Metabolic Control Through the Pgc-1 Review Family of Transcription Coactivators
Abstruse
Chronic disruption of energy balance, where energy intake exceeds expenditure, is a major risk factor for the evolution of metabolic syndrome. The latter is characterized past a constellation of symptoms including obesity, dyslipidemia, insulin resistance, hypertension, and non-alcoholic fatty liver affliction. Altered expression of genes involved in glucose and lipid metabolism too as mitochondrial oxidative phosphorylation has been implicated in the pathogenesis of these disorders. The peroxisome proliferator-activated receptor γ coactivator-1 (PGC-1) family of transcriptional coactivators is emerging as a hub linking nutritional and hormonal signals and free energy metabolism. PGC-1α and PGC-1β are highly responsive to environmental cues and coordinate metabolic gene programs through interaction with transcription factors and chromatin-remodeling proteins. PGC-1α has been implicated in the pathogenic conditions including obesity, blazon two diabetes, neurodegeneration, and cardiomyopathy, whereas PGC-1β plays an important function in plasma lipoprotein homeostasis and serves as a hepatic target for niacin, a stiff hypotriglyceridemic drug. Here, we review recent advances in the identification of physiological and pathophysiological contexts involving PGC-1 coactivators, and also discuss their implications for therapeutic development.
Introduction
The prevalence of contemporary life style, characterized by increased consumption of high-fatty, loftier-fructose food and reduced physical activity, has driven a dramatic increment in the incidence of metabolic syndrome. It has been projected that, by 2025, one in every three American children built-in in the year 2000 volition carry a pregnant lifetime risk of developing type 2 diabetes, and therefore become decumbent to premature cardiovascular disease, incomprehension, kidney failure, and amputations. A fundamental feature of metabolic syndrome is severe obesity, which arises from chronic imbalance between free energy intake and energy consumption. As such, restoration of the free energy residual is a major strategy for the therapy of metabolic disease including obesity, diabetes, hypertension, atherosclerosis, and fatty liver diseases.
Peroxisome proliferator-activated receptor (PPAR) γ coactivator-one (PGC-i) family members are multifunctional transcriptional coregulators that act as 'molecular switches' in many metabolic pathways. PGC-1α and PGC-1β have been shown to regulate adaptive thermogenesis, mitochondrial biogenesis, glucose/fat-acrid metabolism, peripheral circadian clock, cobweb-type switching in skeletal muscle, and centre evolution. Their versatile deportment are accomplished past interacting with different transcription factors in a tissue-specific fashion. The potent furnishings of PGC-i coactivators in analogous various metabolic processes underscore their significant role in the command of energy metabolism as well as their potential as targets for pharmacological intervention.
Structure and part of PGC-1 coactivators
The PGC-1α factor is located on chromosome 5 in mice (chromosome iv in humans) and encodes a protein containing 797 (mouse) or 798 (human) amino acids [1]. Structural and functional studies accept indicated that PGC-1α has a potent transcriptional activation domain at the N terminus, which interacts with several histone acetyltransferase (HAT) complexes including 3'-5'-cyclic adenosine monophosphate (camp) response chemical element-binding protein (CREB)-binding poly peptide, p300, and steroid receptor coactivator-1 [2]. These proteins acetylate histones and remodel chromatin structure into a land that is permissive for transcriptional activation. Adjacent to the North-terminal domain is a regulatory region that roughly spans 200 amino acids. Toward the C terminus, PGC-1α recruits the thyroid receptor-associated protein/vitamin D receptor-interacting protein/mediator complex that facilitates straight interaction with the transcription initiation machinery [iii]. This region besides interacts with the switch/sucrose nonfermentable (SWI/SNF) chromatin-remodeling circuitous through its interaction with BAF60a [4]. The Ser/Arg-rich domain and an RNA-binding domain toward C terminus have been demonstrated to couple pre-mRNA splicing with transcription [5]. As such, PGC-1α serves as a platform for the recruitment and associates of various chromatin-remodeling and histone-modifying enzymes to alter local chromatin country. Importantly, the PGC-1α transcriptional activator circuitous is also able to displace repressor proteins, such equally histone deacetylase and pocket-sized heterodimer partner, on its target promoters, providing an alternative mechanism for gene activation [6]. PGC-1α and PGC-1β share extensive domain similarity and several clusters of conserved amino acids, such every bit the LXXLL motif that interacts with nuclear receptors and host cell factor 1 interacting motif [seven]. The tertiary family member, PRC (PGC-1-related coactivator), too contains the activation domain and RNA-binding domain, just overall has more than express homology to PGC-1α and PGC-1β [eight]. The PGC-1 family unit members are conserved in college vertebrates, including mammals, birds, and fish. Interestingly, a PGC-1 family unit homologue named Spargel was recently identified in Drosophila that could regulate mitochondrial activity and insulin signaling [9].
Both PGC-1α and PGC-1β robustly regulate mitochondrial oxidative metabolism (Fig.1). PGC-1α was initially identified equally a PPARγ-interacting protein from the brown adipose tissue (BAT) that could regulate adaptive thermogenesis in response to cold [i]. Subsequent studies revealed that the core function of PGC-1α was to stimulate mitochondrial biogenesis and oxidative metabolism. PGC-1α is abundantly expressed in tissues with loftier energy demand, including the BAT, centre, skeletal muscle, kidney, and brain [10–12]. In fact, when ectopically expressed in fat or muscle cells, PGC-1α strongly stimulates the programme of nuclear and mitochondrial-encoded mitochondrial genes as well as organelle biogenesis [10]. The stimulatory effects of PGC-1α on mitochondrial genes are achieved through its coactivation of nuclear respiratory factors 1 and ii (NRF1 and NRF2, respectively) and the estrogen-related receptor α (ERRα) [10,13,14]. The induction of NRF1 and NRF2 subsequently leads to the increased expression of mitochondrial transcription factor A (mtTFA) [10] as well as other mitochondrial subunits of the electron transport chain complex such equally β-adenosine-triphosphate (ATP) synthase, cytochrome c, and cytochrome oxidase 4 [15,16]. mtTFA translocates to mitochondrial matrix, where it stimulates mitochondrial DNA replication and mitochondrial factor expression [17].
Figure 1
The working model of PGC-1 coactivators PGC-1α and PGC-1β regulate diverse metabolic programs through coactivating selective transcriptional factors (TF) associated with regulatory elements of target genes. PGC-i recruits HAT, SWI/SNF chromatin-remodeling, Sirt1 deacetylase, and mediator complexes to modulate the epigenetic condition of chromatin.
Figure ane
The working model of PGC-ane coactivators PGC-1α and PGC-1β regulate various metabolic programs through coactivating selective transcriptional factors (TF) associated with regulatory elements of target genes. PGC-i recruits HAT, SWI/SNF chromatin-remodeling, Sirt1 deacetylase, and mediator complexes to modulate the epigenetic condition of chromatin.
