J Appl Physiol 102: 529 –540, 2007;
doi:10.1152/japplphysiol.01126.2005.
Invited Review
Energy sensing and regulation of gene expression in skeletal muscle
Damien Freyssenet
Unité Physiologie et Physiopathologie de l’Exercice et Handicap, EA3062, Université Jean Monnet, Saint-Etienne, France
energy-sensing molecules; endurance training; mitochondria
have maintained a longstanding interest
in bioenergetics, largely because of the exceptional capacity of
skeletal muscle to increase its energy production during exercise. As knowledge of muscle bioenergetics has progressively
increased and numerous training-induced adaptations were
described, many exercise physiologists have focused on dissecting the molecular events and regulatory mechanisms involved in the cellular response. Particular attention has been
paid to exercise-induced extracellular factors, intracellular factors, signaling pathways, and transcription factors, as well as
the regulatory influences responsible for modulating the cellular response (reviewed in Ref. 84). A number of studies have
suggested that the adaptive response of skeletal muscle to
exercise is likely to involve energy metabolism through regulation of key signaling molecules and transcription factors.
Figure 1 illustrates the organizational hierarchy of molecular
components that are believed to be involved in the energy
dependence of nuclear gene expression in skeletal muscle.
The purpose of this review is to examine the role of energy
sensing for regulation of nuclear gene expression (transcription
and translation) in skeletal muscle and to show how energy
sensing may contribute to muscle plasticity. We will first focus
on energy-sensing molecules, which are herein defined as
proteins capable of transducing variations in energy homeostasis into alterations in gene expression. We will also assess how
endurance training can influence the response of energy-sensing molecules and how mitochondria act as sensors for variations in energy homeostasis in skeletal muscle. Specificity of
mitochondrial signaling during skeletal muscle regeneration
will be also developed. Finally, we will discuss some new
EXERCISE PHYSIOLOGISTS
Address for reprint requests and other correspondence: D. Freyssenet, Unité
Physiologie et Physiopathologie de l’Exercice et Handicap, Faculté de Médecine Jacques Lisfranc, 15 rue Ambroise Paré, 42023 Saint-Etienne Cedex 2,
France (e-mail : [email protected]).
http://www. jap.org
candidates that could be tested for potential roles in energy
sensing in skeletal muscle.
ROLE OF ENERGY-SENSING MOLECULES IN REGULATING
GENE EXPRESSION IN SKELETAL MUSCLE
All eukaryotic cells must maintain high ATP-to-ADP ratios.
During skeletal muscle contractions, ATP is hydrolyzed and
free ADP is produced by the myosin ATPase reaction (Fig. 2).
Depending on the extent of ATP demand and the capacity of
the mitochondria to regenerate ATP, energy homeostasis may
be modestly (ATP demand can be matched by mitochondrial
ATP synthesis) or more robustly (ATP demand exceeds mitochondrial ATP synthesis) altered. A lack of balance between
ATP demand and mitochondrial ATP synthesis, if great
enough, will increase lactate concentration and further increase
the free ADP-to-ATP ratio, which will then be amplified into a
much larger increase in the AMP-to-ATP ratio via adenylate
kinase.
The idea that variations in cellular energy status may contribute to the regulation of nuclear gene expression is not new,
but the recent identification of several molecules capable of
sensing and transducing changes in cellular energy status into
altered nuclear gene expression has provided researchers with
a basis for understanding this process in greater detail.
AMP-activated protein kinase. ␤-Guanidinopropionic acid
(␤-GPA) is a creatine analog that depletes high-energy phosphate compounds, including phosphocreatine and ATP, from
skeletal muscle (43). Chronic feeding with ␤-GPA reportedly
stimulates mitochondrial biogenesis in rat skeletal muscle,
particularly in fast-twitch skeletal muscle (44, 137). However,
transgenic mice expressing a dominant-negative mutant of the
AMP-activated protein kinase (AMPK) ␣2-subunit do not
show mitochondrial biogenesis in response to ␤-GPA feeding
(166), suggesting that AMPK is capable of sensing and transducing changes in cellular energy status (i.e., depletion of
8750-7587/07 $8.00 Copyright © 2007 the American Physiological Society
529
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
Freyssenet D. Energy sensing and regulation of gene expression in skeletal muscle.
J Appl Physiol 102: 529 –540, 2007; doi:10.1152/japplphysiol.01126.2005.—Major
modifications in energy homeostasis occur in skeletal muscle during exercise. Emerging evidence suggests that changes in energy homeostasis take part in the regulation of
gene expression and contribute to muscle plasticity. A number of energy-sensing
molecules have been shown to sense variations in energy homeostasis and trigger
regulation of gene expression. The AMP-activated protein kinase, hypoxia-inducible
factor 1, peroxisome proliferator-activated receptors, and Sirt1 proteins all contribute to
altering skeletal muscle gene expression by sensing changes in the concentrations of
AMP, molecular oxygen, intracellular free fatty acids, and NAD⫹, respectively. These
molecules may therefore sense information relating to the intensity, duration, and
frequency of muscle exercise. Mitochondria also contribute to the overall response,
both by modulating the response of energy-sensing molecules and by generating their
own signals. This review seeks to examine our current understanding of the roles that
energy-sensing molecules and mitochondria can play in the regulation of gene expression in skeletal muscle.
Invited Review
530
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
high-energy phosphates) into nuclear gene expression changes.AMPK, an energy-sensing enzyme activated by 5⬘-AMP,
exists as heterotrimeric complexes containing a catalytic
␣-subunit and two regulatory subunits, ␤ and ␥. Binding of
AMP to AMPK stimulates phosphorylation and activation of
AMPK by the protein kinase, LKB1 (also known as STK11)
(56, 133). AMPK activation also occurs directly as a result of
Ca2⫹ signaling in myotubes (45). In tissues other than skeletal
muscle, AMPK may also be phosphorylated and activated by
CaMK kinase (57, 67, 161). The ␤-subunits of AMPK contain
a conserved central domain that binds glycogen (66) and would
be involved in negative regulation of AMPK activity. Since
AMPK has been extensively reviewed elsewhere (22, 54, 55,
76, 80, 157), we will herein focus on a small subset of target
proteins that are particularly relevant to skeletal muscle gene
expression.
Fig. 2. Simplified overview of muscle bioenergetic highlighting messengers involved in
signal transduction. Energy is provided by the
oxidation of glucose, glycogen, and triglycerides. The relative contribution of these substrates depends on exercise intensity and training status. ADP is formed by the myosin
ATPase reaction. Cytosolic creatine kinase
(CK), in conjunction with their mitochondrial
isoenzymes (mt-CK), shuttle ADP from cytosol into the mitochondrial intermembrane
space. Direct diffusion of ADP also participates in the transport of adenylate. In fasttwitch muscles, AMP can be metabolized to
inosine monophosphate (IMP) and ammonia
(NH3) by AMP deaminase (AMPd). In slowtwitch muscles, AMP is predominantly metabolized to adenosine by the 5⬘-nucleotidase (5⬘Nase). G6P, glucose-6-phosphate. LDH, lactate dehydrogenase; Cr, creatine; AK,
adenylate kinase; CrP, creatine phosphate; PPi,
inorganic pyrophosphate.
J Appl Physiol • VOL
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
Fig. 1. Diagram summarizing the theoretical
framework used as the basis of this review.
Listed are messengers and energy sensing
molecules that have been shown to be responsive to variations in energy homeostasis.
Even if not represented, mitochondria are
key contributors in the overall response both
by modulating the response of energy sensing molecules and by generating their own
signaling. AMPK, AMP-activated protein kinase; cGC, cytosolic guanylyl cyclase;
HIF-1, hypoxia-inducible factor-1; MondoA,
basic helix loop helix-leucine zipper partner
for the Max-like protein, Mlx; PPAR, peroxisome proliferator-activated receptors; Sirt1,
sirtuin 1; Sirt3, sirtuin 3.
Invited Review
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
J Appl Physiol • VOL
As exercise intensity increases, muscle cells are subject to
substantial decreases in intracellular PO2 (iPO2) (28, 119).
Although recent evidence suggests that iPO2 does not decrease
linearly with increasing metabolic rate (118), iPO2 falls to a
level of ⬃4 Torr at a maximum work rate (118). It is therefore
possible that HIF-1␣ protein accumulates in response to reduced PO2 in skeletal muscles undergoing high-intensity exercise, suggesting that this oxygen-sensitive pathway could be
involved in muscular adaptation to exercise. Consistent with
this hypothesis, HIF-1␣ protein levels and HIF-1␣ DNA binding activity increases markedly in human skeletal muscle
immediately after a one-legged knee-extension exercise, along
with a concurrent decrease in von Hippel Lindau protein levels
(1), suggesting that the observed increase in HIF-1␣ protein
level is due to protein stabilization. Functional study in HIF-1␣
knockout mice demonstrates that the increased expression of
HIF-1␣-dependent target genes (e.g., vascular endothelial
growth factor, glucose transporter 4) normally observed in
wild-type animals in response to skeletal muscle exercise is
strongly diminished in HIF-1␣ knockout mice, demonstrating
that HIF-1␣ plays an important role in muscle adaptation to
exercise (97).
These data collectively suggest that HIF-1␣ plays a significant role in mediating muscle adaptations to muscle contraction. However, some questions remain unanswered. In particular, the regulation of HIF-1␣ prolyl hydroxylases in response
to endurance exercise has not yet been investigated. It is also
unknown whether O2 is evenly distributed throughout muscle
fibers during exercise or whether O2 microdomains exist for
heterogeneous regulation of HIF-1␣ activity. The regulation of
HIF-1␣ expression in response to acute bouts of exercise is
also poorly understood (94).
