiment 10: 1) WTn, 2) nNOS-/-.

Experiments 11, 12 and 13 (Aim 5) Hypotheses 5a, 5b

L6 and C2C12 were seeded, grown and differentiated in 24-well plates. Cells were then

assigned to one of the following groups and treated for 30 minutes before DAF-FM addition to

the media, and then monitored for 50 minutes (refer to protocol for details) (Experiments 10 and

11, Figure 3-7). Of note, L-NAME and L-NMMA were added to the wells 30-40 minutes before

the addition of AICAR, and subsequent 30 minute-incubation:

Experiment 11: 1) Control, 2) AICAR (1mM), 3) AICAR (3mM), 4) Metformin (2mM)

and 5) MAHMA-NO (1μM) [performed with L6].

Experiment12: 1) Control, 2) AICAR (1mM), 3) L-NAME (1mM), 4)L-NMMA (1mM), 5)

AICAR+L-NAME, 6) AICAR+L-NMMA [performed with C2C12].

In Experiment 13, L6 were seeded, grown and differentiated in 6-well plates, and randomly

assigned to one of the following groups and treated for 16h (mRNA measurements, Figure 3-8):

Experiment 13: 1) Control, 2) AICAR (1mM), 3) L-NAME (1mM), AICAR (1mM) +L-

NAME (1mM).

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Statistical Analysis

Group sample size was determined with power analysis from our preliminary data.

Comparisons between groups were performed using Student t test and one-way ANOVA when

applicable. Fisher LSD test was used post hoc when appropriate. The Levene test for variance

homogeneity was performed a priori, and in cases where group variables presented significantly

different variances, data was log transformed before further analyses were conducted. Group

values, however, are reported on units specified in text and figures. Statistical significance was

established at P<0.05.

Protocols

Experiments 1, 3, 4, 5 and 13

L6 and C2C12 myoblasts were seeded in ~4000cells/cm2 in 10% FBS DMEM containing

5mM glucose. Differentiation was induced by switching 80% confluent cells to medium

containing 2% HoS. Treatments were performed when myotubes were evident (4-5 days of

differentiation in C2C12, and 6-7 days in L6). The doses of AICAR, SNAP, DETA-NO, L-

NAME, L-NMMA, ODQ and Compound C are based upon previous investigations using muscle

cells (36, 89, 105, 113). In groups treated with L-NAME, L-NMMA, ODQ, or Compound C,

cells were pre-incubated with the drug for 30 min before the final treatment. In Experiments 1, 3,

4 and 13, myotubes were treated in 2% HoS from 3hours to 16hours, washed in ice-cold PBS

and harvested in TRizol® for RNA isolation.

For Experiment 5, L6 cells were grown and differentiated as described above. Cells were

treated for 48 hours after 6 days of differentiation. During treatment new medium with

treatments was added every 24 hours. Immediately after the final 24 hours of treatment,

myotubes were washed once in ice-cold PBS containing 1μM Na3VO4, and then harvested in

Non-Denaturing Lysis Buffer (NDLB) containing protease and phosphatase inhibitors as

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previously described (89). As described above, cells were harvested in NDLB and Citrate

Synthase Activity was assessed on the same day (refer to General Methods for details).

Experiments 2 and 8

In experiment 2, C2C12 were grown in 24-well plates and differentiated in similar

conditions as described above for Experiments 1, 3 and 12. When myoblasts became ~90-100%

confluent they were transfected with 0.5µg/well of either the 2kb PGC-1α-luciferase reporter

plasmid (Addgene, Cambridge, MA; plasmid 8887) (50) or the pGL3 Control plasmid (Promega,

Madison, WI; cat E1741). In addition, cells were always cotransfected with 0.01µg/well of the

pRL-CMV (Promega, Madison, WI). Transfections were when C2C12 myoblasts were ~90%

confluent. Lipofectamine reagent was used as the lipid-mediated reagent for transfection

(1μL/well, Invitrogen, Carlsbad, CA; cat. 11668-027), and DNA-Lipofectamine complexes were

formed according to the manufacturer’s instructions. Before transfection, cells were washed in

serum-free Opti-MEM® (Invitrogen, Carlsbad, CA; cat. 31985), and then exposed to DNA-

Lipofectamine complexes also in serum-free Opti-MEM® for 6 hours. During this period, plates

were gently swirled every 2 hours. After that, 12%HoS Opti-MEM® was added in a volume

sufficient to bring the final concentration of serum in wells to 6%HoS. Cells were then kept in

6%HoS Opti-MEM® for 18 hours. At this point cells were switched to 2%HoS DMEM.

Treatments started 48 hours after the beginning of transfection and lasted for 9hours. Cells were

treated in DMEM supplemented with 2% horse serum and 1% penicillin-streptomycin.

Immediately after treatment, cells were harvested in Passive Lysis Buffer after being washed

once in PBS at room temperature and Dual Luciferase Assay was subsequently performed

according to the manufacturer’s instructions (Promega, Madison, WI; cat E1910).

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For Experiment 8, C2C12 will be grown, differentiated and trasnfected as described above,

with the exception that two extra reporter plasmids with site-directed mutation in the PGC-1α

promoter were transfected to independent sets of cells (i.e. ΔMEF2, ΔCRE, Addgene,

Cambridge, MA; plasmids 8888 and 8889). Generation of these plasmids, as well as the

relevance of these binding-sites in the PGC-1α promoter, has been previously documented both

in vitro and in vivo (4, 50). Cells then treated and harvested as described for Experiment 2.

Experiment 6

L6 myoblasts were seeded in 6-well plates, grown and differentiated as previously

mentioned. Treatments started after 2 days of differentiation and were performed in 2%HoS.

Treatments lasted for 12 hours and were performed in 5 consecutive days. Compound C (20μM)

was added to cells always 30-40 min before the addition of DETA-NO (50μM). After each

treatment media was withdrawn and replaced with fresh 2%HoS medium. Cells were harvested

10 hours after the last treatment. At that point cells had been in differentiation medium for 7 days

and myotubes were fully confluent.

Experiment 7

L6 cells were grown and differentiated as described above. Cells were transfected with

siRNA’s for the α1 and α2 subunits of AMPK at 2 days of differentiation (when myotubes

reached 60 to 70% confluence). Transfection was performed exactly as described for

Experiments 2 and 7. The siRNA sequences used were: AMPKα1 (GCA UAU GCU GCA GGU

AGA UdTdT, nucleotides 738 to 756, acc. no. NM019142), AMPKα2 (CGU CAU UGA UGA

UGA GGC UdTdT, nucleotides 865–883, acc. no. NM023991), and unrelated (control siRNA -

AUU CUA UCA CUA GCG UGA CUU). All siRNA oligonucleotides were purchased from

Dharmacon (Lafayette, CO) (83, 113, 156), and transfection efficiency was tested with

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fluorophore-conjugated siRNAs (siGLO® Green, cat. D-001630-01-05). Knockdown

efficiencies at mRNA and protein levels were tested at 48 hours post-transfection. All treatments

aiming at inducing NO/AMPK signaling were performed after 48h of transfection. In the

experiment assessing the phosphorylation status of AMPK and other proteins, cells were kept in

serum-free DMEM (0.5% BSA) for 4 hours before and also during the 1 hour treatments to

control for factors present in the serum that could interfere with basal phosphorylation levels of

these proteins. In the experiment assessing mRNA, cells were treated for 3hours in 2%HoS

DMEM.

Experiments 9 and 10

Young adult female mice (4 weeks old) homozygous for eNOS knockout (eNOS -/-, strain

B6.129P2-NOS3tm1Unc/J) and nNOS knockout (nNOS-/-, strain B6.12954-NOS1tm1P/h/J) allele

genes, as well as their respective wildtypes (eNOS+/+, strain C57BL/6J and nNOS+/+, strain

B6.129SF2/J) were obtained from Jackson Laboratory. Animals were housed for 2 days in the

University of Florida Animal Care Services Center according to the guidelines set forth by the

Institutional Animal Care and Use Committee. During this initial period animals were

maintained on a 12:12 hour light-dark cycle and provided food and water ad libitum. On the third

day animals were brought to the lab, and were sacrificed in the following morning. During this

period animals were kept at the same conditions described above, with the exception that animals

had access only to water within 2 hours before sacrifice. This procedure had the objective to

minimize the variability and therefore the possible confounding effects of different circulating

insulin and energy substrates levels on the basal status of phosphorylation and nuclear

localization of proteins. On the day of tissue collection, animals were anesthetized with 3-5%

Isoflurane, and sacrificed by cervical dislocation. Immediately afterwards, muscles, pancreatic

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fat pad, heart were removed (with careful dissection and isolation of the left ventricle) weighed,

frozen in liquid N2, and kept at −80°C until biochemical analyses were conducted.