As mentioned above, a critical aspect of PGC-1α is that information technology is highly versatile and has the ability to increase the transcriptional activity of many nuclear receptor families, including members of the estrogen, PPAR, retinoid X, mineralocorticoid, glucocorticoid (GR), liver X (LXR), pregnane X, the constitutive androstane, vitamin D, and thyroid hormone receptor families [eighteen,19]. PGC-1α tin can likewise demark to unliganded nuclear receptors, every bit in the example of the orphan hepatic nuclear factor (HNF) 4α, farnesoid X receptor (FXR), and ERRα, suggesting that their conformations are conductive to ligand-contained mechanisms of gene regulation [20]. PGC-1α transcriptional partners are non express to the nuclear receptor superfamily; nevertheless, this coactivator too assembly with a diverse array of other transcription factors, including forkhead/winged helix protein family member FOXO1, as well as a number of zinc-finger proteins identified through a genome-wide coactivation screen [iv,20]. The docking interface for these interacting proteins appears to distribute throughout the length of PGC-1α. In improver, PGC-1α has three functional LXXLL motifs that are responsible for docking nuclear receptors [xviii]. The diversity of PGC-1α-interacting proteins enables PGC-1α to regulate various metabolic processes in a tissue-specific fashion.
Tissue-specific metabolic actions of PGC-1 coactivators
The following section reviews PGC-1 functions in oxidative tissues including the brain, heart, brown fat, skeletal muscle, liver, and pancreatic islets, based on gain and loss-of-function analysis both in cultured cells and in vivo. A summary of tissue-specific PGC-1 functions and the phenotype of PGC-one transgenic mouse models are included in Tables 1 and two, respectively.
Tabular array i
Tissue-specific functions of PGC-1α and PGC-1β
| Tissues | Biological functions | |
|---|---|---|
| PGC-1α | PGC-1β | |
| Brain | Maintenance of mitochondrial function [21], ROS detoxification [22], GABAergic neuronal function [23] | Unknown |
| Heart | Mitochondrial oxidative metabolism [24], fat-acrid β-oxidation [24] | Mitochondrial OXPHOS [25], mediating the furnishings of adrenergic stimulation on heart rate [26] |
| Dark-brown fatty | Mitochondrial biogenesis and fatty oxidation [1]; Adaptive thermogenesis [1] | Brownish adipocyte differentiation [27]; Adaptive thermogenesis [28] |
| Skeletal muscle | Slow-switch muscle fiber [29], mitochondrial biogenesis [29], skeletal muscle atrophy [30] | Type IIX fiber formation [31] |
| Liver | Hepatic fasting response [eighteen,32], homocysteine metabolism [33], integration of cyclic clock and metabolism [34] | Hepatic lipogenesis and VLDL secretion [35], regulation of lipoprotein catabolism [36] |
| Pancreatic islets | Suppression of GSIS and membrane depolarization [37] | Suppression of GSIS [38] |
| Tissues | Biological functions | |
|---|---|---|
| PGC-1α | PGC-1β | |
| Encephalon | Maintenance of mitochondrial part [21], ROS detoxification [22], GABAergic neuronal function [23] | Unknown |
| Eye | Mitochondrial oxidative metabolism [24], fat-acid β-oxidation [24] | Mitochondrial OXPHOS [25], mediating the effects of adrenergic stimulation on heart rate [26] |
| Brownish fat | Mitochondrial biogenesis and fat oxidation [1]; Adaptive thermogenesis [ane] | Brown adipocyte differentiation [27]; Adaptive thermogenesis [28] |
| Skeletal muscle | Dull-switch muscle fiber [29], mitochondrial biogenesis [29], skeletal muscle atrophy [thirty] | Blazon IIX fiber formation [31] |
| Liver | Hepatic fasting response [xviii,32], homocysteine metabolism [33], integration of circadian clock and metabolism [34] | Hepatic lipogenesis and VLDL secretion [35], regulation of lipoprotein catabolism [36] |
| Pancreatic islets | Suppression of GSIS and membrane depolarization [37] | Suppression of GSIS [38] |
VLDL, very-low-density lipoprotein.
Tabular array 1
Tissue-specific functions of PGC-1α and PGC-1β
| Tissues | Biological functions | |
|---|---|---|
| PGC-1α | PGC-1β | |
| Encephalon | Maintenance of mitochondrial function [21], ROS detoxification [22], GABAergic neuronal function [23] | Unknown |
| Middle | Mitochondrial oxidative metabolism [24], fatty-acid β-oxidation [24] | Mitochondrial OXPHOS [25], mediating the furnishings of adrenergic stimulation on heart rate [26] |
| Brown fat | Mitochondrial biogenesis and fat oxidation [1]; Adaptive thermogenesis [1] | Chocolate-brown adipocyte differentiation [27]; Adaptive thermogenesis [28] |
| Skeletal musculus | Irksome-switch muscle cobweb [29], mitochondrial biogenesis [29], skeletal muscle cloudburst [30] | Blazon IIX fiber formation [31] |
| Liver | Hepatic fasting response [eighteen,32], homocysteine metabolism [33], integration of circadian clock and metabolism [34] | Hepatic lipogenesis and VLDL secretion [35], regulation of lipoprotein catabolism [36] |
| Pancreatic islets | Suppression of GSIS and membrane depolarization [37] | Suppression of GSIS [38] |
| Tissues | Biological functions | |
|---|---|---|
| PGC-1α | PGC-1β | |
| Brain | Maintenance of mitochondrial part [21], ROS detoxification [22], GABAergic neuronal function [23] | Unknown |
| Eye | Mitochondrial oxidative metabolism [24], fatty-acid β-oxidation [24] | Mitochondrial OXPHOS [25], mediating the effects of adrenergic stimulation on heart rate [26] |
| Dark-brown fat | Mitochondrial biogenesis and fat oxidation [ane]; Adaptive thermogenesis [ane] | Brown adipocyte differentiation [27]; Adaptive thermogenesis [28] |
| Skeletal musculus | Wearisome-switch muscle fiber [29], mitochondrial biogenesis [29], skeletal musculus atrophy [thirty] | Type IIX fiber germination [31] |
| Liver | Hepatic fasting response [eighteen,32], homocysteine metabolism [33], integration of circadian clock and metabolism [34] | Hepatic lipogenesis and VLDL secretion [35], regulation of lipoprotein catabolism [36] |
| Pancreatic islets | Suppression of GSIS and membrane depolarization [37] | Suppression of GSIS [38] |
VLDL, very-depression-density lipoprotein.
