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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).
23
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
24
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
25
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
26
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.
27
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);
28
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).
29
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).
30
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
31
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).
32
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
33
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
34
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
35
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
36
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).
37
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’.
38
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
39
(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
40
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.
41
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.
42
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).
43
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).
44
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).
45
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.
46
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.
47
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.
48
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.
49
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
50
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).
51
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
52
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).
53
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
54
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
55
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
56
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).
57
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*
58
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.
59
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).
60
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.
61
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.
62
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).
63
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.
64
Figure 4-9. Continued
65
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.
66
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.
67
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.
68
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.
69
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.
70
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.
71
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.
72
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.
73
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.
74
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
80
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
81
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
82
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
83
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
84
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.
85
LIST OF REFERENCES
1. Abdul-Ghani MA, DeFronzo RA. Mitochondrial dysfunction, insulin resistance, and type 2 diabetes mellitus. Curr Diab Rep8:173-178, 2008.
2. Ai H, Ihlemann J, Hellsten Y, Lauritzen HP, Hardie DG, Galbo H, Ploug T. Effect of fiber
type and nutritional state on AICAR- and contraction-stimulated glucose transport in rat muscle. Am J Physiol Endocrinol Metab 282:E1291-E1300, 2002.
3. Akimoto T, Pohnert SC, Li P, Zhang M, Gumbs C, Rosenberg PB, Williams RS, Yan Z.
Exercise Stimulates Pgc-1α Transcription in Skeletal Muscle through Activation of the p38 MAPK Pathway. J Biol Chem 280:19587-19593, 2005.
4. Akimoto T, Sorg BS, Yan Z. Real-time imaging of peroxisome proliferator-activated
receptor-γ coactivator-1α promoter activity in skeletal muscles of living mice. Am J Physiol Cell Physiol 287: C790–C796, 2004.
5. Andersen PH, Lund S, Vestergaard H, Junker S, Kahn BB, Pedersen O. Expression of the
major insulin regulatable glucose transporter (GLUT4) in skeletal muscle of noninsulin-dependent diabetic patients and healthy subjects before and after insulin infusion. J Clin Endocrinol Metab 77:27-32,1993.
6. Balon TW, Jasman AP. Acute exposure to AICAR increases glucose transport in mouse
EDL and soleus muscle. Biochem Biophys Res Commun 282: 1008–1011, 2001. 7. Balon TW, Nadler JL. Nitric oxide release is present from incubated skeletal muscle
preparations. J Appl Physiol 77: 2519–2521, 1994. 8. Barish GD, Narkar VA, Evans RM. PPAR delta: a dagger in the heart of the metabolic
syndrome. J Clin Invest 116:590-597, 2006. 9. Bonawitz ND, Clayton DA, Shadel GS. Initiation and beyond: multiple functions of the
human mitochondrial transcription machinery. Mol Cell 24:813-825, 2006. 10. Bonen A, Clark MG, Henriksen EJ. Experimental approaches in muscle metabolism:
hindlimb perfusion and isolated muscle incubations. Am J Physiol Endocrinol Metab 266:E1-E16, 1994.
11. Bonnard C, Durand A, Vidal H, Rieusset J. Changes in adiponectin, its receptors and
AMPK activity in tissues of diet-induced diabetic mice. Diabetes Metab 34:52-61, 2008. 12. Borniquel S, Valle I, Cadenas S, Lamas S, Monsalve M. Nitric oxide regulates
mitochondrial oxidative stress via the transcriptional coactivator PGC1α. FASEB J 20:E1216-1227, 2006.
86
13. Bradley SJ, Kingwell BA, Canny BJ, McConnell GK. Skeletal muscle neuronal nitric oxide synthase micro protein is reduced in people with impaired glucose homeostasis and is not normalized by exercise training. Metabolism 56:1405-1411, 2007.
14. Bryant NJ, Govers R, James DE. Regulated transport of the glucose transporter GLUT4.
Nat Rev Mol Cell Biol 3:267-277, 2002. 15. Camps M, Costello A, Munoz P, Monfar M, Testar X, Palacin M, Zorzano A.Effect of
diabetes and fasting on GLUT-4 (muscle fat) glucose-transporter expression in insulin-sensitive tissues – heterogeneous response in heart, red and white muscle. Biochem J 282:765-772, 1992.
16. Cartee GD, Young DA, Sleeper MD, Zierath J, Wallberg-Henriksson H, Holloszy JO.
Prolonged increase in insulin-stimulated glucose transport in muscle after exercise. Am J Physiol 258:E390-E393, 1989.
17. Ceddia RB, Sweeney G. Creatine supplementation increases glucose oxidation and AMPK
phosphorylation and reduces lactate production in L6 rat skeletal muscle cells. J Physiol 555: 409-421.
18. Chen ZP, McConell GK, Michell BJ, Snow RJ, Canny BJ, Kemp BE. AMPK signaling in
contracting human skeletal muscle: acetyl-CoA carboxylase and NO synthase phosphorylation. Am J Physiol Endocrinol Metab 279: E1202–E1206, 2000.
19. Chen ZP, Mitchelhill KI, Michell BJ, Stapleton D, Rodriguez-Crespo I, Witters LA, Power
DA, Ortiz de Montellano PR, Kemp BE. AMP-activated protein kinase phosphorylation of endothelial NO synthase. FEBS Lett 443: 285–289, 1999.