Experiments 11 and 12

C2C12 and L6 were seeded in 24-well plates, grown and differentiated as previously

mentioned. After 4 days of differentiation, cells were washed once with phenol-red free and

serum-free DMEM, and were treated on this same medium. L-NAME and L-NMMA were added

to cells 30 minutes before the addition of AICAR. Cells were then incubated for further 30

minutes at 5%CO2 and 37°C. Metformin, another well known activator of AMPK (58, 168), and

MAHMA-NO were also used in the same manner, but in different wells. Immediately

afterwards, 10μM DAF-FM was added to each well while cells were protected from light.

Monitoring of fluorescence started either approximately 5 minutes after the addition of DAF-FM

and continued for 50 minutes (C2C12), or 25 minutes after the addition of DAF-FM and

continued for 20 minutes (L6). Fluorescence was measured in intervals of 5 minutes.

General Methods

Animal Measurements

Body weight and Tissue weight.

NOS knockout animals as well as their respective wildtypes were weighted after being

anesthetized with Isoflurane. Immediately after sacrifice by surgical dislocation, hind-limb

muscles were then removed together with the pancreatic fat pad, and the heart with subsequent

isolation of the left ventricle. All tissues were weighted and immediately frozen in liquid N2, and

then kept in −80°C for further analysis.

Western Blot

Muscle samples were homogenized in 20 mM Tris (pH7.5), 150 mM NaCl, 1 mM EDTA,

1 mM EGTA, 1% Nonidet P-40, 2.5 mM sodium pyrophosphate, 1 mM β-glycerol phosphate, 1

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mM sodium orthovanadate, 1 μg/ml leupeptin, 1 mM PMSF, and 10 μg/ml aprotinin containing

1% vol/vol phosphatase inhibitor (p-5726) from Sigma. Protein concentrations were assessed

using the DC Protein Assay Kit (Bio-Rad Laboratories, Richmond, CA). Aliquots of muscle

homogenates, (50-70μg) were then run in SDS-PAGE gels (either 8% or 4-20% Thermo

Scientific Pierce Precise Protein Gels) for β-actin, eNOS, nNOS, iNOS, GLUT4,

phosphor(Ser79)- and total ACC, phosphor(Thr172)- and total αAMPK, phosphor(Ser498)- and

total HDAC5. Primary antibodies were used as follows: rabbit anti-eNOS (1:500, BD

Biosciences), rabbit anti-nNOS (1:500, BD Biosciences), rabbit anti-iNOS (1:500, BD

Biosciences), rabbit anti-αAMPK and anti-phospho-αAMPK (Thr172) (1:1,500, Cell Signaling),

rabbit anti-ACC (1:1500, Cell Signaling), rabbit anti-phosphoACC (Ser 79) (1:500; Upstate),

goat anti-HDAC5 (1:1,000; Santa Cruz), rabbit anti-phosphoHDAC5 (Ser498) (1:1500; Affinity

Bioreagents), goat anti-GLUT4 (1:1200; Santa Cruz). Total levels of expression of proteins was

normalized to either β-actin (mouse anti-β-actin 1:5000, Abcam), or Ponceau Stain to control for

loading. Reactions were developed by using enhanced chemiluminescence detection reagents

(ECL Plus; Amersham Biosciences, Buckinghamshire, UK), and protein levels were determined

by densitometry (Kodak 1D Image Analysis Software version 3.6).

Nuclear and Cytosolic Fractionation

Fractionation of nuclear and cytosolic proteins was performed only in the Plantaris muscle

of knockouts and wildtypes. Fractionation was performed using a specific kit (NE-PER®,

Thermo Scientific, Pierce Biotechnology) according to the manufaturer’s intructions. Muscles

were homogenized in Cyotoplasmic Extraction Reagent I (CER I,buffer containing 1% vol/vol

protease inhibitors, and 1% vol/vol phosphatase inhibitors). The nuclear isolation buffer

contained protease and phosphatase inhibitors at the same concentrations reported above. The

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DC protein assay kit was used to quantify the protein concentrations of the samples. Appropriate

fractionation of nuclear and cytosolic proteins was confirmed by western blot and subsequent

probing of histone2B (rabbit anti-histone H2B 1:2000, Upstate) and CuZnSOD (rabbit anti-

CuZnSOD, Santa Cruz), respectively. Nuclear samples were run at 34-35μg/gel, whereas

cytosolic fractions were run at 80μg/gel. PGC-1α, αAMPK, and phospho-αAMPK were probed

in both fractions in order to determine whether ablation of either nNOS, or eNOS, would

interfere with the distribution of these proteins in skeletal muscle.

Cell Culture Measurements

Western Blot

For total protein extracts, both C2C12 and L6 cells were washed twice in ice-cold PBS and

harvested in nondenaturing lysis buffer (NDL buffer) containing 1% vol/vol Triton X-100, 0.3 M

NaCl, 0.05 M Tris base, 5 mM EDTA, 3.1 µM NaN3, 95 mM NaF, 22 µM Na3VO4, 0.1% vol/vol

protease inhibitors (p-8340), and 1% vol/vol phosphatase inhibitors (p-5726) from Sigma (St.

Louis, MO). Protein concentrations were assessed using the DC Protein Assay Kit (Bio-Rad

Laboratories, Richmond, CA). Aliquots of cell lysates, (15–30μg) were then run in SDS-PAGE

gels (either 8% or 4-20% Thermo Scientific Pierce Precise Protein Gels) and probed for β-actin,

phospho- and total ACC, phospho- and total αAMPK, and phospho- and total HDAC5 using the

same antibodies as described for the animal/muscle experiments. β-actin blots were used to

control for loading. Reactions were developed by using enhanced chemiluminescence detection

reagents (ECL Plus; Amersham Biosciences, Buckinghamshire, UK), and protein levels were

determined by densitometry (Kodak 1D Image Analysis Software version 3.6).

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Quantitative Real-time PCR

Quantification of mRNA’s from both L6 and C2C12 myotubes were performed with

specific primer and probes using TaqMan® technology designed by Applied Biosystems for

Quantitative real-time PCR as described above. RT was performed using either the Verso cDNA

Kit (Thermo Scientific, UK) or SuperScript III First-Strand Synthesis System for RT-PCR

according to the manufacturers’ instructions (Life Technologies, Carlsbad, CA). Reactions were

carried out using 1–2.5 µg of total RNA.according to the manufacturer’s instructions. Primers

and probes used were: α1AMPK (RefSeq NM_019142.1, assay no. Rn00569558_m1),

α2AMPK (RefSeq NM_023991.1, assay no. Rn00576935_m1), PGC-1α (RefSeq

NM_031347.1, assay no. Rn00580241_m1 [rat] and RefSeq NM_008904.1, assay no.

Mm00447183_m1 [mouse]), F1ATP Synthase (RefSeq NM_134364.1, assay no.

Rn01756310_g1), Citrate Synthase (RefSeq NM_130755.1, assay no. Rn00756225_m1),

GLUT4 (RefSeq NM_012751.1, assay no. Rn00562597_m1 [rat] and RefSeq NM_012751.1,

assay no. Mm00436615_m1 [mouse]), Citochrome C Oxidase (RefSeq NM_009941.2, assay no.

Mm00438289_g1) and HPRT (RefSeq NM_013556.2, assay no. Mm01545399_m1). Primer and

probe were obtained from Applied Biosystems and their sequences are proprietary and therefore

are not reported. Quantitative real-time PCR for the target genes were performed in the ABI

Prism 7700 Sequence Detection System (Applied Biosystems, Foster City, CA), using the 2-

ΔΔCT method, where CT is threshold cycle. Hypoxanthine guanine phosphoribosyl transferase

(HPRT) was used as the control gene for both cells lines. The primer and probe sequences for the

rat HPRT used are: forward, 5’-GTTGGATACAGGCCAGACTTTGT-3’; reverse, 5’-

AGTCAAGGGCATATCCAACAACAA-3’; probe, 5’-ACTTGTCTGGAATTTCA-3’.

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Dual Luciferase Assay

After the 9hr-treatments, C2C12 myotube cultures were washed once in PBS at room

temperature and then lysed by addition of 120 µl passive lysis buffer (PLB, Promega Dual-

Luciferase® Reporter Assay System) as previously described (29). Plates were rocked at room

temperature for 15 min. The lysate was then transferred to microcentrifuge tubes and centrifuged

for 30 seconds at room temperature (2000x g) to sediment cellular debris. Supernatants were

transferred to new tubes for subsequent use in the assay. Firefly luciferase (originating from

transcriptional activity of the PGC-1α, PGC-1αΔCRE, PGC-1αΔMEF2 or pGL3 vectors) and

renilla luciferase activities (originating from the constitutively active uptake-control plasmid;

pRL-CMV) were measured sequentially in the same 10-µl volume of cell lysate. A manual

luminometer (model FB12, Berthold) set to measure average light intensity in relative light units

(RLU) over a 10-s measurement period was used for this assay. The level of transcription activity

of the intact PGC-1α promoter, the mutated ones (PGC-1αΔCRE and PGC-1αΔMEF2), as well

as of pGL3 were considered as the raw firefly luciferase activity (RLU) divided by the renilla

luciferase activity (RLU). Values were always reported as relative to the average obtained for the

control group.