Table 2
Phenotypes of PGC-i knockout and transgenic mouse models
| Models | Manipulations | Phenotypes |
|---|---|---|
| PGC-1α | Whole-body ablation [39] | Hyperactive, cold-sensitive, resistant to diet-induced obesity, lesions in the striatum |
| Whole-body ablation [40] | Reduced muscle performance and exercise capacity, impaired adaptive thermogenesis, hepatic steatosis | |
| Muscle-specific ablation [41] | Impaired glucose tolerance, normal peripheral insulin sensitivity | |
| Muscle-specific overexpression [29] | Switch of type Ii muscle cobweb to blazon IIa and I muscle fibers, resistance to electrically stimulated fatigue | |
| Cardiac-specific overexpression [24] | Loss of sarcomeric structure, dilated cardiomyopathy | |
| Tetracycline-inducible, cardiac-specific overexpression [42] | Increased mitochondrial biogenesis, derangements of mitochondrial ultrastructure, cardiomyopathy | |
| PGC-1β | Whole-torso ablation [28] | Impaired mitochondrial function, reduced body weight and fat mass, increased thermogenesis, blunted chronotropic response to dobutamine in the heart, hepatic steatosis, reduced lipoprotein-associated triglyceride and cholesterol content |
| Whole-body ablation [26] | Decreased action during the dark cycle, abnormal hypothermia and morbidity, hepatic steatosis, increased serum triglyceride and cholesterol | |
| Deletion of exons iii–4 [43] | Mitochondrial dysfunction in liver and skeletal musculus, hepatic steatosis, hepatic insulin resistance | |
| Muscle-specific overexpression [31] | Increased fatty-acid oxidation, hyperphagia, reduced body weight and adipose tissue, increased exercise chapters, increased IIX fiber content | |
| PGC-1α/β | Double knockout [25] | Neonatal lethality, bradycardia, intermittent eye cake, reduced cardiac output, reduced growth, a late fetal arrest in mitochondrial biogenesis |
| Models | Manipulations | Phenotypes |
|---|---|---|
| PGC-1α | Whole-body ablation [39] | Hyperactive, cold-sensitive, resistant to diet-induced obesity, lesions in the striatum |
| Whole-body ablation [40] | Reduced musculus functioning and exercise capacity, impaired adaptive thermogenesis, hepatic steatosis | |
| Musculus-specific ablation [41] | Impaired glucose tolerance, normal peripheral insulin sensitivity | |
| Musculus-specific overexpression [29] | Switch of type Two muscle fiber to type IIa and I musculus fibers, resistance to electrically stimulated fatigue | |
| Cardiac-specific overexpression [24] | Loss of sarcomeric structure, dilated cardiomyopathy | |
| Tetracycline-inducible, cardiac-specific overexpression [42] | Increased mitochondrial biogenesis, derangements of mitochondrial ultrastructure, cardiomyopathy | |
| PGC-1β | Whole-torso ablation [28] | Impaired mitochondrial office, reduced body weight and fatty mass, increased thermogenesis, blunted chronotropic response to dobutamine in the eye, hepatic steatosis, reduced lipoprotein-associated triglyceride and cholesterol content |
| Whole-trunk ablation [26] | Decreased activity during the dark cycle, abnormal hypothermia and morbidity, hepatic steatosis, increased serum triglyceride and cholesterol | |
| Deletion of exons 3–4 [43] | Mitochondrial dysfunction in liver and skeletal muscle, hepatic steatosis, hepatic insulin resistance | |
| Musculus-specific overexpression [31] | Increased fatty-acrid oxidation, hyperphagia, reduced body weight and adipose tissue, increased exercise capacity, increased IIX fiber content | |
| PGC-1α/β | Double knockout [25] | Neonatal lethality, bradycardia, intermittent middle block, reduced cardiac output, reduced growth, a belatedly fetal abort in mitochondrial biogenesis |
Table 2
Phenotypes of PGC-1 knockout and transgenic mouse models
| Models | Manipulations | Phenotypes |
|---|---|---|
| PGC-1α | Whole-body ablation [39] | Hyperactive, cold-sensitive, resistant to diet-induced obesity, lesions in the striatum |
| Whole-torso ablation [40] | Reduced muscle performance and practice capacity, impaired adaptive thermogenesis, hepatic steatosis | |
| Muscle-specific ablation [41] | Impaired glucose tolerance, normal peripheral insulin sensitivity | |
| Musculus-specific overexpression [29] | Switch of type II musculus fiber to type IIa and I muscle fibers, resistance to electrically stimulated fatigue | |
| Cardiac-specific overexpression [24] | Loss of sarcomeric structure, dilated cardiomyopathy | |
| Tetracycline-inducible, cardiac-specific overexpression [42] | Increased mitochondrial biogenesis, derangements of mitochondrial ultrastructure, cardiomyopathy | |
| PGC-1β | Whole-body ablation [28] | Impaired mitochondrial role, reduced trunk weight and fat mass, increased thermogenesis, blunted chronotropic response to dobutamine in the middle, hepatic steatosis, reduced lipoprotein-associated triglyceride and cholesterol content |
| Whole-torso ablation [26] | Decreased action during the dark cycle, abnormal hypothermia and morbidity, hepatic steatosis, increased serum triglyceride and cholesterol | |
| Deletion of exons iii–4 [43] | Mitochondrial dysfunction in liver and skeletal muscle, hepatic steatosis, hepatic insulin resistance | |
| Muscle-specific overexpression [31] | Increased fat-acid oxidation, hyperphagia, reduced torso weight and adipose tissue, increased do capacity, increased IIX fiber content | |
| PGC-1α/β | Double knockout [25] | Neonatal lethality, bradycardia, intermittent centre block, reduced cardiac output, reduced growth, a belatedly fetal abort in mitochondrial biogenesis |
| Models | Manipulations | Phenotypes |
|---|---|---|
| PGC-1α | Whole-body ablation [39] | Hyperactive, cold-sensitive, resistant to nutrition-induced obesity, lesions in the striatum |
| Whole-body ablation [40] | Reduced muscle performance and exercise chapters, impaired adaptive thermogenesis, hepatic steatosis | |
| Muscle-specific ablation [41] | Impaired glucose tolerance, normal peripheral insulin sensitivity | |
| Muscle-specific overexpression [29] | Switch of type II muscle fiber to type IIa and I muscle fibers, resistance to electrically stimulated fatigue | |
| Cardiac-specific overexpression [24] | Loss of sarcomeric construction, dilated cardiomyopathy | |
| Tetracycline-inducible, cardiac-specific overexpression [42] | Increased mitochondrial biogenesis, derangements of mitochondrial ultrastructure, cardiomyopathy | |
| PGC-1β | Whole-torso ablation [28] | Dumb mitochondrial office, reduced body weight and fat mass, increased thermogenesis, blunted chronotropic response to dobutamine in the heart, hepatic steatosis, reduced lipoprotein-associated triglyceride and cholesterol content |
| Whole-torso ablation [26] | Decreased activity during the dark bicycle, abnormal hypothermia and morbidity, hepatic steatosis, increased serum triglyceride and cholesterol | |
| Deletion of exons 3–4 [43] | Mitochondrial dysfunction in liver and skeletal muscle, hepatic steatosis, hepatic insulin resistance | |
| Muscle-specific overexpression [31] | Increased fatty-acrid oxidation, hyperphagia, reduced body weight and adipose tissue, increased do capacity, increased IIX fiber content | |
| PGC-1α/β | Double knockout [25] | Neonatal lethality, bradycardia, intermittent middle cake, reduced cardiac output, reduced growth, a late fetal abort in mitochondrial biogenesis |
Encephalon
PGC-1α cypher mice display spongioform lesions in several encephalon areas, predominantly in the striatum, and exhibit behavioral abnormalities including marked hyperactivity and frequent limb clasping [39]. Recent studies of mice with neuron-specific PGC-1α inactivation back up a crucial role of this cistron in neuronal role and energy balance. Similar brain lesions are observed when PGC-1α is selectively ablated in CaMKIIα-positive neurons, providing direct evidence for its activeness in neurons. Of note, striatal degeneration with hyperactivity is reminiscent of Huntington's illness (HD) in humans, potentially implicating PGC-1α in the selective vulnerability of striatal neurons in HD. To date, the specific role of PGC-1α in linking mitochondrial dysfunction to Hd pathogenesis has been explored. The mutant huntingtin protein accumulated in HD brain interferes with PGC-1α function past repressing its transcription [44]. Dumb PGC-1α expression and mitochondrial function contributes to neurodegeneration in susceptible neurons [21]. In addition, PGC-1α plays an important role in the regulation of genes responsible for the detoxification of reactive oxygen species (ROS), including copper/zinc superoxide dismutase (SOD1), manganese SOD (SOD2), and glutathione peroxidase 1 [22]. In this case, PGC-1α protects dopaminergic neurons from degeneration caused by oxidative stress. Taken together, the finding that PGC-1α expression is impaired in the striatum of Hard disk drive patients raises the possibility that molecules activating PGC-1α may be therapeutically useful.