20. Chen ZP, Stephens TJ, Murthy S, Canny BJ, Hargreaves M, Witters LA, Kemp BE,
McConell GK. Effect of exercise intensity on skeletal muscle AMPK signaling in humans. Diabetes 52: 2205–2212, 2003.
21. Clayton DA. Mitochondrial DNA replication: what we know. IUBMB Life 55:213-217,
2003. 22. Clayton DA. Vertebrate mitochondrial DNA: a circle of surprises. Exp Cell Res 255:4-9,
2000. 23. Connor MK, Irrcher I, Hood DA. Contractile activity-induced transcriptional activation of
cytochrome C involves Sp1 and is proportional to mitochondrial ATP synthesis in C2C12 muscle cells. J Biol Chem 276:15898-15904, 2001.
24. Cushman SW, Wardzala LJ. Potential mechanism of insulin action on glucose transport in
the isolated rat adipose cell. Apparent translocation of intracellular transport systems to the plasma membrane. J Biol Chem 255:4758-4762, 1980.
87
25. Czubryt MP, McAnally J, Fishman GI, Olson EN. Regulation of peroxisome proliferator-activated receptor γ coactivator 1α (PGC1α) and mitochondrial function by MEF2 and HDAC5. Proc Natl Acad Sci USA 100:1711-1716, 2002.
26. Davis BJ, Xie Z, Viollet B, Zou M-H. Activation of AMP-activated protein kinase by
antidiabetes drug Metformin stimulates nitric oxide synthesis in vivo by promoting the association of heat shock protein 90 and endothelial nitric oxide synthase. Diabetes 55:496-505, 2006.
27. DeFronzo RA, Jacot E, Jequier E, Maeder E, Wahren J, Felber JP. The effect of insulin on
the disposal of intravenous glucose. Diabetes 30:1000-1007, 1981. 28. DeFronzo RA. Pathogenesis of type 2 diabetes: metabolic and molecular implications for
identifying diabetes genes. Diabetes Rev 5:177-269, 1997. 29. Drenning JA, Lira VA, Simmons CG, Soltow QA, Sellman JE, Criswell DS. Nitric oxide
facilitates NFAT-dependent transcription in mouse myotubes. Am J Physiol Cell Physiol 294:C1088-1095, 2008.
30. Dufour E, Larsson NG. Understanding aging: revealing order out of chaos. Biochim
Biophys Acta 1658:122-132, 2004. 31. Durante PE, Mustard KJ, Park SH, Winder WW, Hardie DG. Effects of endurance training
on activity and expression of AMP-activated protein kinase isoforms in rat muscles. Am J Physiol Endocrinol Metab 283:E178-E186, 2002.
32. Elia M. Organ and tissue contribution to metabolic rate. In: Kinney JM, Tucker HN, eds.
Energy metabolism: Tissue determinants and cellular corollaries. New York: Raven Express, 62-8, 1992.
33. Engelgau MM, Geiss LS, Saaddine JB, Boyle JP, Benjamin SM, Gregg EW, Tierney EF,
Rios-Burrows M, Mokdad AH, Ford ES, Imperatore G, Narayan KMV. The evolving diabetes burden in the United States. Ann Intern Med 140:945-950, 2004.
34. Evans MJ, Scarpulla RC. Interaction of nuclear factors with multiple sites in the somatic
cytochrome c promoter. Characterization of upstream NRF-1, ATF and intron Sp1 recognition sites. J Biol Chem 264:14361-14368, 1989.
35. Falkenberg M, Gaspari M, Rantanen A, Trifunovic A, Larsson NG, Gustafsson CM.
Mitochondrial transcription factors B1 and B2 active transcription of human mtDNA. Nat Genet 31:289-294, 2002.
36. Fediuc S, Gaidhu MP, Ceddia RB. Regulation of AMP-activated protein kinase and acetyl-
CoA carboxylase phosphorylation by palmitate in skeletal muscle cells. J Lipid Res 47:412-20, 2006.
88
37. Fisher RP, Lisowsky T, Parisi MA, Clayton DA. DNA wrapping and bending by a mitochondrial high mobility group-like transcriptional activator protein. J Biol Chem 267:3358-3367, 1992.
38. Fisher RP, Parisi MA, Clayton DA. Flexible recognition of rapidly evolving promoter
sequences by mitochondrial transcription factor 1. Genes Dev 3:2202-2217, 1989. 39. Friedman JE, Dohm GL, Leggett-Frazier N, Elton CW, Tapscott EB, Pories WP, Caro JF.
Restoration of insulin responsiveness in skeletal muscle of morbidly obese patients after weight loss. J Clin Invest 89:701-705, 1992.
40. Fryer LG, Carling D. AMP-activated protein kinase and the metabolic syndrome. Biochem
Soc Trans 33:362-6, 2005. 41. Fryer LG, Parbu-Patel A, Carling D. The anti-diabetic drugs rosiglitazone and metformin
stimulate AMP-activated protein kinase through distinct pathways. J Biol Chem 277:25226-25232, 2002.
42. Fujimoto M, Shimizu N, Kunii K, Martyn JA, Ueki K, Kaneki M. A role for iNOS in
fasting hyperglycemia and impaired insulin signaling in the liver of obese diabetic mice. Diabetes 54:1340-8, 2005.
