Mitochondrial adaptations in insulin resistant muscle
Citation for published version (APA):Broek, van den, N. M. A. (2010). Mitochondrial adaptations in insulin resistant muscle. Technische Universiteit Eindhoven. https://doi.org/10.6100/IR684831
DOI:
10.6100/IR684831
Document status and date: Published: 01/01/2010
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Mitochondrial adaptations
in insulin resistant muscle
A catalogue record is available from the Eindhoven University of Technology Library ISBN: 978‐90‐386‐2299‐6 Printed by:
Mitochondrial adaptations
in insulin resistant muscle
PROEFSCHRIFT
ter verkrijging van de graad van doctor aan de
Technische Universiteit Eindhoven, op gezag van de
rector magnificus, prof.dr.ir. C.J. van Duijn, voor een
commissie aangewezen door het College voor
Promoties in het openbaar te verdedigen
op donderdag 16 september 2010 om 16.00 uur
door
Nicole Martina Adriana van den Broek
geboren te Tilburg
Copromotor:
dr. J.J. Prompers
Chapter 1 General introduction 1 Chapter 2 Comparison of in vivo post‐exercise phosphocreatine recovery and resting ATP synthesis flux for the assessment of skeletal muscle mitochondrial function 31 Chapter 3 Intersubject differences in the effect of acidosis on phosphocreatine recovery kinetics in muscle after exercise are due to differences in proton efflux rates 53 Chapter 4 Early or advanced stage type 2 diabetes is not accompanied by in vivo skeletal muscle mitochondrial dysfunction 77 Chapter 5 Increased mitochondrial content rescues in vivo muscle oxidative capacity in long‐term high‐fat diet fed rats 99 Chapter 6 Carnitine insufficiency in high‐fat diet fed rats does not contribute to lipid‐induced impairment of skeletal muscle mitochondrial function in vivo 123 Chapter 7 Summary and future perspectives 147 Nederlandse samenvatting Dankwoord List of publications Curriculum Vitae 157
Chapter
What was originally termed adult‐onset diabetes is now one of the most dominant diseases in the world: type 2 diabetes (T2D). The prevalence of diabetes is increasing dramatically. The total number of people with diabetes is projected to rise from 171 million in 2000 to 366 million in 2030 [4]. The majority of all diabetes patients (90%) suffers from T2D. The most important reason for the dramatic increase in prevalence of T2D is the expanding Western lifestyle, which encompasses the combination of an excessive calorie intake and a decreased physical activity, leading to fat accumulation and increased body weight. T2D is characterized by insulin resistance (IR), a disorder in which major metabolic tissues, in particular skeletal muscle, are less sensitive for insulin. A first attempt of the body to compensate for the IR of the tissues is to increase insulin production by the pancreatic ‐ cells. IR is progressive and therefore during the years the need for insulin will keep increasing. After some time the overproduction of insulin will lead to -cell failure, resulting in a diminished insulin secretion and sustained elevated blood glucose levels, a state which is called T2D. After several years, T2D can lead to severe life‐threatening complications like cardiovascular disease, neuropathy, retinopathy and kidney failure.
During the last decade excessive ectopic lipid storage and muscle mitochondrial dysfunction have been associated with the pathogenesis of IR and T2D [5]. However, there is still no consensus about the underlying mechanisms leading to IR and consequently T2D. Therefore further investigation into the etiology of IR and T2D is important to conquer the T2D epidemic. The present chapter provides an overview of the background of this thesis. First the most important cellular events leading to IR and T2D will be explained and the energy‐producing organelles, i.e. the mitochondria, will be introduced. Subsequently, different theories about the mechanism of fatty acid‐induced IR will be summarized and the methods applied in this thesis for measuring muscle lipids and lipid intermediates and muscle mitochondrial function are described. Finally, the outline of the thesis is presented.
Glucose homeostasis, insulin resistance and type 2 diabetes
In healthy subjects blood glucose levels are well controlled. After a meal, glucose is taken up in the blood and transported to tissues, where glucose is used for energy production. Glucose is the most important substrate for the brain. However, skeletal muscle is responsible for as much as 70‐80% of postprandial glucose uptake [6]. When blood glucose levels rise, e.g. after a meal, the ‐cells of the pancreas respond by producing more insulin. Insulin can stimulate the ‘insulin sensitive’ glucose uptake, by binding to cell membrane‐ bound insulin receptors, which activates the insulin signaling cascade. Binding of insulin to its receptor leads to autophosphorylation of its ‐subunits and the phosphorylation of
tyrosine residues in insulin receptor substrates (IRS). IRS activates phosphatidylinositol‐3‐ kinase (PI3K) through its SH2 domain, thus increasing the intracellular concentration of PIP2 and PIP. This, in turn, activates phosphatidylinositol phosphate‐dependent kinase‐1 (PDK‐ 1), which subsequently activates Akt/PKB. This activation cascade results in the translocation of the glucose transporter 4 (GLUT4) from cytoplasmic vesicles to the cell membrane, where GLUT4 facilitates glucose uptake (reviewed in [3]). Besides stimulating the uptake of glucose in muscle, insulin also inhibits glucose production in the liver [7]. As a result of those muscle and liver responses, blood glucose levels will return back to basal values in a relatively short time period. In the muscle, glucose can be used for ATP production. However during rest, when little energy is needed, glucose is converted to glycogen, which is stored in the muscle, thus providing an energy reserve that can quickly be mobilized to meet a sudden need for glucose. In the case of IR, postprandial glucose is still cleared from the blood, but because of the decreased sensitivity of the peripheral tissues for insulin, more insulin is needed for a similar blood glucose lowering effect. Sustained insulin overproduction by pancreatic ‐cells leads to ‐cell failure and decreased insulin production. This results in increased basal glucose levels and increased duration of postprandial blood glucose clearance, which is a condition called T2D. The high blood glucose levels can irreversibly damage tissues, finally leading to well known complications of T2D, e.g. cardiovascular disease, neuropathy, retinopathy and kidney failure. The dramatic rise in T2D is associated with the increased occurrence of obesity and the decrease of physical activity. However, the primary cause of IR and T2D is still not known. During the last decade, skeletal muscle mitochondrial dysfunction has been proposed to play a key role in the development of IR and T2D [8‐13].