PPARs. The PPARs constitute a class of metabolic sensors
within the nuclear receptor family, with three isotypes
(PPAR␣, PPAR␤/␦, and PPAR␥) cloned to date from humans
and rodent (reviewed in Refs. 32, 33). In skeletal muscle, the
relative expression levels of PPARs are, in descending order,
PPAR␤/␦ ⬎ PPAR␣ ⬎ PPAR␥ (48). The PPARs bind DNA at
the PPAR response element as heterodimers with the retinoid
X receptor (48). Either partner can regulate the transcriptional
activity of the DNA-bound complex by interacting with its
cognate ligand either singly or in concert. A large number of
genes encoding proteins involved in mitochondrial metabolism
have been shown to be PPAR responsive in skeletal muscle,
including those encoding proteins responsible for fatty acid
transport and activation, mitochondrial ␤-oxidation, and energy uncoupling (37, 96, 106, 144, 155).
Studies have shown that numerous fatty acids, especially
unsaturated fatty acids, can bind to PPAR␣, PPAR␤/␦, and
PPAR␥ with varying affinities to control target gene transcription (reviewed in Refs. 32, 33), indicating that the
PPARs act as important metabolic sensors. Since oxidation
of triglycerides from extramuscular sources and intramuscular stores is used by skeletal muscle fibers to provide
energy during endurance exercise (81), PPARs may regulate
gene expression in response to variations in the levels of
intramuscular fatty acids during endurance exercise. This
notion is strongly supported by a gain-loss of function study
in mice, showing that targeted expression of an activated
form of PPAR␦ in skeletal muscle increased the number of
type I muscle fiber, the expression of genes involved in
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
AMPK is activated in skeletal muscle during exercise (25,
107, 158) and electrically stimulated contraction (4, 69, 107,
151). In skeletal muscle, AMPK activates expression of peroxisome proliferator-activated receptor (PPAR) ␣ (89),
PPAR␥ coactivator-1␣ (PGC-1␣) (89, 143, 146), and nuclear
respiratory factor 1 (NRF-1) (10). Consistent with its positive
effect on PGC-1␣ and NRF-1 expression, AMPK has also been
shown to stimulate mitochondrial biogenesis in skeletal muscle
fibers (10, 115, 143, 159). The mechanisms by which AMPK
regulates PGC-1␣ expression are currently unknown, but one
possibility is that myocyte enhancer factor-2 and the transcriptional coactivator, p300 histone acetyl transferase, which are
both regulated by AMPK (162, 165), promote transactivation
of the PGC-1␣ gene promoter (53, 114). It is also important to
note that AMPK may not be essential for adaptations of
skeletal muscle to endurance exercise training (62, 77) and that
additional signaling pathways are involved in triggering muscular adaptations to endurance training (recently reviewed in
Ref. 84). Another recently identified target for AMPK in
skeletal muscle is the protein kinase B (Akt)/mammalian target
of rapamycin (mTOR) pathway, which is involved in protein
synthesis and muscle hypertrophy (reviewed in Refs. 49, 75).
AMPK activation has been found to reduce protein synthesis
by 45% in muscle homogenates, in association with decreased
activation of Akt, mTOR, ribosomal protein S6 kinase, and
eukaryotic initiation factor 4E-binding protein (16). In addition, AMPK is believed to promote phosphorylation of tuberous sclerosis complex 2 (TSC2) (72, 145), which forms a
complex with TSC1 to allow conversion of the small G-protein
Ras homolog enriched in brain (Rheb) to a GDP-bound form
that no longer activates mTOR (55). This AMP-mediated
cascade may account for a number of specific muscular adaptations to endurance training (4), including the observation that
endurance training is generally not associated with a gain in
muscle mass. A number of other downstream targets of AMPK
are potentially relevant for skeletal muscle physiology, including the HuR mRNA binding protein, which is involved in
protein stabilization during cytokine-induced skeletal muscle
cachexia (34), as well as the forkhead box O (Foxo) 1a
transcription factor (9) and hypoxia inducible factor-1 (HIF-1)
(70, 88). However, no studies to date have definitively shown
that AMPK activates these target proteins in skeletal muscle.
HIF-1. All mammalian cells have the ability to sense and
respond to changes in O2 concentration; this ability is based on
the action of HIF-1, which functions as a master transcriptional
factor in response to changes in tissue oxygenation. HIF-1 is a
heterodimeric protein composed of HIF-1␣ and HIF-1␤ subunits (154). Under normoxic conditions, the HIF-1␣ protein is
subject to hydroxylation by prolyl hydroxylases (for review,
see Refs. 8, 128), which triggers binding of von Hippel Lindau
protein to HIF-1␣ and targets it for proteasomal degradation. In
contrast, the prolyl hydroxylases do not function under low O2
concentrations, leading to accumulation of HIF-1␣, heterodimerization of HIF-1␣ with HIF-1␤, and subsequent activation of HIF-1-dependent target gene expression. These targets include the gene encoding vascular endothelial growth
factor, as well as genes that enable cells to survive O2 deprivation by providing an O2-independent means of ATP production (glucose transporters and glycolytic enzymes) or by inhibiting hypoxia-induced apoptosis (129, 130).
531
Invited Review
532
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
J Appl Physiol • VOL
of PGC-1␣ by Sirt1 appears to depend on the cellular context.
In cultured hepatocytes, Sirt1-mediated deacetylation of
PGC-1␣ stimulates expression of neoglucogenic genes (122),
whereas it causes a decrease in mitochondrial gene expression
and cellular oxygen consumption in neural cells (109).
Whether Sirt1 activates or inhibits the coactivating function of
PGC-1␣ in skeletal muscle is currently unknown and will
require further study, particularly in the context of exercise.
Sirt1 also regulates the activity of members of the Foxo
family (20, 104, 163), a class of molecules that has been
described to regulate skeletal muscle mass (125, 141). It
remains to be determined whether this function of Sirt1
reported in vitro in nonmyogenic cells is of biological
relevance in skeletal muscle.
It should be noted that NAD⫹ activates AMPK in human
kidney cells (116), suggesting that the NAD⫹-to-NADH
ratio may exert another level of regulatory influence on
energy-sensing molecules. Finally, in Cos-7 cells, Sirt1
deacetylates and activates acetyl-CoA synthase 1 (51),
which is the enzyme responsible for catalyzing the formation of acetyl-CoA from acetate, CoA, and ATP. Regulation
of acetyl-CoA levels is critical for MyoD-dependent transactivation (35) and the availability of free fatty acids.
Although these results should be interpreted within the
context of tissue-specific Sirt1 effects, the studies collectively suggest that Sirt1 may establish regulatory cross-talk
among various energy-sensing molecules. Clearly, a complete characterization of the role of Sirt1 in skeletal muscle,
as well as the identification of its molecular partners, will be
important to draw precise conclusions on its physiological
relevance in skeletal muscle.
REGULATION OF ENERGY-SENSING MOLECULES IN
RESPONSE TO ENDURANCE TRAINING
The concept of the preservation of energy homeostasis in
endurance-trained skeletal muscle, initially explored by Holloszy and Booth (60) and later developed by others (50, 61,
64), is of critical importance when considering the energy
dependence of nuclear gene expression in skeletal muscle.
Central to this concept is the observation that the mitochondrial
content of skeletal muscle increases in response to endurance
training (for review, see Ref. 64). Thus the same cellular rate
of mitochondrial oxidative phosphorylation is attained with a
lower rate of oxidative phosphorylation “per mitochondrion”
for any absolute workload in muscle fibers with elevated
mitochondrial content. ATP and creatine phosphate concentrations decrease to a smaller degree, and the free ADP concentration increases to a lesser extent in cells with elevated
mitochondrial contents at the same absolute work rate (30, 38).
Better preservation of free ADP lowers the rate of AMP
formation, ultimately decreasing the activation of AMPK and
its upstream activators. These hypotheses are supported by
experimental evidences (68, 111, 164), collectively showing
that preservation of energy homeostasis in response to endurance training contributes to the regulation of AMPK activity.
Other well-described metabolic adaptations of skeletal muscle
to endurance training may also participate in the regulation of
energy-sensing molecules. A considerably lower increase in
lactate concentration is observed after endurance training when
exercise is performed at the same submaximal workload (30).
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
mitochondrial biogenesis, and mitochondrial metabolism,
and endurance performance in these mice (155). Similarly,
the mRNA levels of the slow isoform of troponin I, cytochrome c, and subunit IV of cytochrome oxidase are all
upregulated in wild-type mice receiving the PPAR␦-specific
agonist, GW501516. Conversely, PPAR␦-null mice can
only sustain 30 – 40% of the running time and distance of
their wild-type controls (155). Although this study does not
show direct activation of PPAR␦ by fatty acids in response
to endurance exercise or endurance training, the data
strongly support the notion that PPAR␦ activation is an
important mediator of muscular adaptation to endurance
exercise.
Sirtuin 1. Chung et al. (27) originally suggested that the
ratio NAD⫹/NADH might be involved in the regulation of
nuclear gene expression following their observation that the
binding of proteins on an 8-bp response element (called the
REBOX) present in the promoters of the genes encoding
adenine nucleotide translocase and ATP synthase-␤ was
sensitive to variations in NADH concentration. However,
the redox status of NADH was definitively shown to be
involved in nuclear gene expression with the subsequent
discovery that NAD⫹ is an essential substrate for a family of
class III histone deacetylases, collectively called the sirtuins. The sirtuins, which are enzymes capable of coupling
the cleavage of NAD⫹ to nuclear gene expression (reviewed
in Refs. 15, 31), were originally identified in yeast as
NAD-dependent proteins that deacetylate histones and participate in the regulation of longevity (71). In mammals,
Sirtuin 1 (Sirt1) deacetylates histone proteins as well as
some non-histone proteins, including the TAF68 TATA
box-binding protein (108) and p53 (95, 152). This unique
property implies that Sirt1 may function as an energy sensor
linking energy metabolism to transcriptional regulation.
Decreases in the NAD⫹-to-NADH ratio have been observed
during muscle differentiation (47), suggesting that Sirt1 may
participate in the regulation of myogenesis. Consistent with
this notion, inhibition of Sirt1 induces premature differentiation of C2C12 myoblasts, whereas activation of Sirt1 via
elevation of the NAD⫹-to-NADH ratio inhibits muscle cell
differentiation (47). Sirt1 appears to exert this regulatory function by modulating the activity of the myogenic transcription
factor, MyoD (47), and the histone acetylase, p300 (17, 47).
