Mitochondrial function as a target for therapy in type 2 diabetes Wessels, B.

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1 Mitochondrial function as a target for therapy in type 2 diabetes Wessels, B. Published: 01/01/2015 Document Version Publisher s PDF, also known as Version of Record (includes final page, issue and volume numbers) Please check the document version of this publication: A submitted manuscript is the author's version of the article upon submission and before peer-review. There can be important differences between the submitted version and the official published version of record. People interested in the research are advised to contact the author for the final version of the publication, or visit the DOI to the publisher's website. The final author version and the galley proof are versions of the publication after peer review. The final published version features the final layout of the paper including the volume, issue and page numbers. Link to publication Citation for published version (APA): Wessels, B. (2015). Mitochondrial function as a target for therapy in type 2 diabetes Eindhoven: Technische Universiteit Eindhoven General rights Copyright and moral rights for the publications made accessible in the public portal are retained by the authors and/or other copyright owners and it is a condition of accessing publications that users recognise and abide by the legal requirements associated with these rights. Users may download and print one copy of any publication from the public portal for the purpose of private study or research. You may not further distribute the material or use it for any profit-making activity or commercial gain You may freely distribute the URL identifying the publication in the public portal. Take down policy If you believe that this document breaches copyright please contact us (openaccess@tue.nl) providing details. We will immediately remove access to the work pending the investigation of your claim. Download date: 26. Jan. 2019

2 Mitochondrial function as a target for therapy in type 2 diabetes

3 The project described in this thesis was funded by a VIDI grant from the Netherlands Organisation for Scientific Research (NWO), project number A catalogue record is available from the Eindhoven University of Technology Library ISBN: Cover art: Odra Noel Printed by: Ipskamp Drukkers, the Netherlands Copyright 2014 by Bart Wessels All rights reserved. No part of this book may be reproduced, stored in a database or retrieval system, or published, in any form or any way, electronically, mechanically, by print photo print, microfilm, or any other means without prior permission by the author.

4 Mitochondrial function as a target for therapy in type 2 diabetes 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 maandag 26 januari 2015 om uur door Bart Wessels geboren te Eindhoven

5 Dit proefschrift is goedgekeurd door de promotor en de samenstelling van de promotiecommissie is als volgt: Voorzitter: Promotor: Co-promotor: Leden: prof.dr. P.A.J. Hilbers prof.dr. K. Nicolay dr. J.J. Prompers dr. P. Carlier (Institut de Myologie, Paris) prof.dr. C.J.J. Tack (Radboud universiteit Nijmegen) prof.dr. R.J.A. Wanders (AMC-UvA) dr. G.J. Strijkers

6 Table of contents Chapter 1 6 General Introduction Chapter 2 46 O 2 availability does not limit in vivo oxidative capacity in skeletal muscle of healthy and diabetic rats as assessed with 31 P MRS under normoxic conditions Chapter 3 68 Carnitine supplementation in high-fat diet fed rats does not ameliorate lipidinduced skeletal muscle mitochondrial dysfunction in vivo Chapter 4 92 Pioglitazone treatment restores in vivo muscle oxidative capacity in a rat model of diabetes Chapter Metformin impairs mitochondrial function in skeletal muscle of both lean and diabetic rats in a dose-dependent manner Chapter Metformin treatment impairs in vivo skeletal muscle oxidative capacity as well as contractile function in diabetic rats Chapter Summarizing Discussion Summary 155 Dankwoord 162 List of Publications 166 About the Author 168

7 General Introduction Part of the methodological approaches was published as a review article in the International Journal of Biochemistry & Cell Biology (2014) Prompers, J.J., Wessels, B., Kemp, G.J. & Nicolay, K. MITOCHONDRIA: investigation of in vivo muscle mitochondrial function by 31P magnetic resonance spectroscopy. doi: /j.biocel Epub 2014 Feb 22.

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9 Chapter 1 8

10 General Introduction The prevalence of type 2 diabetes (T2D) has grown to epidemic proportions worldwide and is still on the rise in every country 1. The International Diabetes Federation annually releases estimates of the global number of diabetes patients to promote global awareness of the disease. An estimated number of 285 million adults were affected by diabetes in 2010, which increased to 371 million patients only two years later in Based on this rapid rise, the predicted number of diabetes patients for the year 2030 was adjusted from 439 million to 552 million 1,2. The increase in the prevalence of T2D is widely regarded to be linked with the increasing prevalence of obesity, which in turn is associated with the consumption of high-calorie diets and a sedentary lifestyle 1. 1 Patients with T2D are not able to meet the demand for insulin-stimulated glucose disposal, because their major metabolic tissues are less sensitive to insulin and insulin release by the pancreas is reduced. The primary site of glucose metabolism is the skeletal muscle, where 70-80% of the total postprandial glucose disposal occurs. One of the leading hypotheses in the research field of type 2 diabetes is that lipid overload in muscle cells, as marked by the accumulation of intramyocellular lipids (IMCL), leads to impaired insulin signaling 3-9. IMCL probably does not affect insulin resistance directly, but the mechanistic link between IMCL accumulation and insulin resistance is believed to reside in the accumulation of lipid derived intermediates, such as diacylglycerols and ceramides, that lead to impaired insulin signaling 8,10,11. Because of the prominent function of mitochondria in whole-body energy metabolism, these organelles have been implicated to play a pathological role in insulin resistance 12. Mitochondrial dysfunction and a resulting impairment in fatty acid (FA) oxidation have been suggested as causative factors in the excess storage of lipids in insulin-resistant muscle 8,9. However, recent studies have linked insulin resistance to an increased rather than a decreased capacity to oxidize FAs This increased capacity for FA oxidation was shown to be associated with the accumulation of metabolic intermediates of incomplete FA oxidation 17,18, indicating a mismatch between energy supply and demand which may promote mitochondrial oxidative stress. Therefore, although a vicious cycle between lipid accumulation and derangements in mitochondrial metabolism seems to underlie the progression of muscle insulin resistance and T2D, the cause-and-effect relationship between lipid accretion and mitochondrial dysfunction is not yet resolved 19,20. A better understanding of the interaction between lipid accumulation and functioning of mitochondria is therefore imperative in order to make advances in the development of new effective treatment strategies for T2D. 9

11 Chapter 1 Background of pathophysiology and treatment of type 2 diabetes Disturbed glucose homeostasis in type 2 diabetes In a healthy situation, the balance of glucose uptake, oxidation, and storage is tightly controlled in all metabolic tissues and regulated by levels of insulin (Figure 1). After a meal, high levels of blood glucose stimulate the pancreatic β-cells to release insulin, which promotes glucose uptake in skeletal muscle while inhibiting glucose release from the liver. Insulin also plays a central regulatory role in ensuring that excess glucose, which is not immediately utilized, is properly stored. This glucose spill-over is stored in liver and skeletal muscle as glycogen (glyconeogenesis) or, when glycogen storage is exceeded, in adipose tissue where excess glucose is converted into triglycerides (de novo lipogenesis). Food intake liver glycogen glucose digestive sytem glycogen skeletal muscle insulin TG FFA FFA pancreas adipose tissue Figure 1. Regulation of plasma glucose levels by insulin After a meal there is an abundance of carbohydrates which promotes pancreatic β-cells to release insulin. Insulin drives the uptake and conversion of glucose to glycogen in both muscle and the liver, while restricting glucose release from the liver. The uptake of glucose occurs primarily in skeletal muscle. Reversely, dependent on whole-body glucose availability and demand, glucose can again be mobilized from glycogen storage. Glucose taken up in adipose tissue is converted to and stored as triglycerides (TG) and released from adipocytes as free fatty acids (FFA) to be used as an energy source when fuels are scarce. 10

12 General Introduction The insulin signaling pathway in skeletal muscle is activated when insulin binds to the tyrosine kinase receptor (a transmembrane protein), which result in autophosphorylation of specific tyrosine residues of the insulin receptor in the cytoplasmic domain. The catalytic activity of the insulin receptor then creates bindings sites for insulin receptor substrate (IRS) proteins such as IRS-1 11,21. After phosphorylating of IRS-1 tyrosine residues, the IRS proteins then activate phosphatidylinositol 3-kinase (PI3-K), which converts phosphatidylinositol 4,5-biphosphate (PIP 2 ) to phosphatidylinositol 3,4,5-triphosphate (PIP 3 ). PIP 3 activates phosphatidylinositol-dependent kinase (PDK), which then activates protein kinase B (PKB or Akt). Akt targets cytoplasmic vesicles carrying glucose transporter 4 (GLUT-4), which translocate to and fuse with the cell membrane where GLUT-4 facilitates transport of glucose into the skeletal muscle cell (Figure 2). 1 Prior to developing overt type 2 diabetes, high-risk individuals for developing this disease live in a state where major metabolic tissues, such as liver, skeletal muscle and the heart, have turned less sensitive to insulin, a state commonly referred to as insulin resistance. In order to compensate for this loss in insulin sensitivity the pancreas produces more insulin, resulting in a rise of blood insulin levels. The human body can cope with insulin resistance in this manner for years, before the β-cells in the pancreatic islets irrevocably start to fail. At this point insulin levels will drop and the insulinresistant metabolic tissues will not be able to properly respond to rises in blood glucose levels. This leads to an increase in the duration of postprandial blood glucose clearance and elevated levels of blood glucose also in the fasted state, which is a condition referred to as type 2 diabetes. Figure 2. Ge transport activity at the cellular membrane is l Figure 2. GLUT-4 mediated glucose transport into the cell The vesicles containing glucose transporter 4 (GLUT-4) are located within the intracellular compartment during the pre-prandial state; therefore glucose transport activity at the cellular membrane is low. During the post-prandial state blood glucose levels rise with a subsequent increase in blood insulin levels. By binding to the insulin receptor, insulin creates binding sites for insulin receptor substrate 1 (IRS-1), on the cytoplasmic side of the cellular membrane. Next, IRS-1 is phosphorylated, resulting in the binding and activation of phosphatidylinositol 3-kinase (PI3-K). PIP3 activates phosphatidylinositol-dependent kinase (PDK), which then activates protein kinase B (Akt), which mediates the translocation of vesicles containing GLUT-4 to the cell membrane. GLUT-4 then transports glucose molecules across the cell membrane. 11

13 Chapter 1 Link between insulin resistance and lipid accumulation in skeletal muscle Insulin resistance has reportedly been associated with elevated levels of plasma free fatty acids (FFA) and triglycerides (TG) 22,23. However, the inhibition of insulin action was shown to be delayed until 2-4 hours after an acute increase of the plasma FFA concentration, which suggests that the insulin desensitizing effect is not due to a direct effect of circulating FFA 24. Furthermore, the delay in the insulin desensitizing effect indicates that lipids need to accumulate in metabolic tissues first, before they are able to interfere with insulin signaling. A prolonged disturbance in the balance between energy uptake and expenditure, e.g. due to excessive caloric intake in obesity, causes lipid spill-over which provokes a rise in plasma FFA and TG that will end up in metabolic tissues, like the liver, heart and skeletal muscle. The association between intramyocellular lipid (IMCL) accumulation and peripheral insulin resistance is currently well established 3-7. Moreover, insulin resistance resulting from ageing was linked to accumulation of IMCL 25, while low-calorie diets in patients with type 2 diabetes improved insulin sensitivity paralleled with a reduction in IMCL 20,26. These studies all underline the importance of IMCL content in relation to insulin resistance. It is clear though, that IMCL content alone is insufficient to predict insulin resistance, since endurance-trained athletes have IMCL levels close to the reported values for obese and diabetic individuals 7,27,28. These athletes were found to be highly sensitive to insulin, which is why this phenomenon is referred to as the athlete paradox. This led Moro et al. to propose a U-shaped relationship to describe the association between IMCL and insulin sensitivity, with obese, sedentary T2D patients on one end, and lean, endurance-trained athletes on the other end of this U-curve (Figure 3) 28. The athlete paradox may be explained by a higher IMCL turnover within athletes with respect to sedentary T2D patients. The turnover of IMCL is a measure of the dynamic balance between muscle lipolysis and lipid synthesis, which are dependent on mitochondrial fatty acid (FA) oxidation and the availability of plasma FFA. This means that athletes are able to quickly hydrolyze IMCL for utilization, or esterify fatty acids into the IMCL pool by the reverse process when they are not utilized. In patients with T2D however, high levels of IMCL are associated with elevated levels of lipid intermediates, such as diacylglycerol (DAG), acyl-coas and ceramides 11,29,30. Inactivity High calorie diets Exercise Low calorie diet IMCL content obese T2D Low lean Oxidative capacity Insulin sensitivity High athletes Figure 3. Association between IMCL and insulin sensitivity U-curve as a model to describe the association between IMCL content and insulin sensitivity, which is believed to be modulated by oxidative capacity. The effect of lifestyle interventions like exercise therapy and/or low-calorie diets on the IMCL-insulin sensitivity relationship are displayed by the arrows. Modified from Moro et al

14 GLUT-4 General Introduction Long-chain acyl-coas (LC acyl-coas) and DAG can interfere with insulin-stimulated GLUT- 4 translocation by activating novel protein kinases (npkc), which subsequently phosphorylate serine and threonine residues of the insulin receptor and IRS-1, thereby preventing tyrosine phosphorylation. This leads to a dissociation between the insulin receptor and IRS-1 and possibly between IRS-1 and PI3-K, lowering IRS-1 stimulated PI3-K activity 10,11,31. Ceramides are thought to affect GLUT-4 translocation further downstream through the inhibition of Akt phosphorylation. In addition, elevated FA oxidation in the mitochondria raises the production of reactive oxygen species (ROS), which stimulate serine kinases, such as npkc, thereby lowering insulin sensitivity 19,20. The consequence of this cascade of events is that these tissues require higher levels of insulin to reach the same amount of GLUT-4 translocation for glucose transport into the cell, thus leading to insulin resistance (Figure 4). 1 insulin IRS-1 npkc TG LC acyl-coa CD36 FFA DAG Akt Ceramides GLUT-4 ROS glucose G6P glycogen pyruvate Mitochondrion Figure 4. Mechanism of lipid-induced muscle insulin resistance Free fatty acids (FFA) enter the cell via fatty acid protein transporters on the cell surface, primarily via fatty acid translocase (FAT/CD36). Excess supply of FFA lead to elevated levels of lipid intermediates such as diacyglycerol (DAG) and ceramides. DAG can activate serine and threonine kinases such as the novel protein kinases (npkc) that phosphorylate the insulin receptor and insulin receptor substrate 1 (IRS-1). Serine phosphorylation instead of normal insulin-stimulated tyrosine phosphorylation is thought to dissociate IRS-1 from the insulin receptor. Ceramides can inhibit protein kinase B (Akt) which inhibits glucose transporter 4 (GLUT-4) translocation further downstream of the insulin receptor. In addition, a rise of fatty acid oxidation in the mitochondria increases the production of reactive oxygen species (ROS), which stimulate serine kinases, such as npkc, thereby lowering insulin sensitivity. 13

15 Chapter 1 Lipotoxicity and mitochondrial dysfunction Conventionally, mitochondria are described as the powerhouses of the cell, since they are the main providers of adenosine triphosphate (ATP), the universal carrier of energy. Although mostly known for generating ATP through oxidative phosphorylation, mitochondria perform many other metabolic roles such as the generation of a number of metabolites by the tricarboxylic acid (TCA) cycle for cytosolic pathways, oxidative catabolism of amino acids, ketogenesis, as well as the regulation of cytoplasmic calcium to name a few. Although usually represented as small, beanshaped, double-membrane organelles, the mitochondrial structural network actually has a more dynamic nature, a feature that is used to actively move through the cell via interactions with the cytoskeleton. The mitochondrion (Figure 5) is composed of a number of different compartments, each with specialized functions, i.e. the outer mitochondrial membrane, the intermembrane space, the inner mitochondrial membrane and the mitochondrial matrix. The inner membrane is made up of cristae that contain the transmembrane proteins of the electron transport chain and ATPsynthase. The mitochondrial matrix contains all the components of the TCA cycle as well as the β-oxidation pathway. inner membrane pyruvate H + I Q III IV II H + NADH FADH O 2 H 2 O 2 TCA cycle succinate H + cristae c ADP V H + ATP matrix intermembrane space outer membrane Figure 5. Schematic representation of the mitochondrial respiratory pathway Schematic drawing of a mitochondrion, which is composed of a number of different compartments, i.e. the outer mitochondrial membrane, the intermembrane space, the inner mitochondrial membrane and the mitochondrial matrix. The inner membrane is made up of cristae that contain the transmembrane proteins of the electron transport chain and ATP-synthase. NADH and FADH 2 derived from tricarboxylic acid (TCA) cycle activity (or from β-oxidation) feed electrons into the electron transport chain via Complex I and II, respectively. Electrons are transferred from Complex I or II to coenzyme Q (Q), that delivers the electrons to Complex III before reducing O 2 in Complex IV to form water. Translocation of these electrons through the complexes is coupled to proton pumping into the intermembrane space by Complex I, III and IV. The resulting proton motive force, i.e. the ph gradient (DpH) and the membrane potential (DY), drives the proton flux across Complex V (or ATP synthase) back into the mitochondrial matrix, which provides the energy needed to convert ADP to ATP. Both the TCA cycle and β-oxidation pathway produce reducing elements in the form of nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide (FADH 2 ) that enter the electron transport chain (ETC) to generate ATP. Electrons donated from NADH and FADH 2 enter the ETC system via Complex I and Complex II, respectively, which are then transferred through the ubiquinone junction (Q-junction) to Complex III and finally to Complex IV, where the electrons reduce O 2 to form H 2 O. Transfer of electrons through Complex I, III and IV is accompanied by the 14

16 General Introduction pumping of protons across the mitochondrial inner membrane into the intermembrane space, creating an electrochemical gradient. This electrochemical gradient has a chemical contribution, i.e. the ph gradient ( ph), and an electrical contribution, i.e. the membrane potential ( Ψ). The resulting proton motive force is the driving force for ATP-synthase (Complex V) to convert ADP to ATP, while translocating protons back across the inner membrane into the matrix (Figure 5). Before FFA derived substrates enter the β-oxidation pathway, a CoA group is added to the fatty acyl chain by fatty acyl-coa synthase (ACS), resulting in the formation of acyl-coas. Before translocation into the mitochondria, LC acyl-coas are converted to form long-chain acylcarnitines by carnitine palmitoyltransferase 1 (CPT-1) in order to transport the LC acyl-coas across the inner mitochondrial membrane via carnitine-acylcarnitine translocase (CACT) in exchange for carnitine (Figure 6). LC acylcarnitines are converted back to LC acyl-coas at the inner mitochondrial membrane by CPT-2. Short- and medium-chain acyl-coas (SCA-CoA and MCA-CoA, respectively) freely enter the mitochondrion independent of CPT-1. Next, the acyl-coas enter the β-oxidation pathway, producing one acetyl-coa per cycle of the β-oxidation pathway, which then enters the mitochondrial TCA cycle. Besides facilitating transport of FAs into the mitochondria, carnitine is also essential for the efflux of excess intra mitochondrial acyl CoA into the cytosol, and subsequently into the bloodstream. Acylcarnitines are 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 acylcarnitines species with an even number of carbon atoms in their acyl chain reflect incomplete FA oxidation, while odd chain species stem primarily from amino acid catabolism 32. Acetylcarnitine is derived from acetyl CoA, the universal degradation product of all metabolic substrates. 1 Particularly in skeletal muscle, mitochondria are challenged to meet high oxidative energy demands for muscle contraction. Energy turnover in skeletal muscle may increase 400-fold during physical exercise compared with resting conditions 33. In addition to cope with sudden changes in energy demand, muscle cells need metabolic flexibility to rapidly shift between different substrates. The regulation of fuel selection by mitochondria can be described by the Glucose Fatty Acid Cycle or Randle cycle, as proposed by Randle et al. in The Randle cycle describes the competitive relation between glucose and FA oxidation in metabolic tissues, where utilization of one nutrient directly regulates the use of the other, without hormonal intervention. Availability of glucose, i.e. in the fed state, promotes glucose oxidation, while suppressing FA oxidation. Vice versa, supply of tissues with FFA, such as in the fasted state by activation of adipose tissue lipolysis, makes FA the preferred fuel for oxidation and enhances FA oxidation, while oxidation of glucose is suppressed Because skeletal muscle represents a large proportion of total body mass, muscle mitochondrial function therefore has a large impact on total body metabolic function and flexibility. It has been suggested that mitochondria play a prominent role in the pathophysiology of obesity and insulin resistance and that the capacity to oxidize FA is impaired in the insulin resistant state 37,38. A lower capacity of mitochondria to oxidize FA could potentially provoke the accumulation of ectopic lipids and lipid intermediates, thereby inducing insulin resistance. Partially blocking the entry of long-chain FA into mitochondria via inhibition of CPT-1 raises IMCL content, even during low-fat diet conditions, and leads to impaired insulin sensitivity 26. These findings demonstrate that mitochondrial dysfunction can indeed promote ectopic lipid accumulation and result in insulin resistance. Patients with type 2 diabetes were shown to have blunted FA oxidation with 15

17 Chapter 1 respect to body mass index matched control subjects 25 and presented with defects in oxidative metabolism Other studies suggest that a decrease in mitochondrial mass might be a major factor in the pathogenesis of type 2 diabetes, while intrinsic function per mitochondrion is maintained 37,43,44. Furthermore, neither patients nor mouse models with an inherited deficiency in fatty acid oxidation exhibited muscle insulin resistance 45. Therefore the nature of mitochondrial dysfunction and the cause-and-effect relationship between mitochondrial dysfunction and the development of IMCL accretion and insulin resistance still remains elusive 19,46. In fact, recent studies have linked insulin resistance to an increased rather than decreased capacity to oxidize FA 13,14,16. The upregulation of FA oxidation flux is thought to be a mechanism for cells to cope with the excessive influx of FA. However, oxidation of FA substrates is a source of reactive oxygen species (ROS), which promotes mitochondrial uncoupling and could compromise mitochondrial ATP production 19,46,47. Furthermore, it been shown that in insulin-resistant muscle metabolic intermediates of incomplete FA oxidation accumulate 17,28, indicating that FA oxidation flux outpaces the demand of the respiratory system, which may affect mitochondrial redox state and further promote mitochondrial oxidative stress 19,20,47,48. These findings suggest that mitochondrial dysfunction is a consequence rather than a cause of insulin resistance. It does seem evident that a vicious cycle between muscle lipid accumulation and mitochondrial dysfunction underlies the etiology of muscle insulin resistance and T2D 19. However, the exact mechanisms leading to these derangements are poorly understood. Better understanding of the interaction between lipid accretion and functioning of mitochondria is therefore imperative for the development of effective treatment strategies for T2D. 16