Mitochondrial energy production in muscle
Mitochondria are called the ‘powerhouses’ of the cell. Without them, cells would be unable to extract significant amounts of energy from the nutrients, and as a consequence, essentially all cellular functions would fail under high energy demand conditions. The specific structure of the mitochondrion is essential for synthesizing adenosine triphosphate (ATP), the energy source for cells. The organelle is surrounded by two lipid bilayer membranes: an outer and an inner membrane. Many foldings of the inner membrane form cristae, in which oxidative enzymes (NADH dehydrogenase (complex I), cytochrome c reductase (complex III), and cytochrome c oxidase (complex IV)), contributing to the electron transport chain (ETC), and the F1F0‐ATP synthase complex (complex V), responsible
for proton‐driven ATP synthesis, are located (Figure 1). In addition, the inner cavity of the mitochondrion, termed the matrix, contains large quantities of enzymes, which are involved in the oxidation of carbohydrates (CHO) and fatty acids (FAs), i.e. enzymes from the tricarboxylic acid (TCA) cycle and the ‐oxidation pathway, respectively [14].
ATP is used throughout the cell to drive almost all energy‐dependent cellular processes, like active transport of molecules and ions, the synthesis of macromolecules and the performance of mechanical work during muscle contractions and other cellular movements [14]. ATP is therefore also called the energy currency of the cell. ATP is a nucleotide composed of an adenine ring and a ribose sugar (adenosine) and three phosphate groups (triphosphate). The phosphate groups, starting with the group closest to the ribose, are referred to as the alpha (), beta (), and gamma (γ) phosphates. Two phospho‐anhydride bonds (those that connect adjacent phosphates) in ATP are responsible for the high energy content of this molecule. In the context of biochemical reactions, these anhydride bonds are frequently referred to as high‐energy bonds. Energy stored in ATP is released upon hydrolysis of the terminal anhydride bond, thereby forming adenosine diphosphate (ADP) and inorganic phosphate (Pi).
Muscle contraction depends on energy supplied by ATP. The concentration of ATP in the muscle is sufficient to maintain full contraction for only 1 or 2 seconds at most. Therefore, the formed ADP has to be rephosphorylated back to ATP immediately, allowing the muscle to continue its contraction. There are several sources of energy for the rephosphorylation of ADP. The source of energy that is used to reconstitute ATP within the first few seconds of muscle contraction is phosphocreatine (PCr), which carries a high‐energy phosphate bond,
Figure 1. Schematic overview depicting the different components of oxidative phosphorylation, i.e. the
electron transport chain and the F1F0‐ATP synthase in the inner mitochondrial membrane. NADH and
FADH2 donate their electrons to complex I and II, respectively. Electron transfer through complexes I, III
and IV is accompanied by the pumping of protons from the matrix into the intermembrane space, leading to the generation of an electrochemical proton gradient. This gradient is used to drive the translocation of protons through the F1F0‐ATP synthase, which goes along with the formation of ATP.
IMM, inner mitochondrial membrane; complex I, NADH dehydrogenase; complex II, succinate dehydrogenase; complex III, cytochrome bc1 complex; complex IV, cytochrome c oxidase; F1F0 ATP‐ase,
with a slightly higher free energy content than those in ATP. Therefore, phosphocreatine is instantly cleaved and the released energy is used to rephosphorylate ADP to ATP by the creatine kinase (CK) reaction:
MgADP PCr H MgATP Cr Eq. 1
After about five seconds of maximal muscle contraction, PCr is depleted and the next source of energy to reconstitute ATP is addressed, i.e. glycogen. Glycogen has previously been stored in large amounts in muscle cells, thereby providing a rapid source of energy for the muscle in situations of high‐energy demand, i.e. muscle contractions longer than a few seconds. Cleavage of glycogen results in the formation of glucose‐1‐phosphate which, after conversion into glucose‐6‐phosphate, can enter the glycolytic pathway. Via glycolysis, glucose‐6‐phosphate, either formed from glycogen or from glucose directly, is rapidly broken down to pyruvate, thereby forming 2 molecules of ATP. Besides the advantage of fast ATP production, glycolysis has the ability to produce ATP in the absence of oxygen by producing lactate. Unfortunately, however, during anaerobic glycolysis so many end products accumulate that the muscle will lose its capability to sustain maximum contraction after about one minute.