However, it appears as though the role of Sirt1 is not limited to
myogenic differentiation. During muscle contractions, the cytosolic NAD⫹-to-NADH ratio is subject to dynamic fluctuations imposed by the rate of NAD⫹ reduction to NADH, which
is mediated by glyceraldehyde 3-phosphate dehydrogenase,
and the rate of NADH oxidization to NAD⫹, which is collectively governed by lactate dehydrogenase, the glycerol phosphate shuttle, and the malate aspartate shuttle (120). The
mitochondrial monocarboxylate transporter, which mediates
the uptake of pyruvate into mitochondria, also indirectly contributes to the regulation of the NAD⫹-to-NADH ratio. The
lactate-to-pyruvate ratio increases as exercise intensity increases, shifting the lactate dehydrogenase equilibrium and
decreasing the NAD⫹-to-NADH ratio (30), potentially leading
to a decrease in Sirt1 activity. Interestingly, Sirt1 regulates the
activity of PGC-1␣ (109, 122), suggesting that modulation of
Sirt1 activity may participate in the regulation of mitochondrial
biogenesis. However, it is important to note that the regulation
Invited Review
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
Table 1. Examples of genes that are altered in response to
variations in mitochondrial respiratory chain activity
Gene
533
THE MITOCHONDRION: AN ENERGY-SENSING ORGANELLE
THAT PARTICIPATES IN THE REGULATION OF NUCLEAR
GENE EXPRESSION
Cell Type (Ref. No.)
2⫹
C2C12
C2C12
C2C12
C2C12
Tumor invasion and cell cycle
Cathepsin L (1)
p21 (1)
PCNA (2)
TGF-␤ (1)
C2C12 (2, 13)
L6E9 (40)
L6E9 (40)
C2C12 (2, 14)
Apoptosis
Bad (1)
Bc12 (1)
Bid (1)
C2C12 (12, 14)
C2C12 (12, 14)
C2C12 (12, 14)
Gene expression
ATF-2 (1)
cRe1 (1)
1␬B␣ (1)
Myc (1)
Myogenin (2)
NFAT (1)
p50 (1)
p53 (1)
C2C12 (11)
C2C12 (11, 13)
C2C12 (11, 13)
C2C12 (14, 132)
QM7 (121, 132)
C2C12 (11)
C2C12 (11, 13)
C2C12 (14, 132)
(11)
(2, 14)
(11, 13, 14)
(2, 11, 13, 14)
In 1956, Otto Warburg observed that experimentally induced carcinogenesis of rodent cells was accompanied by
reduced respiration-coupled oxidative metabolism and increased glycolysis (156), raising the intriguing possibility that
the original noncarcinogenic phenotype of these mammalian
cells was regulated by mitochondrial oxidative phosphorylation. The existence of mitochondrial regulation of nuclear gene
expression was later demonstrated directly in C2C12 skeletal
myoblast, wherein depletion of mitochondrial DNA (mtDNA)
and subsequent disruption of mitochondrial membrane potential results in enhanced levels of several Ca2⫹-responsive
proteins (calcineurin, nuclear factor of activated T cells) and
Ca2⫹ handling proteins (ryanodine receptor 1, calreticulin)
(11). When the partial depletion of mtDNA is reverted in
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
Ca homeostasis
Calcineurin (1)
Calreticulin (1)
Calsequestrin (1)
RyR 1 (1)
All studies were performed in vitro. Arrows indicate whether gene expression was increased (1) or decreased (2). Decrease in mitochondrial respiratory chain activity was induced by ethidium bromide (2, 11–14), chloramphenicol (121, 132), and respiratory chain-interfering drugs (2, 11–13). Increase in
mitochondrial respiratory chain activity was induced by pyruvate (40). C2C12,
mouse muscle cells; L6E9, rat muscle cells; QM7, quail muscle cells; RyR 1,
ryanodine receptor 1; TGF-␤, transforming growth factor-␤; ATF-2, activating
transcription factor-2; PCNA, proliferating cell nuclear antigen; NFAT, nuclear factor of activated T cells.
Preservation of the pyruvate-to-lactate ratio blunts the decrease
in the NAD⫹-to-NADH ratio, thus potentially contributing to
the regulation of Sirt1 activity. Muscle glycogen stores are
depleted less rapidly during standardized exercise in the trained
state. This glycogen-sparing effect may contribute to reduce
AMPK activation in trained muscles. Indeed, AMPK activity
and high glycogen levels show an inverse relationship in
skeletal muscle (160), but low muscular glycogen levels at the
onset of exercise were shown to enhance AMPK activation vs.
muscles with high glycogen contents (140). Finally, the decrease in the oxidation of carbohydrate during submaximal
exercise in response to endurance training is compensated for
by a proportional increase in fat oxidation (61). This may alter
intramuscular free fatty acid levels during submaximal exercise, suggesting that PPAR transcriptional activity may also be
modified in response to endurance training.
Another important possibility in considering the energy
dependence of nuclear gene expression is that endurance training could alter the expression of energy-sensing molecules.
Increased expression of AMPK isoforms ␥3 (41), ␣1, ␤2 and ␥1
(46), and ␣1 (111) and decreased expression of AMPK ␥3 (46),
have been reported in response to exercise training in rats (41)
and human skeletal muscles (46, 111). Changes in PPAR␣
(124), HIF-1␣, and HIF-1␤ mRNA levels (94) have also been
reported in response to endurance training. Such changes may
alter the sensitivity of energy-sensing molecules to variations
in energy homeostasis.
J Appl Physiol • VOL
Fig. 3. Schematic overview of mitochondrial signaling. Under conditions
where cell energy demand is challenged, mitochondria produce ATP to meet
the energy demand but also initiate and modulate signaling cascades participating in nuclear gene expression. This function is referred to as mitochondrial
signaling or mitochondrial retrograde signaling. The identity of some messengers and downstream effectors is discussed in the text.
102 • FEBRUARY 2007 •
www.jap.org
Invited Review
534
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
direct 0.15– 0.5% of the total O2 consumption to ROS generation (3). Mitochondrially derived ROS function in numerous
signaling processes in a variety of cell types, where they
regulate the redox status of protein kinases (85, 110, 117, 142)
and transcription factors (24, 63, 82, 91, 117). In cardiomyocytes, mitochondrial ROS production is associated with p38
MAPK phosphorylation (85) and JNK activation (36). Myotubes treated with TNF-␣ show increased ROS release from
mitochondria (91), leading to NF-␬B activation and myofibrillar protein loss (92). ROS production may also be involved in
the regulation of myogenic differentiation. Exposure of differentiating myoblasts to hydrogen peroxide almost totally abolished muscle-specific protein expression and myogenic differentiation, and this effect is reversed when cells are incubated
with the ROS scavenger, N-acetyl cysteine (86). The contribution of mitochondrially derived ROS in this situation remains
to be determined.
Contractile activity in skeletal muscle increases the concentration of superoxide anions in both the extracellular (98, 99)
and intracellular compartments (100, 138), suggesting that
muscle exercise may be accompanied by increased mitochondrial ROS production. This raises an important question concerning the relationship between oxidant formation and mitochondrial oxidative phosphorylation rate (debated in Refs. 5,
65, 79, 90). Mitochondria isolated from adult mouse skeletal
Fig. 4. Mechanisms for mitochondrial reactive oxygen species (ROS) generation. Complexes I and III are responsible for most of superoxide anion (O2•⫺)
production. Release of O2•⫺ in the intermembrane space appears to mainly occur at complex III (5, 18, 149). O2•⫺ is rapidly converted to hydrogen peroxide
(H2O2) by manganese superoxide dismutase (MnSOD) on the inner side and copper/zinc superoxide dismutase (CuZnSOD) on the outer side. A number of
regulatory influences may profoundly affect mitochondrial ROS production. An increase in mitochondrial matrix Ca2⫹ concentration activates pyruvate
dehydrogenase and enzymes of tricarboxylic acid (TCA) cycle, isocitrate dehydrogenase (ICDH) and ␣-ketoglutarate dehydrogenase (␣-KGDH) (101, 102), as
well as complex IV of the respiratory chain (79, 93), further contributing to increase mitochondrial ROS production (79, 87, 93). Mitochondrial Ca2⫹ is also a
potent activator of nitric oxide (NO䡠) production by the mitochondrial nitric oxide synthase (mtNOS) (29, 58, 148). NO䡠 can bind iron and copper atoms at the
active site of cytochrome c oxidase (135), thus reversibly inhibiting cytochrome c oxidase activity by reducing the affinity of the enzyme for molecular oxygen
(O2). This ultimately increases the formation of ROS by mitochondria (23, 103). Recent evidences also suggest that ␣-KGDH directly participates in the
mitochondrial production of ROS (139). CoQ, coenzyme Q; Cyt c, cytochrome c; ⌬⌿, mitochondrial membrane potential; FAD, flavine adenine dinucleotide;
i.m., inner membrane; mt-Uni, mitochondrial Ca2⫹ uniporter; NAD⫹, nicotinamide adenine dinucleotide; o.m., outer membrane; I, II, III, and IV refer to the
protein complexes of the electron transport chain.
J Appl Physiol • VOL
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
C2C12 myoblasts, the expression of the nuclear genes encoding these proteins reverts to near control levels, inferring the
direct link between changes in mitochondrial membrane potential and nuclear gene expression. Table 1 shows several
examples of genes that are altered in response to variations in
mitochondrial respiratory chain activity in skeletal muscle
cells. Taken together, these data unambiguously demonstrate
that in addition to generating ATP for energy tasks, mitochondria also initiate and modulate signaling cascades that participate in nuclear gene expression. This function, referred to as
mitochondrial signaling or mitochondrial retrograde signaling
(21), is defined as mitochondrial respiration-dependent pathway of communication between mitochondrion and nucleus
leading to changes in nuclear gene (Fig. 3). Mitochondria may
therefore be viewed as energy-sensing organelles that adapt
nuclear gene expression under conditions where the cell energy
demand is challenged. The cascade of events connecting the
mitochondria to nuclear gene expression is not yet fully understood, but several potential signaling molecules and triggers
have been identified.