Center
Middle is an organ with an extremely high and dynamic need for ATP. Much of this supply comes from fat-acid β-oxidation, though glucose also serves equally fuel source. Several studies have demonstrated that PGC-1α is a crucial regulator of oxidative metabolism in the heart. PGC-1α mRNA levels are strongly induced in the neonatal centre, along with the activation of mitochondrial biogenesis and the metabolic switch from glycolysis to oxidative phosphorylation in cardiac musculus [24]. Overexpression of PGC-1α both in vitro and in vivo powerfully induces mitochondrial gene expression and biogenesis [24]. PGC-1α expression is reduced in several animal models of cardiac dysfunction, which is typically accompanied by a metabolic switch from fat oxidation to glycolysis [45]. PGC-1α null mice exhibit significantly lower cardiac reserve in response to electric or chemical stimulation [46]. Moreover, PGC-1α null mice develop early symptoms of eye failure, such equally activation of the fetal program of cardiac gene expression and a meaning increase in circulating levels of atrial natriuretic peptide, a hallmark of cardiac dysfunction [46]. These mice also exhibit lower treadmill-running capacity and macerated cardiac function afterward practise. Still, information technology should exist noted that superphysiological expression of PGC-1α in the middle leads to robust mitochondrial proliferation and myofibrillar displacement, and dilated cardiomyopathy ensues [24]. Equally such, therapeutic regulation of PGC-1α in eye failure should aim at restoring PGC-1α function in cardiac muscle within a therapeutically benign window.
PGC-1β is also abundantly expressed in the eye [47]. Heart part in PGC-1β-scarce mice is largely unaffected under normal weather condition [26]. Still, PGC-1β ablation reduces mitochondrial content in cardiac muscle and blunts the issue of adrenergic stimulation on heart charge per unit [26]. Remarkably, mice with combined deficiency of PGC-1α and PGC-1β (PGC-1αβ−/−) die before long afterwards nascency with small-scale hearts, bradycardia, intermittent heart block, and a markedly reduced cardiac output [25]. Cardiac-specific ablation of PGC-1β on a PGC-1α-deficient background results in cardiac defects including reduced growth, a late fetal arrest in mitochondrial biogenesis, and persistence of a fetal pattern of gene expression [25]. These observations propose that PGC-1α and PGC-1β collectively are required for the postnatal metabolic and functional maturation of the centre.
Brown fat
In rodents, BAT is the major organ responsible for adaptive thermogenesis during cold exposure. In contrast to white adipose tissue, whose principal physiological function is energy storage, the main function of BAT is energy dissipation, largely in the class of oestrus. The expression of PGC-1α is strongly induced in BAT past cold temperature; PGC-1α is downstream of the β-adrenergic receptor/cAMP pathway and sympathetic nervous organization activeness [1]. In this case, PGC-1α turns on several primal components involved in the adaptive thermogenic program, including the stimulation of fuel uptake, mitochondrial fat-acid β-oxidation, and stimulation of uncoupling protein 1 (UCP1) expression [1]. PGC-1α interacts with other nuclear hormone receptors such as PPARα, retinoic acrid receptor, and thyroid receptor to heighten UCP1 expression. UCP1 dissipates mitochondrial proton gradient by generating heat and uncouples oxidative phosphorylation from ATP product. PGC-1α-deficient mice are unable to defend against cold stress due to thermogenic defects [39].
PGC-1β mRNA is induced during white and brown adipocyte differentiation [27]. Interestingly, while the expression of PGC-1β is not cold inducible, its deficiency too impairs adaptive thermogenesis [28], suggesting that these two coactivators play not-redundant function in fuel oxidation and thermogenic response.
Skeletal muscle
PGC-1α is abundantly expressed in skeletal muscle, particularly slow-twitch myofibers, and is rapidly inducible by do preparation in rodents and humans [48–50]. Information technology is clear that calcium signaling pathways play important roles in the consecration of PGC-1α through calcineurin and calcium-dependent protein kinases and the subsequent activation of CREB and myocyte-enhancing factor 2 [51,52]. In improver, p38 mitogen-activated poly peptide kinase (p38 MAPK) and AMP-dependent kinase (AMPK) are also required for exercise-induced PGC-1α expression [51,53]. Interestingly, muscle-specific overexpression muscle creatine kinase (MCK) of PGC-1α in mice turns white, glycolytic skeletal muscles (fast-twitch muscle fibers) into crimson, oxidative muscles (slow-twitch muscle fibers) with increased mitochondrial biogenesis as well every bit the expression of contractile proteins feature of slow-twitch myofibers [29]. In improver to the regulation of mitochondrial role, PGC-1α increases mRNA content of enzymes involved in fatty metabolism such as fatty-acid translocase/CD36, carnitine palmitoyltransferase I, and medium-chain acyl-coenzyme A dehydrogenase (MCAD) in skeletal musculus [54]. Consequent with the molecular changes, PGC-1α transgenic musculus has increased fatigue resistance post-obit electrical stimulation [29]. In dissimilarity, both whole-trunk and muscle-specific PGC-1α knock out (KO) mice show reduced mRNA and/or protein content of mitochondrial respiratory chain proteins and ATP synthase. They are exercise intolerant and their skeletal muscles are prone to contraction-induced fatigue [40].
In chief cultures of rat muscle cells, PGC-1β increases the expression of glucose transporter 4, myosin heavy chain Ib, and other irksome-twitch musculus markers [55]. While PGC-1β besides stimulates mitochondrial biogenesis in skeletal muscle, it appears to bulldoze a program of gene expression that is reminiscent of blazon IIx fibers [31]. In addition, the expression of PGC-1β, just not PGC-1α, is decreased forth with reduced ERRα action and MCAD expression in skeletal muscle of senescence-accelerated mice [56].
Liver
Hepatic PGC-1α expression reaches its peak during early postnatal menses [vii]. In adults, starvation induces PGC-1α expression in the liver through glucagon and GR signaling [32]. PGC-1α orchestrates a complex program of metabolic changes that occur during the transition of a fed to a fasted liver, including gluconeogenesis, fatty-acid β-oxidation, ketogenesis, heme biosynthesis, and bile-acid homeostasis. These effects of PGC-1α on fasting adaption are achieved by coactivating key hepatic transcription factors, such as HNF4α, PPARα, GR, FOXO1, FXR, and LXR [18]. In accord with these observations, PGC-1α KO mice and RNAi-mediated liver-specific PGC-1α knockdown mice display the impairment of gluconeogenic factor expression and hepatic glucose production [39,57]. These mice develop hypoglycemia and hepatic steatosis upon fasting [xl]. In addition, PGC-1α regulates the genes encoding homocysteine synthesis enzymes in the liver and modulates plasma homocysteine levels [33]. Forced expression of PGC-1α in vivo leads to elevated plasma homocysteine levels.
In mammals, circadian clock regulates major aspects of energy metabolism, including glucose and lipid homeostasis and mitochondrial respiration. Our recent work revealed that PGC-1α is a primal component of the cyclic oscillator that integrates the peripheral clock and energy metabolism [34]. PGC-1α stimulates the expression of Bmal1, a core clock gene, in hepatocytes and muscle cells through coactivation of the ROR family of orphan nuclear receptors. Mice lacking PGC-1α have abnormal diurnal rhythms of activity, body temperature, and metabolic rate. As PGC-1α expression is regulated by nutritional and hormonal cues, it is likely that it links these signals to the clockwork and synchronizes tissue metabolism with circadian pacemaker.