The third source of energy is oxidative metabolism. More than 95% of ATP is formed in the mitochondria via oxidative phosphorylation, by degrading substrates like CHO and FA aerobically. Both substrates have to be converted to the general degradation product acetyl coenzyme A (acetyl‐CoA). Pyruvate, formed via glycolysis, is subsequently transported into the mitochondria and converted to acetyl‐CoA by the pyruvate dehydrogenase complex. Within the outer mitochondrial membrane, the enzyme fatty acyl‐ CoA synthetase (ACS) catalyses the first step in intracellular FA metabolism by converting FAs into fatty acyl‐CoA’s. Upon activation by ACS the resulting fatty acyl‐CoA’s can be transported across the mitochondrial membrane via the carnitine shuttle, which consists of three components: carnitine palmitoyltransferase (CPT) 1 and 2 and carnitine acylcarnitine translocase (CACT). CPT1 executes the initial step in this process by catalyzing the reversible transesterification of long‐chain acyl‐CoA with carnitine. The long‐chain acylcarnitine products of CPT1 are then transported across the inner mitochondrial membrane by CACT. Finally, inside the mitochondrion, CPT2 regenerates acyl‐CoA on the matrix side of the membrane where it can enter the ‐oxidation. Each cycle of the ‐oxidation consists of four reactions, shortening the acyl‐CoA unit by two carbon atoms. Thereby, every cycle one nicotinamide adenine dinucleotide (NADH), one flavin adenine dinucleotide (FADH2) and an
acetyl‐CoA molecule are released. This process continues until the entire acyl chain is cleaved into acetyl‐CoA units. Acetyl‐CoA, derived from CHO via pyruvate or from FAs, then
enters the next step in the oxidative process: the TCA cycle. Acetyl‐CoA transfers its two‐ carbon acetyl group to oxaloacetate to form a six‐carbon compound, i.e. citrate. Citrate then undergoes a series of chemical transformations resulting in oxaloacetate, to which a new acetyl‐CoA can be conjugated to enter the TCA cycle again. During each cycle one ATP, two CO2, one FADH2 and three NADH molecules are released. The electron donors, NADH
and FADH2, transfer their electrons to oxygen, which is reduced to water in the ETC. The
enzymes that catalyze these reactions have the ability to simultaneously create a proton gradient across the inner mitochondrial membrane, which drives the F1F0‐ATP synthase
complex to form ATP from ADP and Pi:
MgATP H O MgADP P H Eq. 2
Figure 2. Schematic overview of FA transport across the mitochondrial membranes by the carnitine
shuttle and the metabolic conversions of FAs and pyruvate to yield ATP. ACS exerts the first step in FA transport by converting FAs into fatty acyl‐CoA’s. CPT1 catalyzes the reversible transesterification of long‐chain acyl‐CoA with carnitine. These long‐chain acylcarnitine species are then transported across the inner mitochondrial membrane by CACT. Inside the mitochondrion, CPT2 regenerates acyl‐CoA on the matrix side of the membrane where it can enter the ‐oxidation, which generates acetyl‐CoA. Pyruvate is transported into the mitochondrial matrix, where it is converted to acetyl‐CoA by PDH. Acetyl‐CoA can enter the TCA cycle. NADH and FADH2, produced in the TCA cycle, enable ATP production
by OXPHOS as explained in more detail in Figure 1. OMM, outer mitochondrial membrane; ACS, acyl‐ CoA synthetase; CPT, carnitine palmitoyl transferase; CACT, carnitine‐acylcarnitine translocase; PDH, pyruvate dehydrogenase; TCA, tricarboxylic acid cycle; OXPHOS, oxidative phosphorylation.
Mechanisms of fatty acid‐induced muscle insulin resistance and
the role of mitochondrial dysfunction
T2D and IR are often associated with hyperlipidemia (excessively high plasma triglyceride and cholesterol levels) as well as elevated levels of plasma FFA (reviewed in [15‐16]). It has been shown that raising plasma FFA levels by intravenously administering IntralipidTM decreases whole body glucose uptake, oxidation and storage [17] and skeletal muscle glucose uptake [18‐19]. Furthermore, lowering of plasma FFA has been shown to reduce the severity of insulin resistance [20]. Plasma FFAs can easily enter cells where they are either oxidized to generate energy in the form of ATP or re‐esterified for storage as triglycerides (TG). Not surprisingly, therefore, raising plasma FFA levels results in intramyocellular accumulation of triglycerides, the so‐called intramyocellular lipids (IMCL) [21], which are correlated to the severity of IR [22]. It was shown that IMCL content is a better predictor for IR than plasma FFA levels [23]. However IMCL is probably not interfering with the insulin signaling pathway directly, but rather represents a surrogate marker of a systemic lipid oversupply, which is believed to cause IR. In this chapter several hypotheses about the underlying mechanism of lipid‐induced IR are summarized.
Inhibition of glucose oxidation
Randle et al. introduced the ‘glucose‐fatty acid cycle’ or ‘Randle cycle’ [2]. In isolated heart and skeletal muscle preparations, Randle et al. demonstrated that the utilization of one nutrient inhibited the use of the other directly and without hormonal mediation. Most emphasis was put on the control of glucose oxidation by FAs. Due to an elevated FA oxidation, the intra‐mitochondrial acetyl‐CoA/CoA ratio increases, resulting in the inhibition of PDH, a regulatory enzyme in glucose metabolism. It has been shown that these changes lead to elevated citrate levels in the cytosol, which inhibit PFK‐1 [24], followed by an increase in glucose‐6‐phosphate, which eventually inhibits hexokinase and therefore diminishes glucose uptake, resulting in elevated plasma glucose levels (Figure 3). Overall, Randle concluded that an increased FA availability leads to increased ‐oxidation and a decrease in muscle glucose uptake and oxidation, and can therefore be seen as a possible cause for elevated plasma glucose levels and skeletal muscle IR [2].
Inhibition of glucose transport and mitochondrial dysfunction
Several years later, the Randle cycle was challenged. A series of MRS studies performed by the group of Shulman pointed out that glucose transport was impaired directly by elevated FFA, without an increase in glucose‐6‐phosphate levels [25]. In addition, others have shown that infusion of lipids induced IR only several hours after it had already decreased glucose oxidation, suggesting that the glucose‐fatty acid cycle may not be responsible for IR [26]. Asan alternative for the glucose‐fatty acid cycle, Shulman proposed a mechanism in which intramyocellular lipids impair glucose uptake by inhibiting the insulin signaling cascade [3, 27]. The proposed mechanism involves the increase in intramyocellular lipid metabolites like diacylglycerol (DAG), long‐chain acyl‐CoA’s and ceramides, due to increased delivery of lipids to muscle and/or a decreased intracellular FA oxidation. These metabolites can activate protein kinase C , leading to the phosphorylation of serine/threonine sites on the insulin receptor substrate 1, which in turn interferes with the activation of phosphatidylinositol 3‐kinase, eventually resulting in a diminished glucose transport activity (Figure 4). Since then, several studies in humans, animals and in vitro preparations have confirmed steps of this mechanism [25, 28‐29]. Figure 3. Mechanism of inhibition of glucose utilization by enhanced fatty acid oxidation as described by Randle et al. [2]. An increase in fatty acid availability promotes fatty acid oxidation. This results in elevated acetyl‐CoA levels, which causes inhibition of PDH activity and reduced pyruvate oxidation. This leads to increased intramitochondrial and consequently intracellular citrate levels. Citrate in turn inhibits PFK‐1, leading to accumulation of G‐6‐P, which inhibits HK and thereby glucose uptake via GLUT4. LC‐FA, long‐chain fatty acid; FAT, fatty acid translocase; CD36, cluster of differentiation 36; LC‐acyl‐CoA, long‐chain acyl‐coenzyme A; ‐ox, ‐oxidation; TCA, tricarboxylicacid cycle; PDH, pyruvate dehydrogenase; PFK‐1, phosphofructokinase‐1; F‐6‐P, fructose‐6‐phosphate; G‐6‐P, glucose‐6‐phosphate; HK, hexokinase; GLUT4, glucose transporter type 4.