43. Garcia-Roves PM, Huss JM, Han DH, Hancock CR, Iglesias-Gutierrez E, Chen M,
Holloszy JO. Raising plasma fatty acid concentration induces increased biogenesis of mitochondria in skeletal muscle. Proc Natl Acad Sci USA 104:10709-10713, 2007.
44. Gleyzer N, Vercauteren K, Scarpulla RC. Control of mitochondrial transcription specificity
factors (TFB1M abd TFB2M) by nuclear respiratory factors (NRF-1 and NRF-2) and PGC-1 family coactivators. Mol Cell Biol 25:1354-1366, 2005.
45. Goodpaster BH, Katsiaras A, Kelley DE. Enhanced fat oxidation through physical activity
is associated with improvements in insulin sensitivity in obesity. Diabetes 52:2191-2197, 2003.
46. Gouill EL, Jimenez M, Binnert C, Jayet P-Y, Thalmann S, Nicod P, Scherrer U,
Vollenweider P. Endothelial nitric oxide synthase (eNOS) knockout mice have defective mitochondrial β-oxidation. Diabetes 56:2690-2696, 2007.
47. Gray S, Feinberg MW, Hull S, Kuo CT, Watanabe M, Sen S, DePina A, Haspel R, Jain
MK. The Kruppel-like factor KLF15 regulates the insulin-sensitive glucose transporter GLUT4. J Biol Chem 277:34322-34328, 2002.
48. Gulick T, Cresci S, Caira T, Moore DD, Kelly DP. The peroxisome proliferator-activated
receptor regulates mitochondrial fatty acid oxidative enzyme gene expression. Proc Natl Acad Sci USA 91:11012-11016, 1994.
89
49. Gulve EA, Spina RJ. Effect of 7-10 days of cycle ergometer exercise on skeletal muscle GLUT4 protein content. J Appl Physiol 79:156201566, 1995.
50. Handschin C, Rhee J, Lin J, Tarr PT, Spiegelman BM. An autoregulatory loop controls
peroxisome proliferator-activated receptor γ coactivator 1α expression in muscle. Proc Natl Acad Sci USA 100:7111-7116, 2003.
51. Handschin C, Spiegelman BM. The role of exercise and PGC1alpha in inflammation and
chronic disease. Nature 454:463-469, 2008. 52. Hardie DG, Carling D, Carlson M. The AMP-activated/SNF1 protein kinase subfamily:
metabolic sensor of eukaryotic cell? Annu Rev Biochem 67: 821–855, 1998. 53. Hardie DG, Hawley SA, Scott JW. AMP-activated protein kinase--development of the
energy sensor concept. J Physiol 574:7-15, 2006. 54. Hardie DG, Hawley SA. AMP-activated protein kinase: the energy charge hypothesis
revisited. Bio essays 23:1112-9, 2001. 55. Hardie DG, Sakamoto K. AMPK: a key sensor of fuel and energy status in skeletal muscle.
Physiology 21:48-60, 2006. 56. Hawley JA. Exercise as a therapeutic intervention for the prevention and treatment of
insulin resistance. Diabetes Metab Res Rev 20:383-393, 2004. 57. Hawley JA, Houmard JA. Introduction-preventing insulin resistance through exercise: a
cellular approach. Med Sci Sports Exerc 36:1187-1190, 2004. 58. Hawley SA, Gadalla AE, Olsen GS, Hardie DG. The anti-diabetic drug metformin activates
the AMP-activated protein kinase cascade via an adenine nucleotide-independent mechanism. Diabetes 51:220-2425, 2002.
59. Helmrich SP, Ragland DR, Leung RW, Paffenbarger RS Jr. Physical activity and reduced
occurrence of non-insulin-dependent diabetes mellitus. N Engl J Med 325:147-152, 1991. 60. Higaki Y, Hirshman MF, Fujii N, Goodyear LJ. Nitric oxide increases glucose uptake
through a mechanism that is distinct from the insulin and contraction pathways in rat skeletal muscle. Diabetes 50:241-247, 2001.
61. Hirschfield W, Moody MR, O’Brien WE, Gregg AR, Bryan RM, Reid MB. Nitric oxide
release and contractile properties of skeletal muscles from mice deficient in type III NOS. Am J Physiol Regul Integrative Comp Physiol 278:R95-R100, 2000.
62. Hogan P, Dall T, Nikolov P. American Diabetes Association. Economic costs of diabetes in
the US in 2002. Diabetes Care 2003; 26:917-32, 2003.
90
63. Holloszy JO. Biochemical adaptations in skeletal muscle. Effects of exercise on mitochondrial O2 uptake and respiratory enzyme activity in skeletal muscle. J Biol Chem 242:2278-2282, 1967.
64. Holloszy JO, Booth FW. Biochemical adaptations to endurance exercise in muscle. Ann
Rev Physiol 38:273-291, 1976. 65. Holloszy JO, Coyle EF. Adaptations of skeletal muscle to endurance exercise and their
metabolic consequences. J Appl Physiol 56:831-839, 1984. 66. Holloszy JO, Nolte LA. Glucose transport in skeletal muscle. In: Muscle Metabolism.
Zierath JR, Wallberg-Henriksson H (Eds.), 2002. 67. Host HH, Hansen PA, Nolte LA, Chen MM, Holloszy JO. Rapid reversal of adaptive
increases in muscle GLUT4 and glucose transport capacity after training cessation. J Appl Physiol 84:798-802, 1998.