PGC-1β expression is increased in response to dietary intake of fats and leads hyperlipidemia through activating hepatic lipogenesis and very-depression-density lipoprotein (VLDL) secretion [35]. Several factors are involved in mediating the effects of PGC-1β on plasma triglyceride metabolism, including sterol response element-binding poly peptide (SREBP), LXR, and Foxa2 [35,58]. Recent studies demonstrated that PGC-1β and its target gene apolipoprotein C3 (ApoC3) are downstream of nicotinic acid, a widely used hypotriglyceridemic drug [36]. Both acute injection and chronic feeding of mice with nicotinic acrid suppress PGC-1β and ApoC3 expression in the liver [36]. These studies illustrated a new role for PGC-1β in modulating lipoprotein catabolism and the relevance of this pathway in therapeutic activity of nicotinic acid. Remarkably, systemic commitment of antisense oligonucleotide targeting PGC-1β improved systemic metabolic homeostasis in the model of fructose-induced insulin resistance [59].
Pancreatic islets
β-Cell dysfunction is cardinal for the development of type 2 diabetes. PGC-1α is expressed in pancreatic β-cells and is elevated in animate being models of type 2 diabetes [37]. Ectopic expression of PGC-1α impairs both early on and delayed glucose-stimulated insulin secretion (GSIS) and suppresses membrane depolarization without affecting baseline insulin secretion. Altered PGC-1α expression is accompanied past increased glucose-6-phosphatase and reduced glucokinase gene expression. Furthermore, UCP2 may exist another effector downstream of PGC-1; UCP2 mediates mitochondrial proton leak, decreases ATP production, and negatively regulates GSIS [lx]. Although the mechanism through which PGC-1α is induced in diabetic animal models is not understood, fatty acids [61] and incomplete inactivation of FOXO1 [62] may contribute to this process. In contrast to animal data, studies in human being type two diabetic islets showed that PGC-1α mRNA expression is markedly reduced and correlated with the reduction in insulin secretion in those islets [63]. Dna methylation of the PGC-1α factor promoter is increased in human diabetic islets. Therefore, the exact function of PGC-1α in pancreatic islets needs further study.
The office of PGC-1β in islets is less studied. A recent study indicated that PGC-1β, in contrast to PGC-1α, straight binds to and acts equally a coactivator of SREBPs and Foxa2 involved in pancreas development and function [38]. The authors also showed that PGC-1β suppresses GSIS via upregulation of UCP2 and granuphilin gene expression in INS-1E cells [38].
Post-translational modifications of PGC-1 coactivators
PGC-1α undergoes extensive post-translational modifications, including acetylation, phosphorylation, methylation, and SUMOylation, in response to nutritional and hormonal signals. These modifications allow fine-tuning of PGC-1α activities in a context-dependent manner. The acetyl transferase general control of amino-acid synthesis 5 acetylates PGC-1α at several lysine residues, alters its localization inside the nucleus, and inhibits its transcriptional activeness [64]. On the contrary, deacetylation of PGC-1α through sirtuin 1 (Sirt1) increases PGC-1α activity on gluconeogenic gene transcription in the liver [65]. PGC-1α is phosphorylated by both p38 MAPK and AMPK in skeletal muscle [66,67], leading to a more stable and agile poly peptide. In contrast, phosphorylation of PGC-1α past Akt/protein kinase B downstream of the insulin signaling cascade in the liver decreased its stability and transcriptional activity [68]. In addition, PGC-1α also undergoes methylation at several arginine residues in the C-terminal region past protein arginine methyltransferase ane [69]. Finally, PGC-1α tin undergo SUMOylation in conserved lysine rest 183 and its transcriptional activity is adulterate [70]. Interestingly, experiments using C2C12 cells accept indicated that AMPK-mediated phosphorylation primes PGC-1α for deacetylation by Sirt1 [71], suggesting that unlike modifications of PGC-1α likely communicate with each other to coordinately regulate its action. PGC-1β is also acetylated at multiple sites [72]; however, the biological significance of these events is less well defined.
PGC-1α and metabolic diseases
Every bit PGC-1α regulates multiple aspects of energy metabolism, it is not surprising that PGC-1α has been found to be dysregulated in several pathological weather. The expression of PGC-1α and its target genes involved in mitochondrial oxidative phosphorylation (OXPHOS) is significantly decreased in the skeletal musculus of patients with type 2 diabetes [73]. Like reduction of PGC-1α expression was also observed in the adipose tissue of insulin-resistant and morbidly obese individuals [74]. Interestingly, thiozolidinedione, an important class of antidiabetic drugs, can heighten the expression of PGC-1α and mitochondrial biogenesis in white adipose tissue [75]. While these observations back up a potentially beneficial function of PGC-1α in insulin resistance and blazon 2 diabetes, several studies suggested singled-out deportment of PGC-1α in other tissues. For example, PGC-1α expression is elevated in the liver of both blazon one and type ii diabetic mouse models [76]. Furthermore, PGC-1α has been shown to stimulate hepatic glucose production and suppress β-prison cell energy metabolism and insulin release in mice [32,37]. Paradoxically, transgenic expression of PGC-1α in skeletal muscle leads to robust mitochondrial biogenesis but also causes insulin resistance, probable the result of imbalance of lipid uptake and oxidation [77]. In addition, a mutual polymorphism of the PGC-1α cistron (Gly482Ser), which apparently reduces PGC-1α action, has been linked to increased risk of type 2 diabetes [78].
In the cardiovascular system, PGC-1α expression is besides decreased in hypertrophic center [45]. PGC-1α null mice display accelerated cardiac dysfunction and clinical signs of heart failure [46]. In contrast, PPARα ligand-dependent transcriptional activity and coactivation by PGC-1α are enhanced in the eye by stress including ischemia and hypoxia [79]. In peripheral vessel tissues, downregulation of PGC-1α expression was observed in vascular smoothen muscle cells (VSMCs) treated by oleic acid [fourscore] and high glucose [81]. Restoration of PGC-1α has beneficial effects on VSMCs and endothelial cells [82]. In this context, PGC-1α appears to play an important part in ROS metabolism and defense against oxidative stress. These observations indicate that PGC-1α is an important factor in the regulation of cardiovascular role.
Abnormalities in mitochondrial function are associated with neurodegenerative disorders including Parkinson's disease, Alzheimer's disease, and HD. Levels of PGC-1α are reduced in the brain of HD patients due to repression of PGC-1α cistron expression by mutant huntingtin, leading to mitochondrial defects and increased oxidative stress [83]. Expression of PGC-1α partially reverses the toxic effects and provides neuroprotection in the HD mutant mouse [44]. In the peripheral nervous system, PGC-1α has been shown to regulate cistron expression at the neuromuscular junction and influences expression of acetylcholine receptors in musculus fibers [84]. In improver, elevated PGC-1α levels protect neural cells in culture from prison cell expiry caused by oxidative-stressor through its consecration of antioxidant genes [22].