Furthermore, it was shown that IMCL, assessed by 1H MRS, is a better predictor for IR than plasma FFA, triglyceride and cholesterol levels, both in adults and in children [23, 30‐33]. Further proof for the relation between IR and IMCL came from Pima Indians in whom IMCL levels were inversely correlated with insulin action [22]. However, it has to be kept in mind that IMCL is probaby not directly affecting insulin sensitivity, but more likely represents the accumulation of lipid intermediates, e.g. DAG, ceramides and long‐chain acyl‐CoA’s, which can on their turn interfere with the insulin signaling cascade [34].
Although many studies strongly suggest that lipid accumulation within muscle tissue is associated with IR, paradoxical results have been presented [35‐37]. Not only within muscle of obese, diabetic subjects, but also within muscle of endurance‐trained, insulin‐sensitive athletes increased IMCL levels have been found. Therefore, the reported correlations between IMCL content and IR do not represent a direct, functional relationship. In addition to this finding, it has been shown that insulin sensitivity and IMCL content are dependent on muscle fiber type. IMCL levels are higher in insulin sensitive oxidative type 1 muscle fibers, compared to the less insulin sensitive, more glycolytic type 2 fibers [37‐39]. These findings together point out that increased lipid content within muscle does not always denote IR and that muscle lipid content should be evaluated within a context of other markers of metabolic capacity.
One such marker is the capacity for lipid oxidation [40], or the capacity for substrate oxidation in general [41]. Several human studies have provided evidence for dysfunctional muscle mitochondria in insulin‐resistant subjects by showing down‐regulation of genes encoding mitochondrial enzymes [42‐43], decreased mitochondrial content and lower mitochondrial respiratory chain activity [44]. Kelly et al. reported a 40% decrease in overall ETC activity (estimated using the activity of rotenone‐sensitive NADH:oxygen oxidoreductase) and smaller, deformed mitochondria in skeletal muscle from T2D patients compared to muscle of healthy volunteers [45]. Moreover, studies in which oxygen consumption was determined in freshly prepared isolated mitochondria or permeabilized muscle fibers showed decreased ADP‐stimulated respiration in diabetic patients compared to BMI‐matched controls [46‐47]. In vivo 31P MRS measurements were also applied to investigate the correlation between mitochondrial function and IR and/or T2D. Resting ATP synthesis flux has been measured by saturation transfer (ST) experiments by the group of Shulman [48‐50]. They showed decreased resting ATP synthesis fluxes in muscle of insulin‐ resistant elderly subjects as well as in insulin‐resistant offspring of type 2 diabetic parents as compared to healthy controls [48‐50]. Another 31P MRS technique to determine mitochondrial function is the measurement of PCr recovery after muscle exercise. A slower PCr recovery, indicating impaired mitochondrial function, was found in overweight diabetic patients as compared to healthy overweight controls [51]. It was hypothesized that inherited or acquired skeletal muscle mitochondrial dysfunction, associated with a reduced
mitochondrial capacity to oxidize FAs, leads to a lipid overload in muscle cells (Figure 4), inducing IR as was explained before.
However, recently more and more studies have been reported in which impaired mitochondrial dysfunction was not observed in insulin‐resistant subjects or patients with type 2 diabetes. PCr recovery rates of early and advanced stage diabetic patients matched for the level of physical activity were not significantly different [52]. Additionally, results from respiration measurements of permeabilized fibers of T2D patients compared to controls are in agreement with the previous results when normalized for mitochondrial DNA content or citrate synthase activity [53]. In other words, the type 2 diabetes patients showed normal intrinsic mitochondrial function, but an impaired oxidative capacity that was entirely attributed to a lower mitochondrial content. Environmental factors play an
Figure 4. Mechanism of inhibition of glucose utilization by enhanced fatty acid uptake and/or
decreased FA oxidation as proposed by Shulman et al. [3]. Increased LC‐FA supply in combination with decreased LC‐FA oxidation leads to an accumulation of lipids and lipid intermediates in muscle. These lipid intermediates (e.g. DAG and ceramides) activate PKC, leading to the phosphorylation of serine/threonine sites on the insulin receptor substrate 1, which in turn interferes with the activation of phosphatidylinositol 3‐kinase, eventually resulting in a diminished GLUT4 translocation to the cell membrane and consequently diminished glucose transport. IMCL, intramyocellular lipids; DAG, diacylglycerol; PKC, protein kinase C; IRS, insulin receptor substrate; tyr, tyrosine; ser, serine; thr, threonine; PI3K, phosphatidylinositol 3‐kinase (for other abbreviation see legend Figure 3).
important role in regulating skeletal muscle oxidative capacity, and the lower mitochondrial content in type 2 diabetes patients might simply be the result of a reduced habitual physical activity level [8, 53‐54]. Furthermore, mitochondrial dysfunction in type 2 diabetes might be secondary to impaired insulin signaling [55‐57] and/or abnormal blood glucose, insulin [58‐59] and FFA [57] levels. Therefore, the debate continues as to whether mitochondrial dysfunction represents either cause or consequence of IR and/or T2D.