Reactive oxygen species. Within the cell, the main sources of
reactive oxygen species (ROS) are the mitochondria, which
produce superoxide anions mainly at complexes I and III of the
respiratory chain (Fig. 4). In skeletal muscle, isolated mitochondria respiring in absence of ADP (state IV of respiration)
Invited Review
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
J Appl Physiol • VOL
ing capacity participates in the regulation of gene expression in
skeletal muscle fibers.
SPECIFICITY OF MITOCHONDRIAL SIGNALING DURING
SKELETAL MUSCLE REGENERATION
The signaling function of mitochondria appears to be particularly relevant to the regulation of myogenic differentiation.
For example, respiration-deficient primary myoblasts depleted
of mitochondrial DNA or myoblasts treated with chloramphenicol and tetracycline fail to differentiate (52, 59, 83, 121),
whereas stimulation of mitochondrial metabolism induces cell
cycle arrest in L6 muscle cells (40), and stimulation of mitochondrial biogenesis by the exogenous expression of p43, a
triiodothyronine-dependent mitochondrial transcription factor,
increases muscle cell differentiation (121, 132). Consistent
with these observations, myogenic differentiation is accompanied by strong stimulation of mitochondrial biogenesis both in
vitro (105) and in vivo during skeletal muscle regeneration
(39). The existence of distinct and opposite metabolic profiles
as a function of cellular state is also illustrated by the observation that quiescent satellite cells have a strong mitochondrial
oxidative potential and a low glycolytic potential when compared with proliferating myoblasts (6, 7). Taken together, these
findings support the notion that mitochondrial metabolism and
mitochondrial content may exert a myogenic influence, and
stimulation of mitochondrial biogenesis is necessary for the
commitment of myoblasts into myotubes. Seyer and colleagues
(132) recently provided new insights into the myogenic influence of mitochondria by showing that stable overexpression of
p43 decreases the mRNA and protein levels of c-myc. Notably,
this is true even in serum-rich medium, further illustrating that
p43 potently blocks proliferation. Overexpression of p43 stimulates expression of the myogenic transcription factor myogenin (121, 132), whereas c-myc overexpression abrogates the
ability of myogenin and MyoD to induce terminal differentiation (132). Therefore, mitochondria appear to exert their myogenic influence by regulating both cell cycle withdrawal and
the expression of myogenic proteins, possibly by regulating
c-myc expression. Future studies will be required to determine
the pathways and target genes involved in the myogenic
influence of mitochondria, as well as to explore the relationship
between the pathways regulating mitochondrial biogenesis and
myogenic differentiation.
FUTURE DIRECTIONS AND CONCLUDING REMARKS
The concept of the energy dependence of nuclear gene
expression in skeletal muscle is still taking shape, but our
current understanding provides a comprehensive and synthetic
view of muscle plasticity and the relationships among various
metabolic and genetic aspects. Additional studies aimed at
identifying the various messengers, energy-sensing molecules,
downstream effector molecules (kinases, phosphatases, transcription and translation factors) and target genes will benefit
our understanding of energy sensing in signaling. For instance,
messengers whose activities are regulated by changes in energy
homeostasis, such as acetyl-CoA (35) and cytochrome c (42),
could be studied in further detail. Additional knowledge regarding the role of amino acids in regulating protein translation
and gene expression will also be useful in this context (for
review, see Ref. 78). Several of the recently identified energy-
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
muscle show significant increases in hydrogen peroxide release
after contractile activity compared with precontraction values
(150), supporting the notion that mitochondria significantly
contribute to exercise-induced ROS production (79). A recent
study further showed that contraction-induced AMPK activity
and phosphorylation were blocked with the ROS scavenger
N-acetylcysteine, suggesting that exercise-induced mitochondrial ROS production may be involved in the regulation of
AMPK-dependent pathway (126). One may also ask the question about the physiological relevance of mitochondrial ROS in
the response of skeletal muscle to endurance training. This
must be critically examined in light of the variations in antioxidant capacity that may occur in response to endurance
training, some reports describing an increase in antioxidant
capacity (74, 131, 147, 153), whereas others report unchanged
antioxidative defense (147, 153). The extent of endurance
training-induced mitochondrial biogenesis has also to be taken
into account. All these parameters will combine to determine
the importance of mitochondrial ROS signaling for a given
exercise intensity after endurance training. Clearly, additional
experiments will be required to determine if and to what degree
mitochondrial ROS production is involved in the adaptive
response of skeletal muscle to endurance exercise and training.
Calcium ion. Recent studies have identified a mitochondrial
component to the regulation of Ca2⫹-dependent signaling
events (2, 11, 13, 14). For example, disruption of the mitochondrial membrane potential triggers a two- to threefold
increase in the cytosolic Ca2⫹ concentration of C2C12 myoblasts, accompanied by changes in the expression levels of
Ca2⫹-responsive proteins (e.g., calcineurin, nuclear factor of
activated T cells) and genes of the NF-␬B pathway (11) (see
also Table 1). In contrast, removal of free Ca2⫹ by chelating
agents reverses the effect of mitochondrial signaling on nuclear
gene expression (2, 13, 14). Furthermore, the activation of
Ca2⫹-responsive factors is abolished in calcineurin A ␤⫺/⫺
cells, as well as in cells treated with the calcineurin inhibitor
FK506, showing that calcineurin may be a downstream effector of mitochondrial signaling (13).
In skeletal muscle fibers, mitochondria actively participate
in the regulation of Ca2⫹ homeostasis by rapidly and efficiently
accumulating Ca2⫹ in the mitochondrial matrix (19). The
Ca2⫹-buffering capacity of mitochondria is tightly linked to the
rate of mitochondrial oxidative phosphorylation. Addition of
respiratory substrates to permeabilized skeletal muscle fibers
greatly diminishes frequency of Ca2⫹ sparks from the sarcoplasmic reticulum (73), which is directly proportional to the
mitochondrial content and to the ability of mitochondria to
accumulate Ca2⫹ in the matrix, due to the close proximity of
some mitochondria to Ca2⫹ release sites (73, 136). In contrast,
addition of a nonmetabolizable substrate or a mitochondrial
inhibitor promotes Ca2⫹ sparks from the sarcoplasmic reticulum (73), illustrating a decrease in the Ca2⫹-buffering capacity
of mitochondria. Clearly, the buffering capacity of mitochondria may be particularly relevant in spatiotemporal regulation
of cytosolic Ca2⫹ concentrations in contracting muscles fibers.
This may contribute to modulating the activity of Ca2⫹-responsive proteins involved in gene expression, such as CaMK,
calcineurin, or Ca2⫹-sensitive protein kinase C (PKC) isoforms
(reviewed in Refs. 26, 84). Additional work will be required to
determine the extent to which the mitochondrial Ca2⫹ buffer-
535
Invited Review
536
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
ACKNOWLEDGMENTS
Thierry Busso is gratefully acknowledged for helpful comments during the
preparation of the manuscript.
GRANTS
This work was supported by a Ligue Contre le Cancer Comité de la Loire
grant to D. Freyssenet.
REFERENCES
1. Ameln H, Gustafsson T, Sundberg CJ, Okamoto K, Jansson E,
Poellinger L, Makino Y. Physiological activation of hypoxia inducible
factor-1 in human skeletal muscle. FASEB J 19: 1009 –1011, 2005.
2. Amuthan G, Biswas G, Zhang SY, Klein-Szanto A, Vijayasarathy C,
Avadhani NG. Mitochondria-to-nucleus stress signaling induces phenotypic changes, tumor progression and cell invasion. EMBO J 20: 1910 –
1920, 2001.
J Appl Physiol • VOL
3. Anderson EJ, Neufer PD. Type II skeletal myofibers possess unique
properties that potentiate mitochondrial H2O2 generation. Am J Physiol
Cell Physiol 290: C844 –C851, 2006.
4. Atherton PJ, Babraj J, Smith K, Singh J, Rennie MJ, Wackerhage
H. Selective activation of AMPK-PGC-1alpha or PKB-TSC2-mTOR
signaling can explain specific adaptive responses to endurance or resistance training-like electrical muscle stimulation. FASEB J 19: 786 –788,
2005.
5. Balaban RS, Nemoto S, Finkel T. Mitochondria, oxidants, aging. Cell
120: 483– 495, 2005.
6. Barani AE, Durieux AC, Sabido O, Freyssenet D. Age-related changes
in the mitotic and metabolic characteristics of muscle-derived cells.
J Appl Physiol 95: 2089 –2098, 2003.
7. Barani AE, Sabido O, Freyssenet D. Mitotic activity of rat muscle
satellite cells in response to serum stimulation: relation with cellular
metabolism. Exp Cell Res 283: 196 –205, 2003.
8. Bardos JI, Ashcroft M. Negative and positive regulation of HIF-1: a
complex network. Biochim Biophys Acta 1755: 107–120, 2005.
9. Barthel A, Schmoll D, Kruger KD, Roth RA, Joost HG. Regulation of
the forkhead transcription factor FKHR (FOXO1a) by glucose starvation
and AICAR, an activator of AMP-activated protein kinase. Endocrinology 143: 3183–3186, 2002.
10. Bergeron R, Ren JM, Cadman KS, Moore IK, Perret P, Pypaert M,
Young LH, Semenkovich CF, Shulman GI. Chronic activation of AMP
kinase results in NRF-1 activation and mitochondrial biogenesis. Am J
Physiol Endocrinol Metab 281: E1340 –E1346, 2001.
11. Biswas G, Adebanjo OA, Freedman BD, Anandatheerthavarada
HK, Vijayasarathy C, Zaidi M, Kotlikoff M, Avadhani NG. Retrograde Ca2⫹ signaling in C2C12 skeletal myocytes in response to mitochondrial genetic and metabolic stress: a novel mode of inter-organelle
crosstalk. EMBO J 18: 522–533, 1999.