68. Host HH, Hansen PA, Nolte LA, Chen MM, Holloszy JO. Glycogen supercompensation
masks the effect of a training-induced increase in GLUT4 on muscle glucose transport. J Appl Physiol 85:133-138, 1998.
69. Houmard JA, Egan PC, Neufer PD, Friedman JE, Wheeler WS, Israel RG, Dohm GL.
Elevated skeletal muscle glucose transporter levels in exercise-trained middle-aged men. Am J Physiol 261:E437-E443, 1991.
70. Houmard JA, Shinebarger MH, Dolan PL, Leggett-Frazier N, Bruner RK, McCammon
MR, Israel RG, Dohm GL. Exercise training increases GLUT-4 protein concentration in previously sedentary middle-aged men. Am J Physiol 264:E896-E901, 1993.
71. Huang S, Czech MP. The GLUT4 glucose transporter. Cell Metab 5:237-252, 2007. 72. Huss JM, Kelly DP. Nuclear receptor signaling and cardiac energetics. Circ Res 95:568-
578, 2004. 73. Huss JM, Kopp RP, Kelly DP. Peroxisome proliferator-activated receptor coactivator-1α
(PGC-1α) coactivates the cardiac-enriched nuclear receptors estrogen-related receptor-α and -γ: identification of novel leucine-rich interaction motif within PGC-1α. J Biol Chem 277:40265-40274, 2002.
74. Irrcher I, Hood DA. Regulation of Egr-1, SRF, and Sp1 mRNA expression in contracting
skeletal muscle cells. J Appl Phyisol 97:2207-2213, 2004. 75. Jager S, Handschin C, St.-Pierre J, Spiegelman BM. AMP-activated protein kinase
(AMPK) action in skeletal muscle via direct phosphorylation of PGC-1α. Proc Natl Acad Sci USA 104:12-17-12022, 2007.
91
76. Ji LL, Gomez-Cabrera MC, Vina J. Exercise and hormesis: activation of cellular antioxidant signaling pathway. Ann NY Acad Sci 1067: 425-435, 2006.
77. Joost HG, Bell GI, Best GD, Birnbaum MJ, Charron MJ, Chen YT, Doege H, James DE,
Lodish HF, Moley KH, Moley JF, Mueckler M, Rogers S, Schurmann A, Seino S, Thorens B. Nomenclature of the GLUT/SLC2A family of sugar/polyol transport facilitators. Am J Physiol Endocrinol Metab 282:E974-E976, 2002.
78. Jorgensen SB, Treebak JT, Viollet B, Schjerling P, Vaulont S, Wojtaszewski JFP, Richter
EA. Role of AMPKα2 in basal, training-, and AICAR-induced GLUT4, hexokinase II, and mitochondrial protein expression in mouse muscle. Am J Physiol Endocrinol Metab 292:E331-E339, 2007.
79. Karnieli E, Armoni M. Transcriptional regulation of the insulin-responsive glucose
transporter GLUT4 gene: from physiology to pathology. Am J Physiol Endocrinol Metab 295:E38-E45, 2008.
80. Katz LD, Glickman MG, Rapoport S, Ferrannini E, DeFronzo RA. Splanchnic and
peripheral disposal of oral glucose in man. Diabetes 32:675-679, 1983. 81. Kelley DE, He J, Menshikova EV, Ritov VB. Dysfunction of mitochondria in human
skeletal muscle in type 2 diabetes. Diabetes 51:2944-2950, 2002. 82. Koh H-J, Arnolds DE, Fujii N, Tran TT, Rogers MJ, Jessen N, Li Y, Liew CW, Ho RC,
Hirshman MF, Kulkarni RN, Kahn CR, Goodyear LJ. Skeletal muscle-selective knockout of LKB1 increases insulin sensitivity, improves glucose homeostasis, and decreases TRB3. Mol Cell Biol 26:8217-8227, 2006.
83. Konrad D, Rudich A, Bilan PJ, Patel N, Richardson C, Witters LA, Klip A. Troglitazone
causes acute mitochondrial membrane depolarisation and an AMPK-mediated increase in glucose phosphorylation in muscle cells. Diabetologia 48:954-66, 2005.
84. Koves TR, Li P, An J, Akimoto T, Slentz D, Ilkayeva O, Dohm GL, Yan Z, Newgard CB,
Muoio DM. Peroxisome proliferator-activated receptor-γ Co-activator 1α-mediated metabolic remodeling of skeletal myocytes mimics exercise training and reverses lipid-induced mitochondrial inefficiency. J Biol Chem 280:33588-33598, 2005.
85. Lau KS, Grange RW, Isotani E, Sarelius IH, Kamm KE, Stull JT. nNOS and eNOS
modulate cGMP formation and vascular response in contracting fast-twitch skeletal muscle. Physiol Genomics 2: 21–27, 2000.
86. Leturque A, Loizeau M, Vaulont S, Salminen M, Girard J. Improvement of insulin action
in diabetic transgenic mice selectively overexpressing GLUT4 in skeletal muscle. Diabetes 45:23-27, 1996.
92
87. Lillioja S, Mott DM, Spraul M, Ferraro R, Foley JE, Ravussin E, Knowler WC, Bennett PH, Bogardus C. Insulin resistance and insulin secretory dysfunction as precursors of non-insulin-dependent diabetes mellitus. Prospective studies of Pima Indians. N Engl J Med 329:1988-1992, 1993.