Incomplete ‐oxidation
The hypothesis that mitochondrial dysfunction induces a decreased capacity to oxidize FAs, which leads to lipid overload and consequently IR, is losing impact. A recent study links IR to an increased capacity to oxidize FAs rather than the reverse [1] (Figure 5). However, the high rates of ‐oxidation in insulin‐resistant states are associated with low rates of complete fat oxidation [1, 60]. High rates of incomplete ‐oxidation occur when carbon flux through the ‐oxidation machinery outpaces the entry of acetyl‐CoA into the TCA cycle. Most of the evidence for the elevated incomplete ‐oxidation in insulin‐resistant states is based on the profiling of acylcarnitines in muscle, blood and urine. The vast majority of acylcarnitines is produced in the mitochondria and can therefore, in combination with measurements of substrate oxidation and mitochondrial function, be interpreted as a measure for (incomplete) ‐oxidation. A growing number of studies reported a negative correlation between circulating and/or tissue‐associated acylcarnitines and glucose tolerance in both animals [1,61] and humans [62‐65]. Several even chain, FA‐derived acylcarnitine intermediates were elevated in muscle of high‐fat diet‐fed rodents, but decreased after a 2‐wk exercise intervention that restored glucose tolerance [60]. Furthermore, in vitro measurement of [14C]‐oleate catabolism revealed disproportionally high rates of incomplete relative to complete fat oxidation in isolated muscle mitochondria from high‐fat diet‐fed compared to lean rodents [61]. From these studies it can be concluded that an excessive lipid load can lead to an increase in incomplete ‐oxidation, followed by the accumulation of lipid intermediates (e.g. acylcarnitines). The accumulation of lipid intermediates reflects a failed attempt to deal with the excess of intra‐ mitochondrial acyl‐CoA’s, causing mitochondrial ‘stress’ (Figure 5). The ‘stress’ is suggested to induce impairments in mitochondrial function and to activate stress kinases, interfering with insulin action [66], both leading to IR.
Current status
Although a lot of research has been performed to investigate the role of skeletal muscle mitochondrial dysfunction in the development of IR, still no consensus has been reached about the exact interplay between mitochondrial function, lipotoxicity and IR and/or T2D.
Aim of the thesis
The aim of this thesis was to study the timing and nature of muscle mitochondrial adaptations during the development of IR, by combining both in vivo and in vitro approaches, in order to gain more insight into the etiology of IR and T2D.
Figure 5. Mechanism of inhibition of glucose utilization by incomplete fatty acid oxidation as
described by Koves et al. [1]. High FA availability increases ‐oxidation, resulting in carbon fluxes through the ‐oxidation machinery which outpace the entry of acetyl‐CoA into the TCA cycle. This results in the accumulation of incompletely oxidized lipid intermediates, which can interfere via PKC with insulin signaling and glucose uptake. CAT, carnitine acyltransferase; CACT, carnitine‐ acylcarnitine translocase; the flash represents the generation of reactive oxygen species (for other abbreviation see legend Figures 3 and 4).
Measurement of muscle lipids and lipid intermediates
In this thesis, the relation between IR and muscle lipid overload was studied by the measurement of IMCL by in vivo 1H MRS and muscle acylcarnitines by in vitro tandem mass spectrometry (MS/MS). These measurement procedures are explained in this paragraph.
IMCL
As was explained in one of the previous paragraphs, the amount of lipids inside the muscle cell, i.e. intramyocellular lipids (IMCL), are strongly correlated with the degree of IR [22]. IMCL is mainly present as liquid droplets in the cytosol of muscle cells. The lipids which are present as subcutaneous or interstitial adipose tissue are referred to as extramyocellular lipids, or EMCL. While EMCL is metabolically relatively inert, there is substantial evidence that IMCL within droplets can be rapidly mobilized and utilized for energy metabolism, particularly as they are primarily located immediately adjacent to mitochondria.
IMCL levels have been determined using a variety of techniques including biochemical extraction [22, 67], Oil red O histochemical staining [36, 68] and electron microscopy morphometry [69] of needle biopsy samples and magnetic resonance imaging (MRI) and computed tomography (CT) [70]. Disadvantages of these techniques are that they are invasive or unable to differentiate between IMCL and EMCL or both. Localized 1H MRS has proven to be a valuable tool for determining IMCL, because 1H MRS is uniquely capable of separately detecting IMCL and EMCL non‐invasively and without the use of any harmful radiation [71]. The IMCL/EMCL peak separation depends on the orientation of the muscle with respect to the magnetic field and amounts to a maximum of circa 0.2 ppm when the muscle is parallel to the magnetic field. In this case the CH2 protons of IMCL and EMCL
resonate around 1.28 and 1.47 ppm, respectively (Figure 6). This chemical shift difference has been shown to originate from bulk magnetic susceptibility (BMS) effects, that are due to the layered ordering of EMCL depots along the main muscle axis, while IMCL is organized in spherical droplets in the cytosol of the muscle cell [71].
Figure 6. Localized 1H MR spectrum measured in tibialis anterior muscle from a Wistar rat fed with a high‐fat diet for 2.5 weeks, showing peaks of total creatine (tCr), extramyocellular lipids (EMCL) and intramyocellular lipids (IMCL).
Thus, 1H MRS based IMCL measurements are very useful for the determination of IMCL levels in longitudinal studies in both humans and animal models during the development of IR and T2D.
Acylcarnitines
The first step for the use of FAs as a substrate for ATP production in the mitochondria is their transport into the mitochondria. A step required for transport across the mitochondrial membrane is the formation of acylcarnitine esters from long‐chain acyl‐ CoA’s by CPT1. Besides fulfilling an important role in mitochondrial import of FAs, the binding to carnitine is also essential for the efflux of excess intra‐mitochondrial acyl‐CoA into the cytosol, and from the cytosol into the bloodstream. Acylcarnitines represent byproducts of substrate catabolism and are formed from their respective acyl‐CoA intermediates by a family of carnitine acyltransferases that reside principally in the mitochondria. Most even chain species reflect incomplete FA oxidation, odd chain species stem primarily from amino acid catabolism, whereas acetylcarnitine is derived from acetyl‐ CoA, the universal degradation product of all metabolic substrates.
Blood acylcarnitine profile analysis is the current standard for the diagnosis of ‐oxidation disorders at the metabolite level [72‐73]. Furthermore, recently muscle acylcarnitine accumulation was related to IR [1, 66, 74]. The acylcarnitine accumulation as was observed in insulin‐resistant skeletal muscle was suggested to reflect a failed attempt to deal with the excess of intra‐mitochondrial acyl‐CoA’s, causing mitochondrial stress. The accumulation of metabolic by‐products (e.g. acylcarnitines) would then activate stress kinases or other signals, interfering with insulin action [66]. However, how and if acylcarnitines directly affect insulin‐mediated glucose uptake is currently not known. The finding that carnitine supplementation improved glucose tolerance while increasing circulating acylcarnitines and leaving muscle acylcarnitines unaffected favors the interpretation that production and efflux of these metabolites is beneficial rather than detrimental. However, the exact physiological relevance of changes in muscle, blood and urine acylcarnitine levels in IR and as a result of carnitine supplementation is still under debate.