18 General Introduction Therapeutic interventions In general, two different additive approaches are implemented for the treatment of T2D, namely lifestyle interventions (diet and exercise therapy) and the use of pharmaceutical agents to lower blood glucose levels 49. Considerable effort is put into the development of insulin sensitizing drugs, like biguanides (such as metformin) and thiazolidinediones (TZDs), to treat or prevent T2D. However the mechanisms behind the therapeutic actions of these agents and their effects on lipid homeostasis and energy metabolism greatly differ and have only partially been resolved. 1 Lifestyle intervention Lifestyle adjustments promoting long-term weight loss, i.e. exercise training and/or dietary changes, have been well documented as effective interventions in the prevention as well as treatment of type 2 diabetes 50,51. The importance of regular physical activity in treatment of diabetes patients is well established In fact, the American Diabetes Association (ADA) recommends a regular moderate-intensity exercise regimen of at least 150 min/week consisting of aerobic exercise activity and/or resistance training, distributed over at least 3 days/week for patients with type 2 diabetes 49. The acute therapeutic efficacy of exercise is credited to the depletion of glycogen stores, insulin-independent activation of glucose transporters via AMP-activated protein kinase (AMPK) 55 and the stimulation of blood flow during and immediately after exercise 54. A number of studies report that short-term endurance training reduces IMCL content in type 2 diabetes 56,57. However, as mentioned earlier, interpretation of IMCL response to exercise intervention is not straightforward as it depends on the metabolic status of the subject. Long-term exercise therapy induces more structural adaptive changes, such as improved insulin sensitivity after prolonged endurance training 58 and increased capacity for glucose disposal as a result of increased skeletal muscle mass after long-term resistance-type exercise training 59. When designing a suitable exercise protocol it is crucial to take the progression of the disease state and body composition of the patient into account. Long-standing diabetes patients may suffer from severe exercise intolerance, as a result of diminished muscle oxidative capacity, muscle weakness, sarcopenia and/or vascular disease. Most exercise intervention programs will be too challenging for this group of patients and their exercise regimen should be adjusted accordingly. Unfortunately, since most patients with T2D have motivational issues, they find it difficult to structurally adhere to lifestyle intervention programs. Therefore they usually have to resort to pharmaceutical intervention strategies to improve their glucose homeostasis. 17

19 Chapter 1 Metformin treatment Metformin is the most commonly prescribed drug to treat type 2 diabetes (T2D), and it has been in clinical use for over 50 years. Metformin is a biguanide and primarily improves insulin sensitivity in the liver, where it inhibits glucose production 60,61, without any marked hypoglycemic effects 62. Moreover, metformin was reported to improve insulin sensitivity in skeletal muscle and stimulate peripheral glucose utilization 63. Its therapeutic effect on glucose homeostasis, its beneficial effects on cardiovascular complications, and the fact that it can easily be combined with other antidiabetic agents, make metformin the first drug of choice for treatment of T2D. In the prevention of type 2 diabetes, metformin was also reported to have potential beneficial effects 64. Recently the mechanism behind metformin s therapeutic actions was clarified 61. Madiraju et al. discovered that the main action of metformin is to block the redox shuttle enzyme mitochondrial glycerophosphate dehydrogenase (mgpd), which decreases the mitochondrial and increases the cellular redox state in the liver. This leads to a reduced conversion of lactate and glycerol to glucose, lowering glucose release by the liver. Treatment with thiazolidinediones Thiazolidinediones (TZDs) were introduced for treatment of T2D in the late 1990s. However, troglitazone was withdrawn from the market 3 years after its introduction in 1997 because of the risk of liver toxicity. Recently, in 2010, rosiglitazone was associated with increased risks for cardiovascular complications 65,66, leading to severe restrictions in its use by the FDA. For this reason pioglitazone is presently the only TZD in use to treat T2D. Aside from its blood glucose lowering effects in the treatment of T2D, a dramatic 72% decline was observed in the risk of developing T2D after treatment with pioglitazone in a prevention study performed in individuals with impaired glucose tolerance. TZDs are activators of the transcription factor peroxisome proliferatoractivated receptor-γ (PPAR-γ) and are known to improve whole-body insulin sensitivity. PPAR-γ is predominantly expressed in white and brown adipose tissue, where it stimulates FA uptake 67,68, β-oxidation, and oxidative phosphorylation 69, and effectively relocates fat from non-adipose to subcutaneous adipose tissue 67. Although PPAR-γ is predominantly expressed in adipose tissue, TZDs are also known to stimulate glucose uptake in skeletal muscle of insulin-resistant patients 70. Insulin therapy When lifestyle interventions and/or oral glucose lowering drugs (i.e. metformin and/or TZDs) fail to reach the recommended glycemic goals (i.e. glycated hemoglobin (HbA1c) values 7.0%), supplementary insulin is administered 28,71,72 to improve metabolic control and prevent microvascular and macrovascular complications 73,74. Moreover, an early start of insulin therapy may protect the β-cells in the pancreas from further damage resulting from long-term hyperglycemia 72,75. The intensity of insulin treatment should be adjusted when glycemic control cannot be maintained due to progressive functional decline of the pancreatic β-cells. Considering the risk reductions of long-term complications as a result of better glycemic control, insulin therapy has proven to be the most effective, tolerable and cost-effective intervention available 76. The main drawbacks of insulin therapy are the risk of hypoglycemia, which is generally smaller in type 2 than in type 1 diabetes 77, and additional weight gain associated with insulin therapy. 18

20 General Introduction The effects of pharmaceutical agents like metformin and TZDs on muscle lipid homeostasis and mitochondrial function are still poorly understood. Because muscle lipid accumulation and mitochondrial dysfunction underlie the etiology of muscle insulin resistance, understanding of the impact of pharmaceutical interventions on these pathological factors is essential to predict their therapeutic efficacy and to develop new anti-diabetic drugs. 1 19

21 Chapter 1 Methodological approaches for the study of muscle lipid homeostasis and mitochondrial function Determination of muscle lipid and lipid intermediate levels In order to study muscle lipid homeostasis, IMCL content and muscle acylcarnitines were determined in this thesis using in vivo 1 H magnetic resonance spectroscopy (MRS) and tandem mass spectrometry (MS/MS) on ex vivo muscle samples, respectively. These measurement techniques will be explained in the following paragraphs. IMCL content IMCL is mainly present as lipid droplets within the cytosol of muscle cells, which can be visualized using electron microscopy 78. Lipids in interstitial adipose tissue are referred to as extramyocellular lipids (EMCL). While EMCL is metabolically relatively inert, IMCL can be rapidly mobilized and utilized for energy metabolism, particularly since the IMCL droplets are primarily located in close contact to the mitochondria in the cell. IMCL levels can be determined using several different techniques including biochemical extraction 71,79, Oil red O histochemical staining 28 and electron microscopy morphometry of needle biopsy samples 56, as well as non-invasive magnetic resonance imaging (MRI) and computed tomography (CT) 57. Disadvantages of these techniques are that they are either invasive, or unable to differentiate between IMCL and EMCL, or both. Magnetic resonance spectroscopy gives a unique view of tissue biochemistry in situ by measuring the concentrations and/or turnover rates of metabolites, and has been widely applied to skeletal muscle H MRS is based on the same principles as the more familiar MRI, which offers excellent soft-tissue contrast and high spatial resolution using signal mainly from tissue water. In a 1 H MR spectrum a range of different metabolites is separately detected, because the different chemical environments of nuclei in different molecules or molecular sites shift their resonance frequencies. IMCL can be distinguished from the EMCL signal dependent on the orientation of the muscle fibers with respect to the magnetic field, with a maximum peak separation of circa 0.2 ppm when the muscle is parallel to the magnetic field. In this case the CH 2 protons of IMCL and EMCL resonate at 1.28 and 1.47 ppm, respectively (Figure 7). This chemical shift difference originates from differences in bulk magnetic susceptibility (BMS) effects, resulting from 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 7. Because 1 H MRS has the capability to non-invasively distinguish IMCL and EMCL, it has proven to be invaluable for the assessment of IMCL 7. However, it is important to note that despite the chemical shift difference between IMCL and EMCL, large fat deposits such as in the adipose tissue should be avoided when planning the voxel localization for the 1 H MRS measurement, to prevent that the IMCL peak will be obscured by the EMCL signal. 20

22 General Introduction Lean rat Diabetic rat IMCL-CH 2 1 tcr-ch 3 IMCL-CH 2 EMCL-CH 2 tcr-ch 3 EMCL-CH 2 IMCL-CH 3 tcr-ch 2 IMCL-CH 3 tcr-ch H chemical shift (ppm) 1 H chemical shift (ppm) FIGURE 7. IMCL content assessed by localized 1 H MRS Spectra were obtained in tibialis anterior muscle of a lean and a diabetic rat at the age of 14 weeks. Peaks of total creatine (tcr), extramyocellular lipids (EMCL) and intramyocellular lipids (IMCL) can be distinguished in these spectra. Acylcarnitine content The current gold-standard for diagnosis of β oxidation disorders at the metabolite level is blood acylcarnitine profile analysis 83. The acylcarnitine accumulation as was observed in insulinresistant skeletal muscle was suggested to reflect a failed attempt to cope with the excess of intra mitochondrial acyl CoAs that causes mitochondrial stress. However, the exact physiological relevance of changes in muscle acylcarnitine levels in insulin resistance is still under debate. Since the vast majority of acylcarnitines is produced in the mitochondria, and acylcarnitine levels can therefore, in combination with measurements of substrate oxidation and mitochondrial function, be interpreted as a measure of β-oxidation flux. Although, measurement of acylcarnitines provides a comprehensive snapshot of intermediary metabolism, it is important to note that the steadystate metabolite concentrations are a representation of the net balance between production, consumption, import and export, and therefore acylcarnitine levels per se do not directly provide information about fluxes through individual metabolic pathways. By means of tandem MS/MS it is possible to analyze 36 independent acylcarnitine species ranging in chain length from 2 to 22 carbons in muscle, blood and urine 84. Acylcarnitines with long chain length (C14-C20) have been reported to be elevated in T2D, while medium-chain (C6-C12) acylcarnitines represent intermediate products of incompletely oxidized FA 85. Furthermore,he shortest acylcarnitine, acetylcarnitine, may reflect the controlling role of acetyl-coa on metabolic flexibility since acetyl- CoA inhibits pyruvate dehydrogenase and the conversion to acetylcarnitine allows transport of acetyl-coa out of the mitochondria. 21

23 Chapter 1 Assessment of muscle mitochondrial function Muscle mitochondrial dysfunction is a hallmark of numerous lifestyle-related diseases, and impairments of mitochondrial function have been reported in heart failure 86, chronic pulmonary disease 87,88 and type 2 diabetes 9,89. A number of factors contribute to the net mitochondrial ATP production in skeletal muscle, such as the cellular ATP demand (driven by the ADP:ATP ratio), intrinsic mitochondrial function, mitochondrial mass, as well as substrate and O 2 availability 82. Different aspects of intrinsic muscle mitochondrial function can be measured in muscle samples ex vivo, such as expression of genes and proteins involved in (regulation of) mitochondrial metabolism or the activity of mitochondrial enzymes 94. Furthermore, O 2 consumption rates or ATP synthesis, normalized to mitochondrial protein mass 95,96, can be measured in isolated mitochondria or permeabilized muscle fibers ex vivo to evaluate the intrinsic functioning of mitochondria, not limited by levels of substrate and O 2 supply 96,97. In order to determine muscle mitochondrial function in vivo, whole-body maximal O 2 consumption can be measured by breath gas analysis, provided a sufficiently large muscle mass is involved during exercise. However, this method tests the entire cardiorespiratory-vascular-muscular system response 98 and therefore does not reflect muscle mitochondrial function alone. Another approach is to measure O 2 consumption of specific muscles by arteriovenous blood sampling, but this is technically complicated and invasive. Finally, oxidative capacity of muscle in vivo can be determined using 31 P MRS by evaluating recovery rates of phosphocreatine (PCr) levels after exercise 99,100. When evaluating oxidative capacity in vivo, aside from mitochondrial content and intrinsic mitochondrial function, also O 2 delivery, delivery of substrates and other factors such as cellular ph, control the rate of oxidative metabolism. Throughout this thesis the main read-out used to determine in vivo mitochondrial function is dynamic 31 P MRS of PCr recovery after muscle exercise. In vivo 31 P MRS is complemented by: 1) ex vivo measurement of the amount of mitochondria (mitochondrial content); 2) detailed characterization of intrinsic mitochondrial function ex vivo using high-resolution respirometry; and 3) in vivo measurement of local changes in skeletal muscle oxygenation during muscle exercise. In vivo 31 P MRS 31 P MRS provides a non-invasive read-out of energy metabolism that is not confounded by motivation or stress, as is the case in the measurement of maximal whole-body O 2 consumption during exercise. 31 P MRS has been widely used to investigate skeletal muscle energy metabolism and its role in the etiology of insulin resistance both in clinical 99 and pre-clinical 100 settings. In the 31 P MR spectrum of skeletal muscle typically five different resonances can be distinguished, representing three different phosphorous containing compounds (Figure 8): inorganic phosphate (P i ), PCr and the α-, β- and γ-phosphate moieties of ATP. Tissue ph can furthermore be determined from the chemical shift of P i, which is typically referenced to the PCr peak that is insensitive to ph At physiological ph, P i is present as both H 2 PO 4- and HPO 4. Because H 2 PO 4 and HPO 4 are in a rapid chemical exchange, only a single P i peak is observed, the frequency of which is dependent on the ph 101. Because 31 P MR spectra of resting skeletal muscle are relatively invariant, even in diseased states, either the magnetic or the chemical equilibrium is usually perturbed to assess muscle mitochondrial defects, as will be described below. 22

24 General Introduction PCr 1 ATP P i ph γ α β P Chemical Shift (ppm) Figure 8. Example of a 31 P MR spectrum from skeletal muscle The 31 P MR spectrum is obtained in skeletal muscle of a lean rat at rest. Peaks from inorganic phosphate (P i ), phosphocreatine (PCr) and the α-, β- and γ-phosphate moieties of ATP can be distinguished in the spectrum. Tissue ph can be determined from the chemical shift difference between P i and PCr. Saturation transfer MRS An elegant feature of MRS is that when nuclei are linked by chemical exchange, the measured signal strength can be sensitized to the rate of exchange even under steady-state conditions, i.e. without changes in metabolite concentrations. In such a magnetization transfer experiment 102,103, we perturb the equilibrium magnetization of one of the nuclei involved in the exchange and measure the effect on the signal strength of its exchange partner. This method has been applied to study ATP turnover in resting muscle. The approach is depicted in Figure 9, in which the red spectrum shows the effect of such a perturbation. The apparent unidirectional P i ATP rate constant is calculated from the reduction in P i signal upon saturation of the γ-atp peak 104 (a variant of magnetization transfer called saturation transfer (ST)) 105, and the P i ATP flux (V ATP ) is obtained by multiplying this by the P i concentration determined from a 31 P MR spectrum without saturation. On the assumption that it predominantly reflects oxidative ATP synthesis by mitochondrial ATP synthase, V ATP has been taken as a measure of mitochondrial function 42, However, the interpretation of ST data is not straightforward The first problem is that ST measures of V ATP in resting skeletal muscle greatly exceed estimates of oxidative ATP synthesis rates by other means Arteriovenous difference measurements of oxygen consumption in human skeletal muscle predict net P i ATP fluxes an order of magnitude lower than the ST estimates 112,113, and while net P i ATP fluxes calculated from 13 C MRS measurements of tricarboxylic acid cycle rate are somewhat higher than those from oxygen consumption measurements, they are far lower than the 31 P ST fluxes Crucially, the P i ATP flux obtained by ST consists of mitochondrial ATP synthase flux plus the flux through any other 23

25 Chapter 1 P i ATP pathway. Of particular relevance are the reactions catalyzed by the glycolytic enzymes glyceraldehyde-3-phosphate dehydrogenase and phosphoglycerate kinase. Although the net glycolytic contribution to ATP production in resting muscle is small 114, these enzymes catalyze a coupled near-equilibrium reaction, and so P i ATP exchange may greatly exceed the net glycolytic flux 104, This glycolytic exchange flux dominates the ST-measured P i ATP flux in resting skeletal muscle A further factor contributing to the overestimation of oxidative ATP synthesis by ST measurements is that at low rates of respiration, such as in resting muscle, the mitochondrial ATP synthase is not operating in a unidirectional manner, and in fact may be near to equilibrium, so that mitochondrial Pi ATP exchange can make a significant contribution to the measured P i ATP flux Finally, it has been suggested that small metabolite pools, including immobilized compounds, which are undetectable in the 31 P MR spectrum, may contribute to the ST effect and therefore lead to an overestimation of the oxidative ATP synthesis flux 109. Figure 9. Measurement of ATP synthesis flux from saturation transfer When nuclei are linked by chemical exchange, the MR signal can be sensitized to the rate of exchange by selectively saturating the equilibrium magnetization of one of the nuclei and measuring the effect on the signal strength of its exchange partner in a saturation transfer experiment. Saturation transfer MRS has been applied to measure ATP synthesis flux in resting skeletal muscle by saturation (i.e. elimination) of the g-atp peak (red spectrum) and by measuring the effect on the inorganic phosphate (P i ) signal, in comparison with a control spectrum with saturation at a downfield frequency, equidistant from P i (black spectrum). The apparent unidirectional P i ATP rate constant (k Pi ATP ) is calculated from the fractional reduction of P i magnetization (DP i ) according to: k Pi ATP = DP i /T 1, where T 1 is the apparent longitudinal relaxation time of P i (measured in a separate experiment). The P i ATP flux (V ATP ) is then determined by multiplying k Pi ATP by the P i concentration at rest, as determined from a 31 P MR spectrum without saturation. The second problem with the interpretation of ST data is that even if V ATP did truly represent oxidative ATP synthesis flux, this still could not be taken as a measure of mitochondrial function Mitochondrial ATP synthesis is a demand-driven process regulated by several feedback-loop error signals, such as the concentrations of ADP and P i. A decreased net ATP synthesis flux in combination with appropriately decreased error signals would simply represent a lower 24

26 General Introduction ATP demand. If mitochondrial function were impaired, the error signals should increase so as to match ATP supply to ATP demand, which in the absence of any other pathology will be normal in mitochondrial disease. While a decreased net ATP synthesis flux with normal or increased error signals might reflect impaired mitochondrial function, in published ST experiments these error signals are usually not reported, and in any case we lack detailed quantitative understanding of the flux-signal relationships around resting rates of ATP turnover 121. Therefore, neither P i ATP exchange flux (measured by ST) or net ATP synthesis rate (measured e.g. indirectly as tricarboxylic acid cycle rate by 13 C MRS 107 ) in resting skeletal muscle has any simple relationship with mitochondrial function Dynamic MRS of PCr recovery As an alternative to the resting state ST experiment, the metabolic steady state of the muscle, i.e. the chemical equilibrium, can be perturbed by inducing muscle contractions, either by letting volunteers perform an exercise protocol or by electrically stimulating the muscle in anaesthetized animals. Cr PCr CK ADP ATP force PCr Cr cytosol CK ADP ATP ATP-synthase mitochondrion Figure 10. PCr energy shuttle mediated by creatine kinase During a sudden increase in energy demand, i.e. at the start of exercise to generate muscle force, creatine kinase (CK) buffers ATP levels by using muscle phosphocreatine (PCr) as an energy source until glycolysis and oxidative phosphorylation are activated. When exercise is halted, PCr is restored to its pre-exercise value. The rate constant of this recovery is a measure of muscle oxidative capacity because, during recovery, PCr is almost exclusively restored from ATP derived from oxidative phosphorylation in muscle mitochondria. 25

27 Chapter 1 The ATP required for muscle contractions is eventually produced by glycolysis and oxidative phosphorylation, but there is a temporary mismatch between ATP demand and supply at the start of a work jump. In order to keep ATP levels constant, this mismatch is met by the breakdown of PCr, a reaction that is catalyzed by creatine kinase (Equation 1; Figure 10). PCr 2- + MgADP - + H + MgATP 2- + Cr (1) This results in an immediate drop of the concentration of PCr, while the P i concentration rises at the start of exercise (Figure 11). After cessation of exercise the PCr level recovers to its pre-exercise value. The rate of PCr resynthesis can be taken as a measure of the suprabasal rate of oxidative ATP synthesis, because, during recovery, PCr is almost exclusively restored from ATP derived from oxidative phosphorylation 124, and because the creatine kinase reaction is much faster than oxidative ATP production 125. A B Figure 11. Changes in metabolite levels during and after exercise using 31 P MRS A) Concentrations of phosphocreatine (PCr; black circles), inorganic phosphate (P i ; grey squares) and ATP (dark grey triangles), assessed by 31 P MRS during 3 minutes of rest, 2 minutes electrical muscle stimulation and 10 minutes recovery protocol in rat tibialis anterior muscle. B) Phosphocreatine concentrations during recovery from muscle activity were fit with a monoexponential function yielding the rate constant of PCr recovery, k PCr. The time resolution of the 31 P MR spectra was 20 seconds. Different approaches have been taken to make inferences about the maximal rate of oxidative ATP synthesis, i.e. the mitochondrial capacity, from PCr recovery data (reviewed in 80,121 ). In the kinetic ADP-control approach, oxidative ATP synthesis rate is described by a hyperbolic 126 or a cooperative, sigmoidal 123 dependence on the ADP concentration, from which the maximal ATP synthesis rate can be extrapolated. The non-equilibrium thermodynamic approach is based on a quasi-linear dependence of the oxidative ATP synthesis rate on the free energy of ATP hydrolysis 127,128, from which the mitochondrial capacity can similarly be inferred. Finally, in what we can broadly term the linear approach (to which the ADP-control and non-equilibrium thermodynamic models both approximate) either the mitochondrial capacity is estimated as the product of the rate constant of PCr recovery (k PCr ; determined by a mono-exponential fit to the PCr recovery data (Equation 2; Figure 11B)) and the resting PCr concentration, or else k PCr itself is used as a relative measure of mitochondrial capacity. PCr(t) = PCr EE - DPCr (1 - e -k PCr t ) (2) 26