88. Lin J, Wu H, Tarr PT, Zhang CY, Wu Z, Boss O, Michael LF, Puigserver P, Isotani E,
Olson EN, Lowell BB, Bassel-Duby R, Spiegelman BM. Transcriptional co-activator PGC-1 alpha drives the formation of slow-twitch muscle fibres. Nature 418:797-801, 2002.
89. Lira VA, Soltow QA, Long JH, Betters JL, Sellman JE, Criswell DS. Nitric oxide increases
GLUT4 expression and regulates AMPK signaling in skeletal muscle. Am J Physiol Endocrinol Metab 293:E1062-8, 2007.
90. Liu ML, Olson AL, Edgington NP, Moye-Rowley WS, Pessin JE. Myocyte enhancer factor
2 (MEF2) binding site is essential for C2C12 myotube-specific expression of the rat GLUT4/muscle-adipose facilitative glucose transporter gene. J Biol Chem 269:2852=14-28521, 1994.
91. Lowell BB, Shulman GI. Mitochondrial dysfunction and type 2 diabetes. Science 307:384-
387, 2005. 92. Mann GE, Yudilevich DL, Sobrevia L. Regulation of amino acid and glucose transporters
in endothelial and smooth cells. Physiol Rev 83:183-252, 2003. 93. Martin BC, Warram JH, Krolewski AS, Bergman RN, Soeldner JS, Kahn CR. Role of
glucose and insulin resistance in development of type 2 diabetes mellitus: results of a 25-year follow-up study. Lancet 340:325-329, 1992.
94. McGee SL, van Denderen BJ, Howlett KF, Mollica J, Schertzer JD, Kemp BE, Hargreaves
M. AMP-activated protein kinase regulates GLUT4 transcription by phosphorylating histone deacetylase 5. Diabetes 57:860-867, 2008.
95. Michael LF, Wu Z, Cheatham RB, Puigserver P, Adelmant G, Lehman JJ, Kelly DP,
Spiegelman BM. Restoration of insulin-sensitive glucose transporter (GLUT4) gene expression in muscle cells by the transcriptional coactivator PGC-1. Proc Natl Acad Sci USA 98:3820-5, 2001.
96. Michel JB, Ordway GA, Richardson JA, Williams RS. Biphasic induction of immediate
early gene expression accompanies activity-dependent angiogenesis and myofiber remodeling of rabbit skeletal muscle. J Clin Invest 94:277-285, 1994.
97. Mokdad AH, Marks LS, Stroup DF, Gerberding JL. Actual causes of death in the United
States, 2000. JAMA 291:1238-1245, 2004.
93
98. Momken I, Fortin D, Serrurier B, Bigard X, Ventura-Clapier R, Veksler V. Endothelial nitric oxide synthase (NOS) deficiency affectsenergy metabolism pattern in murine oxidative skeletal muscle. Biochem J 368:341-347, 2002.
99. Mootha VK, Lindgren CM, Eriksson KF, Subramanian A, Sihag S, Lehar J, Puigserver P,
Carlsson E, Ridderstrale M, Laurila E, Houstis N, Daly MJ, Patterson N, Mesirov JP, Golub TR, Tamayo P, Spiegelman B, Lander ES, Hirschhorn JN, Altshuler D, Groop LC. PGC-1α-responsive genes involved in oxidative phosphorylation are coordinately downregulated in human diabetes. Nat Gen 34:267-273, 2003.
100. Mora S, Pessin JE. The MEF2A isoform is required for striated muscle-specific expression
of the insulin-responsive GLUT4 glucose transporter. J Biol Chem 275:16323-16328, 2000. 101. Moreno H, Serrano AL, Santalucia T, Guma A, Canto C, Brand NJ, Palacin M, Schiaffino
S, Zorzano A. Differential regulation of the muscle-specific GLUT4 enhancer in regenerating and adult skeletal muscle. J Biol Chem 278:40557-40564, 2003.
102. Muoio DM, Newgard CB. Obesity-related derangements in metabolic regulation. Ann Rev
Biochem 75:367-401, 2006. 103. Neufer PD, Carey JO, Dohm GL. Transcriptional regulation of the gene for glucose-
transporter GLUT4 in skeletal muscle – effects of diabetes and fasting. J Biol Chem 268:13824-13829, 1993.
104. Nisoli E, Clementi E, Paolucci C, Cozzi V, Tonello C, Sciorati C, Bracale R, Valerio A,
Francolini M, Moncada S, Carruba MO. Mitochondrial biogenesis in mammals: the role of endogenous nitric oxide. Science 299:896-899, 2003.
105. Nisoli E, Falcone S, Tonello C, Cozzi V, Palomba L, Fiorani M, Pisconti A, Brunelli S,
Cardile A, Francolini M, Cantoni O, Carruba MO, Moncada S, Clementi E. Mitochondrial biogenesis by NO yields functionally active mitochondria in mammals. PNAS 101:16507-16512, 2004.
106. Nisoli E, Tonello C, Cardeile A, Cozzi V, Bracale R, Tedesco L, Falcone S, Valerio A,
Cantoni O, Clementi E, Moncada S, Carruba MO. Calorie restriction promotes mitochondrial biogenesis by inducing the expression of eNOS. Science 310:314-317, 2005.
107. Oshel K, Knight J, Cao K, Thai M, Olson A. Identification of a 30-base pair regulatory
element and novel DNA binding protein that regulates the human GLUT4 promoter in transgenic mice. J Biol Chem 275:23666-23673, 2000.