Energy metabolism in cancer cells differs fundamentally from that in its normal counterparts. In full general, cancer cells have high glycolytic activeness and prefer glucose as a fuel source, a phenomenon known as the Walburg consequence. The switch from OXPHOS to glycolysis occurs fifty-fifty in the presence of sufficient oxygen. This aerobic glycolysis has been postulated to heighten cancer jail cell proliferation and survival. Interestingly, reduced expression of PGC-1α has been observed in human breast cancer [85], colon cancer [86], liver cancer [87], and ovarian cancer [88]. Adenoviral-mediated overexpression of PGC-1α induces E-cadherin expression while decreasing motility of human hepatoma HepG2 cells [89]. Such manipulation too causes jail cell apoptosis in human ovarian cancer cells through a PPARγ-dependent pathway [88]. These findings suggest that PGC-1α is a potentially of import regulator of cancer cell metabolism and contributes to contradistinct metabolic role in cancer cells. A causal relationship between PGC-1α and cancer development, however, remains to exist established.
Summary and perspective
The PGC-1 family of transcriptional coactivators has emerged as a regulatory hub inside the transcriptional networks that maintain metabolic homeostasis. The dynamic regulation of PGC-1 expression and/or post-translational modification in response to nutritional and hormonal signals provides a highly versatile regulatory platform for metabolic control. A major challenge is to map out tissue-specific activities of PGC-1α and PGC-1β as well as the transcriptional partners that mediate PGC-i actions. Of note, recent genome-wide coactivation analyses provide a global view of potential PGC-1α interacting proteins in humans. While the biological office of nuclear receptor targets of PGC-1 is better understood, very little is known almost a big number of zinc-finger proteins that associate with PGC-1α. The identification of PGC-1α splicing isoforms also adds additional complexity [90]. Dysregulation of PGC-1 expression has been observed in a wide variety of pathological weather condition. As such, proper modulation of PGC-1α expression/activity has groovy potential in the prevention and treatment of diseases associated with impaired mitochondrial role and oxidative metabolism. Although targeting a coactivator could exist challenging, certain features of coactivator action tin be exploited for clinical utilise. For case, coactivators have kinetic benefits in decision-making biological programs in that they coordinate different steps in biological programs through the integration of the activeness of various transcription factors. Every bit such, meaning biological effects can exist achieved by quantitatively modulating coactivator office. The diversity of mail-translational modifications of PGC-1 potentially allows targeting specific protein–protein interaction interface. As discussed above, tissue-specific modulation of PGC-one function is essential for metabolic modulation without causing deleterious side furnishings.
Funding
This work was supported past the grants from the National Natural Scientific discipline Foundation of China (30870928), the Research Fund for the Doctoral Program of Higher Teaching of Mainland china (20103207110007), the Fok Ying Tong Education Foundation (121022), the Major Program of Educational Commission of Jiangsu Province (09KJA180004) (to C.L.), and the NIH grant (DK077086) (to J.D.L.).
Acknowledgements
Owing to space limitations, nosotros apologize to those whose publications related to the discussed issues could not exist cited.
References
1
, , , , , .
A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis
,
Cell
,
1998
, vol.
92
(pg.
829
-
839
)
ii
, , , , , , .
Activation of PPARγ coactivator-i through transcription factor docking
,
Science
,
1999
, vol.
286
(pg.
1368
-
1371
)
3
, , , , .
Coordination of p300-mediated chromatin remodeling and TRAP/mediator function through coactivator PGC-1α
,
Mol Jail cell
,
2003
, vol.
12
(pg.
1137
-
1149
)
4
, , , , , , , et al.
Genome-wide coactivation analysis of PGC-1α identifies BAF60a as a regulator of hepatic lipid metabolism
,
Prison cell Metab
,
2008
, vol.
8
(pg.
105
-
117
)
v
, , , , , .
Straight coupling of transcription and mRNA processing through the thermogenic coactivator PGC-1
,
Mol Cell
,
2000
, vol.
6
(pg.
307
-
316
)
half dozen
, , , .
Glucocorticoid signaling is perturbed by the singular orphan receptor and core pressor SHP
,
J Biol Chem
,
2002
, vol.
277
(pg.
49761
-
49766
)
seven
, , , , , , .
PGC-1β in the regulation of hepatic glucose and energy metabolism
,
J Biol Chem
,
2003
, vol.
278
(pg.
30843
-
30848
)
8
, .
Pgc-1-related coactivator, a novel, serum-inducible coactivator of nuclear respiratory factor 1-dependent transcription in mammalian cells
,
Mol Cell Biol
,
2001
, vol.
21
(pg.
3738
-
3749
)
nine
, , , .
The Drosophila PGC-1 homologue Spargel coordinates mitochondrial activity to insulin signaling
,
EMBO J
,
2010
, vol.
29
(pg.
171
-
183
)
x
, , , , , , , et al.
Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1
,
Cell
,
1999
, vol.
98
(pg.
115
-
124
)
11
, , , .
Human peroxisome proliferator activated receptor γ coactivator ane (PPARGC1) gene: cDNA sequence, genomic organization, chromosomal localization, and tissue expression
,
Genomics
,
1999
, vol.
62
(pg.
98
-
102
)
12
, , .
A tissue-specific coactivator of steroid receptors, identified in a functional genetic screen
,
Mol Cell Biol
,
2000
, vol.
20
(pg.
2411
-
2422
)
13
, , , , , , , et al.
Errα and Gabpa/b specify PGC-1α-dependent oxidative phosphorylation cistron expression that is altered in diabetic muscle
,
Proc Natl Acad Sci U.s.a.
,
2004
, vol.
101
(pg.
6570
-
6575
)
14
, , , , , , , et al.
The estrogen-related receptor α (ERRα) functions in PPARγ coactivator 1α (PGC-1α)-induced mitochondrial biogenesis
,
Proc Natl Acad Sci U.s.a.
,
2004
, vol.
101
(pg.
6472
-
6477
)
15
.
Nuclear activators and coactivators in mammalian mitochondrial biogenesis
,
Biochim Biophys Acta
,
2002
, vol.
1576
(pg.
1
-
14
)
xvi
.
Transcriptional activators and coactivators in the nuclear control of mitochondrial function in mammalian cells
,
Gene
,
2002
, vol.
286
(pg.
81
-
89
)
17
, , , , , , , et al.
Mitochondrial transcription factor A is necessary for mtDNA maintenance and embryogenesis in mice
,
Nat Genet
,
1998
, vol.
18
(pg.
231
-
236
)
18
, , .
Metabolic control through the PGC-ane family unit of transcription coactivators
,
Jail cell Metab
,
2005
, vol.
one
(pg.
361
-
370
)
19
, .
PGC-1α: a central regulator of energy metabolism
,
Adv Physiol Educ
,
2006
, vol.
thirty
(pg.
145
-
151
)
twenty
, , , .
PGC-1α: a potent transcriptional cofactor involved in the pathogenesis of type ii diabetes
,
Diabetologia
,
2006
, vol.
49
(pg.
1477
-
1488
)
21
, , , , , , , et al.
Thermoregulatory and metabolic defects in Huntington'southward illness transgenic mice implicate PGC-1α in Huntington'southward disease neurodegeneration
,
Cell Metab
,
2006
, vol.
4
(pg.
349
-
362
)
22
, , , , , , , et al.
Suppression of reactive oxygen species and neurodegeneration by the PGC-1 transcriptional coactivators
,
Cell
,
2006
, vol.
127
(pg.
397
-
408
)
23
, , .
Localization of the transcriptional coactivator PGC-1α to GABAergic neurons during maturation of the rat brain
,
J Comp Neurol
,
2007
, vol.
502
(pg.
1
-
18
)
24
, , , , , .
Peroxisome proliferator-activated receptor γ coactivator-i promotes cardiac mitochondrial biogenesis
,
J Clin Invest
,
2000
, vol.
106
(pg.
847
-
856
)
25
, , , , , , , et al.