28 General Introduction The linear model predicts that k PCr is independent of exercise intensity, which in practice only holds when ph changes are small. Intracellular acidification significantly slows the rate of PCr recovery , which can be explained by the system properties of the creatine kinase equilibrium 132. Unfortunately, the ph-dependence of k PCr differs among subjects, which is related to individual differences in cellular ph control, specifically in proton efflux 132. This implies that correcting k PCr for end-exercise ph using a general formula is not adequate and that, when measuring PCr recovery, care must be taken to prevent muscle acidification. 1 A limitation of all the above approaches is that in conditions of reduced O 2 supply to the muscle, such as in peripheral vascular disease 133, k PCr might not be representative of muscle mitochondrial capacity strictly defined, but rather is determined by extramitochondrial factors. Finally, it has been suggested that during early recovery after intense exercise, PCr recovery is not entirely fueled by oxidative ATP synthesis, but also has a small glycolytic component However, in practice, this fast glycolytic component will not be captured in the analysis of k PCr. There is ample evidence that k PCr, and other measures of mitochondrial capacity based on PCr recovery, correlate well with whole-body maximal O 2 consumption and with ex vivo measurements of mitochondrial function such as muscle mitochondrial content, mitochondrial enzyme activities and maximal O 2 consumption in isolated mitochondria 100, , and that they change appropriately, i.e. in the right direction, with training state , ageing 141,151,152, primary mitochondrial diseases and a variety of secondary causes of mitochondrial dysfunction 133,134, , In contrast, most of the studies directly comparing mitochondrial capacity based on PCr recovery after exercise and V ATP in resting muscle based on ST show no correlation between the two measures 100,143, ; and where such correlations have been observed 108, the considerations explained above mean that the relationship can only be indirect 113. It can be concluded that dynamic MRS of PCr recovery after exercise provides the most reliable method to evaluate in vivo muscle mitochondrial function. However it must be kept in mind that aside from the amount of mitochondria (mitochondrial content) and their intrinsic function (together termed intrinsic mitochondrial capacity), in vivo muscle oxidative capacity also depends on extramitochondrial factors such as the supply of substrates and oxygen (Figure 12). Markers of muscle mitochondrial content There are several ex vivo methods available to determine mitochondrial content in skeletal muscle tissue. The most commonly used markers are citrate synthase (CS) activity, mitochondrial DNA (mtdna) copy number, cytochrome c oxidase content (COX), and cardiolipin content. A recent study investigated which of these markers has the strongest association with the mitochondrial fractional area as determined by transmission electron microscopy imaging (the gold standard for mitochondrial content) 163. Cardiolipin content proved to be the strongest marker of mitochondrial content, followed by citrate synthase activity. Citrate synthase is an enzyme of the tricarboxylic acid (TCA) cycle and it catalyzes the conversion of acetyl-coa and oxaloacetic acid into citric acid. Activity of this enzyme is an exclusive mitochondrial matrix marker and can be determined spectrophotometrically using commercially available assay kits. Although most DNA is packaged as chromosomal DNA in the nucleus, mitochondria also possess a small amount of DNA. Mitochondrial DNA (mtdna) copy number can be measured using real-time Polymerase Chain Reaction (PCR) and, since there are multiple copies of mtdna per cell but only one copy of nuclear DNA, it is expressed relative to nuclear DNA. Interestingly, no correlation was detected 27

29 Chapter 1 between mtdna copy number and mitochondrial fractional area 163. These results indicate that citrate synthase activity is a better marker for mitochondrial content, even though a positive correlation between mtdna and citrate synthase activity has been reported 164. Measurement of intrinsic mitochondrial function ex vivo Intrinsic mitochondrial function of skeletal muscle can be determined ex vivo by measuring O 2 consumption rates (O 2 flux) in isolated mitochondria or permeabilized muscle fibers. Mitochondrial isolation from skeletal muscle involves the homogenization of a fresh muscle sample followed by a series of centrifugation steps to produce a relatively pure mitochondrial fraction. Isolated mitochondria provide a means to specifically study mitochondrial bioenergetics, separate from the influence of other cellular factors, and moreover grant easy access for O 2, substrates and inhibitors. One of the drawbacks of the mitochondrial isolation is, however, that relative large amounts of fresh tissue are needed because the yield at the end of the procedure is relatively low (generally 20-40% of total mitochondrial mass) , which could potentially introduce a bias due to selective isolation from the entire mitochondrial pool 168,169. Other concerns regarding mitochondrial isolation are the disruption of the 3D mitochondrial morphology and, dependent on the experimental aim, the loss of interaction with other cellular compartments 166,167. The permeabilized muscle fiber approach has the advantage that mitochondrial morphology is kept intact as well as interactions between mitochondria and other cellular organelles. The small sample size needed for obtaining permeabilized fibers (3-5 mg wet weight) is another favor over isolated mitochondria ( mg wet weight). Limitations of the permeabilized fiber technique include potential diffusion limitations of O 2, substrates and inhibitors, the tendency for muscle fibers to contract in response to ADP, and complications in normalizing the data to mitochondrial content 167. The latter complication is one of the reasons why we use the isolated mitochondria procedure throughout this thesis. Another reason is that the muscle used for the assessment of in vivo muscle oxidative capacity in this thesis is a mixed fiber type muscle, which means that it contains oxidative as well as glycolytic muscle fibers. Use of the permeabilized muscle fiber technique would introduce uncertainty about the specific fiber types used in each respiration experiment. For this reason we routinely use the entire muscle for mitochondrial isolation. By measuring O 2 consumption rates in isolated mitochondria or permeabilized muscle fibers with different combinations of substrates, inhibitors, and uncouplers, it is possible to attain specific quantitative information on the different mitochondrial respiratory pathways and states (Figure 12). Fueling mitochondria with intermediates from the TCA cycle, such as pyruvate plus malate or succinate, allows estimation of respiratory capacity through complex I and II, respectively. It should be noted that for the specific evaluation of complex II respiration rotenone supplementation is required in order to prevent inhibition of succinate dehydrogenase. Mitochondrial respiration driven by pyruvate plus malate yields NADH, which donates electrons to the ETC complex I, while oxidation of succinate leads to formation of FADH 2, which donates electrons to the ETC cycle via complex II. Electrons from complex I or II are subsequently transferred to complex III and IV where they reduce O 2 to form H 2 O. Protons pumped across the mitochondrial inner membrane via complex I, III and IV provide the proton motive force that drives ATP synthesis in complex V (Figure 5). The capacity of β-oxidation can be determined by fueling mitochondria with fatty acid substrates like palmitoyl-l-carnitine, which can enter mitochondria independent of CPT-1, or palmitoyl-coa plus L-carnitine, which is dependent on CPT-1 for entering the mitochondria, in combination with malate. 28

30 General Introduction CCCP ETS OXP ATP CCCP 1 sub ADP ADP LEAK CAT CCCP OXPHOS LEAK ETS Figure 12. Example of a high-resolution respirometry measurement The blue line represents the O 2 concentration, while the red line represents O 2 flux. The three steady-state fluxes, indicated by the colored rectangles, represent the different respiratory states. After adding substrate (sub; pyruvate plus malate) the mitochondria will respire in a basal resting state. Supplementing the medium with ADP will induce the mitochondria to respire in a fully coupled state of oxidative phosphorylation with maximal ATP synthase activity, the OXPHOS state. Blocking the exchange of ADP and ATP across the mitochondrial inner membrane by ANT with carboxyatractyloside (CAT) effectively halts ATP synthesis and bringing the mitochondria in the LEAK state. Finally, adding the uncoupler carbonyl cyanide 3-chlorophenyl hydrazone (CCCP) uncouples the electron transport system from ATP synthesis, which enables the assessment of the maximal capacity of the electron transport system, the ETS state. In the schematic diagram all the different transitions between respiratory states are presented. Measurement of the oxygen consumption rate of mitochondria can be done with a Clark electrode in a closed system. The different respiratory states are determined by the combination of substrates and inhibitors/uncouplers that are used and they are calculated from the derivative of the oxygen concentration (O 2 flux) in a steady state (Figure 12). The O 2 flux is normalized to the amount of mitochondrial protein and can be used to evaluate intrinsic mitochondrial function. When the mitochondria in a closed system are supplemented with a surplus of substrate, inorganic phosphate, ADP, and O 2, they will respire in a partially coupled state, called the OXPHOS state (classically defined as state 3). In the OXPHOS state, the electrochemical proton gradient generated by proton pumping through complex I, III and IV is used by complex V (ATP synthase) to generate ATP and the OXPHOS state is therefore used as a measure of the capacity for oxidative phosphorylation. By depriving the mitochondria of ADP the system is brought in a nonphosphorylating state of respiration where they continue to respire to compensate for proton leak, which is called the LEAK state (classical state 4). Respiration in the LEAK state is achieved after addition of carboxyatractyloside (CAT), which blocks the exchange of ADP and ATP across the 29

31 Chapter 1 mitochondrial membrane by adenine nucleotide translocase (ANT), thereby effectively preventing ATP synthesis. Finally to assess the maximal capacity of the electron transfer system (ETS), the ETS is uncoupled from ATP synthesis by adding a chemical uncoupler, such as carbonyl cyanide 3-chlorophenyl hydrazone (CCCP). This respiratory state is called the ETS state (classical state U). The ETS state can possibly exceed the OXPHOS capacity, when the ETS system is restricted by the rate of oxidative phosphorylation. Quantitative evaluation of these respiratory states provides a comprehensive evaluation of the intrinsic mitochondrial function. 30

32 General Introduction O 2 intrinsic mitochondrial capacity 1 Mito content X OXPHOS ATP Sub Figure 12. Schematic representation of the network contributing to in vivo muscle oxidative capacity Oxygen (O 2 ) and substrates (Sub) are required for mitochondria to generate energy in the form of ATP during oxidative metabolism (OXPHOS). Therefore O 2 and substrate supply, as well as the amount of mitochondria (Mito content) and their intrinsic function (together termed intrinsic mitochondrial capacity), determine muscle oxidative capacity in vivo. Oxygen supply to skeletal muscle The role of O 2 supply as a modulator and determinant of muscle oxidative capacity has been well documented 20,170,171. Haseler et al. discovered that the training status of an individual influences the matching between O 2 supply and the intrinsic mitochondrial capacity (mitochondrial content and functional capacity per mitochondrion). They reported that muscle oxidative capacity (as determined by the PCr recovery rate constant using 31 P MRS) in sedentary subjects was limited by intrinsic mitochondrial capacity 172, while exercise-trained athletes were limited by O 2 supply 170. In some specific pathologies, such as heart failure or vascular disease, O 2 supply may be more limited than in healthy subjects. Also in the pathogenesis of type 2 diabetes there are findings implying that the delivery of O 2 to the skeletal muscle might be impaired Therefore it is vital to have knowledge on local muscle oxygenation changes when assessing muscle oxidative capacity. Near-infrared spectroscopy Near-infrared spectroscopy (NIRS) is a technique that uses transmission of near-infrared light through a tissue to detect changes in oxygenation. Light of two different wavelengths (766 and 859 nm) is sent through a tissue by placing an optical fiber (optode) directly on the skin while a second optode, placed at some distance (between 3-5 cm for human studies) to the light source, is used to detect the transmitted light. Light penetrates into the skin, subcutaneous fat and muscle and is either scattered or absorbed within the tissue. By differentiating between the two wavelengths, concentration changes of oxy-hemoglobin (O 2 Hb) and deoxy-hemoglobin (HHb) can be monitored locally in muscle tissue. A great advantage of NIRS is that skeletal muscle tissue 31

33 Chapter 1 oxygenation (and hemodynamics) can be monitored real-time and non-invasively during exercise. When combining this technique with 31 P MRS it is possible to monitor the kinetics of intramuscular high-energy phosphates simultaneously with skeletal muscle oxygenation. A limitations of NIRS is dependence of the signal on changes in blood volume (i.e. due to the muscle pump effect at the start of exercise and the hyperemic response at the start of recovery from exercise), which complicates the interpretation of the results. Moreover, NIRS is less suitable for the use in small animals, because of the inferior spatial resolution and the decreased signal-to-noise ratio obtainable in smaller tissue volumes. MR-based oxygenation determination A number of MR-based techniques have been used to non-invasively monitor oxygenation changes in skeletal muscle such as 1 H MRS of deoxymyoglobin 180, arterial spin labelling (ASL) 181,182 and blood-oxygenation level-dependence (BOLD) imaging Myoglobin is exclusively present in muscle tissue. 1 H MRS can be used to detect deoxymyoglobin at 75 ppm downfield from water. The deoxymyoglobin signal can therefore can be used to quantify muscle deoxygenation 180,186,187. A downside of this method is the poor spatial resolution of 1 H MRS of deoxymyoglobin, which leads to signal contamination of adjacent resting muscles. Furthermore, for small animal studies the low myoglobin concentration found in rodents is another drawback of this method. Tissue perfusion is of particular interest for a number of physiological conditions. Some techniques used to measure tissue perfusion require the injection of contrast agents and, because of the required contrast agent clearance, are therefore most useful in steady-state conditions 188. During non-steady-state conditions, ASL can be an attractive alternative, since it does not require the injection of a contrast-agent 182, ASL was first introduced by Detre et al. and is non-invasive, quantitative in absolute terms and spatially and temporally resolved 182,190. By selectively exciting spins in a slice upstream of the target tissue, the spins from the arterial water protons can be used as endogenous contrast. After an evolution time the tagged arterial spins have entered the tissue of interest and will have mixed with the stationary extravascular, tissue-associated spins. The modulation of the tissue magnetization by the arriving arterial spins can be used to estimate perfusion. ASL is able to monitor hyperemic responses over an extended period of time and because of its high sampling rate, is able to capture the rapid and wide range of changes in skeletal muscle perfusion, e.g. during exercise and recovery. Drawbacks of this technique are the low perfusion contrast of ASL, and required knowledge of the water exchange rate in muscle, T 1 and T 2 of blood and muscle and their changes over the capillary path, although the dependence of perfusion values estimated by ASL on these factors is expected to be quite low in vivo 188,191. Furthermore ASL acquisition of muscle perfusion is preferably done during situations where the muscles are motionless, because motion interferences can cause mismatches between the sliceselective inversion and the imaging slice, which makes it a less suitable technique to use during exercise. Finally, analysis of ASL data can be quite complex due to the low contrast-to-noise ratio and inter- and intramuscular differences 188. BOLD is a well-documented contrast mechanism for MRI, which initially was established as a tool to localize oxygenation changes in the brain 183. This technique is based on regional changes in vascular response that can be detected by changes in signal intensity (SI) in T 2 - and T 2 *- weighted MR images. Deoxyhemoglobin is paramagnetic, in contrast to oxyhemoglobin which 32

34 General Introduction is diamagnetic, and affects the magnetic susceptibility of nearby protons consequently lowering the SI. In brain, BOLD signal changes are mainly due to increased blood flow to regions with enhanced neural activity, increasing the local ratio of oxy- to deoxyhemoglobin 184,185,192. A relative drop in local deoxyhemoglobin concentration means reduced intravoxel dephasing, which can thus be observed by an increase in the SI in T 2 *- and T 2 -weighted MR images. Naturally these blood oxygenation changes and consequent effects on the BOLD contrast also occur in skeletal muscle, as was first demonstrated by Toussaint et al. 184 and Lebon et al In contracting skeletal muscle, a negative BOLD effect might be expected as a result of O 2 utilization and subsequent increase in local deoxyhemoglobin levels. However, the negative BOLD effect in muscle during repetitive exercise is hidden by changes in fractional blood volume (the muscle pump effect, capillary recruitment, vascular filling, and vasodilatation 194 ) and intrinsic T 2 increases caused by accumulation of osmolytes that draw water into the cells, leading to an increase in intracellular volume, and by intracellular acidification 194,195. During the recovery from a single, very brief contraction, a transient increase in SI as a result from the hyperemic response can be observed, which has been correlated to NIRS measurements 185,196. Because muscle BOLD MRI needs a high time-resolution, typically sequences based on echo-planar imaging (EPI) are used. Since the O 2 - dependence of blood T 2 and T 2 * are not substantially different, both gradient-echo and spin-echo EPI sequences have been used to sample the BOLD-response in skeletal muscle. The advantage of a gradient-echo compared with a spin-echo sequence is the higher temporal resolution, however this sequence is more sensitive to artefacts. In this thesis a gradient-echo EPI sequence was implemented to monitor local oxygenation changes in skeletal muscle in parallel with the in vivo determination of muscle oxidative capacity. 1 33

35 Chapter 1 Aim of this thesis The aim of this thesis is to determine (1) the pathogenic roles of lipid accumulation and mitochondrial dysfunction in diabetic muscle and (2) the effects of anti-diabetic drug therapies on muscle lipid content and mitochondrial function. To this end, 1 H and 31 P MRS and MRI were used as readouts of in vivo muscle lipid content, mitochondrial oxidative capacity and oxygenation. These in vivo measurements were complemented by ex vivo determination of muscle acylcarnitine levels, markers of mitochondrial content and intrinsic mitochondrial function, to provide a detailed characterization of the interplay between muscle lipid homeostasis and mitochondrial function. Thesis outline Mitochondrial function as a target for therapy in type 2 diabetes Throughout this thesis, dynamic 31 P MRS of PCr recovery after muscle contractions in anesthetized rats is used as a readout of in vivo muscle mitochondrial function. Apart from the number of mitochondria and their intrinsic function, in vivo muscle oxidative capacity also depends on extramitochondrial factors such as the supply of substrates and oxygen. Since oxygen supply is a determinant of muscle oxidative capacity, it is essential that oxygen availability is not a limiting factor to be able to make inferences about in vivo mitochondrial function based on measurements of PCr recovery. In Chapter 2 we determined whether muscle PCr recovery is limited by mitochondrial capacity or by O 2 availability in healthy rats. Furthermore we studied whether an impairment in O 2 availability might play a role in muscle mitochondrial dysfunction in diabetic rats. A vicious cycle between lipid accumulation and impaired mitochondrial oxidative capacity is generally considered to underlie the progression of muscle insulin resistance and T2D. However the cause-and-effect relationship between lipid accretion and mitochondrial dysfunction has not yet been resolved. Accumulation of lipid derived intermediates is thought to lead to insulin resistance and cause mitochondrial stress. In order to gain better understanding of the interaction between lipid accumulation and functioning of mitochondria we first studied treatment strategies aimed at alleviating the lipotoxic effects of these lipid intermediates in diabetic muscle. Carnitine supplementation has been suggested to reduce muscle lipid overload by enhancing fatty acid oxidation and increasing the export of lipid metabolites out of the mitochondria. In Chapter 3, we investigate if carnitine supplementation reduces high-fat diet-induced lipotoxicity, improves in vivo muscle mitochondrial function and ameliorates insulin resistance. Another intervention aimed at reducing lipotoxicity in type 2 diabetes is treatment with thiazolidinediones (TZDs), such as pioglitazone. TZDs promote the relocation of lipids from ectopic sites, such as skeletal muscle, into adipose tissue, thereby reducing the influx of lipids in skeletal muscle. Chapter 4 assesses whether and how pioglitazone treatment affects muscle lipid content and mitochondrial function in lean and diabetic Zucker Diabetic Fatty (ZDF) rats. Metformin is the most commonly prescribed drug worldwide to treat type 2 diabetes and it is reported to inhibit complex I of the respiratory chain. This is surprising, since muscle mitochondrial dysfunction is thought to play a role in the pathogenesis of T2D. Although the specific inhibitory action of metformin on Complex I is well established in vitro, its significance for in vivo skeletal muscle mitochondrial function has yet to be elucidated. In Chapter 5 we determine the effect of different dosages of metformin on in vivo muscle oxidative capacity in a rat model of diabetes. 34

36 General Introduction Next, in Chapter 6, we assess how metformin treatment, aside from its effects on muscle energy metabolism, affects muscle contractile function in diabetic rats. Finally in Chapter 7 a summary and a brief discussion is given on the outcomes of the studies in this thesis, and the chapter is concluded with an outlook for future research. 1 35

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44 General Introduction 167. Perry, C.G., Kane, D.A., Lanza, I.R. & Neufer, P.D. Methods for assessing mitochondrial function in diabetes. Diabetes 62, (2013) Piper, H.M., et al. Development of ischemia-induced damage in defined mitochondrial subpopulations. J Mol Cell Cardiol 17, (1985) Kuznetsov, A.V., et al. Analysis of mitochondrial function in situ in permeabilized muscle fibers, tissues and cells. Nat Protoc 3, (2008) Haseler, L.J., Hogan, M.C. & Richardson, R.S. Skeletal muscle phosphocreatine recovery in exercise-trained humans is dependent on O 2 availability. J Appl Physiol 86, (1999) Butler, A.A. & Kozak, L.P. A recurring problem with the analysis of energy expenditure in genetic models expressing lean and obese phenotypes. Diabetes 59, (2010) Haseler, L.J., Lin, A.P. & Richardson, R.S. Skeletal muscle oxidative metabolism in sedentary humans: 31 P-MRS assessment of O 2 supply and demand limitations. J Appl Physiol 97, (2004) Mac Ananey, O., et al. Cardiac output is not related to the slowed O2 uptake kinetics in type 2 diabetes. Medicine and science in sports and exercise 43, (2011) Hogikyan, R.V., et al. Specific impairment of endothelium-dependent vasodilation in subjects with type 2 diabetes independent of obesity. J Clin Endocrinol Metab 83, (1998) Muniyappa, R. & Sowers, J.R. Role of insulin resistance in endothelial dysfunction. Reviews in endocrine & metabolic disorders 14, 5-12 (2013) Beer, S., et al. Comparison of skin microvascular reactivity with hemostatic markers of endothelial dysfunction and damage in type 2 diabetes. Vascular health and risk management 4, (2008) Reusch, J.E., Bridenstine, M. & Regensteiner, J.G. Type 2 diabetes mellitus and exercise impairment. Reviews in endocrine & metabolic disorders 14, (2013) Sanchez, O.A., et al. Postmaximal contraction blood volume responses are blunted in obese and type 2 diabetic subjects in a muscle-specific manner. Am J Physiol Heart Circ Physiol 301, H (2011) Bauer, T.A., Reusch, J.E., Levi, M. & Regensteiner, J.G. Skeletal muscle deoxygenation after the onset of moderate exercise suggests slowed microvascular blood flow kinetics in type 2 diabetes. Diabetes care 30, (2007) Wang, Z.Y., Noyszewski, E.A. & Leigh, J.S., Jr. In vivo MRS measurement of deoxymyoglobin in human forearms. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 14, (1990) Raynaud, J.S., et al. Determination of skeletal muscle perfusion using arterial spin labeling NMRI: validation by comparison with venous occlusion plethysmography. Magn Reson Med 46, (2001) Williams, D.S., Detre, J.A., Leigh, J.S. & Koretsky, A.P. Magnetic resonance imaging of perfusion using spin inversion of arterial water. Proc Natl Acad Sci U S A 89, (1992) Ogawa, S., Lee, T.M., Kay, A.R. & Tank, D.W. Brain magnetic resonance imaging with contrast dependent on blood oxygenation. Proc Natl Acad Sci U S A 87, (1990) Toussaint, J.F., et al. Perfusion changes in human skeletal muscle during reactive hyperemia measured by echoplanar imaging. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 35, (1996) Damon, B.M., Wadington, M.C., Hornberger, J.L. & Lansdown, D.A. Absolute and relative contributions of BOLD effects to the muscle functional MRI signal intensity time course: effect of exercise intensity. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 58, (2007) Noyszewski, E.A., Chen, E.L., Reddy, R., Wang, Z. & Leigh, J.S. A simplified sequence for observing deoxymyoglobin signals in vivo: myoglobin excitation with dynamic unexcitation and saturation of water and fat (MEDUSA). Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 38, (1997) Ogg, R.J., Kingsley, P.B. & Taylor, J.S. WET, a T1- and B1-insensitive water-suppression method for in vivo localized 1 H NMR spectroscopy. Journal of magnetic resonance. Series B 104, 1-10 (1994) Carlier, P.G., Bertoldi, D., Baligand, C., Wary, C. & Fromes, Y. Muscle blood flow and oxygenation measured by NMR imaging and spectroscopy. NMR in biomedicine 19, (2006) Raynaud, J.S., et al. Characterization of atherosclerotic plaque components by high resolution quantitative MR and US imaging. Journal of magnetic resonance imaging : JMRI 8, (1998). 1 43