By means of tandem MS/MS it is possible to analyze 36 independent acylcarnitine species ranging in size from 2 to 22 carbons in muscle, blood and urine [75]. Important for analyzing acylcarnitine levels is to consider that steady‐state acylcarnitine concentrations in tissues and blood represent the net balance between production, consumption, import and export, and therefore do not directly provide information about fluxes through individual metabolic pathways.
Measurement of muscle mitochondrial function
Data to support the proposed role of skeletal muscle mitochondrial dysfunction in the development of IR and T2D have been obtained with various in vitro methods, including measurements of oxidative enzyme activities [39, 44, 76‐78], mRNA and/or protein expression of oxidative phosphorylation genes [42‐43, 78‐80] as well as mitochondrial content, morphology and respiration [44, 47, 77, 80]. Furthermore, in vivo magnetic resonance spectroscopy (MRS) measurements of basal mitochondrial ATP synthesis flux [48‐49, 81] and PCr [11] and ADP [82] recovery kinetics also point towards a potential role for mitochondrial dysfunction in the etiology of insulin resistance and/or type 2 diabetes. In this thesis in vitro high‐resolution respirometry and in vivo 31P MRS play a dominant role and therefore these techniques are explained in more detail below.
High‐resolution respirometry
Levels of dissolved oxygen in solution can be measured polarographically with a Clark‐type oxygen electrode. Clark electrodes have gold or platinum cathodes and silver or silver/silver chloride anodes, which are connected by a salt bridge and covered by an oxygen‐ permeable membrane. As oxygen diffuses across the membrane, it is reduced by a fixed voltage between the cathode and anode which generates current in proportion to the concentration of oxygen in solution. By calibrating the oxygen electrode with known oxygen concentrations, it is possible to measure the rate of oxygen consumption in a
Figure 7. Typical example of time‐dependent changes in oxygen concentration during a high‐resolution
respiration measurement on a suspension of isolated mitochondria, isolated from tibialis anterior muscle of a Wistar rat. The dark line represents the oxygen concentration in the respiratory chamber, the lighter line represents the slope of the dark line. First, isolated mitochondria were injected in the hermetically closed respiratory chambers (mit). After some time, glucose (gl), hexokinase (hk) and ATP were added, inducing maximal respiration or state 3 respiration. When the oxygen consumption rate reached a steady state, state 4 was induced by adding carboxyatractyloside. Finally, after addition of carbonylcyanide‐3‐chlorophenylhydrazone (CCCP), the maximal oxygen consumption rate in the uncoupled state (state U) was determined.
medium containing actively respiring mitochondria. Since reduction of oxygen is a critical step in the process of mitochondrial electron transport and ATP synthesis, measurement of mitochondrial oxygen consumption provides a convenient way to assess mitochondrial function [83]. Oxygen consumption can be measured in permeabilized muscle fibers and isolated mitochondria. The latter approach has been used in the studies described in this thesis.
Measuring respiration in isolated mitochondria practically means that together with a respiration medium, an aliquot of suspension with isolated mitochondria is added to the closed metabolic chamber. The mitochondria are brought into defined “states” by the sequential addition of substrates or inhibitors. Since the mitochondria consume oxygen, the oxygen concentration drops. This change of oxygen concentration is recorded by the oxygen sensor in the chamber. From the rate of decline in the oxygen concentration (taking into account correction for oxygen diffusion) the respiration rate can be computed for the mitochondria in different states (see Figure 7). State 3, state 4 and state uncoupled (state U) are frequently used to define mitochondrial respiration. State 3 respiration is the maximal respiration, reached when saturating levels of ADP are added to the mitochondria supplemented with oxidizible substrate and excess Pi. State 4 respiration is induced by
addition of carboxyatractyloside, which inhibits the exchange of mitochondrial ATP for extra‐mitochondrial ADP by adenine nucleotide translocase and this way effectively blocks ATP synthesis. The residual oxygen consumption in the absence of ADP phosphorylation is attributable to proton leak across the inner mitochondrial membrane. Thus, carboxyatractyloside inhibited respiration serves as an indicator of the degree of uncoupled respiration or proton leak under these conditions. The respiratory control ratio (RCR), which is calculated by dividing state 3 by state 4 respiration, indicates the tightness of the coupling between respiration and ADP phosphorylation. Stepwise titration of the protonophore carbonylcyanide‐3‐chlorophenylhydrazone (CCCP) induces an uncoupled state by dissipating the proton gradient across the inner mitochondrial membrane (state U). Under these conditions, mitochondrial oxidative capacity can be determined in the absence of the potential control exerted by ATP synthase, adenine nucleotide translocase, or phosphate transporters.
By using different combinations of oxidizible substrates it is possible to quantify specific substrate oxidation capacity in isolated mitochondria. Oxidation of TCA cycle intermediates such as pyruvate plus malate leads to formation of NADH, which can donate electrons to ETC complex I. In turn oxidation of another TCA cycle intermediate succinate leads to formation of FADH2, which bypasses complex I and donates electrons to complex II. By
using fatty acid substrates such as palmitoyl‐L‐carnitine or palmitoyl‐CoA plus L‐carnitine in combination with malate one can assess the capacity of ‐oxidation. The measured oxygen consumption rate is expressed per milligram mitochondrial protein and is used as a measure for the intrinsic mitochondrial function.