Transcriptional coactivators PGC-1α and PGC-1β control overlapping programs required for perinatal maturation of the centre
,
Genes Dev
,
2008
, vol.
22
(pg.
1948
-
1961
)
26
, , , , .
PGC-1β controls mitochondrial metabolism to attune cyclic activity, adaptive thermogenesis, and hepatic steatosis
,
Proc Natl Acad Sci The states
,
2007
, vol.
104
(pg.
5223
-
5228
)
27
, , , , , .
Complementary action of the PGC-1 coactivators in mitochondrial biogenesis and brownish fat differentiation
,
Jail cell Metab
,
2006
, vol.
3
(pg.
333
-
341
)
28
, , , , , , , et al.
Ablation of PGC-1β results in defective mitochondrial activity, thermogenesis, hepatic role, and cardiac performance
,
PLoS Biol
,
2006
, vol.
4
pg.
e369
29
, , , , , , , et al.
Transcriptional coactivator PGC-1α drives the formation of irksome-twitch muscle fibres
,
Nature
,
2002
, vol.
418
(pg.
797
-
801
)
thirty
, , , , , , , et al.
Overexpression of PGC-1α leads to muscle cloudburst with depletion of ATP
,
Am J Pathol
,
2006
, vol.
169
(pg.
1129
-
1139
)
31
, , , , , , , et al.
The transcriptional coactivator PGC-1β drives the formation of oxidative type IIX fibers in skeletal musculus
,
Jail cell Metab
,
2007
, vol.
5
(pg.
35
-
46
)
32
, , , , , , , et al.
Control of hepatic gluconeogenesis through the transcriptional coactivator PGC-1
,
Nature
,
2001
, vol.
413
(pg.
131
-
138
)
33
, , , , , , , et al.
Regulation of homocysteine homeostasis through the transcriptional coactivator PGC-1α
,
Am J Physiol Endocrinol Metab
,
2009
, vol.
296
(pg.
E543
-
E548
)
34
, , , , .
Transcriptional coactivator PGC-1α integrates the mammalian clock and free energy metabolism
,
Nature
,
2007
, vol.
447
(pg.
477
-
481
)
35
, , , , , , , et al.
Hyperlipidemic effects of dietary saturated fats mediated through PGC-1β coactivation of SREBP
,
Cell
,
2005
, vol.
120
(pg.
261
-
273
)
36
, , , , .
Regulation of hepatic ApoC3 expression by PGC-1β mediates hypolipidemic effect of nicotinic acrid
,
Cell Metab
,
2010
, vol.
12
(pg.
411
-
419
)
37
, , , , , , , et al.
Suppression of β cell energy metabolism and insulin release by PGC-1α
,
Dev Cell
,
2003
, vol.
5
(pg.
73
-
83
)
38
, , , , .
Transcriptional coactivator peroxisome proliferator-activated receptor (PPAR) γ coactivator-1β is involved in the regulation of glucose-stimulated insulin secretion in INS-1E cells
,
J Mol Med
,
2009
, vol.
87
(pg.
299
-
306
)
39
, , , , , , , et al.
Defects in adaptive free energy metabolism with CNS-linked hyperactivity in PGC-1α null mice
,
Cell
,
2004
, vol.
119
(pg.
121
-
135
)
40
, , , , , , , et al.
PGC-1α deficiency causes multisystem free energy metabolic derangements: muscle dysfunction, abnormal weight control and hepatic steatosis
,
PLoS Biol
,
2005
, vol.
three
pg.
e101
41
, , , , , , , et al.
Abnormal glucose homeostasis in skeletal musculus-specific PGC-1α knockout mice reveals skeletal muscle-pancreatic beta cell crosstalk
,
J Clin Invest
,
2007
, vol.
117
(pg.
3463
-
3474
)
42
, , , , , , , et al.
Cardiac-specific induction of the transcriptional coactivator PGC-1α promotes mitochondrial biogenesis and reversible cardiomyopathy in a developmental stage-dependent manner
,
Circ Res
,
2004
, vol.
94
(pg.
525
-
533
)
43
, , , , , , , et al.
Hypomorphic mutation of PGC-1β causes mitochondrial dysfunction and liver insulin resistance
,
Prison cell Metab
,
2006
, vol.
iv
(pg.
453
-
464
)
44
, , , , , .
Transcriptional repression of PGC-1α by mutant huntingtin leads to mitochondrial dysfunction and neurodegeneration
,
Cell
,
2006
, vol.
127
(pg.
59
-
69
)
45
, .
Mitochondrial energy metabolism in eye failure: a question of balance
,
J Clin Invest
,
2005
, vol.
115
(pg.
547
-
555
)
46
, , , , , , , et al.
Transcriptional coactivator PGC-1α controls the free energy state and contractile role of cardiac muscle
,
Prison cell Metab
,
2005
, vol.
1
(pg.
259
-
271
)
47
, , , , .
Peroxisome proliferator-activated receptor gamma coactivator 1β (PGC-1β), a novel PGC-1-related transcription coactivator associated with host prison cell factor
,
J Biol Chem
,
2002
, vol.
277
(pg.
1645
-
1648
)
48
, , , , , , , et al.
Adaptations of skeletal muscle to practice: rapid increase in the transcriptional coactivator PGC-ane
,
FASEB J
,
2002
, vol.
xvi
(pg.
1879
-
1886
)
49
, , , , , , .
cDNA cloning and mRNA analysis of PGC-1 in epitrochlearis muscle in swimming-exercised rats
,
Biochem Biophys Res Commun
,
2000
, vol.
274
(pg.
350
-
354
)
50
, , , , , .
PGC-1α mRNA expression is influenced past metabolic perturbation in exercising homo skeletal muscle
,
J Appl Physiol
,
2004
, vol.
96
(pg.
189
-
194
)
51
, , , , , , , et al.
Practise stimulates PGC-1α transcription in skeletal muscle through activation of the p38 MAPK pathway
,
J Biol Chem
,
2005
, vol.
280
(pg.
19587
-
19593
)
52
, , , , , , .
Regulation of mitochondrial biogenesis in skeletal musculus by CaMK
,
Scientific discipline
,
2002
, vol.
296
(pg.
349
-
352
)
53
, , , , , , .
AMP kinase is required for mitochondrial biogenesis in skeletal muscle in response to chronic energy deprivation
,
Proc Natl Acad Sci U.s.
,
2002
, vol.
99
(pg.
15983
-
15987
)
54
, , .
PGC-1α-mediated adaptations in skeletal muscle
,
Pflugers Arch
,
2010
, vol.
460
(pg.
153
-
162
)
55
, , , , .
PGC-1α and PGC-1β have both similar and distinct effects on myofiber switching toward an oxidative phenotype
,
Am J Physiol Endocrinol Metab
,
2006
, vol.
291
(pg.
E807
-
E816
)
56
, , , , , , , et al.
PGC-1β downward-regulation is associated with reduced ERRα activity and MCAD expression in skeletal muscle of senescence-accelerated mice
,
J Gerontol A Biol Sci Med Sci
,
2006
, vol.
61
(pg.
773
-
780
)
57
, , , , , , , et al.
PGC-1 promotes insulin resistance in liver through PPAR-α-dependent induction of TRB-3
,
Nat Med
,
2004
, vol.
10
(pg.
530
-
534
)
58
, .
Coactivation of Foxa2 through PGC-1β promotes liver fat acid oxidation and triglyceride/VLDL secretion
,
Cell Metab
,
2006
, vol.
iii
(pg.