Fluorescence-Based Real-Time Measurement of NO Production

Intracellular NO was monitored with DAF-FM (Invitrogen, Carlsbad, CA), a pH-

insensitive fluorescent dye that emits increased fluorescence after reaction with an active

intermediate of NO formed during the spontaneous oxidation of NO to NO2– (110, 115). C2C12

myotubes were incubated at 37°C for 30 min in phenol red-free, serum-free DMEM containing

10 µM of DAF-FM diacetate. After loading was completed, cells were rinsed three times with

phenol red-free, serum-free DMEM and then placed in a SpectraMax M5 multi-detection reader

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(Molecular Devices, Sunnyvale, CA) for fluorometric analysis of live cells. NO fluorescence was

measured every 5 minutes using excitation and emission wavelengths of 488 and 520 nm,

respectively. Results consist in the difference between the fluorescence seen at a given minute

minus the background (fluorescence obtained on the first reading).

Citrate Synthase Assay

Citrate Synthase activity was assayed using a modified protocol from Srere (134). The

assay buffer (230 μL final volume) contained monobasic and dibasic potassium phosphate

buffers (36.5 and 63.5 mM, respectively), EDTA (10mM), DTNB (0.2mM), acetyl-CoA

(0.1mM) and Triton X-100 (0.05% v/v). The reaction was initiated by the addition of 2 μL of

cell lysate and of 10μL of oxaloacetate. Absorbance at 412 nm (25°C) was measured during 5

minutes. Values were then normalized to protein content assessed with the DC Protein Assay Kit

as previously mentioned.

Cell Respiration

L6 cells were grown, differentiated, as described for Experiment 5 in the Protocols section.

Harvesting and cell respiration measurements were performed as previously described (84).

Basically, approximately 10 hours after the last treatment, myotubes were washed once in PBS,

trypsinized, and centrifuged at 800xg for 3 minutes. Cell pellets were then washed twice and

resuspended in 400μL of supplemented PBS containing 10mM glucose, 10mM HEPES, 0.2%

BSA and pH 7.45 (10, 84). Cells were maintained at 37°C thereafter. First 300μL of

supplemented PBS was added to the oxymeter chamber (Hansatech Instruments, Norfolk, UK)

and equilibrated for 1 min. Immediately afterwards, 350 μL of the cell suspension solution was

added to the chamber. Basal respiration rate was recorded for approximately 4½ min. At this

point, Oligomycin, an inhibitor of F1/F0 ATP Synthase was added to a final concentration of

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3μg/mL and non-mitochondrial respiration was monitored for approx. 2 mins. Subsequently,

uncoupled respiration, or maximal respiration, was then monitored for 1½min. after the addition

of 6μM FCCP. The highest respiration rates (observed in 20 sec-intervals) were considered as

representative of basal respiration (initial phase of the protocol) and uncoupled respiration (after

the addition of FCCP). The most constant rates (observed in 10 sec-intervals) during the

Oligomycin incubation period were considered as representative of non-mitochondrial

respiration and these values were subtracted from basal and uncoupled respiration rates before

further analyses. Results were then normalized to protein concentration measured in the aliquot

of the cell suspension solution remaining after the respiration assay. Basically cells were lysed

by 3 consecutive freezing and thaw cycles in liquid N2. No debris was observed even after a 5

min centrifugation at 350xg. Alliquots of these cell lysates were used to measured protein

concentrations using the DC assay protein kit as described above.

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Figure 3-1. Experimental designs to address whether nitric oxide (NO) transcriptionally regulates PGC-1α expression. These experiments were used to test Hypotheses 1a and 1b from Aim 1. A) Experiment 1, both L6 and C2C12 were used, and cells were treated with either 25μM SNAP or 50μM DETA-NO, B) Experiment 2, a separate set of cells was transfected with the control vector pGL3 (not shown in the figure) and exposed to the same treatments.

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Figure 3-2. Experimental designs to address whether nitric oxide (NO) and cGMP upregulate PGC-1α mRNA through AMPK activation. These experiments were used to test Hypothesis 2a from Aim 2. A) Experiment 3, L6 myotubes were treated with DETA-NO (50μM) and the Guanylate Cyclase inhibitor (ODQ, 1μM), B) Experiment 4, L6 were treated with DETA-NO (50μM), the cGMP analog (8-Br-cGMP, referred as cGMP, 2mM) and a specific AMPK inhibitor (Compound C, referred as Cop C, 40μM).

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Figure 3-3. Experimental designs to address whether nitric oxide (NO) upregulates mitochondrial function through AMPK activation. These experiments were used to test Hypothesis 2b from Aim 2. A) Experiment 5, L6 were treated with the NO-donor SNAP and the cGMP analog (8-Br-cGMP, referred as cGMP), B) Experiment 6, L6 were treated with the NO-donor DETA-NO (50μM) alone or in combination with the AMPK inhibitor Compound C (referred as Cop C, 20μM).

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Figure 3-4. Experimental design to address which isoform of the α catalytic subunit of AMPK is required for the NO-induced regulation of PGC-1α mRNA, ACC phosphorylation and HDAC5 phosphorylation. This Experiment 7 was used to test Hypothesis 2c from Aim 2. Treatments with 50μM DETA-NO were started 48h after the beginning of siRNA transfection. Treatments lasted for 1h (protein extraction) or for 3h (mRNA isolation).

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Figure 3-5. Experimental design to test whether both MEF2- and CRE-binding sites in the PGC-1α promoter are required for the NO-induced upregulation of PGC-1α. Experiment 8 was used to test Hypothesis 3 from Aim 3. Treatments consisted of 1mM AICAR and 50μM DETA-NO. A separate set of cells was transfected with the control vector pGL3 (not shown in the figure) and exposed to the same treatments.

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Figure 3-6. Experimental designs to test whether basal levels of activation of AMPK, as well as phosphorylation, expression and localization of some of its downstream targets are altered in muscles from eNOS and nNOS knockout mice in comparison to their respective wildtypes. These experiments were used to test Hypothesis 4 from Aim 4. A) Experiment 9, muscles from eNOS(-/-) mice and their wildtype, B) Experiment 10, muscles from nNOS(-/-) mice and their wildtype.

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Figure 3-7. Experimental designs to test whether AMPK activation causes an increase in endogenous NO production. These experiments were used to test Hypothesis 5a from Aim 5. A) Experiment 11, L6 were treated with either AICAR, Metformin, or MAHMA-NO at the concentrations shown, B) Experiment 12, C2C12 were treated with AICAR (1mM), L-NAME (1mM), and L-NMMA (1mM) either alone or in combination, C) Protocol for experiments involved addition of inhibitors (L-NAME or L-NMMA) 30 minutes before the addition of AICAR to the media. DAF-FM (10μM) was added only 30 minutes after AICAR or Metformin.

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Figure 3-8. Experimental design to test whether endogenous NO production is required for AMPK-dependent upregulation of PGC-1α mRNA and the mitochondrial genes F1ATP Synthase and Citrate Synthase. Cells were treated with AICAR and L-NAME (both at 1mM) either alone or in combination. Experiment 13 was used to test Hypothesis 5b from Aim 5.

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CHAPTER 4 RESULTS

Nitric Oxide, AMPK, PGC-1α and GLUT4 Expression in Skeletal Muscle

Nitric Oxide Increases αAMPK Phosphorylation and Upregulates PGC-1α and GLUT4 Gene Expression in Myotubes.

Treatments of 1 hour with low levels of the NO-donors, SNAP and DETA-NO, caused an

increase in αAMPK phosphorylation in both L6 and C2C12 myotubes. L6 myotubes were more

sensitive to the NO-mediated effect than C2C12, with both NO donors causing a ~2-fold

increase in αAMPK phosphorylation in L6 versus a ~1.3- to 1.5-fold in C2C12 (Figure 4-1).

This trend was also observed in relation to PGC-1α mRNA, with the L6 and C2C12 presenting

2- to 3.5-fold and 1.7- to 1.9-fold increases, respectively, in response to 3 hour treatments with

the NO-donors. Although upregulation of PGC-1α mRNA was only statistically significant in

L6, in both cell types PGC-1α mRNA levels tended to remain elevated with up to 16 hours of

treatments (Figures 4-2 and 4-3). Since the responses to either SNAP or DETA-NO were very

similar in terms of αAMPK activation and induction of gene expression in both cell types, for

simplicity we elected to pool those samples for analysis of GLUT4 mRNA. A significant 5-fold

increase in GLUT4 mRNA was only observed with 16 hours of treatments (Figure 4-4). Both

PGC-1α and GLUT4 mRNA levels were back to control levels after 48 hours of treatments in

both cell types (data not shown).