45 Chapter Detre, J.A., Leigh, J.S., Williams, D.S. & Koretsky, A.P. Perfusion imaging. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 23, (1992) Frank, L.R., Wong, E.C., Haseler, L.J. & Buxton, R.B. Dynamic imaging of perfusion in human skeletal muscle during exercise with arterial spin labeling. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 42, (1999) Noseworthy, M.D., Bulte, D.P. & Alfonsi, J. BOLD magnetic resonance imaging of skeletal muscle. Seminars in musculoskeletal radiology 7, (2003) Lebon, V., Brillault-Salvat, C., Bloch, G., Leroy-Willig, A. & Carlier, P.G. Evidence of muscle BOLD effect revealed by simultaneous interleaved gradient-echo NMRI and myoglobin NMRS during leg ischemia. Magn Reson Med 40, (1998) Duteil, S., et al. Influence of vascular filling and perfusion on BOLD contrast during reactive hyperemia in human skeletal muscle. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 55, (2006) Patten, C., Meyer, R.A. & Fleckenstein, J.L. T2 mapping of muscle. Semin Musculoskelet Radiol 7, (2003) Sanchez, O.A., Copenhaver, E.A., Elder, C.P. & Damon, B.M. Absence of a significant extravascular contribution to the skeletal muscle BOLD effect at 3 T. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 64, (2010). 44

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47 O 2 availability does not limit in vivo oxidative capacity in skeletal muscle of healthy and diabetic rats as assessed with 31 P MRS under normoxic conditions Bart Wessels 1, Henk M. De Feyter 2, Klaas Nicolay 1 and Jeanine J. Prompers 1 1 Biomedical NMR, Department of Biomedical Engineering, Eindhoven University of Technology, the Netherlands, 2 Department of Diagnostic Radiology, Yale University School of Medicine, New Haven, Connecticut, USA.

48 2

49 Chapter 2 48

50 effect of O 2 availability on muscle oxidative capacity Abstract Aim: Measurement of the rate constant of phosphocreatine (PCr) recovery after exercise by 31 P magnetic resonance spectroscopy (MRS) is a powerful method for the non-invasive determination of muscle oxidative capacity. However, apart from the number of mitochondria and their intrinsic function, in vivo muscle oxidative capacity also depends on extramitochondrial factors such as the supply of O 2. The aim of this study was to determine whether muscle PCr recovery in healthy rats is limited by mitochondrial capacity or by O 2 availability and whether an impairment in O 2 availability plays a role in muscle mitochondrial dysfunction in diabetic rats. Methods: Experiments were performed in lean, healthy (fa/+) and obese, diabetic (fa/fa) 2 Zucker diabetic fatty rats (n=6 per group), whilst breathing (1) normoxic (fraction of inspired O 2 (Fi O2 )=0.21) or (2) hyperoxic (Fi O2 =1.00) air under anesthesia. Blood O 2 saturation levels were monitored throughout the measurements. 31 P MR spectra were measured in the tibialis anterior muscle during and after muscle stimulation, using a minimally invasive stimulation protocol. In addition, local changes in blood oxygenation during muscle stimulation were assessed using T 2 * weighted magnetic resonance imaging (MRI). Results: Both in lean and diabetic rats, average blood O 2 saturation levels were lower when the animals were breathing Fi O2 =0.21 compared with Fi O2 =1.00. Furthermore, at Fi O2 =0.21 muscle stimulation induced a decrease in blood oxygenation in muscle of both lean and diabetic rats, which was not apparent when the animals were breathing Fi O2 =1.00. PCr recovery was slower in diabetic rats compared with lean rats; however, it was unaffected by supplemental O 2 in both groups of animals. Conclusion: Under normoxic conditions muscle oxidative capacity in both lean and diabetic rats is limited by mitochondrial capacity rather than O 2 availability. Therefore, O 2 availability does not play a role in the observed impairment of in vivo muscle oxidative capacity in diabetic rats. 49

51 Chapter 2 Introduction Diseases that affect skeletal muscle mitochondrial function are rapidly gaining socio-economic impact. This is particularly due to the increasing prevalence of conditions involving secondary causes of mitochondrial dysfunction, such as heart failure 1, chronic pulmonary disease 2,3 and type 2 diabetes 4,5. Accurate assessment of muscle mitochondrial function is therefore essential for the development and evaluation of (therapeutic) interventions for these diseases. Different aspects of muscle mitochondrial function can be measured in muscle samples ex vivo, such as expression of genes and proteins involved in (regulation of) mitochondrial metabolism 6-9, mitochondrial mass 10, activity of mitochondrial enzymes 11, or mitochondrial respiratory capacity 12,13. However, these parameters cannot easily be translated to in vivo mitochondrial function, which arises from an integrated network involving both intra- and extramitochondrial factors. 31 P magnetic resonance spectroscopy (MRS) is a powerful non-invasive technique for the in vivo read-out of muscle oxidative capacity 14. In this method, 31 P MR spectra are recorded during recovery after muscle exercise to assess the rate of phosphocreatine (PCr) resynthesis, which is directly related to the (suprabasal) rate of mitochondrial ATP production 15,16. The maximal rate of oxidative ATP synthesis, i.e. oxidative capacity, is then inferred from the rate constant of PCr recovery (k PCr ) 14. Apart from the number of mitochondria and their intrinsic function, in vivo muscle oxidative 17 capacity also depends on extramitochondrial factors such as the supply of substrates and O 2. In order to make inferences about intrinsic mitochondrial capacity from measurements of PCr recovery, it is therefore essential that O 2 availability is not a limiting factor. Haseler et al. showed that in sedentary humans under normoxic conditions, k PCr is not limited by O 2 availability and can therefore be taken as a measure of mitochondrial capacity 18. However, in exercise-trained humans breathing normoxic air, k PCr was determined by O 2 availability and not mitochondrial metabolic limits 19. In animals, O 2 delivery and mitochondrial capacity are supposedly more tightly matched than in humans 20, but the effect of O 2 availability on PCr recovery kinetics has, to the best of our knowledge, never been investigated in animals. In diseased states, O 2 supply may be more limited than in healthy conditions. For example, diabetes has been associated with impairments in muscle blood flow 21,22 as a result of both structural and functional alterations in the microvasculature 23, which may hamper the delivery of O 2 to the mitochondria. The role of mitochondrial dysfunction in type 2 diabetes has been widely debated and therefore the effect of O 2 availability on mitochondrial function in diabetic muscle warrants further investigation. The aim of this study was to determine whether muscle PCr recovery is limited by mitochondrial capacity or by O 2 availability in healthy rats. Furthermore we studied whether an impairment in O 2 availability might play a role in muscle mitochondrial dysfunction in diabetic rats. To this end, 31 P MRS measurements were performed on hindlimb muscles in combination with a minimally invasive protocol to electrically stimulate the dorsal flexor muscles in anesthetized rats, whilst breathing (1) normoxic (21% O 2 ) or (2) hyperoxic (100% O 2 ) air. In addition, we assessed local changes in blood oxygenation during muscle stimulation using T * 2 weighted magnetic resonance imaging (MRI). 50

52 effect of O 2 availability on muscle oxidative capacity Research Design and Methods Ethics statement All experimental procedures were reviewed and approved by the Animal Experimental Committee of Maastricht University (permit number: ). Surgery, MRS experiments, blood sampling and termination were performed using isoflurane (IsoFlo; Abbott Laboratories Ltd, Maidenhead, Berkshire, UK) anesthesia (2-3%) with additional pain relief by buprenorphine (Temgesic; Schering- Plough BV, Houten, The Netherlands), and all efforts were made to minimize animal suffering. Animals Validation of the minimally invasive muscle stimulation protocol was performed in Wistar rats (n=5). The effect of O 2 availability on muscle oxidative capacity was studied in lean, non-diabetic fa/+ (body weight 312±6 g; n=6) and obese, diabetic fa/fa (body weight 369±17 g, P<0.001 when compared with lean animals; n=6) male Zucker diabetic fatty (ZDF) rats of 12 weeks of age. All animals were purchased from Charles River Laboratories (Sulzfield, Germany). The animals were housed pairwise, in a controlled environment (20 C and 50% relative humidity on a 12-h light-dark cycle) and given ad libitum access to water and chow (Wistar rats: R/M-H diet, Ssniff Spezialdiäten GmbH, Soest, Germany; ZDF rats: Purina Formula 5008, Bioservices, the Netherlands). 2 Minimally invasive stimulation of dorsal flexor muscles The rats were anaesthetized and positioned supine in a cradle. A hind limb support and footplate were constructed in such a way that the angle of the hip, the knee and the ankle was ~90. Two small incisions in the skin were made (~1.5 mm wide) along the distal trajectory of the N. peroneus communis to induce muscle contractions in the dorsal flexor muscles, i.e. the tibialis anterior (TA), extensor digitorum longus (EDL), peroneus longus (PL) and peroneus brevis (PB) muscles. Two skin pockets were made by blunt preparation, an electrode was inserted, and the wounds were closed with a single stitch (5.0, Prolene, Ethicon, Inc. NJ, USA). The stimulation electrodes were built in-house and were made of copper strips (2 8 mm 2 ) that were soldered to flexible multistranded copper wire (Cooner Wire, AS , Chatsworth, Ca, USA). After finishing the MR experiments, the electrodes were removed and the incisions were closed with a single stitch. Validation of minimally invasive muscle stimulation protocol using muscle functional MRI All MR experiments were performed using a horizontal 6.3-T MR scanner (Bruker, Ettlingen, Germany). T 2 weighted images were acquired with a Helmholtz 1 H coil using a multi-echo spinecho sequence with 12 echo times (TE) in a range of 7-85 ms (repetition time (TR) = 2000 ms, field of view (FOV) = mm 2, matrix = , slice thickness = 2 mm, averages = 2, acquisition time = 12.9 min). Two T 2 weighted data sets were acquired: (1) Before electrical stimulation, and (2) during electrical stimulation of the dorsal flexor muscles. The stimulation protocol consisted of block pulses with a length of 10 ms, applied every 400 ms during the relaxation delay in the spinecho sequence, with a voltage of 4 V, and was carried out until the end of the image acquisition. Regions of interest (ROIs) representing different muscles of the lower hind limb (TA; EDL; PL and PB; soleus; flexor digitorum longus, flexor hallucis longus and tibialis posterior; gastrocnemius lateralis; plantaris; and gastrocnemius medialis; Figures 1A and 1B) were drawn by hand on a short echo-time image (TE = 7.1 ms) using a custom-built program (Mathematica 7.0, Wolfram Research, Champaign, IL, USA). For each muscle group, T 2 was then determined based on the averaged ROI 51

53 Chapter 2 signals from the 6 even echoes (TE in a range of ms). Effect of O 2 availability on muscle oxidative capacity Lean and diabetic ZDF rats were subjected to MR measurements twice, on separate days, 2-3 days apart: (1) During isoflurane anesthesia with medical air as carrier gas (fraction of inspired O 2 (Fi O2 )=0.21), and (2) during isoflurane anesthesia with 100% O 2 as carrier gas (Fi O2 =1.00). Systemic blood O 2 saturation was monitored using a pulse oximeter (Nonin, Minnesota), which was attached to the foot of the hind limb that was not electrically stimulated. At the end of the MR measurements, a blood sample was taken from the vena saphena and the blood hemoglobin concentration was determined using a Hemocue Hb 201+ blood analyzer (HemoCue, Zoetermeer, the Netherlands). To assess local blood oxygenation changes in the TA muscle during muscle stimulation, T * 2 weighted images were acquired with a circular 1 H surface coil ( 40 mm) using single-shot echo planar imaging (EPI) with fat suppression (TR/TE = 1000/30 ms, FOV = mm 2, matrix = , slice thickness = 1 mm, averages = 1, acquisition time = 1 s per image) during a resting period of 3 min, 2 min of electrical stimulation, and 10 min of recovery. Stimulation pulses were applied every second (during the relaxation delay of the MR sequence) with a stimulation pulse length of 100 ms and a frequency of 80 Hz. The voltage of the stimulation pulses (3-5 V) was optimized at the start of the protocol to reach maximal contractile force, as measured by a custom made force transducer. To quantify the changes in EPI signal intensity (SI EPI ) in the TA muscle during stimulation and recovery, an ROI was manually drawn on a high-resolution gradient echo image and then superimposed on the T * 2 weighted images (Figures 3A and 3B) using Matlab 2010b (Mathworks, Natick, MA, USA). The average SI EPI of the ROI was normalized relative to the mean baseline SI EPI determined during the 3 min of rest. The initial drop in SI EPI at the start of muscle stimulation, defined as the area of the drop below baseline (Figure 3C), was taken as a measure of the muscle blood oxygenation level-dependent (BOLD) effect (AUC BOLD ). In vivo oxidative capacity of the TA muscle was assessed using dynamic 31 P MRS with an ellipsoid (10/18 mm) 31 P surface coil, as described previously 27,28. A fully relaxed spectrum (repetition time = 20 s, 32 averages) was recorded first, followed by a time series of spectra (repetition time = 5 s, 4 averages) obtained during a resting period of 3 min, 2 min of electrical stimulation, and 10 min of recovery, identical to the stimulation protocol for the T * 2 weighted imaging. MR spectra were fitted in the time domain using a nonlinear least squares algorithm (advanced method for accurate, robust, and efficient spectral fitting; AMARES) in the jmrui software package 29 as described previously 27. In short, spectral analysis of the 31 P MR spectra was done by fitting the PCr peak to Lorentzian and the inorganic phosphate (P i ) as well as the α-, β- and γ-atp peaks to Gaussian line shapes. Intracellular ph was calculated from the chemical shift difference between the P i and PCr resonances 28,30. For the time series, the concentrations of PCr determined during recovery were fit to a mono-exponential function using Matlab yielding the rate constant k PCr. For each rat, results from two time series with end-stimulation ph values higher than 6.9 were averaged

54 effect of O 2 availability on muscle oxidative capacity Statistical analysis Data are presented as means ± SD. Statistical significance of the effects of electrical stimulation on muscle T 2 relaxation times were assessed using two-sided paired t-tests. For the determination of the effects of O 2 availability, statistical analysis was performed using a 2x2 mixed design ANOVA with one within-subjects factor (Fi O2 ) and one between-subjects factor (genotype) in the IBM SPSS 20 statistical package (SPSS Inc., Chicago, IL, USA). In case the interaction between Fi O2 and genotype was significant, the differences were evaluated in more detail by separately analyzing the effects of Fi O2 and genotype using Bonferroni-corrected two-sided paired and unpaired t-tests, respectively. The level of statistical significance was set at P<

55 Chapter 2 Results Validation of minimally invasive muscle stimulation protocol using muscle functional MRI The T 2 weighted images acquired during electrical stimulation showed an area of homogeneous signal enhancement with respect to the images before stimulation, which appeared confined to the dorsal flexor muscles (Figures 1C and 1D). The specificity of the stimulation procedure for the dorsal flexor muscles was confirmed by the analysis of the T 2 values for the different muscle groups. An increase in T 2 was only detected in the TA (29 ± 5 %, P<0.001), EDL (25 ± 4 %, P<0.001), PL and PB (21 ± 6 %, P<0.001) muscles, whereas the T 2 values of the other muscles of the lower hind limb were not affected by the stimulation protocol (Figure 1E). Figure 1. Muscle functional MRI during minimally invasive muscle stimulation A) Short echo-time transversal spin-echo image of a Wistar rat lower hind leg (TR/TE = 2000/7.1 ms). B) ROI delineations of separate muscle groups of the rat lower hind leg manually drawn on the short echo-time image: 1 = tibialis anterior, 2 = extensor digitorum longus, 3 = peroneus longus and peroneus brevis, 4 = soleus, 5 = flexor digitorum longus, flexor hallucis longus and tibialis posterior, 6 = gastrocnemius lateralis, 7 = plantaris, 8 = gastrocnemius, 9 = gastrocnemius medialis. Longer echo-time images (TR/TE = 2000/35.3 ms) acquired before (C) and during (D) electrical stimulation of the dorsal flexor muscles. E) T 2 relaxation times for the different muscle groups of the rat lower hind leg before (pre-stim) and during (post-stim) electrical stimulation of the dorsal flexor muscles. T 2 was determined based on the averaged ROI signals from the spin-echo images, only including the 6 even echoes (TE in a range of ms). Data are represented as means ± SD (n=5). *** P<0.001 when compared with pre-stim in the same muscle group. 54

56 effect of O2 availability on muscle oxidative capacity A B C D E 40 T 2 (ms) 35 *** *** *** pre-stim post-stim ROI number 55

57 Chapter 2 Effect of O 2 availability on muscle oxidative capacity Whole-body blood oxygenation parameters Blood hemoglobin concentrations were slightly higher in diabetic rats than in lean rats, independent of Fi O2 (P<0.01; Figure 2A). When breathing Fi O2 =0.21, average blood O 2 saturation levels during the MR measurements were 13% lower in lean rats and 25% lower in diabetic rats when compared with Fi O2 =1.00 (P<0.001; Figure 2B). Moreover, for Fi O2 =0.21 blood O 2 saturation was 16% lower in diabetic rats than in lean rats (P<0.001), while for Fi O2 =1.00 this difference was only 2% (P<0.05; Figure 2B). Fi O 2 = 0.21 A 12 Fi O 2 = 1.00 B 100 # *** *** [Blood Hb] (mm) O 2 saturation (%) ### 0 Lean Diabetic 0 Lean Diabetic Figure 2. Whole-body blood oxygenation Whole-body blood oxygenation parameters in lean and diabetic ZDF rats during anesthesia under normoxic (Fi O2 =0.21) and hyperoxic (Fi O2 =1.00) conditions. A) Concentration of hemoglobin in blood ([Blood Hb]) and B) average blood O 2 saturation level during the MR measurements. Data are represented as means ± SD (n=6 per group). The hemoglobin concentration was significantly higher in diabetic rats compared with lean rats, independent of Fi O2 : P<0.01 (two-way ANOVA). For blood O 2 saturation, the interaction between Fi O2 and genotype was significant and a pairwise analysis of differences of Fi O2 and genotype is provided by Bonferroni-corrected two-sided paired and unpaired t-tests, respectively: *** P<0.001 when compared with animals of the same genotype breathing Fi O2 =0.21; # P<0.05, ### P<0.001 when compared with lean animals breathing the same Fi O2. * T 2 weighted imaging of local blood oxygenation in the muscle Both in lean and diabetic rats, a distinct drop in SI EPI was detected at the start of muscle stimulation when the animals were breathing Fi O2 =0.21 (Figure 3C shows an example for a lean rat). However, when the animals were breathing Fi O2 =1.00 this drop in SI EPI was barely detectable, both in lean and diabetic rats (Figure 3D shows an example for a lean rat). The muscle BOLD effect was estimated from the area of the drop in SI EPI below baseline (AUC BOLD ). AUC BOLD was higher for Fi O2 =0.21 than for Fi O2 =1.00, both in lean (P<0.01) and in diabetic (P<0.001) animals (Figure 3E). In addition, AUC BOLD was higher in diabetic rats compared with lean rats when the animals were breathing Fi O2 =0.21 (P<0.05). AUC BOLD was significantly correlated with blood O 2 saturation levels (R = 0.869, P<0.001; Figure 3F). 31 P MRS measurement of in vivo muscle oxidative capacity PCr and P i concentrations and intracellular ph measured with 31 P MRS in TA muscle at rest and at the end of muscle stimulation are listed in Table 1. End-stimulation ph was slightly lower in diabetic rats breathing Fi O2 =1.00 compared with diabetic rats breathing Fi O2 =0.21 (P<0.05) and compared with lean rats breathing Fi O2 =1.00 (P<0.05). However, end-stimulation ph was higher 56