108. Olsen RH, Krogh-Madsen R, Thomsen C, Booth FW, Pedersen BK. Metabolic responses to
reduced daily steps in healthy nonexercising men. JAMA 299:1261-1263, 2008.
94
109. Patti ME, Butte AJ, Crunkhorn S, Cusi K, Berria R, Kashyap S, Miyazaki Y, Kohane I, Costello M, Saccone R, Landaker EJ, Goldfine AB, Mun E, DeFronzo R, Finlayson J, Kahn CR, Mandarino LJ. Coordinated reduction of genes of oxidative metabolism in humans with insulin resistance and diabetes: potential role of PGC1 and NRF1. Proc Natl Acad Sci 100:8466-8471, 2003.
110. Patwell DM, McArdle A, Mogan JE, Patridge TA, Jackson MJ. Release of reactive oxygen
and nitrogen species from contracting skeletal muscle cells. Free Rad Biol Med 37: 1064–1072, 2004.
111. Pedersen O, Bak JF, Andersen PH, Lund S, Moller DE, Flier JS, Kahn BB. Evidence
against altered expression of GLUT1 or GLUT4 in skeletal muscle of patients with obesity or NIDDM. Diabetes 39:865-870, 1990.
112. Petersen KF, Dufour S, Befroy D, Garcia R, Shulman GI. Impaired mitochondrial activity
in the insulin-resistant offspring of patients with type 2 diabetes. N engl J Med 350:664-671, 2004.
113. Pilon G, Dallaire P, Marette A. Inhibition of inducible nitric-oxide synthase by activators
of AMP-activated protein kinase: a new mechanism of action of insulin-sensitizing drugs. J Biol Chem 279:20767-20774, 2004.
114. Putman CT, Martins KJB, Gallo ME, Lopaschuk GD, Pearcey JA, MacLean IM,
Saranchuk RJ, Pette D. α-Catalytic subunits of 5’AMP-activated protein kinase display fiber-types specific expression and are upregulated by chronic low-frequency stimulation in rat muscle. Am J Physiol Regul Integr Comp Physiol 293:R1325-R1334, 2007.
115. Pye D, Palomero J, Kabayo T, Jackson MJ. Real-time measurement of nitric oxide in single
mature mouse skeletal muscle fibres during contractions. J Physiol 581:309-18, 2007. 116. Reaven GM. Why Syndrome X? From Harold Himsworth to insulin resistance syndrome.
Cell Metabol 1:9-14, 2005. 117. Rector RS, Thyfault JP, Laye MJ, Morris RT, Borengasser SJ, Uptergrove GM,
Chakravarthy MV, Booth FW, Ibdah JA. Cessation of daily exercise dramatically alters precursors of hepatic steatosis in Otsuka Long-Evans Tokushima Fatty (OLETF) rats. J Physiol 586:4241-4249, 2008.
118. Reid MB. Role of nitric oxide in skeletal muscle: synthesis, distribution and functional
importance. Acta Physiol Scand 162: 401–409, 1998. 119. Ren JM, Marshall BA, Mueckler MM, McCaleb M, Amatruda JM, Shulman GI.
Overexpression of Glut4 protein in muscle increases basal and insulin-stimulated whole body glucose disposal in conscious mice. J Clin Invest 95:429-432, 1995.
95
120. Ren JM, Semenkovich CF, Gulve EA, Gao J Holloszy JO. Exercise induces rapid increases in GLUT4 expression, glucose transport capacity and insulin-stimulated glycogen storage in muscle. J Biol Chem 269:14396-14401, 1994.
121. Richter EA, Garetto LP, Goodman MN, Ruderman NB. Muscle glucose metabolism
following exercise in the rat. Increased sensitivity to insulin. J Clin Invest 69:785-793, 1982.
122. Saltiel AR, Kahn CR. Insulin signaling and the regulation of glucose and lipid metabolism.
Nature 414:799-806, 2001. 123. Santalucia T, Moreno H, Palacin M, Yacoub MH, Brand NJ, Zorzano A. A novel
functional co-operation between MyoD, MEF2 and TRalpha1 is sufficient for the induction of GLUT4 gene transcription. J Mol Biol 314:195-204, 2001.
124. Scarpulla RC. Transcriptional paradigms in mammalian mitochondrial biogenesis and
function. Physiol Rev 88:611-638, 2008. 125. Schiaffino S, Sandri M, Murgia M. Activity-dependent signaling pathways controlling
muscle diversity and plasticity. Physiology 22:269-78, 2007. 126. Schon EA, Manfredi G. Neuronal degeneration and mitochondrial dysfunction. J Clin
Invest 111:303-312, 2003. 127. Schrauwen-Hinderling VB, Kooi ME, Hesselink MK, Jeneson JA, Backes WH, van
Echteld CJ, van Engelshoven JM, Mensink M, Schrauwen P. Impaired in vivo mitochondrial function but similar intramyocellular lipid content in paitents with type 2 diabetes mellitus and BMI-matched control subjects. Diabetologia 50:113-120, 2007.
128. Shankar RR, Wu Y, Shen H, Zhu J-S, Baron AD. Mice with gene disruption of both
endothelial and neuronal nitric oxide synthase exhibit insulin resistance. Diabetes 49:684-687, 2000.