To confirm that the effect of NO on PGC-1α mRNA was due to upregulation of

transcription, we transfected C2C12 with a 2kb-promoter of PGC-1α driving luciferase (45) and

performed reporter gene assays as described in the Methods section. The NO donor DETA-NO

at two different concentrations (i.e. 25μM and 50μM) was effective in upregulating PGC-1α

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promoter activity by ~ 1.8-fold. We did not observe any effect of NO treatment on the control

vector pGL3 (Figure 4-5).

Both Nitric Oxide and cGMP Upregulate PGC-1α mRNA and Mitochondrial Enzyme Activity in Myotubes.

First, we wanted to confirm that the NO-dependent effects on PGC-1α and mitochondrial

function are related to activation of soluble Guanylate Cyclase (sGC) and upregulation of

intracellular cGMP levels. Both DETA-NO and the cGMP analog (8-Br-cGMP) induced a small,

but significant, increase in PGC-1α mRNA. The DETA-NO effect was prevented by co-

treatment with the sGC inhibitor ODQ (Figure 4-6). In addition, 48-hour treatments with either

low levels of the NO donor (SNAP) or cGMP induced an approximate 40% increase in maximal

Citrate Synthase activity (Figure 4-7).

Nitric Oxide-Dependent Upregulation of PGC-1α Gene Expression and Mitochondrial Function Requires AMPK Activation.

To test whether the NO-dependent effect on PGC-1α gene expression and mitochondrial

function was AMPK-dependent, we manipulated AMPK activity both pharmacologically and

genetically.

Pharmacological Evidence

First, we observed that co-treatment of L6 myotubes with DETA-NO and the

pharmacological inhibitor of AMPK Compound C totally prevented the effect on PGC-1α

mRNA seen with the NO-donor alone (Figure 4-6). Second, DETA-NO treatment for 5 days

(12h/day) induced ~40% increase in L6 basal respiration rates, and a 15 to 20% increase in

maximal (uncoupled) respiration rates in these cells. However, co-treatment with Compound C

for the same period of time completely ablated the positive effect of low levels of chronic NO

supplementation on the mitochondrial function of these cells (Figure 4-8).

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Genetic Evidence

To inhibit the activity of the catalytic subunit of AMPK (i.e. αAMPK) we used siRNAs

for both α1AMPK and α2AMPK (α1 siRNA and α2 siRNA, respectively), which sequences had

been previously reported (83, 113, 156). The first set of experiments refers to the optimization of

transfection and knocking down of αAMPK isoforms assessed by mRNA and protein

expression. In the first experiment, we transfected L6 myotubes with a fluorophore-conjugated

siRNA and observed an approximate 90% uptake efficiency 24 hours after transfection (Figure

4-9, A). Then, we optimized the concentration of each siRNA by assessing their effect on the

mRNA levels of each isoform of αAMPK 48 hours after transfections. As shown in Figure 4-9

(B) we observed that α1 siRNA was effective in knocking down α1AMPK mRNA by 50% to

70% at concentrations ranging from 120nM to 240nM. Surprisingly, α2 siRNA also caused a

~10% to 30% reduction in α1AMPK mRNA at concentrations ranging from 120nM to 200nM.

At 240nM, however, α2 siRNA actually increased α1AMPK mRNA by 30%. To what concerns

α2AMPK mRNA, α1 siRNA had little effect on its level of expression, except at the highest

dose, which significantly reduced the expression of α2AMPK mRNA by 60% (Figure 4-9, C).

The α2 siRNA caused a 95% or higher reduction in α2AMPK mRNA in all doses tested. In the

following experiment, we tested the effect of both siRNAs used together, but at different

concentrations (Figure 4-9, D). We observed that all combinations were effective in significantly

knocking down α2AMPK mRNA, but only the combination between 120nM of α1 siRNA and

200nM of α2 siRNA was effective in knocking down α1AMPK mRNA levels. Therefore, these

concentrations were used in the subsequent experiments. In the following experiment, we tested

at which time point after transfection αAMPK protein level was reduced the most. We observed

a 60% to 70% reduction in total αAMPK protein expression at 48 h, 56h, and 64h after

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transfection (Figure 4-9, E). Based on the results of mRNA and protein expression we decided to

perform the experiments involving NO treatments starting only 48 hours after transfection. Since

our main goal was to test whether the NO-dependent upregulation of PGC-1α required

functional αAMPK subunits, we tested the isolated effect of the αAMPK siRNAs on PGC-1α

gene expression. As shown in Figure 4-9 (F), the siRNAs either together or alone did not cause

significant effects on basal PGC-1α mRNA.

In the following experiments we were then able to test whether both αAMPK isoforms

were required for the NO-dependent induction of PGC-1α. As shown in Figure 4-10 (A and B) a

significant reduction in total αAMPK expression was only observed in cells treated with either

α1siRNA, or both α1 and α2 siRNAs. This observation suggested that in L6 α1AMPK is the

mostly abundant isoform at the protein level. In fact, although expression of both genes was

easily detected by real-time PCR, we observed a ~50% higher level of expression of α1AMPK

mRNA in comparison to α2AMPK mRNA (Figure 4-10, A[inset]). However, regardless of the

degree of total αAMPK expression, DETA-NO treatment caused a 2- to 2.5-fold increase in

αAMPK phosphorylation (Figure 4-10, C). Interestingly, only when total αAMPK expression

was unaffected by siRNA (i.e. unrelated siRNA and α2siRNA) was DETA-NO able to induce a

significant increase in αAMPK activity, indirectly assessed by ACC phosphorylation (Figure 4-

10, D). NO treatments did not affect HDAC5 phosphorylation (Figure 4-10, E). Finally,

consistent with the effects of NO on AMPK activation and activity, we observed a 2-fold

increase in PGC-1α gene expression only in cells which total αAMPK levels were not reduced

(i.e. Unrel siRNA and α2 siRNA groups) (Figure 4-10, F).

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Presence of Either the CRE or the MEF2 site is Sufficient for the Nitric Oxide-Dependent Upregulation of PGC-1α Promoter Activity

It has been previously reported that AICAR, an AMP analog, at concentrations as low as

0.5mM induces αAMPK activation and subsequent upregulation of the PGC-1α promoter

activity in C2C12 (75). Therefore, we sought to test whether upregulation of PGC-1α promoter

activity mediated by specific αAMPK activation, via AICAR, and by NO would be affected

similarly by different mutations at the promoter. As shown in Figure 4-11, both AICAR

(0.65mM) and DETA-NO (50µM) increased gene reporter activity of the intact promoter by 65%

and 80%, respectively. Both treatments were again effective in increasing the activity of the

ΔCRE promoter, but the magnitude of the effect was slightly higher (i.e. 2.1- and 2-5-fold

respectively). However, AICAR did not induce an increase in the activity of the PGC-1α

promoter lacking the MEF2 binding site, whereas DETA-NO caused an approximate 75%

upregulation in the activity of this promoter.

Observations in eNOS(-/-) and nNOS(-/-) Mice

Since our cell-based experiments indicated that NO transcriptionally regulates PGC-1α

expression and that normal αAMPK expression and activity is required for this pathway to be

functional, we tested whether AMPK-dependent signaling, as well as PGC-1α and GLUT4

expression would be affected at basal conditions in mice harboring ablation of either the eNOS

or the nNOS gene.

Our first observation was that eNOS(-/-) mice did not present any clear difference in whole

body mass as well as in the mass of several skeletal muscles, heart, and pancreatic fat pad, when

compared to their wildtype (eWT, Table 4-1). Conversely, nNOS(-/-) mice presented a 27.5%

larger pancreatic fat pad and lower mass for the Soleus, Plantaris and EDL muscles by 34%,

13.8%, and 15.7%, respectively, compared to their wildtype (nWT). Next, we performed specific

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measurements of protein levels in both knockouts and compared their expression with the

respective wildtypes. The ablation of eNOS in eNOS(-/-) mice and nNOS in nNOS(-/-) mice was

confirmed by western blots in both the EDL and Soleus muscles (Figure 4-12). No over-

compensation in the expression of the other constitutive NOS isoform was observed in either

knockout.

When examining each skeletal muscle more closely we observed very few differences

between the knockouts and their wildtypes (Figures 4-13, 4-14, 4-15, 4-16, and 4-17). In the

EDL muscle, GLUT4 expression was reduced by approximately 30% in the nNOS knockouts

only, although PGC-1α expression was normal in both knockouts (Figure 4-14, A and B).