12. Biswas G, Anandatheerthavarada HK, Avadhani NG. Mechanism of
mitochondrial stress-induced resistance to apoptosis in mitochondrial
DNA-depleted C2C12 myocytes. Cell Death Differ 12: 266 –278, 2005.
13. Biswas G, Anandatheerthavarada HK, Zaidi M, Avadhani NG.
Mitochondria to nucleus stress signaling: a distinctive mechanism of
NFkappaB/Rel activation through calcineurin-mediated inactivation of
IkappaBbeta. J Cell Biol 161: 507–519, 2003.
14. Biswas G, Guha M, Avadhani NG. Mitochondria-to-nucleus stress
signaling in mammalian cells: nature of nuclear gene targets, transcription regulation, and induced resistance to apoptosis. Gene 354: 132–139,
2005.
15. Blander G, Guarente L. The Sir2 family of protein deacetylases. Annu
Rev Biochem 73: 417– 435, 2004.
16. Bolster DR, Crozier SJ, Kimball SR, Jefferson LS. AMP-activated
protein kinase suppresses protein synthesis in rat skeletal muscle through
down-regulated mammalian target of rapamycin (mTOR) signaling.
J Biol Chem 277: 23977–23980, 2002.
17. Bouras T, Fu M, Sauve AA, Wang F, Quong AA, Perkins ND, Hay
RT, Gu W, Pestell RG. SIRT1 deacetylation and repression of p300
involves lysine residues 1020/1024 within the cell cycle regulatory
domain 1. J Biol Chem 280: 10264 –10276, 2005.
18. Brand MD, Affourtit C, Esteves TC, Green K, Lambert AJ, Miwa S,
Pakay JL, Parker N. Mitochondrial superoxide: production, biological
effects, and activation of uncoupling proteins. Free Radic Biol Med 37:
755–767, 2004.
19. Brini M, De Giorgi F, Murgia M, Marsault R, Massimino ML,
Cantini M, Rizzuto R, Pozzan T. Subcellular analysis of Ca2⫹ homeostasis in primary cultures of skeletal muscle myotubes. Mol Biol Cell
8: 129 –143, 1997.
20. Brunet A, Sweeney LB, Sturgill JF, Chua KF, Greer PL, Lin Y, Tran
H, Ross SE, Mostoslavsky R, Cohen HY, Hu LS, Cheng HL,
Jedrychowski MP, Gygi SP, Sinclair DA, Alt FW, Greenberg ME.
Stress-dependent regulation of FOXO transcription factors by the SIRT1
deacetylase. Science 303: 2011–2015, 2004.
21. Butow RA, Avadhani NG. Mitochondrial signaling: the retrograde
response. Mol Cell 14: 1–15, 2004.
22. Carling D. The AMP-activated protein kinase cascade—a unifying
system for energy control. Trends Biochem Sci 29: 18 –24, 2004.
23. Carreras MC, Clara Franco M, DPC, Finocchieto P, Galli S, Jose
Poderoso J. Cell H2O2 steady-state concentration and mitochondrial
nitric oxide. Methods Enzymol 396: 399 – 414, 2005.
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
sensing molecules, such as Sirt1 (discussed above), Sirt3,
cytosolic guanylyl cyclases (cGCs) and MondoA, could be
studied in greater detail. Localized inside the mitochondrion,
Sirt3, like Sirt1, is a NAD⫹-dependent protein deacetylase.
Expression of Sirt3 in HB1B adipocytes enhances expression
of PGC-1␣ and uncoupling protein 1 (134). This effect involves the phosphorylation of cAMP response element binding
protein (CREB) in the cytosol, suggesting that Sirt3 expression
triggers a mitochondrial signaling response. This is strengthened by the observation that Sirt3 regulates the amount of
acetyl-CoA available for the tricarboxylic acid cycle (51), thus
contributing to the regulation of mitochondrial respiration. The
GCs are nitric oxide (NO䡠) “receptors” that convert GTP to
cGMP, a second messenger that activates G family kinases.
Activation of cGMP-dependent pathways by NO䡠 activates
Sirt1 expression (113) and stimulates mitochondrial biogenesis
in skeletal muscle (112). cGC is inhibited by the binding of
ATP on an allosteric purinergic site of the enzyme (123).
Notably, this inhibitory effect tightly depends on the production of ATP by mitochondrial oxidative phosphorylation,
showing that cGC has the capability to modulate nuclear gene
expression by sensing local variations in ATP concentration.
Another molecule whose activity is presumably regulated by
energy homeostasis is MondoA, a bHLH-leucine zipper partner for the Max-like protein, Mlx. MondoA is a nuclear
transcription factor that shuttles between mitochondria and the
nucleus in skeletal muscle to regulate expression of glycolytic
enzymes, such as lactate dehydrogenase, hexokinase, and
phosphofructokinase (127). Given the role of mitochondria in
intracellular bioenergetics, it has been proposed that MondoA
communicates information about the energy state between the
mitochondria and the nucleus to activate the transcription of
glycolytic genes (127). Overall these data are providing multiples avenues for further study of the regulation of muscle
gene expression by energy-sensing molecules. The development of knockout mice for energy-sensing molecules, as well
as the use of drugs or diets that markedly alter the energetic
status of skeletal muscle, will also be helpful for testing the
responses of various energy sensing molecules. Finally, the
role of acute and chronic muscle contractile activity in triggering and modifying the process of energy sensing should be
evaluated. Clarifying the exact mechanisms and regulatory
influences among energy-sensing molecules will be essential
for a comprehensive analysis of skeletal muscle cell plasticity
in response to altered functional demand.
Invited Review
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
J Appl Physiol • VOL
47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
66.
67.
68.
69.
regulated by endurance training in human skeletal muscle. Am J Physiol
Endocrinol Metab 286: E411–E417, 2004.
Fulco M, Schiltz RL, Iezzi S, King MT, Zhao P, Kashiwaya Y,
Hoffman E, Veech RL, Sartorelli V. Sir2 regulates skeletal muscle
differentiation as a potential sensor of the redox state. Mol Cell 12:
51– 62, 2003.
Gilde AJ, Van Bilsen M. Peroxisome proliferator-activated receptors
(PPARS): regulators of gene expression in heart and skeletal muscle.
Acta Physiol Scand 178: 425– 434, 2003.
Glass DJ. Molecular mechanisms modulating muscle mass. Trends Mol
Med 9: 344 –350, 2003.
Gollnick PD, Saltin B. Significance of skeletal muscle oxidative enzyme
enhancement with endurance training. Clin Physiol 2: 1–12, 1982.
Hallows WC, Lee S, Denu JM. Sirtuins deacetylate and activate
mammalian acetyl-CoA synthetases. Proc Natl Acad Sci USA 103:
10230 –10235, 2006.
Hamai N, Nakamura M, Asano A. Inhibition of mitochondrial protein
synthesis impaired C2C12 myoblast differentiation. Cell Struct Funct 22:
421– 431, 1997.
Handschin C, Rhee J, Lin J, Tarr PT, Spiegelman BM. An autoregulatory loop controls peroxisome proliferator-activated receptor gamma
coactivator 1alpha expression in muscle. Proc Natl Acad Sci USA 100:
7111–7116, 2003.
Hardie DG. New roles for the LKB1 3 AMPK pathway. Curr Opin Cell
Biol 17: 167–173, 2005.
Hardie DG, Sakamoto K. AMPK: a key sensor of fuel and energy status
in skeletal muscle. Physiology 21: 48 – 60, 2006.
Hawley SA, Boudeau J, Reid JL, Mustard KJ, Udd L, Makela TP,
Alessi DR, Hardie DG. Complexes between the LKB1 tumor suppressor, STRADalpha/beta and MO25alpha/beta are upstream kinases in the
AMP-activated protein kinase cascade. J Biol 2: 28, 2003.
Hawley SA, Pan DA, Mustard KJ, Ross L, Bain J, Edelman AM,
Frenguelli BG, Hardie DG. Calmodulin-dependent protein kinase kinase-beta is an alternative upstream kinase for AMP-activated protein
kinase. Cell Metab 2: 9 –19, 2005.
Haynes V, Elfering S, Traaseth N, Giulivi C. Mitochondrial nitricoxide synthase: enzyme expression, characterization, and regulation.
J Bioenerg Biomembr 36: 341–346, 2004.
Herzberg NH, Zwart R, Wolterman RA, Ruiter JP, Wanders RJ,
Bolhuis PA, van den Bogert C. Differentiation and proliferation of
respiration-deficient human myoblasts. Biochim Biophys Acta 1181:
63– 67, 1993.
Holloszy JO, Booth FW. Biochemical adaptations to endurance exercise
in muscle. Annu Rev Physiol 38: 273–291, 1976.
Holloszy JO, Coyle EF. Adaptations of skeletal muscle to endurance
exercise and their metabolic consequences. J Appl Physiol 56: 831– 838,
1984.
Holmes BF, Lang DB, Birnbaum MJ, Mu J, Dohm GL. AMP kinase
is not required for the GLUT4 response to exercise and denervation in
skeletal muscle. Am J Physiol Endocrinol Metab 287: E739 –E743, 2004.
Hongpaisan J, Winters CA, Andrews SB. Calcium-dependent mitochondrial superoxide modulates nuclear CREB phosphorylation in hippocampal neurons. Mol Cell Neurosci 24: 1103–1115, 2003.
Hood DA. Contractile activity-induced mitochondrial biogenesis in skeletal muscle. J Appl Physiol 90: 1137–1157, 2001.
Hood DA, Irrcher I, Ljubicic V, Joseph AM. Coordination of metabolic plasticity in skeletal muscle. J Exp Biol 209: 2265–2275, 2006.
Hudson ER, Pan DA, James J, Lucocq JM, Hawley SA, Green KA,
Baba O, Terashima T, Hardie DG. A novel domain in AMP-activated
protein kinase causes glycogen storage bodies similar to those seen in
hereditary cardiac arrhythmias. Curr Biol 13: 861– 866, 2003.