In vivo
31P MRS
31
P MRS offers a non‐invasive approach of recording concentrations of phosphorylated metabolites and intracellular pH. 31P MRS has widely been used to study skeletal muscle metabolism in living mammalian tissues, and is also one of the major types of measurement methods used in this thesis. 31P MR spectra of skeletal muscle typically show five major resonances: an inorganic phosphate (Pi) peak, a phosphocreatine (PCr) peak and
three ATP peaks, from the , and phosphate groups of ATP (Figure 8). In some cases also peaks from phosphomonoesters (PME) and phosphodiesters (PDE) are visible. Additionally, metabolic information can be derived indirectly from the 31P MR spectra, i.e. the intracellular pH and the free concentration of ADP. Tissue pH can be deduced from the chemical shift of the Pi peak [84], which actually originates from both H2PO4
‐
and HPO4 2‐
. However, as these compounds are in rapid chemical exchange, a single peak is observed, with a chemical shift dependent on the ratio of the H2PO4‐ and HPO42‐ concentrations,
which in this way serves as an indicator of intracellular pH. Free levels of ADP in the cell can be calculated indirectly by use of the CK equilibrium (Eq. 1) [85]). The concentration of ADP in healthy, resting or moderately active muscle is typically in the tens of micromolar range, which is too low to allow direct detection with 31P MRS. The knowledge of ADP levels is relevant, because ADP is an important regulator of the mitochondrial ATP synthesis flux. 31P MR spectra of resting skeletal muscle are relatively constant, even in diseased states, and to assess impairments in mitochondrial energy production one needs to perturb either the chemical or the magnetic equilibrium. Most important 31P MR techniques in studies on the role of mitochondrial function in IR and T2D, including those in this thesis, are saturation transfer (ST) MRS, to measure (resting) creatine kinase and ATP synthesis fluxes, and the dynamic measurement of post‐exercise recovery of PCr.
Figure 8. 31P MR spectrum measured in resting tibialis anterior muscle of a Wistar rat, showing the peaks of inorganic phosphate (Pi), phosphocreatine
(PCr) and the three (, and ) phosphate groups in ATP.
Saturation transfer
Magnetization transfer experiments allow to determine fluxes between compounds that are in chemical exchange, as long as the system is in a metabolic steady state. One variant of the magnetization transfer technique relies on the selective saturation of the magnetization of one of the reactants with a long, frequency‐selective RF pulse, i.e. a saturation transfer (ST) experiment. Due to the chemical exchange of the reactants, this results in a decrease of magnetization of the exchange partner. With equation 3 the exchange rate constant kAB between reactants A and can be determined. 1 , , Eq. 3
M’A and M0A are the magnitudes of magnetization of compound A, when compound B is
selectively saturated and when compound B is not saturated, respectively. The apparent longitudinal relaxation time of compound A (T1A’) can be determined from an inversion
recovery experiment with saturation of B prior to and during the inversion time. To calculate the flux from compound A to B, kAB has to be multiplied by the concentration of compound A. One of the chemical exchange processes most‐studied using ST, concerns the flux through the creatine kinase (CK) reaction [86]. CK catalyzes a phosphate exchange reaction, in which a phosphate moiety is exchanged between the position of ATP to PCr and vice versa. As was described earlier in this chapter, the CK reaction is of high metabolic relevance, since PCr is an important energy buffer for keeping ATP levels constant. The CK flux can be determined by complete saturation of the ‐ATP resonance and the quantification of the reduction in the signal intensity of PCr. Measurements of CK flux in skeletal muscle as a function of workload have shown that CK kinetics is rather insensitive to alterations in ATP demand and that CK flux greatly exceeds maximal ATP turnover rates [87‐88].
31
P ST MRS has also been used to measure the mitochondrial ATP synthesis flux in rodent and human muscle, under resting conditions [48‐50, 89‐92]. Instead of the signal intensity of PCr, the magnetization of Pi has to be quantified to calculate the ATP synthesis flux.
Assuming that the muscle ATP synthesis flux, as measured by 31P ST MRS, is predominantly reflecting oxidative ATP synthesis by the F1F0‐ATP synthase in the mitochondria, muscle ATP
synthesis flux has been taken as a measure for mitochondrial function. Studies have shown
decreased resting ATP synthesis fluxes in skeletal muscle of insulin‐resistant subjects and insulin‐resistant animal models.
However, the interpretation of 31P ST MRS data, particularly at rest, is not straightforward. The lower ATP synthesis flux in resting muscle of insulin‐resistant subjects could actually reflect a normal regulatory response to a lower energy demand, e.g. caused by impaired insulin signaling, rather than an impairment of intrinsic mitochondrial function [55‐56, 93]. In order to interpret the ST data in terms of mitochondrial function it is necessary to take the error signals, i.e. the concentrations of ADP and Pi, into account. Moreover, the Pi
ATP fluxes obtained from 31P ST measurements are comprised of both mitochondrial F1F0‐
ATP synthesis flux and flux through other Pi ATP pathways, in particular the reactions
catalyzed by the glycolytic enzymes glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) and phosphoglycerate kinase (PGK) [88, 92, 94‐95]. Although the net glycolytic contribution to the production of ATP (via GAPDH and PGK) versus that of oxidative phosphorylation is small [96], these enzymes catalyze a coupled near‐equilibrium reaction, and consequently, exchange between Pi and ATP may greatly exceed the net glycolytic flux [92]. The
interpretation of resting ATP synthesis flux as a measure for mitochondrial function is therefore relatively complicated. In order to create better conditions for detecting a defect in mitochondrial oxidative phosphorylation, the ST experiment would need to be done in exercising muscle [88].
Post‐exercise PCr recovery
31P MRS provides the opportunity of acquiring data during skeletal muscle exercise and
recovery in a non‐invasive manner and with a time resolution of seconds. Figure 9 shows the concentrations of PCr, Pi and ATP derived from a time series of 31P MR spectra acquired
during rest, exercise and subsequent recovery in rat tibialis anterior (TA) muscle. When muscle contraction starts, an immediate decrease in PCr concentration and increase in Pi
concentration can be observed (Figure 9), whereas ATP remains constant, although ATP is used for the muscle contractions. After the muscle contractions stop, the PCr buffer is restored and Pi levels normalize. The ATP used for resynthesis of PCr is originating
predominantly from oxidative phosphorylation in the mitochondria [97‐98]. Because the CK reaction is much faster than the ATP synthase reaction [87‐88], the rate of PCr recovery mirrors the rate of oxidative ATP synthesis and therefore the oxidative capacity of the mitochondria. A short PCr recovery time constant, corresponding to a fast recovery, reflects a high oxidative capacity, while slow recovery kinetics may indicate impaired oxidative capacity. PCr recovery is determined by mitochondrial content, intrinsic mitochondrial function and other factors like the supply of oxygen and substrates [99‐100]. This implies that additional (in vitro) measurements are necessary to identify the origin of changes in oxidative capacity measured with PCr recovery.