Levels of αAMPK were slightly lower in eNOS(-/-) mice, but no difference in the expression of

its surrogate ACC was observed (Figure 4-14, C and E, respectively). Phospho-to-total

αAMPK was 32% lower in nNOS(-/-) mice, almost reaching statistical significance (P=0.14,

Figure 4-14, D). This observation was paralleled by a trend of 35% reduction in ACC

phosphorylation (P=0.30, Figure 4-14, F). In the Soleus muscle no difference in GLUT4 or

PGC-1α was observed in both knockouts (Figure 4-15, A and B). The expression of αAMPK

was slightly reduced in nNOS(-/-) mice, but phospho-to-total αAMPK presented a 2.2-fold

increase (Figure 4-15, C and D). ACC expression was similar between knockouts and

wildtypes (Figure 4-15, E). Phospho-to-total ACC, however, presented a slight trend for

reduction only in the eNOS(-/-) mice (Figure 4-15, F).

It has been previously shown that both PGC-1α and αAMPK translocate to the nucleus

when activated (89, 114, 141, 159). Therefore, we tried to measure the distribution of these

proteins across nuclear and cytosolic compartments in the plantaris muscle in order to test

whether basal levels of activation of these proteins were altered in the knockout animals (Figures

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4-16 and 4-17). GLUT4 expression in the knockouts, assessed in the cytosolic fraction only, was

very similar to wildtypes (Figure 4-17, A and E). In addition, no differences in αAMPK and

phospho-to-total αAMPK were seen between knockouts and wildtypes both in the nuclear and

cytosolic fractions (Figure 4-17, C, D, G and H). Total and cytosolic PGC-1α levels were

elevated in nNOS(-/-) mice only (Figure 4-17, B and F).

AMPK Activation Increases NO Production in Both L6 and C2C12 Myotubes

In the first experiment, which was performed in L6, the specificity of the DAF-FM probe

for nitric oxide was confirmed by the fact that the highest levels of fluorescence were observed

in cells treated with the NO-donor MAHMA-NO (Figure 4-18). In addition, AICAR at 1mM and

3mM, as well as Metformin (2mM), another AMPK activator, induced significant increases in

DAF-FM fluorescence at 35 and 40 minutes of incubation with the probe. In the second

experiment, we sought to test whether the same phenomenon would be present in C2C12, and

also whether co-treatment of cells with the NOS inhibitors L-NAME and L-NMMA, both at

1mM, would prevent the AICAR-induced increase in DAF-FM fluorescence. However, in order

to avoid missing any response to DAF-FM in the first minutes following its addition to the

medium, we monitored DAF-FM fluorescence continuously during 45 minutes. As seen in

Figure 4-19, AICAR treatment induced significantly higher DAF-FM fluorescence than

AICAR+L-NMMA starting at 30 minutes of incubation. Further, DAF-FM fluorescence in

response to AICAR treatment was significantly higher than all other groups from the minute 35

until the last minute monitored (i.e. 45th minute).

AMPK-Dependent Induction of PGC-1α and Mitochondrial Genes Requires Endogenous NO Production in L6 Myotubes.

We observed that L6 myotubes treated for 16h with AICAR presented a ~10-fold increase

in PGC-1α and F1ATP Synthase mRNA levels. At the same conditions, Citrate Synthase

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mRNAs increased 4-fold. Co-treatment with the NOS inhibitor L-NAME blunted the AICAR

effect on both PGC-1α and ATP Synthase gene expression, whereas it completely prevented the

AICAR effect on Citrate Synthase mRNA (Figure 4-20).

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Table 4-1. Body weight and anatomical characteristics of eNOS(-/-) and nNOS(-/-) mice, as well as of their respective wildtypes (eWT and nWT). The average weight of left and right hindlimb muscles was considered as representative of each animal. Values are presented as mean±SE calculated within groups. Values for percent differences (% differ.) represent percent differences between values observed in knockout animals versus wildtypes for any given variable.* Significantly different from nWT.

Variable eWT eNOS(-/-) % differ. P nWT nNOS(-/-) % differ. P

Body Mass (g) 16.382±0.36 15.937±0.25 − 2.7 0.340 17.297±0.48 16.983±0.38 − 1.8 0.620 Pancreatic Fat Pad (mg) 84.283±4.48 85.367±5.14 + 1.3 0.880 76.180±5.15 97.150±6.35 + 27.5 0.028*Heart (mg) 72.017±2.11 69.967±1.41 − 2.9 0.440 70.100±3.92 68.350±3.33 − 2.5 0.740 Left Ventricle (mg) 29.033±2.07 29.117±2.09 + 0.3 0.980 25.767±2.45 29.283±1.15 + 13.7 0.220 Soleus (mg) 4.675±0.17 4.500±0.12 − 3.7 0.420 4.100±0.21 2.708±0.11 − 33.9 0.001*Plantaris (mg) 9.167±0.38 9.017±0.34 − 1.6 0.770 10.083±0.51 8.692±0.37 − 13.8 0.051 EDL (mg) 6.258±0.20 5.908±0.19 − 5.6 0.220 5.842±0.26 4.925±0.21 − 15.7 0.001*

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Figure 4-1. αAMPK phosphorylation in response to NO treatments of 1 hour in L6 and C2C12 myotubes. A) αAMPK phosphorylation in L6 myotubes, B) αAMPK phosphorylation in C2C12 myotubes. In both figures αAMPK phosphorylation is normalized to total αAMPK expression. * P<0.05 in comparison to untreated (Control) cells.

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Figure 4-2. PGC-1α mRNA expression in L6 myotubes treated with NO donors for 3 hours, 8.5 hours and 16 hours. Values represent fold changes (mean±SE) versus untreated (Control) cells (n=4-7/group). * P<0.05 in comparison to untreated (Control) cells.

Figure 4-3. PGC-1α mRNA expression in C2C12 myotubes treated with NO donors for 3 hours, 8.5 hours and 16 hours. Values represent fold changes (mean±SE) versus untreated (Control) cells (n=4-8/group). There was a trend for upregulation of PGC-1α mRNA in the NO treated groups in comparison to untreated (Control) cells at 3 hours (P<0.08).

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Figure 4-4. GLUT4 mRNA expression in C2C12 myotubes treated with NO donors for 3 hours, 8.5 hours and 16 hours. NO donors values represent fold changes (mean±SE) of samples treated with either SNAP (25μM) or DETA-NO (50μM) versus untreated (Control) cells (n=7-13/group). * P<0.05 compared to Control at the same time point.

Figure 4-5. NO induces PGC-1α promoter activity in C2C12 myotubes. Cells were treated for 9 hours with different concentrations of the NO donor DETA-NO. Values represent fold changes (mean±SE) versus untreated (Control) cells (n=4-12/group). *P<0.05 compared to Control.

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Figure 4-6. PGC-1α mRNA expression in L6 myotubes treated for 3 hours. Values represent fold changes (mean±SE) versus untreated (Control) cells (n=4-5/group). * P<0.05 in comparison to all groups except 8-Br-cGMP and ODQ+DETA-NO, #P<0.05 in comparison to all groups except DETA-NO.

Figure 4-7. Chronic NO and cGMP treatments increase maximal Citrate Synthase activity in L6 myotubes. Cells were treated for 48 hours. Values represent enzyme activity normalized to protein concentration (mean±SE, n=6/group). * P<0.05 compared to Control.

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Figure 4-8. AMPK inhibition prevents chronic NO-dependent upregulation of basal and maximal mitochondrial respiration in L6 myotubes. Values represent respiration rates normalized to protein concentration (mean±SE, n=5/group). Respiration rates observed in the presence of Oligomycin were subtracted from both basal and uncoupled respiration before protein normalization. *P<0.05 in comparison to all groups except Compound C, #P<0.05 in comparison to DETA-NO + Compound C (P=0.054 in comparison to Control).

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Figure 4-9. Effect of α1AMPK siRNA and α2AMPK siRNA on αAMPK and PGC-1α gene expression. A) Pictures of L6 myotubes treated with Lipofectamine (control vehicle) or transfected with siGLO Green siRNA for 24 hours (Merged pictures are shown, Blue - depicts nuclei stained with DAPI, Green - depicts siRNA, magnification =200x), B) Effect of either α1AMPK siRNA or α2AMPK siRNA on α1AMPK mRNA expression (n=4/group, *P<0.05 in comparison to Control [open bar], #P<0.05 in comparison to other concentrations of α2AMPK siRNA), C) Effect of either α1AMPK siRNA or α2AMPK siRNA on α2AMPK mRNA expression (n=3-4/group, *P<0.05 in comparison to Control [open bar], # P<0.05 in comparison to other concentrations of α1AMPK siRNA), D) Effect of different concentrations of α1AMPK siRNA and α2AMPK siRNA used in combination on α1AMPK mRNA and α2AMPK mRNA expression (n=4/group, *P<0.05 in comparison to Control and Lipofectamine), E) Effect of α1AMPK siRNA and α2AMPK siRNA used in combination on αAMPK protein expression at different time points after transfection (n=2/group,*P<0.05 in comparison to Control), F) Effect of α1AMPK siRNA and α2AMPK siRNA used either in combination or separately on PGC-1α mRNA expression (n=3-4/group). Except for A, in all other experiments samples were harvested 48 hours after transfection.