58 effect of O 2 availability on muscle oxidative capacity than 6.95 for all animals and therefore muscle ph is not expected to affect PCr recovery kinetics. The end-stimulation concentration of P i was higher in muscle of diabetic rats compared with lean rats, independent of Fi O2 (P<0.001), but the percentage of PCr depletion was not different among groups. Table 1. Metabolite concentrations and ph in TA muscle measured by 31 P MRS of lean and diabetic ZDF rats under normoxic (Fi O2 =0.21) and hyperoxic (Fi O2 =1.00) conditions. Lean Diabetic Fi O2 =0.21 Fi O2 =1.00 Fi O2 =0.21 Fi O2 =1.00 Rest ph (-) 7.16 ± ± ± ± [PCr] (mm) 34.6 ± ± ± ± 1.2 [P i ] (mm) 2.1 ± ± ± ± 0.5 End-stimulation ph (-) 7.05 ± ± ± ± 0.03*,# [PCr] (mm) 13.9 ± ± ± ± 2.0 [P i ] (mm) 26.5 ± ± ± ± 2.2 PCr (%) 63.9 ± ± ± ± 3.1 Data are represented as means ± SD (n=6 per group). End-stimulation [P i ] was significantly higher for diabetic rats compared with lean animals, independent of Fi O2 (two-way ANOVA: P<0.001). For end-stimulation ph, the interaction between Fi O2 and genotype was significant and a pairwise analysis of differences of Fi O2 and genotype is provided by Bonferroni-corrected two-sided paired and unpaired t-tests, respectively: * P<0.05 when compared with animals of the same genotype breathing Fi O2 =0.21; # P<0.05 when compared with lean animals breathing the same Fi O2. 57

59 Chapter 2 A B C D Rest Stim Recovery Rest Stim Recovery E 2.0 # F 1.5 AUC BOLD (-) Fi O 2 = ** *** Fi O 2 = 1.00 Lean Diabetic 58

60 effect of O 2 availability on muscle oxidative capacity 2 Figure 3. T 2 * weighted imaging during muscle stimulation in lean and diabetic ZDF rats A) High-resolution gradient echo image in which an ROI was manually drawn in the TA muscle. B) Example of an EPI image with the ROI drawn in the high-resolution image superimposed. C) Typical example of EPI signal intensities (SI EPI ) determined in the ROI in the TA muscle during 3 min of rest, 2 min of electrical stimulation, and 10 min of recovery in a lean rat breathing Fi O2 =0.21 conditions. SI EPI was normalized relative to the mean baseline SI EPI determined during the 3 min of rest. The initial drop in SI EPI at the start of muscle stimulation, defined as the area of the drop below baseline (denoted in red), was quantified and taken as a measure of the muscle blood oxygenation level-dependent (BOLD) effect (AUC BOLD ). D) Typical example of SI EPI during rest, electrical stimulation, and recovery in a lean rat breathing Fi O2 =1.00 air. AUC BOLD is barely detectable under these conditions. E) AUC BOLD in lean and diabetic rats under Fi O2 =0.21 and Fi O2 =1.00 conditions. Data are represented as means ± SD (n=6 per group). The interaction between Fi O2 and genotype was significant and a pairwise analysis of differences of Fi O2 and genotype is provided by Bonferroni-corrected two-sided paired and unpaired t-tests, respectively: ** P<0.01, *** P<0.001 when compared with animals of the same genotype breathing Fi O2 =0.21; # P<0.05 when compared with lean animals breathing the same Fi O2 ) Correlation between AUC BOLD and the average blood O2 saturation level during the MR measurements. 59

61 Chapter 2 The PCr recovery rate constant, k PCr, was 23% lower in diabetic rats compared with lean rats, independent of Fi O2 (0.51 ± 0.03 versus 0.66 ± 0.06 min -1, P<0.001; Figure 4). However, increasing Fi O2 from 0.21 to 1.00 did not affect k PCr in either lean or diabetic animals, indicating that under normoxic conditions muscle oxidative capacity in both lean and diabetic rats is limited by mitochondrial capacity and not O 2 availability. Lean Diabetic 0.7 k PCr (min -1 ) Fi O 2 = 0.21 Fi O 2 = 1.00 Fi O 2 = 0.21 Fi O 2 = 1.00 Figure 4. PCr recovery measurements Rate constants of PCr recovery (k PCr ) after electrical stimulation measured in TA muscle of lean and diabetic ZDF rats during anesthesia under normoxic (Fi O2 =0.21) and hyperoxic (Fi O2 =1.00) conditions. Data are represented as means ± SD (n=6 per group). k PCr was significantly lower in diabetic rats compared with lean rats, independent of Fi O2 : P<0.001 (two-way ANOVA). 60

62 effect of O 2 availability on muscle oxidative capacity Discussion We aimed to determine to what extent the rate constant of PCr recovery, determined in vivo in anesthetized lean and diabetic rats under normoxic conditions using 31 P MRS, is limited by mitochondrial capacity or by O 2 availability. Both in lean and diabetic rats, average blood O 2 saturation levels were lower when the animals were breathing Fi O2 =0.21 compared with Fi O2 =1.00. Furthermore, at Fi O2 =0.21 muscle stimulation induced a decrease in muscle BOLD contrast in both lean and diabetic rats, which was not apparent when the animals were breathing Fi O2 =1.00. PCr recovery was slower in diabetic rats compared with lean rats; however, it was unaffected by supplemental O 2 in both groups of animals. Together, these findings indicate that under normoxic conditions muscle oxidative capacity in both lean and diabetic rats is limited by mitochondrial capacity rather than O 2 availability. 2 We presented a minimally invasive and MR-compatible method to induce muscle contractions in vivo in rat skeletal muscle. The stimulation method requires only minor surgery involving subcutaneous implantation of two electrodes, which can easily be repeated within the same animal in longitudinal study designs. We induced electrical stimulation through the N. peroneus communis, the nerve that innervates the dorsal flexor muscles. Muscle activation increases the T 2 of muscle water, which is employed in T 2 weighted, muscle functional MRI to characterize spatial patterns and intensity of muscle activation 32. The increase in T 2 upon muscle contractions is thought to be caused by the accumulation of osmolytes that draw water into the cells, leading to an increase in intracellular volume, and by intracellular acidification 33,34. The specificity of the stimulation procedure for the dorsal flexor muscles was confirmed by the quantification of muscle T 2 values before and after muscle stimulation. An increase in T 2 was only detected in the TA, EDL, PL and PB muscles, whereas the other muscles of the lower hind limb did not show any change in T 2 upon stimulation. Moreover, activation of the N. peroneus communis by the use of subcutaneously implanted electrodes led to a very homogeneous stimulation of the dorsal flexor muscles and highly reproducible levels of muscle activation among different animals, which is less easy to accomplish with completely non-invasive, transcutaneous stimulation protocols 35. When breathing normoxic air, blood O 2 saturation levels were significantly lower in anesthetized lean and diabetic rats when compared with hyperoxic conditions. To quantify local changes in blood oxygenation in the muscle during electrical stimulation, we performed T 2 * weighted imaging. During muscle stimulation, decreases in blood volume and oxygenation due to increased intramuscular pressure (leading to ejection of blood) and O 2 extraction, respectively, lower both T 2 and T 2 *, which is known as the muscle BOLD effect However, metabolic activity during muscle contractions leads to an osmotically-driven increase in intracellular volume and to intracellular acidification, causing an increase in T 2 33,34. Therefore, the contribution of the actual BOLD effect to changes in signal intensity in the T 2 * weighted images is hard to estimate. Nevertheless, the BOLD contrast is increasingly being employed to assess changes in, usually post-contractile, muscle blood oxygenation 21, However, in protocols with longer periods of muscle stimulation (3 min in our case) the post-contractile T 2 * weighted signal is heavily affected by changes in T 2 unrelated to the BOLD phenomenon and therefore we quantified the initial drop in T 2 * weighted signal intensity at the start of muscle stimulation as a measure of the muscle BOLD effect 45. Besides the BOLD effect, this early dip is thought to be determined by the alkalotic effect of the creatine kinase reaction (tending to decrease T 2 ) and the osmotic effects of PCr hydrolysis (tending to increase T 2 ) 36. However, the rate of PCr hydrolysis and the intracellular ph during the first 60 seconds of muscle stimulation as determined from the 31 P MR spectra did not differ among groups (data not 61

63 Chapter 2 shown), indicating that differences in the decrease in T * 2 weighted signal intensity between lean and diabetic rats or between the Fi O2 =0.21 and Fi O2 =1.00 conditions predominantly represent BOLD contrast. Under normoxic conditions, there was a significant decrease in T * 2 weighted signal intensity at the start of muscle stimulation (AUC BOLD ), both in lean and diabetic rats, which was however almost completely blunted with supplemental O 2. Decreases in blood volume due to an increase in intramuscular pressure at the start of muscle stimulation are not expected to be modulated by Fi O2. Moreover, the intensity of muscle stimulation was kept similar between animals to ensure it would not affect the BOLD contrast 36. Therefore, the larger AUC BOLD under normoxic conditions can be interpreted as a larger decrease in muscle blood oxygenation during muscle contractions as compared with hyperoxic conditions. Even though supplemental O 2 increased average blood O 2 saturation levels and blunted the stimulation-induced decrease in muscle blood oxygenation, it did not affect k PCr in either lean or diabetic rats. These results are in line with findings in sedentary young and elderly human subjects 18,46,47 and indicate that in these species under normoxic conditions muscle oxidative capacity is determined by mitochondrial metabolic limits and not by O 2 availability. In contrast, it was shown that in exercise-trained humans k PCr increased upon O 2 supplementation, suggesting that in that case muscle oxidative capacity is limited by O 2 availability in normoxia 19, which can be explained by the higher mitochondrial density in exercise-trained individuals. Earlier studies in trained subjects demonstrated that also during maximal exercise muscle oxidative capacity is limited by O 2 supply 48,49. Moreover, it was recently shown that in untrained human subjects O 2 delivery is inadequate to match muscle mitochondrial capacity during exercise engaging a large proportion of muscle mass; however, this was not the case during exercise involving a small muscle mass 20. Taken together, it appears that O 2 delivery differentially affects muscle oxidative capacity depending on the individual s physical fitness and the proportion of muscle mass which has to be supplied. In our study, the dorsal flexor muscles of one hind limb represent only a minor proportion of whole-body muscle mass and therefore cardiac output limits are not expected to play a role. It should be noted that although overall we did not observe a significant effect of supplemental O 2 on k PCr, in two lean rats k PCr increased substantially when the animals were breathing Fi O2 =1.00 compared with Fi O2 =0.21 (Figure 4). Surprisingly, these two animals had a relatively slow PCr recovery at Fi O2 =0.21, which normalized to the level of the other lean animals when given supplemental O 2. Unlike in trained subjects, the effect of supplemental O 2 in these two lean rats can therefore not be explained by a higher mitochondrial capacity compared with the other lean rats. In agreement with previous studies 28,50, we observed that k PCr was 23% lower in diabetic rats compared with lean controls. Additionally, we found that this difference was independent of Fi O2, which suggest that O 2 availability does not play a role in the impairment of in vivo muscle oxidative capacity in diabetic rats. In our earlier studies we showed that ex vivo mitochondrial respiratory capacity, mtdna copy number and citrate synthase activity were not different between muscle of lean and diabetic rats, implying that neither a lower mitochondrial content nor an impairment of their intrinsic function can account for the lower k PCr in diabetic rats 28,50. Instead, we demonstrated that the impairment of in vivo muscle oxidative capacity in diabetic rats is likely the result of lipid-induced mitochondrial uncoupling and/or inhibition 50, which is no longer apparent in ex vivo preparations. 62

64 effect of O 2 availability on muscle oxidative capacity The lack of effect of supplemental O 2 on k PCr in diabetic rats has to be interpreted with caution. Under normoxic conditions, blood O 2 saturation was 16% lower and AUC BOLD was 150% higher in diabetic rats than in lean rats, indicating that muscle blood oxygenation was compromised in the diabetic animals. However, supplemental O 2 might not have an effect on muscle blood oxygenation when the delivery of blood to the muscle is impaired. Evidence of restricted blood flow in diabetic muscle has grown in recent years 21,22 and has been related to both structural and functional deficiencies of the microvasculature 23,51,52. Functional deficiencies include diminished vasodilation and capillary recruitment in response to classic endothelium-dependent vasodilators and hyperemia 53,54, and a lower sensitivity to insulin-induced vasodilation and capillary recruitment 52, Structural impairments of the diabetic microvasculature encompass a decreased capillary density, structural remodeling and increased atherosclerosis However, in our study there is no evidence of a blood flow limitation in diabetic muscle, as supplemental O 2 decreased AUC BOLD to the same level seen in lean rats. Therefore, the lack of effect of supplemental O 2 on k PCr in diabetic rats cannot be explained by an impaired delivery of O 2 to the muscle, but truly represents a limitation of the mitochondrial machinery. 2 63

65 Chapter 2 In conclusion, we demonstrated that under normoxic conditions muscle stimulation induced a larger decrease in muscle blood oxygenation in anesthetized lean and diabetic rats as compared with hyperoxic conditions. However, supplemental O 2 did not affect the rate constant of PCr recovery after muscle stimulation, showing that under normoxic conditions muscle oxidative capacity in both lean and diabetic rats is limited by mitochondrial capacity rather than O 2 availability. Moreover, our findings demonstrate that O 2 availability does not play a role in the observed impairment of in vivo muscle oxidative capacity in diabetic rats. Acknowledgements We thank Leonie Niesen and David Veraart for their assistance in animal handling, and Larry de Graaf for the construction of the stimulation electrodes. 64

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67 Chapter Patti, M.E. & Corvera, S. The role of mitochondria in the pathogenesis of type 2 diabetes. Endocrine reviews 31, (2010). 26. Dumas, J.F., Simard, G., Flamment, M., Ducluzeau, P.H. & Ritz, P. Is skeletal muscle mitochondrial dysfunction a cause or an indirect consequence of insulin resistance in humans? Diabetes Metab 35, (2009). 27. De Feyter, H.M., et al. Increased intramyocellular lipid content but normal skeletal muscle mitochondrial oxidative capacity throughout the pathogenesis of type 2 diabetes. FASEB J. 22, (2008). 28. Wessels, B., Ciapaite, J., van den Broek, N.M., Nicolay, K. & Prompers, J.J. Metformin impairs mitochondrial function in skeletal muscle of both lean and diabetic rats in a dose-dependent manner. PloS one 9, e (2014). 29. Vanhamme, L., van den Boogaart, A. & Van Huffel, S. Improved method for accurate and efficient quantification of MRS data with use of prior knowledge. J. Magn. Reson. 129, (1997). 30. Taylor, D.J., Bore, P.J., Styles, P., Gadian, D.G. & Radda, G.K. Bioenergetics of intact human muscle. A 31P nuclear magnetic resonance study. Mol Biol Med 1, (1983). 31. van den Broek, N.M., De Feyter, H.M., de Graaf, L., Nicolay, K. & Prompers, J.J. Intersubject differences in the effect of acidosis on phosphocreatine recovery kinetics in muscle after exercise are due to differences in proton efflux rates. Am J Physiol Cell Physiol 293, C (2007). 32. Meyer, R.A. & Prior, B.M. Functional magnetic resonance imaging of muscle. Exercise and sport sciences reviews 28, (2000). 33. Patten, C., Meyer, R.A. & Fleckenstein, J.L. T 2 mapping of muscle. Seminars in musculoskeletal radiology 7, (2003). 34. Damon, B.M., et al. Intracellular acidification and volume increases explain R 2 decreases in exercising muscle. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 47, (2002). 35. Giannesini, B., et al. New experimental setup for studying strictly noninvasively skeletal muscle function in rat using 1 H-magnetic resonance (MR) imaging and 31 P-MR spectroscopy. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 54, (2005). 36. Damon, B.M., Wadington, M.C., Hornberger, J.L. & Lansdown, D.A. Absolute and relative contributions of BOLD effects to the muscle functional MRI signal intensity time course: effect of exercise intensity. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 58, (2007). 37. Lebon, V., Carlier, P.G., Brillault-Salvat, C. & Leroy-Willig, A. Simultaneous measurement of perfusion and oxygenation changes using a multiple gradient-echo sequence: application to human muscle study. Magnetic resonance imaging 16, (1998). 38. Noseworthy, M.D., Bulte, D.P. & Alfonsi, J. BOLD magnetic resonance imaging of skeletal muscle. Seminars in musculoskeletal radiology 7, (2003). 39. Duteil, S., et al. Influence of vascular filling and perfusion on BOLD contrast during reactive hyperemia in human skeletal muscle. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 55, (2006). 40. Damon, B.M., Hornberger, J.L., Wadington, M.C., Lansdown, D.A. & Kent-Braun, J.A. Dual gradient-echo MRI of post-contraction changes in skeletal muscle blood volume and oxygenation. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 57, (2007). 41. Towse, T.F., Slade, J.M., Ambrose, J.A., DeLano, M.C. & Meyer, R.A. Quantitative analysis of the postcontractile blood-oxygenation-level-dependent (BOLD) effect in skeletal muscle. J Appl Physiol (1985) 111, (2011). 42. Meyer, R.A., et al. BOLD MRI mapping of transient hyperemia in skeletal muscle after single contractions. NMR in biomedicine 17, (2004). 43. Andreisek, G., et al. T 2* -weighted and arterial spin labeling MRI of calf muscles in healthy volunteers and patients with chronic exertional compartment syndrome: preliminary experience. AJR. American journal of roentgenology 193, W (2009). 44. Jacobi, B., et al. Skeletal muscle BOLD MRI: From underlying physiological concepts to its usefulness in clinical conditions. Journal of Magnetic Resonance Imaging 35, (2012). 66

68 effect of O 2 availability on muscle oxidative capacity 45. Schmid, A.I., et al. Exercising calf muscle T 2 * changes correlate with ph, PCr recovery and maximum oxidative phosphorylation. NMR in biomedicine 27, (2014). 46. Haseler, L.J., Lin, A., Hoff, J. & Richardson, R.S. Oxygen availability and PCr recovery rate in untrained human calf muscle: evidence of metabolic limitation in normoxia. American journal of physiology. Regulatory, integrative and comparative physiology 293, R (2007). 47. Layec, G., Haseler, L.J. & Richardson, R.S. Reduced muscle oxidative capacity is independent of O 2 availability in elderly people. Age (Dordr) 35, (2013). 48. Knight, D.R., et al. Effects of hyperoxia on maximal leg O 2 supply and utilization in men. J Appl Physiol (1985) 75, (1993). 49. Richardson, R.S., et al. Evidence of O 2 supply-dependent VO 2 max in the exercise-trained human quadriceps. J Appl Physiol (1985) 86, (1999). 50. Wessels, B., et al. Pioglitazone treatment restores in vivo muscle oxidative capacity in a rat model of diabetes. Diabetes Obes Metab (2014). 51. Reusch, J.E., Bridenstine, M. & Regensteiner, J.G. Type 2 diabetes mellitus and exercise impairment. Reviews in endocrine & metabolic disorders 14, (2013). 52. de Jongh, R.T., Serne, E.H., RG, I.J., de Vries, G. & Stehouwer, C.D. Impaired microvascular function in obesity: implications for obesity-associated microangiopathy, hypertension, and insulin resistance. Circulation 109, (2004). 53. Hogikyan, R.V., et al. Specific impairment of endothelium-dependent vasodilation in subjects with type 2 diabetes independent of obesity. J Clin Endocrinol Metab 83, (1998). 54. Steinberg, H.O., et al. Obesity/insulin resistance is associated with endothelial dysfunction. Implications for the syndrome of insulin resistance. J Clin Invest 97, (1996). 55. Laakso, M., Edelman, S.V., Brechtel, G. & Baron, A.D. Decreased effect of insulin to stimulate skeletal muscle blood flow in obese man. A novel mechanism for insulin resistance. J Clin Invest 85, (1990). 56. Barrett, E.J., et al. The vascular actions of insulin control its delivery to muscle and regulate the rate-limiting step in skeletal muscle insulin action. Diabetologia 52, (2009). 57. Muniyappa, R. & Sowers, J.R. Role of insulin resistance in endothelial dysfunction. Reviews in endocrine & metabolic disorders 14, 5-12 (2013). 58. Frisbee, J.C. Hypertension-independent microvascular rarefaction in the obese Zucker rat model of the metabolic syndrome. Microcirculation 12, (2005). 59. Stepp, D.W. Impact of obesity and insulin resistance on vasomotor tone: nitric oxide and beyond. Clin Exp Pharmacol Physiol 33, (2006). 60. Williams, I.L., Wheatcroft, S.B., Shah, A.M. & Kearney, M.T. Obesity, atherosclerosis and the vascular endothelium: mechanisms of reduced nitric oxide bioavailability in obese humans. Int J Obes Relat Metab Disord 26, (2002). 2 67

69 Carnitine supplementation in high-fat diet fed rats does not ameliorate lipid-induced skeletal muscle mitochondrial dysfunction in vivo Bart Wessels 1, Nicole M.A. van den Broek 1, Jolita Ciapaite 1,*, Sander M. Houten 2,#, Ronald J. A. Wanders 2, Klaas Nicolay 1, Jeanine J. Prompers 1 1 Biomedical NMR, Department of Biomedical Engineering, Eindhoven University of Technology, Eindhoven, the Netherlands, 2 Laboratory Genetic Metabolic Diseases, Departments of Pediatrics and Clinical Chemistry, Academic Medical Center, Amsterdam, the Netherlands, *Current affiliation: Center for Liver, Digestive and Metabolic Diseases, Department of Pediatrics, University of Groningen, University Medical Center Groningen, Groningen, the Netherlands, # Current affiliation: Department of Genetics and Genomic Sciences, Icahn Institute for Genomics and Multiscale Biology, Icahn School of Medicine at Mount Sinai, New York, USA

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72 Carnitine insufficiency and mitochondrial function Abstract Aim: Muscle lipid overload and the associated accumulation of lipid intermediates plays an important role in the development of insulin resistance. Carnitine insufficiency is a common feature of insulin-resistant states and might lead to incomplete fatty acid oxidation and impaired export of lipid intermediates out of the mitochondria. The aim of the present study was to test the hypothesis that carnitine supplementation reduces high-fat diet-induced lipotoxicity, improves muscle mitochondrial function and ameliorates insulin resistance. Methods: Wistar rats were fed either normal chow or a high-fat diet for 15 weeks. One group of high-fat diet fed rats was supplemented with L-carnitine during the last 8 weeks. Muscle mitochondrial function was measured in vivo by 31 P magnetic resonance spectroscopy (MRS) and ex vivo by high-resolution respirometry. Muscle lipid status was determined by 1 H MRS (intramyocellular lipids) and tandem mass spectrometry (acylcarnitines). Results: High-fat diet feeding induced insulin resistance, was associated with decreases in muscle and blood free carnitine, elevated levels of muscle lipids and acylcarnitines, and an increased number of muscle mitochondria that showed an improved capacity to oxidize fat-derived substrates when tested ex vivo. This was however not accompanied by an increase in muscle oxidative capacity in vivo, indicating that in vivo mitochondrial function was compromised. Despite partial normalization of muscle and blood free carnitine content, carnitine supplementation did not induce improvements in muscle lipid status, in vivo mitochondrial function or insulin sensitivity. Conclusion: Carnitine insufficiency does not play a major role in high-fat diet induced muscle mitochondrial dysfunction in vivo. 3 71