Hurley RL, Anderson KA, Franzone JM, Kemp BE, Means AR,
Witters LA. The Ca2⫹/calmodulin-dependent protein kinase kinases are
AMP-activated protein kinase kinases. J Biol Chem 280: 29060 –29066,
2005.
Hurst D, Taylor EB, Cline TD, Greenwood LJ, Compton CL, Lamb
JD, Winder WW. AMP-activated protein kinase kinase activity and
phosphorylation of AMP-activated protein kinase in contracting muscle
of sedentary and endurance-trained rats. Am J Physiol Endocrinol Metab
289: E710 –E715, 2005.
Hutber CA, Hardie DG, Winder WW. Electrical stimulation inactivates muscle acetyl-CoA carboxylase and increases AMP-activated protein kinase. Am J Physiol Endocrinol Metab 272: E262–E266, 1997.
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
24. Carriere A, Fernandez Y, Rigoulet M, Penicaud L, Casteilla L.
Inhibition of preadipocyte proliferation by mitochondrial reactive oxygen
species. FEBS Lett 550: 163–167, 2003.
25. 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.
26. Chin ER. Role of Ca2⫹/calmodulin-dependent kinases in skeletal muscle
plasticity. J Appl Physiol 99: 414 – 423, 2005.
27. Chung AB, Stepien G, Haraguchi Y, Li K, Wallace DC. Transcriptional control of nuclear genes for the mitochondrial muscle ADP/ATP
translocator and the ATP synthase beta subunit. Multiple factors interact
with the OXBOX/REBOX promoter sequences. J Biol Chem 267:
21154 –21161, 1992.
28. Connett RJ, Gayeski TE, Honig CR. Lactate accumulation in fully
aerobic, working, dog gracilis muscle. Am J Physiol Heart Circ Physiol
246: H120 –H128, 1984.
29. Dedkova EN, Ji X, Lipsius SL, Blatter LA. Mitochondrial calcium
uptake stimulates nitric oxide production in mitochondria of bovine
vascular endothelial cells. Am J Physiol Cell Physiol 286: C406 –C415,
2004.
30. Denis C, Dormois D, Linossier MT, Geyssant A, Lacour JR. Total
muscle NADH during 30-min submaximal exercise: effects of endurance
training. Acta Physiol Scand 142: 301–302, 1991.
31. Denu JM. The Sir 2 family of protein deacetylases. Curr Opin Chem
Biol 9: 431– 440, 2005.
32. Desvergne B, Michalik L, Wahli W. Transcriptional regulation of
metabolism. Physiol Rev 86: 465–514, 2006.
33. Desvergne B, Wahli W. Peroxisome proliferator-activated receptors:
nuclear control of metabolism. Endocr Rev 20: 649 – 688, 1999.
34. Di Marco S, Mazroui R, Dallaire P, Chittur S, Tenenbaum SA,
Radzioch D, Marette A, Gallouzi IE. NF-␬B-mediated MyoD decay
during muscle wasting requires nitric oxide synthase mRNA stabilization, HuR protein, and nitric oxide release. Mol Cell Biol 25: 6533– 6545,
2005.
35. Dilworth FJ, Seaver KJ, Fishburn AL, Htet SL, Tapscott SJ. In vitro
transcription system delineates the distinct roles of the coactivators pCAF
and p300 during MyoD/E47-dependent transactivation. Proc Natl Acad
Sci USA 101: 11593–11598, 2004.
36. Dougherty CJ, Kubasiak LA, Frazier DP, Li H, Xiong WC, Bishopric NH, Webster KA. Mitochondrial signals initiate the activation of
c-Jun N-terminal kinase (JNK) by hypoxia-reoxygenation. FASEB J 18:
1060 –1070, 2004.
37. Dressel U, Allen TL, Pippal JB, Rohde PR, Lau P, Muscat GE. The
peroxisome proliferator-activated receptor beta/delta agonist,
GW501516, regulates the expression of genes involved in lipid catabolism and energy uncoupling in skeletal muscle cells. Mol Endocrinol 17:
2477–2493, 2003.
38. Dudley GA, Tullson PC, Terjung RL. Influence of mitochondrial
content on the sensitivity of respiratory control. J Biol Chem 262:
9109 –9114, 1987.
39. Duguez S, Feasson L, Denis C, Freyssenet D. Mitochondrial biogenesis
during skeletal muscle regeneration. Am J Physiol Endocrinol Metab
282: E802–E809, 2002.
40. Duguez S, Sabido O, Freyssenet D. Mitochondrial-dependent regulation of myoblast proliferation. Exp Cell Res 299: 27–35, 2004.
41. Durante PE, Mustard KJ, Park SH, Winder WW, Hardie DG.
Effects of endurance training on activity and expression of AMPactivated protein kinase isoforms in rat muscles. Am J Physiol Endocrinol Metab 283: E178 –E186, 2002.
42. Fernando P, Kelly JF, Balazsi K, Slack RS, Megeney LA. Caspase 3
activity is required for skeletal muscle differentiation. Proc Natl Acad Sci
USA 99: 11025–11030, 2002.
43. Fitch CD, Jellinek M, Mueller EJ. Experimental depletion of creatine
and phosphocreatine from skeletal muscle. J Biol Chem 249: 1060 –1063,
1974.
44. Freyssenet D, Berthon P, Barthelemy JC, Busso T, Geyssant A, Denis
C. Energetic status and mitochondrial oxidative capacity of rat skeletal
muscle in response to creatine analogue ingestion. Biochim Biophys Acta
1228: 211–215, 1995.
45. Freyssenet D, Irrcher I, Connor MK, Di Carlo M, Hood DA. Calciumregulated changes in mitochondrial phenotype in skeletal muscle cells. Am J
Physiol Cell Physiol 286: C1053–C1061, 2004.
46. Frosig C, Jorgensen SB, Hardie DG, Richter EA, Wojtaszewski JF.
5⬘-AMP-activated protein kinase activity and protein expression are
537
Invited Review
538
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
J Appl Physiol • VOL
94. Lundby C, Gassmann M, Pilegaard H. Regular endurance training
reduces the exercise induced HIF-1alpha and HIF-2alpha mRNA expression in human skeletal muscle in normoxic conditions. Eur J Appl
Physiol 96: 363–369, 2006.
95. Luo J, Nikolaev AY, Imai S, Chen D, Su F, Shiloh A, Guarente L, Gu
W. Negative control of p53 by Sir2alpha promotes cell survival under
stress. Cell 107: 137–148, 2001.
96. Mascaro C, Acosta E, Ortiz JA, Marrero PF, Hegardt FG, Haro D.
Control of human muscle-type carnitine palmitoyltransferase I gene
transcription by peroxisome proliferator-activated receptor. J Biol Chem
273: 8560 – 8563, 1998.
97. Mason SD, Howlett RA, Kim MJ, Olfert IM, Hogan MC, McNulty
W, Hickey RP, Wagner PD, Kahn CR, Giordano FJ, Johnson RS.
Loss of skeletal muscle HIF-1alpha results in altered exercise endurance.
PLoS Biol 2: e288, 2004.
98. McArdle A, Pattwell D, Vasilaki A, Griffiths RD, Jackson MJ.
Contractile activity-induced oxidative stress: cellular origin and adaptive
responses. Am J Physiol Cell Physiol 280: C621–C627, 2001.
99. McArdle A, van der Meulen J, Close GL, Pattwell D, Van Remmen
H, Huang TT, Richardson AG, Epstein CJ, Faulkner JA, Jackson
MJ. Role of mitochondrial superoxide dismutase in contraction-induced
generation of reactive oxygen species in skeletal muscle extracellular
space. Am J Physiol Cell Physiol 286: C1152–C1158, 2004.
100. McArdle F, Pattwell DM, Vasilaki A, McArdle A, Jackson MJ.
Intracellular generation of reactive oxygen species by contracting skeletal
muscle cells. Free Radic Biol Med 39: 651– 657, 2005.
101. McCormack JG, Denton RM. Signal transduction by intramitochondrial Ca2⫹ in mammalian energy metabolism. News Physiol Sci 9: 71–76,
1994.
102. McCormack JG, Halestrap AP, Denton RM. Role of calcium ions in
regulation of mammalian intramitochondrial metabolism. Physiol Rev
70: 391– 425, 1990.
103. Moncada S, Erusalimsky JD. Does nitric oxide modulate mitochondrial
energy generation and apoptosis? Nat Rev Mol Cell Biol 3: 214 –220,
2002.
104. Motta MC, Divecha N, Lemieux M, Kamel C, Chen D, Gu W,
Bultsma Y, McBurney M, Guarente L. Mammalian SIRT1 represses
forkhead transcription factors. Cell 116: 551–563, 2004.
105. Moyes CD, Mathieu-Costello OA, Tsuchiya N, Filburn C, Hansford
RG. Mitochondrial biogenesis during cellular differentiation. Am J
Physiol Cell Physiol 272: C1345–C1351, 1997.
106. Muoio DM, MacLean PS, Lang DB, Li S, Houmard JA, Way JM,
Winegar DA, Corton JC, Dohm GL, Kraus WE. Fatty acid homeostasis and induction of lipid regulatory genes in skeletal muscles of peroxisome proliferator-activated receptor (PPAR) alpha knock-out mice.
Evidence for compensatory regulation by PPAR delta. J Biol Chem 277:
26089 –26097, 2002.
107. Musi N, Hayashi T, Fujii N, Hirshman MF, Witters LA, Goodyear
LJ. AMP-activated protein kinase activity and glucose uptake in rat
skeletal muscle. Am J Physiol Endocrinol Metab 280: E677–E684, 2001.
108. Muth V, Nadaud S, Grummt I, Voit R. Acetylation of TAF(I)68, a
subunit of TIF-IB/SL1, activates RNA polymerase I transcription. EMBO
J 20: 1353–1362, 2001.
109. Nemoto S, Fergusson MM, Finkel T. SIRT1 functionally interacts with
the metabolic regulator and transcriptional coactivator PGC-1alpha.