129. Short KR, Vittone JL, Bigelow ML, Proctor DN, Rizza RA, Coenen-Schimke JM, Nair KS.
Impact of aerobic exercise traning on age-related changes in insulin sensitivity and muscle oxidative capacity. Diabetes 52:1888-1896, 2003.
130. Silveira LR, Pereira-Da-Silva L, Juel C, Hellsten Y. Formation of hydrogen peroxide and
nitric oxide in rat skeletal muscle cells during contractions. Free Radic Biol Med 35: 455–464, 2003.
131. Simoneau JA, Kelley DE. Altered glycolytic and oxidative capacities of skeletal muscle
contribute to insulin resistance in NIDDM. J Appl Physiol 83:166-171, 1997.
96
132. Smith AC, Bruce CR, Dyck DJ. AMP-kinase activation with AICAR simultaneously increases fatty acid and glucose oxidation in resting rat soleus muscle. J Physiol 565: 547–553, 2005.
133. Smith LW, Smith JD, Criswell DS. Involvement of nitric oxide synthase in skeletal muscle
adaptation to chronic overload. J Appl Physiol 92:2005-2011, 2002. 134. Sparks LM, Xie H, Koza RA, Mynatt R, Hulver MW, Bray GA, Smith SR. A high-fat diet
coordinately downregulates genes required for mitochondrial oxidative phosphorylation in skeletal muscle. Diabetes 54:1926-33, 2005.
135. Srere PA. Citrate Synthase. Methods Enzymol 13:3-5, 1969. 136. Sriwijitkamol A, Coletta DK, Wajcberg E, Balbontin GB, Reyna SM, Barrientes J, Eagan
PA, Jenkinson CP, Cersosimo E, DeFronzo RA, Sakamoto K, Musi N. Effect of acute exercise on AMPK signaling in skeletal muscle of subjects with type 2 diabetes: a time-course and dose-response study. Diabetes 56:836-848, 2007.
137. Stamler JS, Meissner G. Physiology of nitric oxide in skeletal muscle. Physiol Rev 81:
209–237, 2001. 138. Steensberg A, Keller C, Hillig T, Frosig C, Wojtaszewski JFP, Pedersen BK, Pilegaard H,
Sander M. Nitric oxide production is a proximal signaling event controlling exercise-induced mRNA expression in human skeletal muscle. FASEB J 21:2683-2694, 2007.
139. Steinberg GR, Michell BJ, van Denderen BJW, Watt MJ, Carey AL, Fam BC,
Andrikopoulos S, Proietto J, Gorgun CZ, Carling D, Hotamisligil GS, Febbraio MA, Kay TW, Kemp BE. Tumor necrosis factor α-induced skeletal muscle insulin resistance involves suppression of AMP-kinase signaling. Cell Metab 4:465-474, 2006.
140. Stoppani J, Hildebrandt AL, Sakamoto K, Cameron-Smith D, Goodyear LJ, Neufer PD.
AMP-activated protein kinase activates transcription of the UCP3 and HKII genes in rat skeletal muscle. Am J Physiol Endocrinol Metab 283: E1239–E1248, 2002.
141. Suzuki A, Okamoto S, Lee S, Saito K, Shiuchi T, Minokoshi Y. Leptin stimulates fatty acid
oxidation and peroxisome-proliferator activated receptor alpha gene expression in mouse C2C12 myoblasts by changing the subcellular localization of the alpha2 form of AMP-activated protein kinase. Mol Cell Biol 27:4340-4346, 2007.
142. Thomson DM, Herway ST, Filmore N, Kim H, Brown JD, Barrow JR, Winder WW. AMP-
activated protein kinase phosphorylates transcription factors of the CREB family. J Appl Physiol 104:429-438, 2008.
143. Thong FS, Dugani CB, Klip A. Turning signals on and off: GLUT4 traffic in the insulin-
signaling highway. Physiology (Bethestha) 20:271-284, 2005.
97
144. Thyfault JP, Cree MG, Zheng D, Zwetsloot JJ, Tapscott EB, Koves TR, Ilkayeva O, Wolfe RR, Muoio DM, Dohm GL. Contraction of insulin-resistant muscle normalizes insulin action in association with increased mitochondrial activity and fatty acid catabolism. Am J Physiol Cell Physiol 292:C729-C739, 2007.
145. Thyfault JP. Setting the stage: possible mechanisms by which acute contraction restores
insulin sensitivity in muscle. 294:R1103-R1110, 2008. 146. Toledo FG, Menshikova EV, Ritov VB, Azuma K, Radikova Z, DeLany J, Kelley DE.
Effects of physical activity and weight loss on skeletal muscle mitochondria and relationship with glocse control in type 2 diabetes. Diabetes 56:2142-2147, 2007.
147. Toledo FG, Watkins S, Kelley DE. Changes induced by physical activity and weight loss in
the morphology of intermyofibrillar mitochondria in obese men and women. J Clin Endocrinol Metab 91:3224-3227, 2006.
148. Tsao TS, Burcelin R, Katz EB, Huang L, Charron MJ. Enhanced insulin action due to
targeted GLUT4 overexpression exclusively in muscle. Diabetes 45:28-36, 1996. 149. Valerio A, Cardile A, Cozzi V, Bracale R, Tedesco L, Pisconti A, Palomba L, Cantoni O,
Clementi E, Moncada S, Carruba MO, Nisoli E. TNF-α downregulates eNOS expression and mitochondrial biogenesis in fat and muscle of obese rodents. J Clin Invest 116:2791-2798, 2006.