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Figure 4-9. Continued

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Figure 4-10. Knockdown of α1AMPK prevents NO-dependent increase in PGC-1α mRNA. A) Representative blots of proteins assessed and their phosphorylation status (Inset depicts the comparison between α1AMPK mRNA and α2AMPK mRNA in untreated cells [n=8]), B) αAMPK expression (*P<0.05 in comparison to Control, Unrel. siRNA and α2AMPK siRNA, P=0.09 between Control and α1AMPK siRNA +α2AMPK siRNA groups), C) Phospho-to-total αAMPK ratio, D)Phospho-to-total ACC ratio (*P<0.05 in comparison to Control and α1AMPK, P=0.07 between either Unrel.siRNA or α2AMPK siRNA and α1AMPK siRNA +α2AMPK siRNA groups), E) Phospho-HDAC5 expression, F) PGC-1α mRNA (*P<0.05 in comparison to Control, α1AMPK siRNA and α1AMPK siRNA +α2AMPK siRNA groups). Except for F, in all other experiments cells were treated for 1 hour after 48 hours of transfection. PGC-1α mRNA was measured after a 3h treatment with DETA-NO after 48 hours of transfection.

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Figure 4-10. Continued

Figure 4-11. NO induction of PGC-1α promoter activity does not require intact CRE and MEF2 sites in C2C12 myotubes. Cells were treated for 9 hours with either AICAR (0.65mM) or DETA-NO (50μM). Values represent fold changes (mean±SE) versus untreated (control) cells (n=12-15/group in pGL3 control & PGC-1α experiments; n=8-12 in ΔCRE & ΔMEF2 experiments). *P<0.05 compared to Control, #P<0.05 compared to Control and AICAR.

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Figure 4-12. eNOS and nNOS expression in EDL and Soleus muscles from eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes. A) Representative blots of eNOS, nNOS and Ponceau stain for EDL and Soleus muscles from eNOS(-/-) and respective wildtype, B) Representative blots of eNOS, nNOS and Ponceau stain for EDL and Soleus muscles from nNOS(-/-) and respective wildtype.

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Figure 4-13. Representative blots of proteins assessed in EDL and Soleus muscles of eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes. A) Representative blots of proteins assessed in the EDL, B) Representative blots of proteins assessed in the Soleus.

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Figure 4-14. Protein expression of proteins related to AMPK signaling in the EDL muscle from eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes. A) GLUT4 expression, B) PGC-1α expression, C) αAMPK expression, D) phospho-to-total αAMPK ratio (P=0.14 between WT and nNOS(-/-)), E) ACC expression, F) phospho-to-total ACC ratio (P=0.3 between WT and nNOS(-/-)). All proteins were normalized to Ponceau stain. *P<0.05 in comparison to respective wildtype.

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Figure 4-15. Protein expression of proteins related to AMPK signaling in the Soleus muscle from eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes. A) GLUT4 expression, B) PGC-1α expression, C) αAMPK expression, D) phospho-to-total αAMPK ratio, E) ACC expression, F) phospho-to-total ACC ratio. All proteins were normalized to Ponceau stain. *P<0.05 in comparison to respective wildtype.

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Figure 4-16. Representative blots of proteins assessed in nuclear and cytosolic fractions of the Plantaris muscle of eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes. A) Representative blots of proteins assessed in eNOS(-/-) mice and their respective wildtypes, B) Representative blots of proteins assessed in nNOS(-/-) mice and their respective wildtypes, C) Ponceau stain and representative blots of CuZnSOD and Histone H2B demonstrating successful fractionation of samples.

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Figure 4-17. Protein expression of proteins related to AMPK signaling in nuclear and cytosolic fractions of the Plantaris muscle from eNOS(-/-) and nNOS(-/-) mice and their respective wildtypes. A, B, C and D) Expression of GLUT4, PGC-1α, αAMPK, and phospho-to-total αAMPK ratio, respectively, in muscles from eNOS(-/-) mice and their respective wildtypes; E, F, G, and H) Expression of GLUT4, PGC-1α, αAMPK, and phospho-to-total αAMPK ratio, respectively, in muscles from nNOS(-/-) mice and their respective wildtypes. All proteins were normalized to Ponceau stain. *P<0.05 in comparison to respective wildtype.

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Figure 4-17. Continued

Figure 4-18. AMPK activation stimulates NO production in L6 myotubes. Values represent the fluorescence at each 5 minute-interval minus the background fluorescence assessed at 20 minutes (Δ fluorescence) and are presented as mean±SE of (n=6/group). *P<0.05 MAHMA-NO in comparison to Control, #P<0.05 AICAR (1mM and 3mM) and Metformin (2mM) in comparison to Control.

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Figure 4-19. AMPK activation stimulates NO production in C2C12 myotubes. Values represent the fluorescence at each 5 minute-interval minus the background fluorescence assessed at minute zero (Δ fluorescence) and are presented as mean±SE of (n=6/group). *P<0.05 AICAR in comparison to AICAR+L-NMMA, #P<0.05 AICAR in comparison to all other groups.

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Figure 4-20. NOS inhibition blunts AMPK-dependent upregulation of PGC-1α, F1ATP Synthase and Citrate Synthase mRNAs in L6 myotubes. Values represent fold change versus untreated (Control) cells (mean±SE, n=5-7/group). A) Effect of AICAR treatment alone or together with L-NAME on PGC-1α mRNA, B) Effect of AICAR treatment alone or together with L-NAME on F1ATP Synthase mRNA, C) Effect of AICAR treatment alone or together with L-NAME on Citrate Synthase mRNA. *P<0.05 in comparison to all other groups.

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CHAPTER 5 DISCUSSION

Several studies have shown that NO induces mitochondrial biogenesis in different tissues,

including skeletal muscle, and that PGC-1α upregulation is involved in this process (104-106).

These reports provide strong evidence for the influence of NO, and eNOS expression, on

mitochondrial function. However, they provide limited information on the mechanisms of PGC-

1α regulation in response to NO. Our lab has recently shown that NO regulates GLUT4

expression via AMPK stimulation in skeletal muscle (89). Since GLUT4 expression can also be

regulated by PGC-1α (95), and AMPK-dependent signaling induces PGC-1α expression and

activation (75, 170), our central hypothesis was that NO regulates mitochondrial biogenesis and

GLUT4 expression via AMPK activation, and subsequent induction of PGC-1α.

We addressed our central hypothesis with 5 Specific Aims, encompassing several

experiments that involved L6 and C2C12 myotubes, and adult skeletal muscle of eNOS(-/-) and

nNOS(-/-) mice as well. The reasons for studying both L6 and C2C12 are two-fold. First, L6 cells

have been frequently used in investigations involving skeletal muscle metabolic adaptations,

especially in relation to NO and AMPK signaling (83, 89, 105, 156). Second, the mechanisms

involved in PGC-1α transcription regulation have been mostly studied with gene reporter assays

using the mouse PGC-1α promoter transfected to C2C12 cells (50, 75, 95). Therefore, the use of

both cell types was intended to improve our capacity of discussion and comparison of results

with previous studies investigating similar signaling pathways.

In Specific Aim 1 we addressed whether NO would upregulate PGC-1α gene expression in

both L6 and C2C12, and whether the NO-dependent regulation of the gene would occur at the

transcription level. The first set of experiments demonstrated that NO induces PGC-1α in both

L6 and C2C12 within 3 hours of treatments, and that AMPK activation occurs very rapidly (i.e.