J Biol Chem 280: 16456 –16460, 2005.
110. Nemoto S, Takeda K, Yu ZX, Ferrans VJ, Finkel T. Role for
mitochondrial oxidants as regulators of cellular metabolism. Mol Cell
Biol 20: 7311–7318, 2000.
111. Nielsen JN, Mustard KJ, Graham DA, Yu H, MacDonald CS,
Pilegaard H, Goodyear LJ, Hardie DG, Richter EA, Wojtaszewski
JF. 5⬘-AMP-activated protein kinase activity and subunit expression in
exercise-trained human skeletal muscle. J Appl Physiol 94: 631– 641,
2003.
112. 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. Proc Natl Acad Sci USA
101: 16507–16512, 2004.
113. Nisoli E, Tonello C, Cardile 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.
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
70. Hwang JT, Lee M, Jung SN, Lee HJ, Kang I, Kim SS, Ha J.
AMP-activated protein kinase activity is required for vanadate-induced
hypoxia-inducible factor 1alpha expression in DU145 cells. Carcinogenesis 25: 2497–2507, 2004.
71. Imai S, Armstrong CM, Kaeberlein M, Guarente L. Transcriptional
silencing and longevity protein Sir2 is an NAD-dependent histone
deacetylase. Nature 403: 795– 800, 2000.
72. Inoki K, Zhu T, Guan KL. TSC2 mediates cellular energy response to
control cell growth and survival. Cell 115: 577–590, 2003.
73. Isaeva EV, Shirokova N. Metabolic regulation of Ca2⫹ release in
permeabilized mammalian skeletal muscle fibres. J Physiol 547: 453–
462, 2003.
74. Itoh H, Ohkuwa T, Yamamoto T, Sato Y, Miyamura M, Naoi M.
Effects of endurance physical training on hydroxyl radical generation in
rat tissues. Life Sci 63: 1921–1929, 1998.
75. Jackman RW, Kandarian SC. The molecular basis of skeletal muscle
atrophy. Am J Physiol Cell Physiol 287: C834 –C843, 2004.
76. Jorgensen SB, Richter EA, Wojtaszewski JF. Role of AMPK in
skeletal muscle metabolic regulation and adaptation in relation to exercise. J Physiol 574: 17–31, 2006.
77. Jorgensen SB, Wojtaszewski JF, Viollet B, Andreelli F, Birk JB,
Hellsten Y, Schjerling P, Vaulont S, Neufer PD, Richter EA, Pilegaard H. Effects of alpha-AMPK knockout on exercise-induced gene
activation in mouse skeletal muscle. FASEB J 19: 1146 –1148, 2005.
78. Jousse C, Averous J, Bruhat A, Carraro V, Mordier S, Fafournoux
P. Amino acids as regulators of gene expression: molecular mechanisms.
Biochem Biophys Res Commun 313: 447– 452, 2004.
79. Kadenbach B. Intrinsic and extrinsic uncoupling of oxidative phosphorylation. Biochim Biophys Acta 1604: 77–94, 2003.
80. Kahn BB, Alquier T, Carling D, Hardie DG. AMP-activated protein
kinase: ancient energy gauge provides clues to modern understanding of
metabolism. Cell Metab 1: 15–25, 2005.
81. Kiens B. Skeletal muscle lipid metabolism in exercise and insulin
resistance. Physiol Rev 86: 205–243, 2006.
82. Kietzmann T, Fandrey J, Acker H. Oxygen radicals as messengers in
oxygen-dependent gene expression. News Physiol Sci 15: 202–208, 2000.
83. Korohoda W, Pietrzkowski Z, Reiss K. Chloramphenicol, an inhibitor
of mitochondrial protein synthesis, inhibits myoblast fusion and myotube
differentiation. Folia Histochem Cytobiol 31: 9 –13, 1993.
84. Koulmann N, Bigard AX. Interaction between signalling pathways
involved in skeletal muscle responses to endurance exercise. Pflügers
Arch 452: 125–139, 2006.
85. Kulisz A, Chen N, Chandel NS, Shao Z, Schumacker PT. Mitochondrial ROS initiate phosphorylation of p38 MAP kinase during hypoxia in
cardiomyocytes. Am J Physiol Lung Cell Mol Physiol 282: L1324 –
L1329, 2002.
86. Langen RC, Schols AM, Kelders MC, Van Der Velden JL, Wouters
EF, Janssen-Heininger YM. Tumor necrosis factor-alpha inhibits myogenesis through redox-dependent and -independent pathways. Am J
Physiol Cell Physiol 283: C714 –C721, 2002.
87. Lee I, Bender E, Kadenbach B. Control of mitochondrial membrane
potential and ROS formation by reversible phosphorylation of cytochrome c oxidase. Mol Cell Biochem 234 –235: 63–70, 2002.
88. Lee M, Hwang JT, Lee HJ, Jung SN, Kang I, Chi SG, Kim SS, Ha J.
AMP-activated protein kinase activity is critical for hypoxia-inducible
factor-1 transcriptional activity and its target gene expression under
hypoxic conditions in DU145 cells. J Biol Chem 278: 39653–39661,
2003.
89. Lee WJ, Kim M, Park HS, Kim HS, Jeon MJ, Oh KS, Koh EH, Won
JC, Kim MS, Oh GT, Yoon M, Lee KU, Park JY. AMPK activation
increases fatty acid oxidation in skeletal muscle by activating PPARalpha
and PGC-1. Biochem Biophys Res Commun 340: 291–295, 2006.
90. Leeuwenburgh C, Heinecke JW. Oxidative stress and antioxidants in
exercise. Curr Med Chem 8: 829 – 838, 2001.
91. Li YP, Atkins CM, Sweatt JD, Reid MB. Mitochondria mediate tumor
necrosis factor-alpha/NF-kappaB signaling in skeletal muscle myotubes.
Antioxid Redox Signal 1: 97–104, 1999.
92. Li YP, Schwartz RJ, Waddell ID, Holloway BR, Reid MB. Skeletal
muscle myocytes undergo protein loss and reactive oxygen-mediated
NF-kappaB activation in response to tumor necrosis factor alpha. FASEB
J 12: 871– 880, 1998.
93. Ludwig B, Bender E, Arnold S, Huttemann M, Lee I, Kadenbach B.
Cytochrome c oxidase and the regulation of oxidative phosphorylation.
Chembiochem 2: 392– 403, 2001.
Invited Review
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
J Appl Physiol • VOL
134. Shi T, Wang F, Stieren E, Tong Q. SIRT3, a mitochondrial sirtuin
deacetylase, regulates mitochondrial function and thermogenesis in
brown adipocytes. J Biol Chem 280: 13560 –13567, 2005.
135. Shiva S, Oh JY, Landar AL, Ulasova E, Venkatraman A, Bailey SM,
Darley-Usmar VM. Nitroxia: the pathological consequence of dysfunction in the nitric oxide-cytochrome c oxidase signaling pathway. Free
Radic Biol Med 38: 297–306, 2005.
136. Shkryl VM, Shirokova N. Transfer and tunneling of Ca2⫹ from sarcoplasmic reticulum to mitochondria in skeletal muscle. J Biol Chem 281:
1547–1554, 2006.
137. Shoubridge EA, Challiss RA, Hayes DJ, Radda GK. Biochemical
adaptation in the skeletal muscle of rats depleted of creatine with the
substrate analogue beta-guanidinopropionic acid. Biochem J 232: 125–
131, 1985.
138. 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.
139. Starkov AA, Fiskum G, Chinopoulos C, Lorenzo BJ, Browne SE,
Patel MS, Beal MF. Mitochondrial alpha-ketoglutarate dehydrogenase
complex generates reactive oxygen species. J Neurosci 24: 7779 –7788,
2004.
140. Steinberg GR, Watt MJ, McGee SL, Chan S, Hargreaves M, Febbraio MA, Stapleton D, Kemp BE. Reduced glycogen availability is
associated with increased AMPK␣2 activity, nuclear AMPK␣2 protein
abundance, and GLUT4 mRNA expression in contracting human skeletal
muscle. Appl Physiol Nutr Metab 31: 302–312, 2006.
141. Stitt TN, Drujan D, Clarke BA, Panaro F, Timofeyva Y, Kline WO,
Gonzalez M, Yancopoulos GD, Glass DJ. The IGF-1/PI3K/Akt pathway prevents expression of muscle atrophy-induced ubiquitin ligases by
inhibiting FOXO transcription factors. Mol Cell 14: 395– 403, 2004.
142. Storz P, Doppler H, Toker A. Protein kinase D mediates mitochondrionto-nucleus signaling and detoxification from mitochondrial reactive oxygen
species. Mol Cell Biol 25: 8520 – 8530, 2005.
143. Suwa M, Nakano H, Kumagai S. Effects of chronic AICAR treatment
on fiber composition, enzyme activity, UCP3, and PGC-1 in rat muscles.
J Appl Physiol 95: 960 –968, 2003.
144. Tanaka T, Yamamoto J, Iwasaki S, Asaba H, Hamura H, Ikeda Y,
Watanabe M, Magoori K, Ioka RX, Tachibana K, Watanabe Y,
Uchiyama Y, Sumi K, Iguchi H, Ito S, Doi T, Hamakubo T, Naito M,
Auwerx J, Yanagisawa M, Kodama T, Sakai J. Activation of peroxisome proliferator-activated receptor delta induces fatty acid beta-oxidation in skeletal muscle and attenuates metabolic syndrome. Proc Natl
Acad Sci USA 100: 15924 –15929, 2003.
145. Tee AR, Manning BD, Roux PP, Cantley LC, Blenis J. Tuberous
sclerosis complex gene products, Tuberin and Hamartin, control mTOR
signaling by acting as a GTPase-activating protein complex toward Rheb.
Curr Biol 13: 1259 –1268, 2003.
146. Terada S, Goto M, Kato M, Kawanaka K, Shimokawa T, Tabata I.
Effects of low-intensity prolonged exercise on PGC-1 mRNA expression
in rat epitrochlearis muscle. Biochem Biophys Res Commun 296: 350 –
354, 2002.