150. Valverde AM, Benito M, Lorenzo M. The brown adipose cell: a model for understanding
the molecular mechanisms of insulin resistance. Acta Physiol Scand 183:59-73, 2005. 151. Virbasius JV, Scarpulla RC. Transcriptional activation through ETS domain binding sites
in the cytochrome c oxidase subunit IV gene. Mol Cell Biol 11:5631-5638, 1991. 152. Wadley GD, Choate J, McConell GK. NOS isoform-specific regulation of basal but not
exercise-induced mitochondrial biogenesis in mouse skeletal muscle. J Physiol 585:253-262, 2007.
153. Wallace DC. A mitochondrial paradigm of metabolic and degenerative diseases, aging,
cancer: a dawn for evolutionary medicine. Annu Rev Genet 39:359-407, 2005. 154. Wallberg-Henriksson H, constable SH, Young DA, Holloszy JO. Glucose tansport into rat
skeletal muscle: interaction between exercise and insulin. J Appl Physiol 65:909-913, 1988. 155. Watson RT, Kanzaki M, Pessin JE. Regulated membrane trafficking of the insulin-
responsive glucose transporter 4 in adipocytes. Endocr Rev 25:177-204, 2004. 156. Wijesekara N, Tung A, Thong F, Klip A. Muscle cell depolarization induces a gain in
surface GLUT4 via reduced endocytosis independently of AMPK. Am J Physiol Endocrinol Metab 290:E1276-86, 2006.
98
157. Winder WW, Hardie DG. AMP-activated protein kinase, a metabolic master switch:
possible roles in type 2 diabetes. Am J Physiol Endocrinol Metab 277: E1–E10, 1999. 158. Winder WW, Holmes BF, Rubink DS, Jensen EB, Chen M, Holloszy JO. Activation of
AMP-activated protein kinase increases mitochondrial enzymes in skeletal muscle. J Appl Physiol 88:2219-2226, 2000.
159. Wright DC, Dong-Ho H, Garcia-Roves PM, Geiger PC, Jones TE, Holloszy JO. Exercise-
induced mitochondrial biogenesis begins before the increase in muscle PGC-1α expression. J Biol Chem 282:194-199, 2007.
160. Wu H, Naya FJ, McKinsey TA, Mercer B, Shelton JM, Chin ER, Simard AR, Michel RN,
Bassel-Duby R, Olson EN, Williams RS. MEF2 respondes to multiple calcium-regulated signals in the control of skeletal muscle fiber type. EMBO J 19:1963-1973, 2000.
161. Wu Z, Puigserver P, Andersson U, Zhang C, Adelmant G, Mootha V, Troy A, Cinti S,
Lowell B, Scarpulla RC, Spiegelman BM. Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1. Cell 98:115-24, 1999.
162. Yamamoto DL, Hutchinson DS, Bengtsson T. Beta(2)-Adrenergic activation increases
glycogen synthesis in L6 skeletal muscle cells through a signalling pathway independent of cyclic AMP. Diabetologia 50:158-67, 2007.
163. Yan Z, Li P, Akimoto T. Transcriptional control of the Pgc-1alpha gene in skeletal muscle
in vivo. Exerc Sport Sci Rev 35:97-101, 2007. 164. Young DA, Wallberg-Henriksson H, Sleeper M, Holloszy JO. Reversal of the exercise-
induced increase in muscle permeability to glucose. Am J Physiol 253:E331-E335, 1987. 165. Young ME, Leighton B. Evidence for altered sensitivity of the nitric oxide/cGMP signaling
cascade in insulin-resistant skeletal muscle. Biochem J 329:73-79, 1998. 166. Zhang P, Engelgau MM, Norris SL, Gregg EW, Narayan KMV. Application of economic
analysis to diabetes and diabetes care. Ann Intern Med 140:972-977, 2004. 167. Zheng D, MacLean PS, Pohnert SC, Knight JB, Olson AL, Winder WW, Dohm GL.
Regulation of muscle GLUT-4 transcription by AMPactivated protein kinase. J Appl Physiol 91: 1073–1083, 2001.
168. Zhou G, Myers R, Li Y, Chen Y, Shen X, Fenyk-Melody J, Wu M, Ventre J, Doebber T,
Fujii N, Musi N, Hirshman MF, Goodyear LJ, Moller DE. Role of AMP-activated protein kinase in mechanism of metformin action. J Clin Invest 108:1167-1174, 2001.
99
169. Zimmet P, Alberti KGMM, Shaw J. Global and societal implications of diabetes epidemic. Nature 414:782-787, 2001.
170. Zong H, Ren JM, Young LH, Pypaert M, Mu J, Birnbaum MJ, Shulman GI. AMP kinase is
required for mitochondrial biogenesis in skeletal muscle in response to chronic energy deprivation. Proc Natl Acad Sci USA 99: 15983–15987, 2002.
171. Zorzano A, Palacin M, Guma A. Mechanisms regulating GLUT4 glucose transporter
expression and glucose transport in skeletal muscle. Acta Physiol Scand 183:43-58, 2005. 172. Zorzano A, Sevilla L, Tomas E, Guma A, Palacin M, Fischer Y. GLUT4 trafficking in
cardiac and skeletal muscle: isolation and characterization of distinct intracellular GLUT4-containing vesicle populations. Biochem Soc Trans 25:968-974, 1997.
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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.