One of the first who proved that PCr recovery kinetics provides information about oxidative ATP synthesis was Mahler et al., who reported that post‐exercise PCr resynthesis is inversely proportional to the rate of oxygen consumption [101]. This finding was
strengthened by the observation that the PCr concentration does not restore during ischemic recovery [97‐98]. Furthermore, the correlations of post‐exercise PCr recovery kinetics with citrate synthase activity [102], succinate dehydrogenase activity and peak oxygen consumption (VO2peak) [103] are additional findings demonstrating that post‐
exercise PCr resynthesis changes reflects aerobic metabolism.
An important complication in interpreting post‐exercise PCr recovery data is that strenuous exercise‐induced acidosis negatively affects PCr resynthesis [104‐114]. Therefore it has been recommended to avoid low end‐exercise pH values, which is difficult to achieve in combination with a significant PCr depletion. Other strategies to deal with the acidification problem is to aim at similar end‐exercise pH values within different groups or applying general correction factors for the influence of pH [106, 109, 111].
Several MR compatible exercise set‐ups have been built for humans, most often for voluntary arm or leg exercise. In anaesthetized animals, muscle contractions are usually induced by electrical stimulation. As an alternative to the invasive, direct nerve‐stimulation method, a minimally invasive electrical stimulation method was developed that allows longitudinal studies in rats [115]. By subcutaneously implanting electrodes along the nerve trajectory of the N. Peroneus Communis, highly specific dorsal flexor muscle contractions can be induced. This procedure was also used in the present thesis. Figure 9. Peak concentrations of phosphocreatine (black), inorganic phosphate (light gray) and ATP (dark gray), as measured using 31P MRS during a 3‐min rest, 2‐min electrical stimulation and 10‐min recovery protocol in rat tibialis anterior muscle. Time resolution was 20 seconds.
Outline of this thesis
Although a lot of research has been performed to investigate the role of skeletal muscle mitochondrial function and lipid accumulation in the development of IR, still no consensus is reached about the exact mechanism(s) leading to IR. The aim of this thesis was to study the timing and nature of muscle mitochondrial adaptations during the development of IR, by using both in vivo and in vitro measurement techniques. 31P MRS plays an important role in these studies. The first part of this thesis deals with methodological aspects of the 31P
MRS techniques, whereas the second part describes research on mitochondrial adaptations in both patients with T2D and animal models of IR.
Both 31P MRS PCr recovery after muscle contraction and resting ATP synthesis flux have been used to determine mitochondrial function in vivo. However these methods have provided ambiguous results. Therefore, PCr recovery and resting ATP synthesis flux were compared in a rat model of known mitochondrial dysfunction in order to establish which method is most appropriate to assess in vivo skeletal muscle mitochondrial function in
chapter 2. Mitochondrial dysfunction was induced in rats by daily subcutaneous injections
with diphenyleneiodonium (DPI), which irreversibly inhibits complex I (NADH‐ubiquinone reductase) of the respiratory chain. 31P MRS PCr recovery after exercise is applied in every chapter of this thesis as a measure for in vivo muscle oxidative capacity. However, cytosolic pH has a strong influence on the kinetics of PCr recovery and thereby complicates the interpretation of PCr recovery data. It has been suggested that PCr recovery should be normalized for end‐exercise pH. However a general correction can only be applied if there are no intersubject differences in the pH dependence of PCr recovery. In chapter 3 we investigated the pH dependence of PCr recovery on a subject‐by‐subject basis in vastus lateralis muscle of healthy humans. Furthermore, we determined the relation between the pH dependence of PCr recovery and the kinetics of proton efflux at the start of recovery. To get more insight in the role of muscle mitochondrial dysfunction and lipotoxicity in the etiology of IR and T2D, we performed three studies in which we determined muscle lipid content and mitochondrial function. In a cross‐sectional study, described in chapter 4, we examined in vivo skeletal muscle mitochondrial function in early and advanced stages of T2D in human subjects. Long‐standing, insulin‐treated type 2 diabetes patients, subjects with impaired fasting glucose, impaired glucose tolerance and/or recently diagnosed type 2 diabetes, and healthy, normoglycaemic controls, matched for age and body composition and with low habitual physical activity levels were studied. In vivo mitochondrial function of the vastus lateralis muscle was evaluated from post‐exercise PCr recovery kinetics using 31P MRS. IMCL content was assessed in the same muscle using single‐voxel 1H MRS. Chapter 5 describes a longitudinal study with the aim to gain more insight into the timing and nature of mitochondrial adaptations during the development of high‐fat diet‐induced IR in a well known rodent model of IR. Adult Wistar rats were fed a high‐fat diet or normal chow for 2.5 and 25 wk. IMCL was measured with in vivo 1H MRS and acylcarnitine levels were quantified
in vitro using tandem MS/MS. Muscle oxidative capacity was assessed in TA muscle in vivo
using 31P MRS PCr recovery and in vitro by measuring mitochondrial DNA copy number and oxygen consumption in isolated mitochondria. Currently, it is not clear whether mitochondrial dysfunction is (1) a cause of lipid accumulation due to a decreased capacity to oxidize FAs or (2) a consequence of the accumulation of lipid intermediates as a result of increased incomplete FA oxidation. Carnitine is known to stimulate FA oxidation and export. The aim of the study described in chapter 6 was to test the hypothesis that carnitine supplementation reduces high‐fat diet‐induced lipotoxicity, improves in vivo muscle mitochondrial function and ameliorates insulin resistance. Wistar rats were fed either normal chow or a high‐fat diet for 15 wk and one group of high‐fat diet fed rats was supplemented with L‐carnitine during the last 8 wk. Muscle mitochondrial function was measured in vivo by 31P MRS (PCr recovery and resting ATP synthase and creatine kinase fluxes) and in vitro by high‐resolution respirometry. Muscle lipid levels were determined by
1H MRS (IMCL) and tandem mass spectrometry (acylcarnitines).
Chapter 7 provides a summary with the main outcomes of the studies described in this
thesis. The results are compared to the current hypotheses and future perspectives for research in the field of muscle mitochondrial function in relation to the development of IR are discussed.