73 Chapter 3 Introduction Diabetes has reached epidemic proportions worldwide 1. Type 2 diabetes accounts for 85-95% of all diabetes cases and is characterized by insulin resistance in major metabolic tissues such as skeletal muscle 2. One of the leading hypotheses in the research field of type 2 diabetes is that lipid overload in muscle cells, supposedly as a result of a reduced mitochondrial capacity to oxidize fatty acids (FAs), leads to impaired insulin signaling 3,4. However, recent studies have linked insulin resistance to an increased rather than a decreased capacity to oxidize FAs 5-9. The increased FA oxidation capacity in insulin-resistant states was shown to be associated with the accumulation of intermediates of incomplete FA oxidation 10-12, indicating that FA oxidation flux outpaces the demand of the respiratory system. This mismatch between FA substrate supply and demand may promote mitochondrial oxidative stress, which is thought to contribute to the development of insulin resistance 11,13,14. Carnitine is an essential nutrient with multiple functions. Its major role is in the formation of acylcarnitines from long-chain FAs, which is required for the transport of acyl moieties into the mitochondrial matrix for β-oxidation 15,16. A second role for carnitine is to increase acyl and acetyl group efflux out of the mitochondria and into the plasma 17. Moreover, carnitine stimulates the oxidation of pyruvate by lowering the mitochondrial acetyl-coa/coa ratio in a reaction catalyzed by carnitine acetyltransferase (CrAT), which converts acetyl-coa into acetylcarnitine 18. Carnitine insufficiency is a common feature of insulin-resistant states and it has been shown that muscle free carnitine negatively correlates with insulin resistance 12. Therefore, it has been suggested that carnitine supplementation might be an effective treatment for type 2 diabetes 19,20. Indeed, intravenous infusion of carnitine during a hyperinsulinaemic-euglycaemic clamp has been shown to increase whole-body glucose disposal in both healthy subjects and type 2 diabetes patients Moreover, carnitine supplementation has been shown to improve whole-body glucose tolerance in insulin-resistant human subjects and rodent models of metabolic disease 12,26,27. The positive effect of carnitine on insulin sensitivity can be explained by different mechanisms, depending on its concentration. In the perfused isolated working rat heart, the addition of carnitine to the perfusion medium increased glucose oxidation by lowering the concentration of acetyl-coa in the mitochondrial matrix through CrAT 28. However, at the lower concentrations achievable in vivo, carnitine has been shown to stimulate FA oxidation 29,30 through a mass-action effect on the transport of long-chain FAs into the mitochondrial matrix 31. At the same time, carnitine increases the efflux of acylcarnitines from muscle tissue 12. Therefore, it has been proposed that carnitine supplementation ameliorates insulin resistance by reducing lipotoxicity both through increased oxidation and increased export of muscle lipid metabolites 19. In a previous study, we showed that in long-term high-fat diet fed rats in vivo muscle mitochondrial function is compromised by mitochondrial lipid overload 8. The aim of the present study 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 weeks and one group of high-fat diet fed rats was supplemented with L-carnitine (300 mg/kg body weight/day) during the last 8 weeks. In vivo muscle mitochondrial function was measured by 31 P magnetic resonance spectroscopy (MRS) and ex vivo mitochondrial function was determined by measuring oxygen consumption in isolated mitochondria. Furthermore, intramyocellular lipid levels were determined by in vivo 1 H MRS and free carnitine and acylcarnitine levels were determined upon sacrifice in muscle tissue, 72

74 Carnitine insufficiency and mitochondrial function and in blood and urine, using tandem mass spectrometry. Materials and Methods Animals Adult male Wistar rats (14 weeks of age, n=30; Charles River Laboratories, The Netherlands) were housed in pairs at 20 C and 50% humidity, with a 12-h light-dark cycle. Ad libitum food and water was provided during a period of 15 weeks. The rats were divided into three groups (n=10 per group): A control group receiving normal chow (NC; 9% calories from fat, 67% calories from carbohydrate, 24% calories from protein; R/M-H diet, Ssniff Spezialdiäten GmbH, Soest, Germany), a group receiving a high-fat diet (HFD; 45% calories from fat (predominantly lard), 35% calories from carbohydrate, 20% calories from protein; D12451, Research Diet Services, Wijk bij Duurstede, the Netherlands), and a group receiving the same high-fat diet, supplemented with 300 mg/kg body weight/day L-carnitine in their drinking water for the last 8 weeks (HFDC). Body weight and food and water intake were determined weekly. Two days after the in vivo MRS measurements, rats were sacrificed by incision of the inferior vena cava under anesthesia. One tibialis anterior (TA) muscle was used for isolation of mitochondria. The other TA was frozen in liquid nitrogen and stored at -80 C for acylcarnitine content and mitochondrial DNA (mtdna) copy number determinations. All experimental procedures were reviewed and approved by the Animal Experimental Committee of Maastricht University. 3 Oral glucose tolerance test An oral glucose tolerance tests (OGTT) was performed after 15 weeks of diet, three to five days before the in vivo measurements. After a four-hour fast, rats received an oral glucose bolus of 1 g/kg body weight. Blood samples were taken without anesthesia from the vena saphena just before and at 15, 30, 60, 90 and 120 min after the glucose bolus. Plasma glucose concentration was determined using an automatic glucometer (FreeStyle, Abbott, IL, USA). Plasma insulin concentration was determined using an ultrasensitive rat insulin ELISA kit (Mercodia, Uppsala, Sweden). Areas under the OGTT curves for both glucose (AUC g ) and insulin (AUC i ) were calculated. Magnetic resonance spectroscopy All magnetic resonance spectroscopy (MRS) measurements were performed on a 6.3 Tesla horizontal Bruker MR system (Bruker, Ettlingen, Germany). Animals were anaesthetized using isoflurane (Forene ) (1.5-2%) with medical air (0.6 L/min) and body temperature was maintained at 36 ± 0.5 C using heating pads. Respiration was monitored using a pressure sensor registering thorax movement (Rapid Biomedical, Rimpar, Germany). Intramyocellular lipid (IMCL) content in TA was measured using single-voxel localized 1 H MRS. Voxels of mm 3 were measured in the medial part of the TA, close to the tibia bone, with a circular 1 H surface coil (Ø 40 mm) and using the PRESS sequence (repetition time TR = 1.5 s, echo time TE = 9.4 ms). One spectrum was acquired without water suppression (16 averages) and one with water suppression (VAPOR water suppression, 512 averages). 1 H MR spectra were fit in the time domain using the advanced method for accurate, robust, and efficient spectral fitting (AMARES) in the jmrui software package (jmrui V2.1) 32 as described previously 9. IMCL was expressed as a percentage of the non-suppressed water signal measured in the same voxel. 73

75 Chapter 3 The rate constant of phosphocreatine (PCr) recovery (k PCr ) and creatine kinase flux (V CK ) were measured in the TA by 31 P MRS to determine in vivo mitochondrial function. 31 P MRS was performed using a combination of a circular 1 H surface coil (Ø 40 mm) for shimming and an ellipsoid 31 P MRS surface coil (10/18 mm), positioned over the TA. 31 P MR spectra were acquired applying an adiabatic excitation pulse with a flip angle of 90. A fully relaxed (TR = 25 s, 48 averages) spectrum was measured at rest, followed by the saturation transfer (ST) experiment in resting TA muscle to determine V CK. Two spectra (TR = 10.4 s) were acquired for the ST experiments: A spectrum with frequency selective saturation of the γ-atp peak yielding the steady state PCr magnetization in the presence of saturation ( M PCr, 2*64 averages), and a reference spectrum with saturation at a downfield frequency, equidistant from PCr, yielding the equilibrium PCr magnetization (M 0,PCr, 64 averages). For all experiments the saturation pulse length was set to 10 s. The apparent longitudinal relaxation time of PCr ( T 1, PCr ) was determined by performing a 7-point inversion recovery experiment with an adiabatic full passage pulse for inversion and with γ-atp saturation prior to (10 s) and during the inversion delay (inversion times = 0.01, 1, 2, 4, 6.5, 10.5 and 17 s, 32 averages). After the ST experiments, time series of 31 P MR spectra (TR = 5 s, 4 averages) were acquired during 3 min of rest, 2 min of muscle stimulation and 10 min of recovery, as described previously 33,34. Muscle contractions were induced by electrical stimulation of the TA, via subcutaneously implanted electrodes positioned along the distal N. peroneus communis 33. Stimulation pulse length was 100 ms, frequency was 80 Hz and stimulation voltage varied between 2.5 and 4 V, to reach similar levels of PCr depletion. 31 P MR spectra were fit using AMARES in jmrui as described previously 35. Concentrations of PCr and inorganic phosphate (P i ) were determined relative to the ATP concentration, which was assumed to be 8.2 mm in resting TA muscle 36. Intracellular ph was calculated from the chemical shift difference between the P i and PCr resonances 37. T 1, PCr was determined by fitting the inversion recovery data with a 3-parameter mono-exponential function using Matlab (version R2010b, Mathworks, Natick, MA, USA). The PCr ATP exchange rate constant, k PCr ATP, was calculated according to: k PCr ATP = 1 MPCr M 0,PCr 1,PCr T The creatine kinase flux, V CK, was then calculated by multiplying k PCr ATP with the resting PCr concentration. The data of PCr recovery were fit to a mono-exponential function using Matlab yielding the rate constant of PCr recovery, k PCr. Results from two time series with end-stimulation ph values higher than 6.9 were averaged 38. Measurement of oxygen consumption Mitochondria were isolated from one whole TA muscle through a differential centrifugation procedure as described previously 8,39. Protein content was determined using the BCA protein assay kit (Pierce, Thermo Fisher Scientific Inc., Rockford, IL, USA). Oxygen consumption rates were measured at 37 C using a two-channel high-resolution Oroboros oxygraph-2k (Oroboros, Innsbruck, Austria). Mitochondria were incubated in the assay medium containing 110 mm KCl, 20 mm Tris, 2.3 mm MgCl 2, 5 mm KH 2 PO 4 and 1 mg/ml BSA, ph 7.3. All measurements were performed in 1 ml of assay medium containing 0.15 mg/ml of mitochondrial protein. Three different 74

76 Carnitine insufficiency and mitochondrial function combinations of substrates were used to assess mitochondrial capacity to oxidize tricarboxylic acid cycle (TCA) and β-oxidation substrates: (i) 5 mm pyruvate plus 5 mm malate (TCA cycle), (ii) 25 µm palmitoyl-l-carnitine plus 2.5 mm malate (β-oxidation and TCA cycle), and (iii) 25 µm palmitoyl-coa plus 2.5 mm L-carnitine plus 2.5 mm malate (carnitine palmitoyltransferase 1 (CPT1), β-oxidation and TCA cycle). An ADP-regenerating system consisting of excess hexokinase (4.8 U/ ml) and glucose (12.5 mm) was used to maintain steady-state oxygen consumption rates. Maximal ADP-stimulated oxygen consumption rate, i.e. the OXPHOS state 40, was initiated by addition of 1 mm of ATP. Maximal oxygen consumption rate in the uncoupled state, i.e. the ETS state, was determined after addition of 1 µm carbonyl cyanide 3-chlorophenyl hydrazone (CCCP). The oxygen consumption rate due to proton leak across the mitochondrial inner membrane, i.e. the LEAK state, was measured after fully blocking ATP synthesis with 1.25 µm carboxyatractyloside (CAT). The sensitivity of the basal proton leak rate to FA, which reflects activation of the uncoupling proteins (UCPs) 41, was determined by measuring stimulation of oxygen consumption rate in the LEAK state after addition of 90 µm of palmitic acid (C16:0). The signals from the oxygen electrode were recorded at 0.5 s intervals. Data acquisition and analysis was performed using Oxygraph-2k- DatLab software version 4.2 (Oroboros, Innsbruck, Austria). 3 Determination of the mtdna copy number Genomic DNA was isolated from a ~25 mg transversal slice of mid-belly TA using GenElute Mammalian Genomic DNA Miniprep Kit (Sigma-Aldrich, Zwijndrecht, The Netherlands). The mtdna copy number was assessed by determining the copy number of mitochondrial genome-encoded ATP synthase subunit 6 gene (mt-atp6) relative to a single copy nuclear peroxisome proliferatoractivated receptor-γ coactivator 1α (PGC-1α) gene using quantitative PCR as described in 42. Primer sequences were: mt-atp6 forward 5 -ACACCAAAAGGACGAACCTG-3, mt-atp6 reverse 5 -ATGGGGAAGAAGCCCTAGAA-3, and PGC-1α forward 5 -ATGAATGCAGCGGTCTTAGC-3, PGC-1α reverse 5 -AACAATGGCAGGGTTTGTTC-3. Determination of acylcarnitine content The content of free carnitine and acylcarnitines was determined in freeze-dried TA muscle, blood spots and urine samples, by tandem mass spectrometry as described previously Statistical analysis Data are presented as means ± SD. The listed n values represent the number of animals used for a particular experiment. Statistical significance of the differences was assessed by applying a oneway analysis of variance (ANOVA) using Tukey HSD post-hoc analyses in the SPSS 20.0 statistical package (SPSS Inc., Chicago, IL, USA). The level of statistical significance was set at P<

77 Chapter 3 Results Animal model Animal characteristics are summarized in Table 1. Before the start of the diets, body weight was significantly higher for the HFDC group compared with the HFD group (P<0.05). After 15 weeks of diet, body weight and body weight gain were higher in both HFD and HFDC rats compared with NC controls (P<0.001). However, body weight gain was lower in the HFDC group than in the HFD group (P<0.05). Average energy intake did not differ between groups. After 15 weeks of diet, HFD and HFDC rats had higher fasting plasma glucose (P<0.001) and insulin (P<0.01) levels than NC rats. Moreover, AUC g, AUC i and AUC g *AUC i from the OGTT were significantly higher for both HFD and HFDC rats when compared with NC controls (P<0.01 or P<0.001), while there were no differences between HFD and HFDC groups. Table 1. Animal characteristics. Animal characteristics of Wistar rats after fed for 15 weeks with normal chow (NC), a high-fat diet (HF) and a HF diet plus 8 weeks of carnitine supplementation (HFDC). NC HFD HFDC Body weight (g), t = ± ± ± 12 Body weight (g), t = ± ± 30*** 538 ± 23*** Delta body weight (g) 76 ± ± 37*** 180 ± 21***, Food intake (kj/week) 2247 ± ± ± 187 Fasting glucose (mm) 4.3 ± ± 0.7*** 5.8 ± 0.5*** AUC g (mm h) 9.4 ± ± 1.2*** 12.7 ± 0.7*** Fasting insulin (pm) 230 ± ± 207*** 521 ± 160** AUC i (pm h) 759 ± ± 370** 1534 ± 465*** AUC g *AUC i (mm h pm h) 7005 ± ± 4739** ± 6319*** Data are from n=10 NC, n=9 HFD and n=10 HFDC animals after 15 weeks of diet (except for baseline body weight at t=0) and are expressed as means ± SD. NC, normal chow; HFD, high-fat diet; HFDC, high-fat diet supplemented with carnitine; AUC g and AUC i, area under the glucose and insulin curve from the oral glucose tolerance test, respectively; ** P<0.01, *** P<0.001 when compared with NC, P<0.05 when compared with HFD. 76

78 Carnitine insufficiency and mitochondrial function IMCL content In vivo 1 H MRS was applied to evaluate the effects of 15 weeks of high-fat diet and 15 weeks of high-fat diet in combination with 8 weeks of carnitine supplementation on IMCL levels in TA muscle (Figure 1A). IMCL content was approximately 10-fold higher in HFD and HFDC rats as compared with NC controls (P<0.001; Figure 1B). However, 8 weeks of carnitine supplementation did not affect IMCL content in high-fat diet fed rats. A B IMCL-CH *** *** tcr-ch 3 tcr-ch 2 IMCL-CH 3 HFDC HFD IMCL (% of water signal) NC HFD HFDC NC H chemical shift (ppm) Figure 1. IMCL content assessed by 1 H MRS in TA muscle (A) Representative examples of 1 H MR spectra (512 averages) from the medial part of the TA muscle of a NC, HFD and HFDC rat. (B) IMCL content in the medial part of the TA muscle of NC (n=10), HFD (n=9) and HFDC (n=10) rats. IMCL content was expressed as a percentage of the water signal. *** P<0.001 when compared with NC. 77

79 Chapter 3 Acylcarnitine content For a more detailed characterization of changes in lipid metabolism, we determined the content of different acylcarnitine species in TA muscle, blood and urine (Figures 2 and 3). After 15 weeks of diet, free carnitine (C0) levels were lower in muscle (P<0.001) and blood (P<0.001) of HFD rats compared with NC controls (Figures 2A and 2B). A 1400 Free carnitine and acetylcarnitine (pmol/mg wet weight) * * * * NC HFD HFDC 0 C0 C2 B Free carnitine and acetylcarnitine (mm) * * * * NC HFD HFDC 0 C0 C2 C * Free carnitine and acetylcarnitine (nmol/mmol creatinine) C0 C2 * NC HFD HFDC Figure 2. Free carnitine and acetylcarnitine content in TA muscle (A), blood (B) and urine (C) of NC (n=8 for muscle, n=10 for blood and urine), HFD (n=9 for muscle, blood and urine) and HFDC (n=6 for muscle, n=10 for blood and urine) rats. * P<0.05 when compared with NC, P<0.05 when compared with HFD 78

80 Carnitine insufficiency and mitochondrial function However, carnitine supplementation in rats fed a high-fat diet partially normalized free carnitine in muscle and blood to the level of NC controls. Muscle acetylcarnitine (C2) levels were lower in HFD and HFDC rats (P<0.01), while acetylcarnitine levels in blood were higher in HFD and HFDC rats (P<0.001) compared with NC controls. Moreover, blood acetylcarnitine concentration was 2-fold higher in the HFDC group than in the HFD group (P<0.001). Carnitine supplementation resulted in massive increases in urine free carnitine and acetylcarnitine levels (P<0.001; Figure 2C). Medium- and long-chain acylcarnitine species are almost absent in urine and are therefore not shown. Muscle medium- and long-chain acylcarnitine levels were significantly elevated in HFD and HFDC rats compared with NC controls (P<0.05), but did not differ between HFD and HFDC groups (Figures 3A and 3B). Most medium- and long-chain acylcarnitines in blood were lowered after 15 weeks of high-fat diet (P<0.05), but they were normalized to the level of NC controls upon carnitine supplementation (Figures 3C and 3D). 3 A 1.4 NC HFD HFDC B 30 Acylcarnitine (pmol/mg wet weight) * * * * * * * * Acylcarnitine (pmol/mg wet weight) * * * * * * * * * * 0.0 C4 C6 C8 C10 C12 0 C14 C16 C18 C16:1 C18:1 C D 1.0 NC HFD * HFDC * * * Acylcarnitine (mm) * Acylcarnitine (mm) 2 * * * * * * * * C4 C6 C8 C10 C * * * * C14 C16 C18 C16:1 C18:1 Figure 3. Short- and medium-chain acylcarnitine content in TA muscle (A) and blood (C), and long-chain acylcarnitine content in TA muscle (B) and blood (D) of NC (n=8 for muscle, n=10 for blood), HFD (n=9 for muscle and blood) and HFDC (n=6 for muscle, n=10 for blood) rats. * P<0.05 when compared with NC, P<0.05 when compared with HFD. 79

81 Chapter 3 Mitochondrial function in vivo In vivo 31 P MRS was used to assess in vivo muscle mitochondrial function. Concentrations of metabolites and ph obtained from the 31 P MR spectra in resting TA muscle are summarized in Table 2. Resting ph was slightly lower in HFD (P<0.001) and HFDC (P<0.01) rats compared with NC controls, but concentrations of PCr and P i did not differ among groups. Table 2. Metabolite concentrations and ph in TA measured by in vivo 31 P MRS in Wistar rats after fed for 15 weeks with normal chow (NC), a high-fat diet (HF) and a HF diet plus 8 weeks of carnitine supplementation (HFDC). NC HFD HFDC Rest ph 7.21 ± ± 0.02*** 7.19 ± 0.01** [PCr] (mm) 32.2 ± ± ± 2.9 [P i ] (mm) 2.6 ± ± ± 0.3 End-stimulation ph 7.00 ± ± ± 0.04 [PCr] (mm) 13.8 ± ± 2.1* 15.7 ± 1.4 [P i ] (mm) 20.1 ± ± ± 1.3 PCr (%) 59.8 ± ± 4.0* 55.5 ± 2.9 Data are from n=10 NC, n=9 HFD and n=10 HFDC animals and are expressed as means ± SD. NC, normal chow; HFD, high-fat diet; HFDC, high-fat diet supplemented with carnitine; PCr, phosphocreatine; P i, inorganic phosphate. * P<0.05, ** P<0.01, *** P<0.001 when compared with NC. From the 31 P ST spectra (Figure 4A), the ratio of PCr magnetization with and without selective saturation of γ-atp ( MPCr M0, PCr ) was determined, which did not differ between groups (Table 3). The apparent longitudinal relaxation time of PCr in the presence of γ-atp saturation ( T 1, PCr ) determined from the 7-point inversion recovery experiment was higher in HFD and HFDC rats than in NC controls (P<0.001; Table 3). However the PCr ATP exchange rate constant, k PCr ATP, and the creatine kinase flux, V CK, did not differ between NC, HFD, and HFDC groups (Table 3). A B Figure 4. Representative examples of 31 P MR spectra from TA muscle of a NC rat (A) 31 P saturation transfer spectra (saturation pulse length = 10 s, TR = 10.4 s, 64 averages) with saturation on g-atp (black) and with saturation at a downfield frequency, equidistant from PCr (grey). (B) 31 P MR spectrum in resting muscle (black; TR = 5 s, 32 averages) and a dynamic representation of the PCr peak during muscle stimulation and recovery (grey; TR = 5 s, 4 averages). 80