147. Tonkonogi M, Walsh B, Svensson M, Sahlin K. Mitochondrial function and antioxidative defence in human muscle: effects of endurance
training and oxidative stress. J Physiol 528: 379 –388, 2000.
148. Traaseth N, Elfering S, Solien J, Haynes V, Giulivi C. Role of calcium
signaling in the activation of mitochondrial nitric oxide synthase and
citric acid cycle. Biochim Biophys Acta 1658: 64 –71, 2004.
149. Turrens JF. Mitochondrial formation of reactive oxygen species.
J Physiol 552: 335–344, 2003.
150. Vasilaki A, Mansouri A, Remmen H, van der Meulen JH, Larkin L,
Richardson AG, McArdle A, Faulkner JA, Jackson MJ. Free radical
generation by skeletal muscle of adult and old mice: effect of contractile
activity. Aging Cell 5: 109 –117, 2006.
151. Vavvas D, Apazidis A, Saha AK, Gamble J, Patel A, Kemp BE,
Witters LA, Ruderman NB. Contraction-induced changes in acetylCoA carboxylase and 5⬘-AMP-activated kinase in skeletal muscle. J Biol
Chem 272: 13255–13261, 1997.
152. Vaziri H, Dessain SK, Ng Eaton E, Imai SI, Frye RA, Pandita TK,
Guarente L, Weinberg RA. hSIR2(SIRT1) functions as an NADdependent p53 deacetylase. Cell 107: 149 –159, 2001.
153. Walsh B, Tonkonogi M, Sahlin K. Effect of endurance training on
oxidative and antioxidative function in human permeabilized muscle
fibres. Pflügers Arch 442: 420 – 425, 2001.
102 • FEBRUARY 2007 •
www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
114. Puigserver P, Adelmant G, Wu Z, Fan M, Xu J, O’Malley B,
Spiegelman BM. Activation of PPARgamma coactivator-1 through
transcription factor docking. Science 286: 1368 –1371, 1999.
115. Putman CT, Kiricsi M, Pearcey J, MacLean IM, Bamford JA,
Murdoch GK, Dixon WT, Pette D. AMPK activation increases uncoupling protein-3 expression and mitochondrial enzyme activities in rat
muscle without fibre type transitions. J Physiol 551: 169 –178, 2003.
116. Rafaeloff-Phail R, Ding L, Conner L, Yeh WK, McClure D, Guo H,
Emerson K, Brooks H. Biochemical regulation of mammalian AMPactivated protein kinase activity by NAD and NADH. J Biol Chem 279:
52934 –52939, 2004.
117. Ranganathan AC, Nelson KK, Rodriguez AM, Kim KH, Tower GB,
Rutter JL, Brinckerhoff CE, Huang TT, Epstein CJ, Jeffrey JJ,
Melendez JA. Manganese superoxide dismutase signals matrix metalloproteinase expression via H2O2-dependent ERK1/2 activation. J Biol
Chem 276: 14264 –14270, 2001.
118. Richardson RS, Newcomer SC, Noyszewski EA. Skeletal muscle
intracellular PO2 assessed by myoglobin desaturation: response to graded
exercise. J Appl Physiol 91: 2679 –2685, 2001.
119. Richardson RS, Noyszewski EA, Kendrick KF, Leigh JS, Wagner
PD. Myoglobin O2 desaturation during exercise. Evidence of limited O2
transport. J Clin Invest 96: 1916 –1926, 1995.
120. Robergs RA, Ghiasvand F, Parker D. Biochemistry of exerciseinduced metabolic acidosis. Am J Physiol Regul Integr Comp Physiol
287: R502–R516, 2004.
121. Rochard P, Rodier A, Casas F, Cassar-Malek I, Marchal-Victorion
S, Daury L, Wrutniak C, Cabello G. Mitochondrial activity is involved
in the regulation of myoblast differentiation through myogenin expression and activity of myogenic factors. J Biol Chem 275: 2733–2744,
2000.
122. Rodgers JT, Lerin C, Haas W, Gygi SP, Spiegelman BM, Puigserver
P. Nutrient control of glucose homeostasis through a complex of PGC1alpha and SIRT1. Nature 434: 113–118, 2005.
123. Ruiz-Stewart I, Tiyyagura SR, Lin JE, Kazerounian S, Pitari GM,
Schulz S, Martin E, Murad F, Waldman SA. Guanylyl cyclase is an
ATP sensor coupling nitric oxide signaling to cell metabolism. Proc Natl
Acad Sci USA 101: 37– 42, 2004.
124. Russell AP, Feilchenfeldt J, Schreiber S, Praz M, Crettenand A,
Gobelet C, Meier CA, Bell DR, Kralli A, Giacobino JP, Deriaz O.
Endurance training in humans leads to fiber type-specific increases in
levels of peroxisome proliferator-activated receptor-gamma coactivator-1 and peroxisome proliferator-activated receptor-alpha in skeletal
muscle. Diabetes 52: 2874 –2881, 2003.
125. Sandri M, Sandri C, Gilbert A, Skurk C, Calabria E, Picard A,
Walsh K, Schiaffino S, Lecker SH, Goldberg AL. Foxo transcription
factors induce the atrophy-related ubiquitin ligase atrogin-1 and cause
skeletal muscle atrophy. Cell 117: 399 – 412, 2004.
126. Sandstrom ME, Zhang SJ, Bruton J, Silva JP, Reid MB, Westerblad
H, Katz A. Role of reactive oxygen species in contraction-mediated
glucose transport in mouse skeletal muscle. J Physiol 575: 251–262,
2006.
127. Sans CL, Satterwhite DJ, Stoltzman CA, Breen KT, Ayer DE.
MondoA-Mlx heterodimers are candidate sensors of cellular energy
status: mitochondrial localization and direct regulation of glycolysis. Mol
Cell Biol 26: 4863– 4871, 2006.
128. Schofield CJ, Ratcliffe PJ. Signalling hypoxia by HIF hydroxylases.
Biochem Biophys Res Commun 338: 617– 626, 2005.
129. Semenza GL. Hydroxylation of HIF-1: oxygen sensing at the molecular
level. Physiology 19: 176 –182, 2004.
130. Semenza GL. Targeting HIF-1 for cancer therapy. Nat Rev Cancer 3:
721–732, 2003.
131. Servais S, Couturier K, Koubi H, Rouanet JL, Desplanches D,
Sornay-Mayet MH, Sempore B, Lavoie JM, Favier R. Effect of
voluntary exercise on H2O2 release by subsarcolemmal and intermyofibrillar mitochondria. Free Radic Biol Med 35: 24 –32, 2003.
132. Seyer P, Grandemange S, Busson M, Carazo A, Gamaleri F, Pessemesse L, Casas F, Cabello G, Wrutniak-Cabello C. Mitochondrial
activity regulates myoblast differentiation by control of c-Myc expression. J Cell Physiol 207: 75– 86, 2006.
133. Shaw RJ, Kosmatka M, Bardeesy N, Hurley RL, Witters LA, DePinho RA, Cantley LC. The tumor suppressor LKB1 kinase directly
activates AMP-activated kinase and regulates apoptosis in response to
energy stress. Proc Natl Acad Sci USA 101: 3329 –3335, 2004.
539
Invited Review
540
ENERGY SENSING AND SKELETAL MUSCLE GENE EXPRESSION
154. Wang GL, Jiang BH, Rue EA, Semenza GL. Hypoxia-inducible factor
1 is a basic-helix-loop-helix-PAS heterodimer regulated by cellular O2
tension. Proc Natl Acad Sci USA 92: 5510 –5514, 1995.
155. Wang YX, Zhang CL, Yu RT, Cho HK, Nelson MC, BayugaOcampo CR, Ham J, Kang H, Evans RM. Regulation of muscle fiber
type and running endurance by PPARdelta. PLoS Biol 2: e294, 2004.
156. Warburg O. On the origin of cancer cells. Science 123: 309 –314, 1956.
157. Winder WW. Energy-sensing and signaling by AMP-activated protein
kinase in skeletal muscle. J Appl Physiol 91: 1017–1028, 2001.
158. Winder WW, Hardie DG. Inactivation of acetyl-CoA carboxylase and
activation of AMP-activated protein kinase in muscle during exercise.
Am J Physiol Endocrinol Metab 270: E299 –E304, 1996.
159. 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.
160. Wojtaszewski JF, Jorgensen SB, Hellsten Y, Hardie DG, Richter
EA. Glycogen-dependent effects of 5-aminoimidazole-4-carboxamide
(AICA)-riboside on AMP-activated protein kinase and glycogen synthase
activities in rat skeletal muscle. Diabetes 51: 284 –292, 2002.
161. Woods A, Dickerson K, Heath R, Hong SP, Momcilovic M, Johnstone SR, Carlson M, Carling D. (Ca2⫹)/calmodulin-dependent protein
162.
163.
164.
165.
166.
kinase kinase-beta acts upstream of AMP-activated protein kinase in
mammalian cells. Cell Metab 2: 21–33, 2005.
Yang W, Hong YH, Shen XQ, Frankowski C, Camp HS, Leff T.
Regulation of transcription by AMP-activated protein kinase: phosphorylation of p300 blocks its interaction with nuclear receptors. J Biol Chem
276: 38341–38344, 2001.
Yang Y, Hou H, Haller EM, Nicosia SV, Bai W. Suppression of
FOXO1 activity by FHL2 through SIRT1-mediated deacetylation.
EMBO J 24: 1021–1032, 2005.
Yu M, Stepto NK, Chibalin AV, Fryer LG, Carling D, Krook A,
Hawley JA, Zierath JR. Metabolic and mitogenic signal transduction in
human skeletal muscle after intense cycling exercise. J Physiol 546:
327–335, 2003.
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.
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.
Downloaded from http://jap.physiology.org/ by 10.220.32.247 on June 16, 2017
J Appl Physiol • VOL
102 • FEBRUARY 2007 •
www.jap.org