99
-
110
)
59
, , , , , , , et al.
The part of peroxisome proliferator-activated receptor gamma coactivator-1 beta in the pathogenesis of fructose-induced insulin resistance
,
Cell Metab
,
2009
, vol.
ix
(pg.
252
-
264
)
60
, , , , , , , et al.
Peroxisome proliferator-activated receptor gamma coactivator-ane-dependent uncoupling poly peptide-2 expression in pancreatic islets of rats: a novel pathway for neural command of insulin secretion
,
Diabetologia
,
2003
, vol.
46
(pg.
1522
-
1531
)
61
, , , , , , .
Gratis fatty acids increase PGC-1α expression in isolated rat islets
,
FEBS Lett
,
2005
, vol.
579
(pg.
1446
-
1452
)
62
, , , , , , , et al.
The forkhead transcription factor Foxo1 links insulin signaling to Pdx1 regulation of pancreatic β cell growth
,
J Clin Invest
,
2002
, vol.
110
(pg.
1839
-
1847
)
63
, , , , , , , et al.
Epigenetic regulation of PPARGC1A in man type 2 diabetic islets and effect on insulin secretion
,
Diabetologia
,
2008
, vol.
51
(pg.
615
-
622
)
64
, , , , , .
GCN5 acetyltransferase complex controls glucose metabolism through transcriptional repression of PGC-1α
,
Cell Metab
,
2006
, vol.
iii
(pg.
429
-
438
)
65
, , , , , .
Nutrient command of glucose homeostasis through a circuitous of PGC-1α and SIRT1
,
Nature
,
2005
, vol.
434
(pg.
113
-
118
)
66
, , , , , , , et al.
Cytokine stimulation of energy expenditure through p38 MAP kinase activation of PPARγ coactivator-one
,
Mol Jail cell
,
2001
, vol.
8
(pg.
971
-
982
)
67
, , , .
AMP-activated poly peptide kinase (AMPK) action in skeletal muscle via direct phosphorylation of PGC-1α
,
Proc Natl Acad Sci USA
,
2007
, vol.
104
(pg.
12017
-
12022
)
68
, , , .
Akt/PKB regulates hepatic metabolism by directly inhibiting PGC-1α transcription coactivator
,
Nature
,
2007
, vol.
447
(pg.
1012
-
1016
)
69
, , , , .
Activation of nuclear receptor coactivator PGC-1α by arginine methylation
,
Genes Dev
,
2005
, vol.
xix
(pg.
1466
-
1473
)
seventy
, .
SUMOylation attenuates the function of PGC-1α
,
J Biol Chem
,
2009
, vol.
284
(pg.
26184
-
26193
)
71
, , , , , , , et al.
AMPK regulates energy expenditure by modulating NAD+ metabolism and SIRT1 activity
,
Nature
,
2009
, vol.
458
(pg.
1056
-
1060
)
72
, , , , .
GCN5-mediated transcriptional command of the metabolic coactivator PGC-1β through lysine acetylation
,
J Biol Chem
,
2009
, vol.
284
(pg.
19945
-
19952
)
73
, , , , , , , et al.
Coordinated reduction of genes of oxidative metabolism in humans with insulin resistance and diabetes: potential function of PGC1 and NRF1
,
Proc Natl Acad Sci USA
,
2003
, vol.
100
(pg.
8466
-
8471
)
74
, , , , , , , et al.
Expression of the thermogenic nuclear hormone receptor coactivator PGC-1α is reduced in the adipose tissue of morbidly obese subjects
,
Int J Obes Relat Metab Disord
,
2004
, vol.
28
(pg.
176
-
179
)
75
, , , , , , , et al.
Mitochondrial remodeling in adipose tissue associated with obesity and treatment with rosiglitazone
,
J Clin Invest
,
2004
, vol.
114
(pg.
1281
-
1289
)
76
, .
Peroxisome proliferator-activated receptor-γ coactivator 1α (PGC-1α): transcriptional coactivator and metabolic regulator
,
Endocr Rev
,
2003
, vol.
24
(pg.
78
-
xc
)
77
, , , , , , , et al.
Paradoxical furnishings of increased expression of PGC-1α on muscle mitochondrial role and insulin-stimulated muscle glucose metabolism
,
Proc Natl Acad Sci U.s.
,
2008
, vol.
105
(pg.
19926
-
19931
)
78
, , , , , , , et al.
Mutation analysis of peroxisome proliferator-activated receptor-gamma coactivator-ane (PGC-one) and relationships of identified amino acid polymorphisms to type II diabetes mellitus
,
Diabetologia
,
2001
, vol.
44
(pg.
2220
-
2226
)
79
, .
PPAR transcriptional activator complex polymorphisms and the promise of individualized therapy for heart failure
,
Eye Fail Rev
,
2010
, vol.
15
(pg.
197
-
207
)
eighty
, , , , , , , et al.
PGC-1α inhibits oleic acid induced proliferation and migration of rat vascular smooth muscle cells
,
PLoS Ane
,
2007
, vol.
two
pg.
e1137
81
, , , , , , , et al.
PGC-1α is a primal regulator of glucose-induced proliferation and migration in vascular smooth musculus cells
,
PLoS One
,
2009
, vol.
iv
pg.
e4182
82
, , , , .
PGC-1α regulates the mitochondrial antioxidant defense arrangement in vascular endothelial cells
,
Cardiovasc Res
,
2005
, vol.
66
(pg.
562
-
573
)
83
, , , , .
PGC-1α as modifier of onset age in Huntington disease
,
Mol Neurodegener
,
2009
, vol.
four
pg.
ten
84
, , , , , .
PGC-1α regulates the neuromuscular junction plan and ameliorates Duchenne muscular dystrophy
,
Genes Dev
,
2007
, vol.
21
(pg.
770
-
783
)
85
, , , .
The localization and reduction of nuclear staining of PPARγ and PGC-ane in human breast cancer
,
Oncol Rep
,
2004
, vol.
12
(pg.
483
-
488
)
86
, , , , .
Peroxisome proliferator-activated receptors (PPARs) and associated transcription factors in colon cancer: reduced expression of PPARγ-coactivator one (PGC-1)
,
Cancer Lett
,
2004
, vol.
203
(pg.
25
-
33
)
87
, , , .
Down-regulation of PGC-1α expression in homo hepatocellular carcinoma
,
Zhonghua Zhong Liu Za Zhi
,
2008
, vol.
30
(pg.
593
-
597
)
88
, , , , , , , et al.
PGC-1α induces apoptosis in human epithelial ovarian cancer cells through a PPARγ-dependent pathway
,
Cell Res
,
2007
, vol.
17
(pg.
363
-
373
)
89
, , , , .
PPARγ/PGC-1α pathway in E-cadherin expression and movement of HepG2 cells
,
Anticancer Res
,
2009
, vol.
29
(pg.
5057
-
5063
)
90
, , , , , , , et al.
Alternative mRNA splicing produces a novel biologically agile brusque isoform of PGC-1α
,
J Biol Chem
,
2009
, vol.
284
(pg.
32813
-
32826
)
© The Writer 2011. Published past ABBS Editorial Office in association with Oxford University Press on behalf of the Constitute of Biochemistry and Jail cell Biology, Shanghai Institutes for Biological Sciences, Chinese University of Sciences.
Source: https://academic.oup.com/abbs/article/43/4/248/884
0 Response to "Metabolic Control Through the Pgc-1 Review Family of Transcription Coactivators"
Post a Comment