82 Carnitine insufficiency and mitochondrial function Table 3. Parameters determined from 31 P saturation transfer MRS in Wistar rats after fed for 15 weeks with normal chow (NC), a high-fat diet (HF) and a HF diet plus 8 weeks of carnitine supplementation (HFDC). NC HFD HFDC M PCr / M 0,PCr 0.56 ± ± ± 0.02 T 1,PCr (s) 1.95 ± ± 0.08*** 2.08 ± 0.07*** k PCr ATP (min -1 ) 13.6 ± ± ± 0.8 V CK (mm/min) 438 ± ± ± 35 Data are from n=10 NC, n=9 HFD and n=9 HFDC animals and are expressed as means ± SD. NC, normal chow; HFD, high-fat diet; HFDC, high-fat diet supplemented with carnitine; M PCr, magnetization of PCr with saturation of γ-atp; M 0,PCr, magnetization of PCr with saturation at a downfield frequency, equidistant from PCr; T 1, PCr, apparent longitudinal relaxation time of PCr with saturation of γ-atp; k PCr ATP, PCr ATP exchange rate constant; V CK, resting creatine kinase flux. *** P<0.001 when compared with NC. 3 Dynamic 31 P MRS measurements were performed during and after recovery from electrical stimulation of the TA muscle to determine the rate constant of PCr recovery, k PCr, after contractions (Figure 4B). Concentrations of PCr and P i and the intracellular ph at the end of stimulation were not different between groups, with the exception of the end-stimulation concentration of PCr in HFD rats, which was slightly higher than in NC controls (P<0.05; Table 2). However, k PCr did not differ among NC, HFD, and HFDC groups (Figure 5A). A B mtdna copy no (-) Figure 5. Rate constant of PCr recovery (k PCr ) after muscle stimulation (A) and relative mitochondrial DNA (mtdna) copy number (B) measured in TA muscle of NC (n=7 for k PCr, n=8 for mtdna), HFD (n=9 for k PCr and mtdna) and HFDC (n=10 for k PCr and mtdna) rats. * P<0.05, ** P<0.01 when compared with NC. 81

83 Chapter 3 Mitochondrial function ex vivo Table 4 summarizes the effects of 15 weeks of high-fat diet feeding and 15 weeks of high-fat diet feeding in combination with 8 weeks of carnitine supplementation on intrinsic mitochondrial function ex vivo, represented by oxygen consumption rates in isolated TA mitochondria oxidizing glucose- or fat-derived substrates in different respiratory states. Both high-fat diet feeding and high-fat diet feeding in combination with carnitine supplementation had no effect on pyruvate plus malate-driven oxygen consumption rates in the OXPHOS and ETS states, although oxygen consumption was slightly higher in the LEAK state of HFDC rats compared with NC controls (P<0.001). Interestingly, palmitoyl-l-carnitine plus malate driven oxygen consumption rate in the OXPHOS and ETS states was (or tended to be) ~15% higher in mitochondria from HFD and HFDC rats compared with NC controls (P<0.05 or 0.05 P<0.1). A similar effect was observed when palmitoyl-coa plus L-carnitine plus malate was used as the oxidizible substrate. However, respiratory capacity in the OXPHOS and ETS states did not differ between HFD and HFDC groups for either palmitoyl-l-carnitine plus malate or palmitoyl-coa plus L-carnitine plus malate. Oxygen consumption in the LEAK state was similar in all three groups for both fat-derived substrates. Table 4. Oxygen consumption rates in isolated TA mitochondria oxidizing different substrates in different metabolic states. Pyruvate plus malate NC HFD HFDC OXPHOS (nmol O 2 min -1 mg protein -1 ) 533 ± ± ± 74 LEAK (nmol O 2 min -1 mg protein -1 ) 30 ± 2 34 ± 3 38 ± 5*** ETS (nmol O 2 min -1 mg protein -1 ) 644 ± ± ± 85 Pyruvate plus malate plus palmitic acid LEAK (nmol O 2 min -1 mg protein -1 ) 195 ± ± 22** 237 ± 56 # Palmitoylcarnitine plus malate OXPHOS (nmol O 2 min -1 mg protein -1 ) 147 ± ± 20* 166 ± 20 # LEAK (nmol O 2 min -1 mg protein -1 ) 32 ± 1 31 ± 2 32 ± 3 ETS (nmol O 2 min -1 mg protein -1 ) 225 ± ± 22 # 254 ± 29* PalmitoylCoA plus L-carnitine plus malate OXPHOS (nmol O 2 min -1 mg protein -1 ) 146 ± ± 14 # 167 ± 26 # LEAK (nmol O 2 min -1 mg protein -1 ) 31 ± 1 31 ± 2 31 ± 5 ETS (nmol O 2 min -1 mg protein -1 ) 232 ± ± 15* 263 ± 31* Data are from n=10 NC, n=9 HFD and n=10 HFDC animals and are expressed as means ± SD. NC, normal chow; HFD, highfat diet; HFDC, high-fat diet supplemented with carnitine; OXPHOS, maximal ADP-stimulated oxygen consumption; LEAK, oxygen consumption in the absence of ATP synthesis; ETS, oxygen consumption after uncoupling. * P<0.05, ** P<0.01, *** P<0.001 when compared with NC, # 0.05 P<0.1 (trend) when compared with NC. 82

84 Carnitine insufficiency and mitochondrial function Table 4 also shows the effect of 90 µm of palmitic acid on the oxygen consumption rate in the LEAK state in isolated mitochondria respiring on pyruvate plus malate. Addition of palmitic acid caused a significantly larger increase in oxygen consumption in the HFD group than in the NC group (P<0.01), indicating increased FA-induced mitochondrial uncoupling. The effect of palmitic acid in the HFDC group was similar to that in the HFD group, but it only tended to differ from the effect in the NC group (P=0.074). The effect of 15 weeks of high-fat diet feeding and 15 weeks of high-fat diet feeding in combination with 8 weeks of carnitine supplementation on mitochondrial biogenesis was assessed by determining mtdna copy number in TA muscle. Relative mtdna copy number was significantly higher in HFD (P<0.01) and HFDC (P<0.05) rats compared with NC controls, whereas no difference was observed between HFD and HFDC groups (Figure 5B). 3 83

85 Chapter 3 Discussion We aimed to determine whether carnitine supplementation reduces lipotoxicity, improves in vivo muscle mitochondrial function and ameliorates insulin resistance in high-fat diet fed rats. Fifteen weeks of high-fat diet feeding in rats resulted in insulin resistance. In agreement with previous findings 5,6,8,11,12,46, this was associated with decreases in muscle and blood free carnitine, increased IMCL and muscle acylcarnitine levels, an increased capacity to oxidize fat-derived substrates ex vivo, and an elevated number of muscle mitochondria. The increase in mitochondrial content was however not accompanied by an increase in muscle oxidative capacity in vivo, suggesting that under in vivo conditions the function of individual mitochondria was compromised 8. Despite the partial normalization of muscle free carnitine content, carnitine supplementation in high-fat diet fed rats did not induce improvements in muscle lipid status, in vivo mitochondrial function or insulin sensitivity. Insulin resistance has previously been linked to an increased capacity to oxidize FAs in order to cope with the high lipid loads 5-9. The results of our ex vivo measurements, showing upregulation of FA oxidation capacity in isolated mitochondria of HFD rats, are in line with these findings. Our results are also in agreement with other studies which have shown that high-fat diet feeding does not affect respiration of rat muscle mitochondria when using substrates other than fatty acids, i.e. pyruvate/glutamate plus malate or succinate 6,47,48, implying that the intrinsic functioning of the mitochondria is not impaired when probed ex vivo. The observed 30% increase in mtdna copy number in HFD rats is in agreement with our previous study, in which it was shown that long-term high-fat diet feeding causes an increase in PGC-1α expression with a concomitant increase in mtdna copy number and citrate synthase activity compared with normal chow fed controls 8. Furthermore, a number of other studies have likewise shown that high-fat diet feeding in rats induces increased biogenesis of mitochondria 5,6,46,49-51 as an adaptive response to the higher mitochondrial FA load. However, despite the 30% increase in mitochondrial content, the in vivo PCr recovery rate constant (k PCr ) was similar for HFD and NC rats. These data indicate that an increased number of mitochondria with normal or even improved function ex vivo is required to maintain normal muscle oxidative capacity in vivo in HFD rats 8. To exclude that k PCr in HFD rats was affected by impairments in the creatine kinase system, we determined the creatine kinase flux (V CK ) from 31 P ST experiments and we showed that V CK was similar for NC, HFD and HFDC rats. In a previous study, V CK was found to be increased in skeletal muscle of high-fat diet fed rats compared with controls, which was explained as a compensatory mechanism in response to impaired mitochondrial function 52. However, this explanation seems unlikely, because in normal conditions V CK is already much higher than oxidative ATP synthesis flux 53. HFD rats had a 12-fold higher IMCL content compared with NC controls. The acylcarnitine profile likewise showed that muscle of HFD rats was overloaded with lipids. The vast majority of acylcarnitines is produced in the mitochondria, and acylcarnitine levels can therefore, in combination with measurements of substrate oxidation and mitochondrial function, be interpreted as a measure of β-oxidation flux. In agreement with previous studies, even, medium-chain (C6- C12) acylcarnitine intermediates, which represent incompletely oxidized FAs, were elevated in muscle of HFD rats compared with NC controls Although the measurement of acylcarnitines provides a comprehensive snapshot of intermediary metabolism, it is important to emphasize that steady-state metabolite concentrations represent the net balance between production, 84

86 Carnitine insufficiency and mitochondrial function consumption, import and export. However, the combination of increased ex vivo FA oxidation capacity and increased levels of medium-chain acylcarnitines in muscle of HFD animals, suggests that incomplete β-oxidation is elevated in HFD animals. It has been shown that elevated levels of FAs may impair mitochondrial ATP production through a number of mechanisms. Long-chain acyl-coa esters may inhibit the mitochondrial adenine nucleotide translocator leading to impaired exchange of cytosolic ADP for mitochondrial ATP 54. Moreover, increased availability of FAs without a concomitant rise in energy demand could lead to increased expression 6,8 as well as activation of UCP3 41, which in turn may diminish the efficiency of ATP synthesis by increasing mitochondrial uncoupling 55. In order to test the latter mechanism, we measured LEAK state oxygen consumption rates in isolated mitochondria respiring on pyruvate plus malate in the presence of palmitic acid. In this experiment, we observed significantly higher oxygen consumption rates in mitochondria isolated from HFD rats compared with NC rats, indicating increased mitochondrial uncoupling due to the presence of FAs. Together with the finding that muscle medium- and long-chain acylcarnitines were increased in HFD rats compared with NC controls, these data strongly suggest that FA-induced mitochondrial uncoupling may contribute to the observed mitochondrial functional impairment in muscle of HFD rats in vivo, similar to our earlier findings in diabetic and long-term high-fat diet fed rats 8,9. 3 In parallel with the increase in muscle acylcarnitine levels, a decrease was observed in free carnitine levels in muscle and blood of HFD animals. It has been shown that whole-body carnitine insufficiency is a common feature in insulin-resistant states such as advanced age, genetic diabetes and dietinduced obesity 12. This can be explained by a decreased biosynthesis in the liver 11,12,56,57, but also by increased sequestration of carnitine in the muscle acylcarnitine pool 11,12. The insulin resistance related decline in free carnitine has been associated with impaired mitochondrial function and an imbalance between complete and incomplete fat oxidation 12,19. Most direct evidence linking carnitine insufficiency to mitochondrial dysfunction comes from juvenile visceral steatosis (jvs) mice, a genetic model of carnitine deficiency, showing marked derangements in mitochondrial biogenesis and morphology, as well as whole-body lipid and energy metabolism 58,59. It has been hypothesized that carnitine supplementation ameliorates mitochondrial function and insulin resistance by reducing lipotoxicity both through the increased oxidation and increased export of muscle lipid metabolites 19. In a study by Noland et al., it was shown that 8 weeks of carnitine supplementation in long-term high-fat diet fed rats restored the ratio of complete to incomplete fat oxidation and increased efflux of muscle acylcarnitine intermediates, while improving glucose tolerance 12. To study the effects of carnitine supplementation on muscle lipid status, muscle mitochondrial function and insulin sensitivity in high-fat diet fed rats, we supplemented them with 300 mg/ kg body weight/day carnitine during the last 8 weeks of the 15 weeks diet. Although carnitine supplementation partially normalized free carnitine in muscle and blood to the level of NC controls, no differences in IMCL levels were observed between HFD and HFDC rats. Similar levels of muscle acylcarnitines in HFD and HFDC rats provided further evidence of unchanged lipid levels inside the muscle, despite the increase in acylcarnitines levels in blood upon carnitine supplementation. The effects of carnitine supplementation on acylcarnitine profiles in muscle, blood and urine are in agreement with the results of Noland et al. 12, with the exception of free carnitine in muscle and blood, which was completely restored to the level of normal chow fed controls in the study of Noland et al. whereas it was only partially normalized in the current study. 85

87 Chapter 3 In line with the lack of effects on muscle lipid status, carnitine supplementation also did not affect muscle mitochondrial function or insulin sensitivity in high-fat diet fed rats. Oxygen consumption rates of muscle mitochondria respiring on glucose- or fat-derived substrates were similar for HFD and HFDC rats. Addition of palmitic acid had the same effect on mitochondria from HFD and HFDC rats, both showing increased FA-induced mitochondrial uncoupling compared with mitochondria from NC controls. Both in HFD and HFDC rats muscle mitochondrial content was about 30% higher than in NC controls, but in both high-fat diet fed groups this did not result in an increased muscle oxidative capacity in vivo as determined from k PCr, indicating that in vivo mitochondrial function was similarly impaired. Our results on muscle mitochondrial function and insulin sensitivity are in contrast with the findings of Noland et al., who showed that exactly the same regimen of carnitine supplementation (i.e. 300 mg/kg body weight/day during 8 weeks) improved whole-body glucose tolerance and reversed mitochondrial abnormalities in rats fed with a high-fat diet 12. A difference between the two studies is the duration of the high-fat diet, i.e. 15 weeks in the current study versus 12 months in the study by Noland et al. However, the 15 weeks and 12 months of high-fat diet feeding led to similar decreases in muscle free carnitine and comparable increases in muscle acylcarnitines. Another difference between the two studies concerns the methods to assess muscle mitochondrial function. Noland et al. measured the ratio of complete to incomplete oleate oxidation in isolated mitochondria, which was decreased in high-fat diet fed rats, but completely restored after carnitine supplementation 12. In the current study we have no data on the ratio of complete to incomplete FA oxidation, but we have shown that carnitine supplementation does not improve in vivo muscle mitochondrial function. Reduction of the free carnitine pool in response to high-fat diet feeding might have a negative effect on mitochondrial FA oxidation only if free carnitine is depleted to an extent that it becomes limiting for CPT1 activity, resulting in a decreased entry of long-chain acyl-coas into the mitochondrial matrix. The absence of an effect of 8 weeks of carnitine supplementation in high-fat diet fed animals regarding muscle mitochondrial function suggests that this was not the case in the present study. Moreover, our observation that OXPHOS state oxygen consumption rates with palmitoyl-coa plus L-carnitine plus malate and palmitoyl-l-carnitine plus malate as the oxidizable substrates were similar in HFD and HFDC groups suggests that neither palmitoyl-coa transport nor β-oxidation capacity was limited by carnitine insufficiency in high-fat diet fed rats. In conclusion, we showed that high-fat diet feeding induces carnitine insufficiency and muscle lipid overload, which was accompanied by an increased number of muscle mitochondria with an improved capacity to oxidize FAs ex vivo, but without a concomitant increase in muscle oxidative capacity in vivo. We provided evidence that this impairment in in vivo mitochondrial function in high-fat diet fed rats is caused by elevated levels of lipid intermediates, leading to increased FAinduced mitochondrial uncoupling and therefore less efficient ATP synthesis. Despite a partial normalization of free carnitine in muscle and blood, carnitine supplementation did not induce improvements in muscle lipid status, in vivo mitochondrial function or insulin sensitivity. These results suggest that carnitine insufficiency does not play a major role in high-fat diet induced muscle mitochondrial dysfunction in vivo. 86

88 Carnitine insufficiency and mitochondrial function Acknowledgements We are grateful to sigma-tau (Utrecht, the Netherlands) for providing L-carnitine. The work of J.C. is financed by the Netherlands Consortium for Systems Biology (NCSB) which is part of the Netherlands Genomics Initiative / Netherlands Organization for Scientific Research. J.J.P. and S.M.H. are supported by VIDI grants from the Netherlands Organization for Scientific Research (VIDI grant numbers and , respectively). 3 87

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91 Chapter Nemeth, P.M., Rosser, B.W., Choksi, R.M., Norris, B.J. & Baker, K.M. Metabolic response to a high-fat diet in neonatal and adult rat muscle. Am J Physiol 262, C (1992). 52. Chanseaume, E., et al. Impaired resting muscle energetics studied by 31 P-NMR in diet-induced obese rats. Obesity (Silver Spring) 16, (2008). 53. Vicini, P. & Kushmerick, M.J. Cellular energetics analysis by a mathematical model of energy balance: estimation of parameters in human skeletal muscle. Am J Physiol Cell Physiol 279, C (2000). 54. Lerner, E., Shug, A.L., Elson, C. & Shrago, E. Reversible inhibition of adenine nucleotide translocation by long chain fatty acyl coenzyme A esters in liver mitochondria of diabetic and hibernating animals. J Biol Chem 247, (1972). 55. Brand, M.D. Uncoupling to survive? The role of mitochondrial inefficiency in ageing. Experimental gerontology 35, (2000). 56. Mogensen, M., et al. Mitochondrial respiration is decreased in skeletal muscle of patients with type 2 diabetes. Diabetes 56, (2007). 57. Mikhail, M.M. & Mansour, M.M. The relationship between serum carnitine levels and the nutritional status of patients with schistosomiasis. Clin Chim Acta 71, (1976). 58. Kaido, M., et al. Mitochondrial abnormalities in a murine model of primary carnitine deficiency. Systemic pathology and trial of replacement therapy. Eur Neurol 38, (1997). 59. Tein, I. Carnitine transport: pathophysiology and metabolism of known molecular defects. J Inherit Metab Dis 26, (2003). 90

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93 Pioglitazone treatment restores in vivo muscle oxidative capacity in a rat model of diabetes Bart Wessels 1, Jolita Ciapaite 2, Nicole M.A. van den Broek 1, Sander M. Houten 3,*, Klaas Nicolay 1 and Jeanine J. Prompers 1 1 Biomedical NMR, Department of Biomedical Engineering, Eindhoven University of Technology, the Netherlands, 2 Center for Liver, Digestive and Metabolic Diseases, Department of Pediatrics, University of Groningen, University Medical Center Groningen, Groningen, the Netherlands, 3 Laboratory Genetic Metabolic Diseases, Departments of Pediatrics and Clinical Chemistry, Academic Medical Center, Amsterdam, the Netherlands, *Current affiliation: Department of Genetics and Genomic Sciences, Icahn Institute for Genomics and Multiscale Biology, Icahn School of Medicine at Mount Sinai, New York, USA Published in Diabetes, obesity and metabolism (2014) doi: /dom Epub 2014 Oct 6.