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within 1 hour of treatments). Although not statistically significant, PGC-1α mRNA tended to

remain increased even with longer treatments of 8.5 and 16 hours. We are unaware of other

studies looking at short-term regulation of PGC-1α mediated by NO. However, Nisoli et al. (105)

reported a comparable increase in PGC-1α mRNA (~80%) in response to a 6-day treatment with

50µM DETA-NO in L6 cells. In addition, we also demonstrated that NO donors were able to

upregulate GLUT4 mRNA with 16 hours of treatments in C2C12, similarly to what was

previously observed in L6 (89). These experiments served as proof of principle that similar

mechanisms of regulation of PGC-1α and GLUT4 gene expression were in place in these two

cell types. Of note, we used two different NO donors, namely SNAP (5h half-life at 37°C and pH

7.4) and DETA-NO (20h half-life at 37°C and pH 7.4) (12). DETA-NO releases 2 moles of NO

per mole of compound, whereas SNAP releases 1 mole of NO per mole of compound. The

combination of NO-release kinetics and initial concentrations of these two donors (i.e.25µM of

SNAP and 50µM DETA-NO) should result in very similar NO release during the first hour (~3.3

and 3.4µM at 1h, respectively) and second hour of treatments (6.1 and 6.8µM at 2h,

respectively). However, after 16 hours of treatments, 25μM SNAP will have released

approximately half as much NO as 50μM DETA-NO (i.e. 22.3 and 42.6 µM, respectively).

Therefore, considering that the overall pattern of induction of PGC-1α, upregulation of GLUT4

and activation of AMPK was very similar between NO donors, these data suggest that these

cellular responses are related to events occurring early (within 1 to 2 hours of treatments).

Further, the fact that NO upregulated PGC-1α much more rapidly than GLUT4, suggests that

increased expression of PGC-1α may be required for the NO-dependent upregulation of GLUT4

in myotubes. However, further studies are required to appropriately address this question. In

addition, our gene reporter assays showed a ~1.8-fold stimulation of the intact 2kb promoter of

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PGC-1α in response to NO treatments, which was consistent with the effects observed at the

mRNA level, also in C2C12 myotubes.

In Specific Aim 2 we tested whether the NO-dependent regulation of PGC-1α and

mitochondrial function required functional AMPK. In agreement with previous reports, we

observed that both NO and cGMP increased PGC-1α mRNA, whereas AMPK pharmacological

inhibition prevented the NO-dependent upregulation of the gene. We have recently reported that

Compound C has the same effect on the NO-dependent upregulation of GLUT4 mRNA (89).

Altogether these findings support the notion that NO upregulates PGC-1α and GLUT4 mRNAs

via AMPK activation. Our results also provide clear evidence not only for the NO- and cGMP-

dependent regulation of Citrate Synthase activity, but also for the dependence on functional

AMPK for the NO-mediated improvement in mitochondrial function in skeletal muscle

myotubes. The ~40% increase in the activity of a mitochondrial enzyme with a 48hour treatment

with either a NO donor, or a cGMP analog, is consistent with: a) ~40% increase in basal

respiration rates of L6 with a NO treatment for 5 days (12h/day), and b) the magnitude of

increase in PGC-1α mRNA. Interestingly, Nisoli et al (105) observed a 140% increase in basal

respiration rates of L6 treated with a cGMP analog continuously for 6 days. It is possible that a

continuous manipulation of a more distal player in the signal (i.e. cGMP versus NO) may induce

more pronounced adaptations. However, Nisoli et al. (105) did not report when in the

differentiation process of L6 their treatments started, making it difficult to draw any conclusion.

Koves et al. (84) reported a ~150% increase in mitochondrial respiration rates of L6 cells

transfected with an adenovirus containing the mouse PGC-1α. Therefore, our results seem to be

consistent with a lower, but physiological, degree of stimulation of this adaptation pathway.

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To gain further insight on the AMPK role, and more specifically on the role of αAMPK

catalytic subunits, in the NO-dependent regulation of PGC-1α, we performed experiments with

siRNAs for both α1AMPK and α2AMPK. A surprising finding was that although both isoforms

of αAMPK were detectable at the mRNA level, and that both siRNAs were effective in reducing

the expression of their target mRNAs, when used alone only α1siRNA caused a reduction in total

αAMPK protein expression. These findings suggest that α1AMPK is the main isoform expressed

at protein levels in L6 myotubes. Previous studies using the same siRNA sequences reported

similar reductions in total αAMPK protein expression (83, 113, 156), but to our knowledge this

is the first study to try to knockdown each αAMPK separately. Therefore, comparisons between

their results and ours are not possible at this point. Although protein measurements of each

αAMPK would be ideal to confirm these results, this is consistent with our observation of a

higher level of α1AMPK mRNA in these cells. Indeed, others have reported the expression of

both αAMPK proteins in L6, but their level of expression was not compared (17).

Since we were not able to reduce total αAMPK protein expression with the α2si RNA

alone, we were not able to test whether the α2AMPK isoform is involved in the NO-dependent

signaling. However, our results clearly show that α1AMPK is required for the NO-dependent

regulation of AMPK activity and PGC-1α expression. This is in agreement with the observation

that sodium nitroprusside preferentially activates α1AMPK in isolated rat EDL muscles (60).

Another interesting observation was that DETA-NO caused very similar levels of AMPK

activation across siRNA treated groups, but a measurable increase in AMPK activity, indirectly

assessed by phospho-to-total ACC quantification, was only induced when total αAMPK

expression remained at control levels. This observation suggests that: a) reductions of ~40-60%

of αAMPK expression are sufficient to physiologically compromise the enzyme’s function in

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these cells, even with normal activation by upstream signals taking place, and b) total αAMPK

expression should also be considered when one is assessing AMPK-dependent signaling in

skeletal muscle. Indeed, a recent report shows that α1AMPK and α2AMPK are expressed at

higher levels in the more oxidative and metabolically active type IIa fibers, and that chronic

electrical stimulation of the TA muscle increases their expression (114). Finally, in contrast with

the increase in PGC-1α mRNA seen in groups displaying normal AMPK expression that were

treated with NO, we failed to detect a parallel increase in HDAC5 phosphorylation. This finding

suggests that the NO-mediated regulation of PGC-1α may not rely on HDAC5 phosphorylation

and liberation of MEF2 trans-activity, but rather on AMPK direct phosphorylation of PGC-1α

instead (75).

In Specific Aim 3 we assessed whether the MEF2 or CRE sites at the PGC-1α promoter

would be required for the NO-dependent upregulation of PGC-1α. Contrary to what we initially

hypothesized, NO was capable of inducing reporter activity of the intact PGC-1α promoter, and

also of both ΔCRE- and ΔMEF2-PGC-1α promoters, whereas AICAR was ineffective in

increasing luciferase activity of the ΔMEF2-PGC-1α promoter. This finding is consistent with

the observation that NO treatment does not seem to be involved in significant upregulation of

HDAC5 phosphorylation, and subsequent increase in MEF2 availability. To our knowledge this

is the first study to examine the requirement of these binding sites of the PGC-1α promoter for

NO- and AMPK-dependent induction of PGC-1α. Spiegelman’s group has shown that calcium-

calmodulin dependent protein kinase (CAMKIV) and calcineurin (CaN) induction of PGC-1α

require both sites to be intact, whereas PGC-1α may regulate its own expression through

interaction with MEF2 at its own promoter (50). Akimoto et al. (3, 4) also reported that both sites

are required for the exercise-mediated induction of PGC-1α in adult skeletal muscle in vivo. In

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this context the fact that NO requires either one of these sites is unique. We can not offer a

definitive explanation for this phenomenon at this moment. As previouosly mentioned, it was

recently demonstrated that AMPK can directly phosphorylate PGC-1α, increasing its activity,

which further stimulates PGC-1α transcription (75). In addition, AMPK is known to

phosphorylate and activate CREB in skeletal muscle (142). Therefore, it is possible that our

results reflect different degrees of AMPK activation (i.e lower levels of AMPK activation with

0.65mM AICAR versus 50µM DETA-NO). In addition, it is also possible that NO-dependent

regulation of PGC-1α involves other mechanisms that are not dependent on AMPK signaling.

Further studies are required to confirm these possibilities and also test the effect of NO

supplementation, or NOS inhibition, on CREB-related adaptations in skeletal muscle.

In Specific Aim 4 we ascertained whether genetic ablation of either the eNOS or the nNOS

gene would affect PGC-1α and GLUT4 expression, as well as basal levels of AMPK-dependent

signaling in skeletal muscle. We did not observe an overcompensation of the other constitutive

NOS isoform in the muscles of either knockout. Recently, Wadley et al. (152) reported an

overcompensation of eNOS in nNOS(-/-) mice and of nNOS in eNOS(-/-) mice only in the EDL

muscle. Although we can not provide a clear reason for the differences between their results and

ours, it is possible that the age of the animals may have an influence. We studied very young

mice (4-weeks old), whereas Wadley et al. used 16-week old mice. It is possible that

compensations become more evident once these animals become adults, as a result of a longer

challenge to their normal systemic physiology imposed by the lack of either the nNOS or the

eNOS genes.

The most striking findings, however, were the lower GLUT4 expression observed in the

EDL muscle, the higher pancreatic fat pad depot, and the lower mass of the EDL, Plantaris and

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Soleus muscles in nNOS(-/-) mice. The reduction in muscle mass is corroborated by Wadley et al.

(152). We are unaware of previous studies reporting an increase in visceral fat in such an early

age in these mice. The lower expression of GLUT4 in a predominantly fast-twitch muscle is

consistent with nNOS being more abundant in glycolytic skeletal muscle (85, 152). This

observation supports our previous findings where inhibition of nNOS and iNOS prevented the

AMPK-dependent upregulation of GLUT4 in the plantaris muscle in vivo (89). Although PGC-

1α protein levels were not altered in the EDL muscle of these animals, both phospho-to-total

αAMPK as well as phospho-to-total ACC showed a trend for reduction. Whether this trend is

exacerbated as the animals age, deserves further investigation. However, nNOS knockouts

display signs of peripheral insulin resistance (128). Therefore, it is possible that the increase in

visceral fat accumulation, as well as lower expression of GLUT4 in glycolytic fibers, may

contribute to this phenomenon. On the other hand, mitochondrial dysfunction is observed

preferentially in eNOS knockout mice (46, 98, 104, 106, 149), consistent with the observation

that these animals develop both central and peripheral insulin resistance (128). However, in

agreement with Wadley et al (152), we failed to detect any reduction in basal PGC-1α expression

in both knockouts. In addition, we also did not observe a significant difference in nuclear

localization of PGC-1α in the plantaris muscle of these animals. Considering that in slow-twitch

muscle the array of signals contributing to PGC-1α and GLUT4 regulation is constantly more

active than in fast-twitch muscle, one could expect that inhibition of one of the positive

regulators (i.e. NO production) would have a larger impact in fast glycolytic muscle. Therefore,

although we observed that at a very early age nNOS gene ablation causes structural metabolic

changes in glycolytic muscle only, represented by a significant reduction in GLUT4, we interpret

these results as evidence for a role of NO in the maintenance of normal GLUT4 levels in skeletal

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muscle. This interpretation should be confirmed in models where normal physiology would be

more severely challenged, such as in response to chronic high-fat diet. Future studies addressing

this issue in nNOS knockout mice are still lacking and will definitely contribute for a better

understanding of the interaction between NO- and AMPK-dependent signaling in skeletal

muscle.

In our final Specific Aim we addressed whether AMPK-dependent regulation of PGC-1α

and mitochondrial genes would be affected by inhibition of NO production. Similarly to our

observations with GLUT4 (89), NOS inhibition blunted AMPK induction of PGC-1α, F1ATP

Synthase, and Citrate Synthase mRNA in L6 myotubes. This observation is consistent with the

findings that AMPK activation, via AICAR and Metformin, increase NO production in L6.

In general our results support our central hypothesis that NO regulates PGC-1α and

GLUT4 expression via AMPK activation. We demonstrated for the first time that NO regulates

PGC-1α transcriptionally and that normal αAMPK function is required for the NO-dependent

increase in mitochondrial respiration in skeletal muscle cells. More specifically, we provide

evidence for an important role of α1AMPK expression in the NO-mediated regulation of PGC-

1α in L6 myotubes. Regarding the regulation of PGC-1α by NO, our observations suggest that

NO does not require intact CRE and MEF2 sites at the PGC-1α promoter to induce its activity. In

addition, we also observed that ablation of the nNOS gene results in lower expression of GLUT4

in the glycolytic EDL muscle only, paralleled by a trend towards reduced markers of AMPK-

dependent signaling. Finally, we demonstrated that endogenous NO production is required for

AMPK-dependent upregulation of PGC-1α, a marker of mitochondrial biogenesis, as well as of

F1ATP Synthase and Citrate Synthase in L6 myotubes. Altogether these results corroborate our

proposed model that NO and AMPK interact through a positive feedback loop that impact

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metabolic adaptations in skeletal muscle (89). According to this model, low and basal levels of

NO production are required for normal AMPK activity, and stimulation of AMPK causes small

increases in NO production from the constitutive NOS isoforms in skeletal muscle. Future

studies to test whether this signaling mechanism is involved in skeletal muscle adaptations to

metabolic challenging conditions, such as cold exposure, exercise and caloric restriction are still

required.

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BIOGRAPHICAL SKETCH

The experience I have been accumulating since I completed my bachelor’s degree in

physical education continues to influence my goals for the future as a professional. Therefore,

throughout this text I would like to briefly allude to a couple of facts in my career and discuss

how they helped to shape what I now see as my professional future.

My interest and curiosity for the effects of exercise on wellbeing and prevention of certain

diseases started when I was still pursuing my undergraduate degree. Back then, I was very

interested in knowing how exercise could positively affect the immune system, and whether that

could be beneficial for HIV-positive patients. I was fortunate to be able to perform a relatively

long-term study with sixteen patients that were randomly assigned to the control or the exercise

group. I personally was responsible for monitoring and developing the training sessions of the

exercising group (8 patients), which occurred three times a week for six months, and consisted of

low- to moderate-intensity endurance exercises. Before the training program started and every

two moths thereafter we performed physical fitness and wellbeing assessments, and also assessed

white blood cell counts, as a rough measure of immune function. By the end of the 6-month

program of exercise we observed no clear immune improvement and only a mild increase in

general physical fitness (i.e. aerobic capacity, flexibility and strength all slightly improved).

However, the most striking result was the dramatic decrease in anxiety, depression and stress

levels in the exercise group. This rich experience taught me that exercise definitely added “life”

to those 6 months for each of the patients involved in the training program. This wonderful

experience served as a great motivation for me to try to understand even further how changes in

activity levels induce adaptations in skeletal muscle.

My next step as a professional was to pursue a master’s degree with emphasis in physical

fitness assessment. During my master’s I worked on the development of a test called Sitting-

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Rising test, which basically grades the actions of sitting and rising from the ground,

independently, using a scale from 0 to 4. The idea was to develop an evaluation tool that could

assess functional fitness in a very simple way, and therefore have the potential of broad use with

different populations (e.g. obese individuals, elderly) and in different settings (e.g. on the field,

rehab centers, clinics, etc.). Interestingly, the scores in those two very simple tasks correlated

very well with leg strength and flexibility, and were adversely affected by body mass index

(BMI) and adiposity (body fat %) in people of different ages and fitness levels. Of note, for some

individuals safely sitting and rising from the ground, without the help of hands or arms for

support, were very challenging tasks. This has clearly drawn my attention to the fact that if these

simple tasks were so challenging for some individuals, exercise at a level that could bring about

important improvements in muscle function would probably be as well. Actually this is the case

for some elderly, and individuals affected by muscle and joint problems, neurological disorders,

as well as those with extremely high body fat. As I started to understand more and more about

physiology in general, I also began to realize that for one to have a palpable notion of the

exercise effects on certain organs or tissues (e.g. skeletal muscle) it is important to be trained in

more specific measurements in humans, but also understand and use animal models to address

some questions in further detail.

These observations led me to search for a better education and training on the cellular and

molecular mechanisms by which exercise can improve muscle function. My expectation was that

once having this kind of training I would be able to work on the identification of proteins (e.g.

enzymes) and/or molecules that could serve as pharmacological targets for the induction of some

of the exercise-induced adaptations. This could be of great help in the treatment of obesity,

diabetes and also for the elderly. As a result, I applied to the Ph.D program in Exercise

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Physiology here at the Department of Applied Physiology and Kinesiology of the College of

Health and Human Performance of the University of Florida.

Since the beginning of my Ph.D program I have been learning about the mechanisms

involved in exercise-induced adaptations in skeletal muscle, and also about some molecular

biology tools that can be used in this type of study. This is mainly because of the interaction with

Dr. Criswell, but it is also because of the healthy environment and the great amount of

information that we are exposed to in the Center of Exercise Science. In Dr. Criswell’s lab we

have been performing experiments with mice and rats, muscles in baths, as well as muscle cell

culture to investigate some of skeletal muscle adaptations to different stimuli. More specifically,

my project focus on the role of nitric oxide, a free radical produced at higher levels in skeletal

muscle during contraction, in the regulation of mitochondrial function and expression of the

major glucose transporter in muscle (i.e. GLUT4). This study is directed towards the

development of possible treatment alternatives for chronic diseases related to over nutrition.

Diabetes, obesity and coronary heart disease are all related to deficient glucose transport and

mitochondrial function in skeletal muscle.

In the near future I intend to pursue post-doctoral training focusing on the mechanisms

involved in the improvement of metabolic function in skeletal muscle mediated by exercise.

Ultimately, I intend to pursue a career in academia, where I would like to be involved with both

teaching and research. I enjoy doing experiments with animal and cell culture models to address

some adaptations in skeletal muscle. However, I also would like to collaborate with other

investigators that perform human studies. I believe this type of interaction is critical for a more

prompt development of better strategies for treatment and prevention of chronic diseases such as

obesity and diabetes.