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96 Effect of pioglitazone on muscle mitochondrial function Abstract Aim: Pioglitazone improves peripheral insulin sensitivity in diabetes patients by redistributing lipids from muscle into adipose tissue. Mitochondria are thought to play a crucial role in the etiology of muscle insulin resistance. We aimed to determine the effect of pioglitazone treatment on in vivo and ex vivo muscle mitochondrial function. Materials and Methods: Lean, healthy and obese, diabetic Zucker diabetic fatty rats were treated with either pioglitazone (30 mg/kg/day) or water as a control (n=6 per group), for 2 weeks. In vivo 1 H and 31 P magnetic resonance spectroscopy were performed on skeletal muscle to assess intramyocellular lipids (IMCL) and muscle oxidative capacity, respectively. Ex vivo muscle mitochondrial respiratory capacity was evaluated using high-resolution respirometry. In addition, several markers of mitochondrial content were determined. Results: IMCL content was 14-fold higher and in vivo muscle oxidative capacity was 26% lower in diabetic rats compared with lean rats, which was however not caused by impairments of ex vivo mitochondrial respiratory capacity or a lower mitochondrial content. Pioglitazone treatment restored in vivo muscle oxidative capacity in diabetic rats to the level of lean controls. This amelioration was not accompanied by an increase in mitochondrial content or ex vivo mitochondrial respiratory capacity, but rather was paralleled by an improvement in lipid homeostasis, i.e. lowering of plasma triglycerides and muscle lipid and long-chain acylcarnitine content. Conclusion: Diminished in vivo muscle oxidative capacity in diabetic rats results from mitochondrial lipid overload and can be alleviated by redirecting the lipids from the muscle into adipose tissue using pioglitazone treatment. 4 95

97 Chapter 4 Introduction Thiazolidinediones (TZDs) improve whole-body insulin sensitivity and blood glucose levels in patients with type 2 diabetes. TZDs activate the transcription factor peroxisome proliferatoractivated receptor-γ (PPAR-γ), which is predominantly expressed in adipose tissue 1,2. This leads to upregulation of genes encoding for mitochondrial proteins and enhanced fatty acid (FA) uptake and oxidation in adipose tissue 3-6, and, as a result, relocation of fat from ectopic sites, such as skeletal muscle, into adipose tissue 5. Moreover, TZDs stimulate insulin-stimulated glucose uptake in skeletal muscle 7-9. Skeletal muscle is the primary site for postprandial, insulin-stimulated glucose disposal. A strong association has been established between insulin resistance and the accumulation of intramyocellular lipids (IMCL) The mechanistic link between IMCL overload and insulin resistance is believed to reside in the accumulation of lipid derived intermediates, such as diacylglycerols and ceramides, leading to impaired insulin signaling 15. Mitochondrial dysfunction and a resulting impairment in FA oxidation have been suggested as causative factors in the excess storage of lipids in insulin-resistant muscle 16. However, the nature of mitochondrial dysfunction and the cause and effect relationship between mitochondrial dysfunction and the development of IMCL accretion and insulin resistance remain elusive In fact, recent studies have linked insulin resistance to an increased rather than a decreased capacity to oxidize FAs Oxidation of FA substrates is a source of reactive oxygen species (ROS) and promotes mitochondrial uncoupling, which could lead to compromised mitochondrial ATP production 18,19,23. Moreover, it has been shown that in insulin-resistant muscle metabolic intermediates of incomplete FA oxidation accumulate 24, indicating that FA oxidation flux outpaces the demand of the respiratory system, which may affect mitochondrial redox state and further promote mitochondrial oxidative stress 25,26. A number of studies have shown that TZDs, in particular pioglitazone, reduce IMCL content in insulin-resistant states 9,27,28, which is the most likely mechanism for TZDs to improve peripheral insulin sensitivity. However, the effects of TZDs on mitochondrial function in skeletal muscle are less clear. TZDs have been reported to inhibit Complex I of the mitochondrial electron transport chain and, consequently, treatment with TZDs may induce mitochondrial dysfunction However, it has also been suggested that TZDs may improve muscle mitochondrial function in patients with type 2 diabetes 32,33. Expression of PPAR-γ coactivator 1α (PGC-1α) and PPAR-β/δ were found to be upregulated in skeletal muscle of type 2 diabetes patients after treatment with rosiglitazone 34, suggesting that TZDs have a direct effect on mitochondrial gene expression, not only in adipose tissue but also in skeletal muscle. Alternatively, the stimulating effect of TZDs on PGC-1α expression could be mediated by circulating adiponectin 35, which has been shown to increase upon treatment with TZDs 36. Moreover, the lipid-lowering effect of TZDs could potentially improve muscle mitochondrial function in insulin-resistant states through a decreased formation of ROS and a reduced accumulation of FA-derived metabolites. Therefore, the effect of TZDs on muscle mitochondrial function and the underlying mechanism warrant further investigation. The aim of this study was to determine the effect of pioglitazone treatment on in vivo muscle mitochondrial function in a rat model of diabetes. The readout of in vivo mitochondrial function was complemented by measurements of ex vivo mitochondrial function and content, to investigate if pioglitazone affects intrinsic mitochondrial properties. Lean and diabetic Zucker diabetic fatty (ZDF) rats were dosed orally with pioglitazone (30 mg/kg body weight/day) for 2 weeks. In vivo muscle mitochondrial function was assessed using 31 P magnetic resonance spectroscopy (MRS). 96

98 Effect of pioglitazone on muscle mitochondrial function Ex vivo mitochondrial respiratory capacity was determined using high-resolution respirometry of isolated muscle mitochondria with tricarboxylic acid (TCA) cycle and FA oxidation substrates. Additionally, IMCL content was determined by 1 H MRS. 4 97

99 Chapter 4 Materials and methods Animals Male lean, healthy fa/+ and obese, diabetic fa/fa ZDF rats (12 weeks of age; Charles River Laboratories, Sulzfield, Germany) were housed in pairs in a controlled environment (20 C and 50% relative humidity, 12-h light-dark cycle) and given ad libitum access to water and standardized chow for ZDF rats (19% calories from fat, 54% calories from carbohydrates and 27% calories from protein; Purina Formula 5008, Bioservices, the Netherlands). Animals were dosed for 15 days via oral gavage with pioglitazone (30 mg/kg body weight/day ) in water (pioglitazone; n=6 per genotype), or only water as a control (water; n=6 per genotype). Dosing was performed once daily between 4 and 6 pm. In vivo MRS experiments were performed at day 15 before pioglitazone dosing (two animals of one group per day). Following the MRS measurements, animals were administered with the last dose of pioglitazone. The following day, between 8 and 10 am, the animals were sacrificed under anesthesia by incision of the vena cava. One tibialis anterior (TA) muscle (fresh) was used for isolation of mitochondria. The other TA was frozen in liquid nitrogen and stored at -80 C. All experimental procedures were reviewed and approved by the Animal Experimental Committee of Maastricht University. Plasma parameters After 13 days of therapy, a blood sample was taken in conscious animals from the vena saphena after a 4 hour fast. Plasma glucose concentrations were determined using an automatic glucometer (Freestyle, Abbott, IL, USA). Plasma insulin concentrations were determined with an ultrasensitive rat insulin ELISA kit (Mercodia, Uppsala, Sweden). Free fatty acids (FFA) and triglyceride (TG) concentrations were determined in plasma samples taken in anesthetized animals in the fed state after 15 days of therapy, using a NEFA-HR(2) kit (Wako chemicals, Neuss, Germany) and a serum triglyceride determination kit (Sigma-Aldrich, Zwijndrecht, the Netherlands), respectively. Magnetic Resonance Spectroscopy MRS measurements were performed on a 6.3-T horizontal Bruker MR scanner (Bruker, Ettlingen, Germany) using an ellipsoid (10/18 mm) 31 P coil combined with a circular (40 mm diameter) 1 H surface coil, which were positioned over the TA. The animals were anaesthetized using isoflurane (2-3%). Intramyocellular lipid (IMCL) content was determined in the TA muscle using localized 1 H MRS. A voxel of 3x3x3 mm 3 was selected in the medial part of the TA, close to the tibia bone. 1 H MR spectra were acquired with a PRESS sequence (repetition time = 1.5 s, echo time = 9.4 ms, VAPOR water suppression, 256 averages). Unsuppressed water spectra (16 averages) were recorded in the same voxel and were used as internal reference. MR spectra were fit in the time domain using a nonlinear least squares algorithm (advanced method for accurate, robust, and efficient spectral fitting; AMARES) in the jmrui software package 40. In the water-suppressed 1 H MR spectra, the central peak of the tcr-ch 3 triplet, the IMCL-CH 2 peak (1.28 ppm) and the extramyocellular lipid (EMCL)-CH 2 peak were fitted to Gaussian line shapes. The tcr and IMCL linewidths were constrained as described previously 41. IMCL content was expressed relative to the water signal measured in the same voxel without water suppression, which was fitted to a Lorentzian line shape. 98

100 Effect of pioglitazone on muscle mitochondrial function 31 P MRS was applied to assess in vivo oxidative capacity of the TA muscle, as described previously 42,43. A fully relaxed spectrum (repetition time = 20 s, 32 averages) was recorded first, followed by a time series of spectra (repetition time = 5 s, 4 averages) obtained during a resting period of 3 min, 2 min of electrical stimulation and 10 min of recovery. The TA muscle was stimulated via subcutaneously implanted electrodes positioned along the distal nerve trajectory of the N. peroneus communis. The stimulation pulse length was 100 ms with a frequency of 80 Hz. Pulses were applied every second and a stimulation voltage of approximately 3-4 V was used to reach similar levels of phosphocreatine (PCr) depletion for the different animals. The 31 P MR measurements were gated to the stimulation pulses. MR spectra were fitted in the time domain using a nonlinear least squares algorithm (advanced method for accurate, robust, and efficient spectral fitting; AMARES) in the jmrui software package 40 as described previously 42. In short, spectral analysis of the 31 P MR spectra was done by fitting the PCr peak to Lorentzian and the inorganic phosphate (P i ) as well as the α-, β- and γ-atp peaks to Gaussian line shapes. Intracellular ph was calculated from the chemical shift difference between the Pi and PCr resonances 44. For the time series, the concentrations of PCr determined during recovery were fit to a mono-exponential function using Matlab (version R2010b, Mathworks, Natick, MA, USA) yielding a rate constant, k PCr, which is a measure of in vivo muscle mitochondrial oxidative capacity. For each rat, results from two time series with end-stimulation ph values higher than 6.9 were averaged High-resolution respirometry Mitochondria were isolated from one whole TA muscle using a differential centrifugation procedure as described previously 22. Protein content was determined using a BCA protein assay kit (Pierce, Thermo Fisher Scientific Inc., Rockfort, IL, USA). Intrinsic mitochondrial function was evaluated ex vivo by measuring O 2 consumption rates (O 2 flux) at 37 C using a 2-channel high-resolution oxygraph-2k (Oroboros, Innsbruck, Austria). Isolated mitochondria (0.15 mg/ml) were incubated in 1 ml of assay medium (110 mm KCl, 20 mm Tris, 2.3 mm MgCl 2, 5 mm KH 2 PO 4 and 1 mg/ml BSA, ph=7.2 at 37 C). O 2 flux was fueled with two TCA cycle substrates: i) 5 mm pyruvate plus 5 mm malate (Complex I-dependent respiration), and ii) 5 mm succinate plus 1 µm rotenone (Complex II-dependent respiration), and a substrate fueling β-oxidation: 25 µm palmitoyl-l-carnitine plus 2.5 mm malate. The maximal rate of oxygen consumption coupled to ATP synthesis, i.e. the OXPHOS state 46, was determined after adding an ADP-regenerating system consisting of excess hexokinase (4.8 U/ml), glucose (12.5 mm) and ATP (1 mm). Oxygen consumption rate due to proton leak across the mitochondrial inner membrane, i.e. the LEAK state, was assessed after addition of 1.25 µm carboxyatractyloside (CAT). The maximal capacity of the electron transfer system (ETS), i.e. the ETS state, was determined by uncoupling the ETS from ATP synthesis with the addition of 1 µm carbonyl cyanide 3-chlorophenyl hydrazone (CCCP). The signals from the oxygen electrodes were recorded at 0.5-s intervals and all respiration measurements were performed in duplicate. Data acquisition and analysis was performed using Oxygraph-2k-Datlab software (Oroboros, Innsbruck, Austria). Determination of muscle acylcarnitine content The content of acylcarnitines was determined in freeze-dried TA muscle by tandem mass spectrometry as described previously

101 Chapter 4 Determination of markers of muscle mitochondrial biogenesis and content Determination of PGC-1α protein expression by immunoblotting Equal amounts (15 µg) of total TA protein were resolved with SDS-PAGE (10% gel) and transferred to nitrocellulose membranes using Trans-Blot Turbo Midi Nitrocellulose Transfer Packs, and Trans-Blot Turbo Transfer Starter System (Bio-Rad Laboratories Inc., Hercules, CA, USA). After blocking with TBS containing 0.1% Tween (TBST) and 5% skim milk powder for 1 h at room temperature, the membranes were incubated overnight at 4 C with rabbit polyclonal anti-pgc- 1α antibody (1:1000; cat. no. sc-13067, Santa Cruz Biotechnology, Santa Cruz, CA, USA) or mouse monoclonal anti-glyceraldehyde 3-phosphate dehydrogenase (GAPDH) antibody (1:5000, cat. no. CB1001, Calbiochem, Espoo, Finland). Next, membranes were washed 3 5 min with TBST and incubated with a horse-radish peroxidase-conjugated goat anti-rabbit secondary antibody for 1 h at room temperature. After the final wash of 3 5 min with TBST and 1 5 min with TBS, the immunocomplexes were detected using SuperSignal West Dura Extended Duration Substrate (Pierce, Thermo Fisher Scientific Inc., Rockford, IL, USA), visualized using ChemiDoc XRS+ imaging system and quantified using Image Lab analysis software version 3.0 (Bio-Rad Laboratories Inc., Hercules, CA, USA). All data were normalized to GAPDH expression level and expressed relative to the lean water-treated controls. Determination of mtdna copy number Genomic DNA was isolated from ~20 mg of TA tissue using GenElute Mammalian Genomic DNA Miniprep Kit (Sigma-Aldrich, Zwijndrecht, The Netherlands). The mtdna copy number was assessed by determining the copy number of mitochondrial genome-encoded ATP synthase subunit 6 gene (mt-atp6) relative to a single copy nuclear PGC-1α gene using quantitative PCR as described in 48. Real-time PCR was performed in MicroAmp optical 96-well plates using StepOne Real-Time PCR system (Applied Biosystems). Reaction volume of 15 µl contained 10 ng of genomic DNA, forward and reverse primers (0.25 µm each) and 1 SensiMix SYBR Hi-ROX mastermix (cat. no. QT605-05, Bioline). Primer sequences were: mt-atp6 forward 5 - ACACCAAAAGGACGAACCTG-3, mt-atp6 reverse 5 - ATGGGGAAGAAGCCCTAGAA-3, and PGC-1α forward 5 - ATGAATGCAGCGGTCTTAGC-3, PGC-1α reverse 5 - AACAATGGCAGGGTTTGTTC-3. Citrate synthase activity was measured in TA muscle homogenate as described in 49. Statistical analysis Data are presented as means±sd. Statistical significance of genotype and treatment effects was assessed by applying a two-way analysis of variance (ANOVA) in the SPSS 20.0 statistical package (SPSS Inc., Chicago, IL, USA). In case the interaction between genotype and treatment was significant (P<0.05) or borderline significant (P<0.1), the differences were evaluated in more detail by separately analyzing the effects of genotype and treatment using Bonferroni-corrected two-sided unpaired t-tests. The level of statistical significance was set at P<

102 Effect of pioglitazone on muscle mitochondrial function Results Animal characteristics Body weight (P<0.001), fasting plasma glucose (P<0.001) and fasting plasma insulin (P<0.01) were higher in diabetic rats compared with lean rats, both for water- and pioglitazone-treated groups (Table 1). Treatment with pioglitazone did not affect whole-body glucose homeostasis in lean rats. In diabetic rats, 2 weeks of treatment with pioglitazone lowered fasting plasma glucose levels compared with water-treated diabetic rats (P<0.01), but plasma glucose remained higher than in lean rats (P<0.001). Plasma FFA concentrations were similar between all groups, whereas plasma TG levels were higher in water-treated diabetic rats compared with water-treated lean rats (P<0.001). Treatment with pioglitazone lowered plasma TG in diabetic rats by 50% (P<0.01). Table 1 Animal characteristics of lean and diabetic ZDF rats after 2 weeks of treatment with either pioglitazone or water as a control. ANOVA Lean rats Diabetic rats Genotype Treatment Interaction Water Pioglitazone Water Pioglitazone Body weight (g) ± ± ± ± 30 Fasting glucose (mm) ± ± ± 0.9 ### 12.6 ± 4.7**,### Fasting insulin (pm) ± ± ± ± 101 Triglycerides (mm) ± ± ± 2.6 ### 2.8 ± 0.4** 4 FFA (mm) 0.7 ± ± ± ± 0.2 Data are represented as means ± SD (n=6 per group). + P<0.05, ++ P<0.01, +++ P<0.001: effects of genotype, treatment, and the interaction between genotype and treatment, using a two-way ANOVA. If the interaction between genotype and treatment was significant, a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: ** P<0.01, when compared with water-treated animals of the same genotype, ### P<0.001 when compared with lean animals of the same treatment regimen. 101

103 Chapter 4 IMCL content 1 H MRS was applied to assess the effect of pioglitazone treatment on IMCL levels. Representative examples of water-suppressed 1 H MR spectra obtained from the TA muscle of a lean rat and a diabetic rat are presented in Figure 1A. IMCL content was 14-fold higher in water-treated diabetic rats compared with water-treated lean controls (P<0.001; Figure 1B). In lean rats, pioglitazone treatment did not affect IMCL content. In contrast, treatment with pioglitazone lowered IMCL content in diabetic rats by 43% (P<0.01), yet IMCL levels remained 8-fold higher compared with lean controls (P<0.001). A Lean rat Diabetic rat IMCL EMCL-CH 2 tcr-ch 3 IMCL-CH 2 EMCL-CH 2 H 2 O tcr-ch 3 H 2 O tcr-ch 2 tcr-ch Chemical Shift (ppm) Chemical Shift (ppm) 1 0 B ### IMCL / H 2 O (%) ** ### LW LP DW DP 0.0 Figure 1. IMCL content assessed by 1 H MRS in TA muscle. Representative examples of 1 H MR spectra (256 averages) from the medial part of the TA muscle of a water-treated lean and diabetic rat (A). IMCL content expressed as a percentage of the water signal (B). Data of lean water-treated (LW), lean pioglitazone-treated (LP), diabetic water-treated (DW) and diabetic pioglitazone-treated (DP) ZDF rats are represented as means ± SD (n=6 per group). The interaction between genotype and treatment was significant (P<0.01) and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: ** P<0.01 when compared with watertreated animals of the same genotype, ### P<0.001 when compared with lean animals of the same treatment regimen. EMCL, extramyocellular lipid; tcr, total creatine. Muscle acylcarnitine content Levels of different acylcarnitine species in TA muscle were determined to examine whether treatment with pioglitazone affects the accumulation of FA intermediates. Free carnitine (P<0.001), acetylcarnitine (P<0.01), and most short-, medium- and long-chain acylcarnitines 102

104 Effect of pioglitazone on muscle mitochondrial function A 1200 Free carnitine and acetylcarnitine (pmol/mg dry weight) * ### C0 C2 LW LP DW DP B 20 Acylcarnitine (pmol/mg dry weight) ### * LW LP DW DP 4 0 C4 C5 C6 C8 C10 C12 C Acylcarnitine (pmol/mg dry weight) ### * # ### ** # ### * #, LW LP DW DP 0 C14 C16 C18 C16:1 C18:1 C18:2 Figure 2. Free carnitine and acetylcarnitine content (A) and levels of short-, medium- (B) and long-chain (C) acylcarnitine species in TA muscle of lean water-treated (LW), lean pioglitazone-treated (LP), diabetic water-treated (DW) and diabetic pioglitazone-treated (DP) ZDF rats. Data are represented as means ± SD (n=6 per group). General effects of genotype: P<0.05, P<0.01 (two-way ANOVA) when compared with lean animals, independent of treatment regimen. General effects of treatment: P<0.05 (two-way ANOVA) when compared with water-treated animals, independent of genotype. In case the interaction between genotype and treatment was significant (P<0.05; C0, C5) or borderline significant (P<0.1; C16, C18, C18:1) a pairwise analysis of differences is provided by Bonferronicorrected two-sided unpaired t-tests: * P<0.05, ** P<0.01 when compared with water-treated animals of the same genotype, # P<0.05, ### P<0.001 when compared with lean animals of the same treatment regimen. 103

105 Chapter 4 (P<0.05) were higher in muscle of diabetic rats compared with lean control animals (Figure 2). Muscle free carnitine was also elevated in lean rats treated with pioglitazone compared with water-treated lean animals (P<0.05). In diabetic animals, pioglitazone treatment lowered C5, C16, C18, C18:1 and C18:2 acylcarnitine species (P<0.05), leading to a partial normalization of muscle acylcarnitines to the level of lean controls. A PCr B ATP PCr ATP P i γ α β P i γ α β P chemical shift (ppm) 31 P chemical shift (ppm) C D Stimulation 0.8 * ## [PCr] (mm) k PCr (min -1 ) LW LP DW DP Time (min) Figure 3. In vivo oxidative capacity of TA muscle assessed by 31 P MRS. Representative examples of 31 P MR spectra obtained during rest with 32 averages (A) and at the end of the electrical-stimulation protocol with 4 averages (B). (C) Representative example of PCr concentrations determined during the rest-stimulation-recovery protocol. A mono-exponential function (solid line) was fit to the PCr concentrations during recovery. (D) Rate constants of PCr recovery (k PCr ) after electrical stimulation measured in TA muscle of lean water-treated (LW), lean pioglitazone-treated (LP), diabetic water-treated (DW) and diabetic pioglitazone-treated (DP) ZDF rats. Data are represented as means ± SD (n=6 per group). The interaction between genotype and treatment was significant (P<0.05) and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: * P<0.05 when compared with water-treated animals of the same genotype, ## P<0.01 when compared with lean animals of the same treatment regimen. 0.0 In vivo mitochondrial oxidative capacity 31 P MR spectroscopy was performed to determine whether pioglitazone-treatment affects in vivo muscle oxidative capacity. A representative example of a 31 P MR spectrum at rest is shown in Figure 3A, while a typical spectrum acquired after 2 minutes of muscle stimulation is depicted in Figure 3B. Concentrations of PCr and inorganic phosphate (P i ), and intracellular ph in TA muscle determined at rest and at the end of muscle stimulation are presented in Table 2. At rest, inorganic phosphate (P i ) levels were higher in muscle of diabetic rats compared with lean rats, independent 104

106 Effect of pioglitazone on muscle mitochondrial function of treatment regimen (P<0.05; Table 2). In addition, the concentration of PCr was lower in resting muscle of water-treated diabetic rats compared with water-treated lean rats (P<0.05), but this was normalized after treatment with pioglitazone. At the end of the 2 min of electrical stimulation of the TA muscle, P i levels were somewhat lower in diabetic rats compared with lean rats (P<0.05), but PCr concentrations, the amount of PCr depletion (ΔPCr) and end-exercise ph did not differ among groups. Table 2 Metabolite concentrations and ph measured using 31 P MRS in TA muscle of lean and diabetic ZDF rats after 2 weeks of treatment with either pioglitazone (PIO) or water as a control. Lean Diabetic water PIO water PIO Rest parameters ph 7.16 ± ± ± ± 0.04 [PCr] (mm) 35.5 ± ± ± 2.3 # 35.1 ± 0.7* [P i ] (mm) 2.4 ± ± ± ± 0.5 End-stimulation parameters ph 7.03 ± ± ± ± 0.08 [PCr] (mm) 19.0 ± ± ± ± 4.5 [P i ] (mm) 20.9 ± ± ± ± 2.5 ΔPCr (%) 54 ± 9 56 ± 7 52 ± 3 49 ± 8 Data is represented as mean ± SD (n=6 per group). At rest [P i ] was significantly higher and at the end of stimulation [P i ] was significantly lower in diabetic animals compared with lean animals, independent of treatment regimen (two-way ANOVA: P<0.05). * P<0.05 when compared with water-treated animals of the same genotype. # P<0.05 when compared with lean animals of the same treatment regimen. 4 The PCr recovery rate constant (k PCr ), which is a measure of in vivo muscle oxidative capacity, was obtained by fitting a mono-exponential function through the PCr concentrations during the recovery phase (Figure 3C). In water-treated diabetic rats, muscle oxidative capacity was impaired with respect to water-treated lean rats (P<0.01; Figure 3D). Pioglitazone treatment did not affect muscle oxidative capacity in lean rats. In contrast, in diabetic animals treatment with pioglitazone restored muscle oxidative capacity (P<0.05) to the level of lean controls. 105

107 Chapter 4 Ex vivo mitochondrial function High-resolution respirometry was performed to determine oxygen consumption rates (O 2 flux) of mitochondria isolated from TA muscle using TCA cycle or β-oxidation substrates in different metabolic states (Figure 4). O 2 fluxes fueled by the TCA cycle substrates pyruvate (Complex I-dependent respiration; Figure 4A) and succinate (Complex II-dependent respiration; Figure 4B) were similar in lean and diabetic rats, irrespective of respiratory state, and were not affected by pioglitazone treatment. Figure 4. Ex vivo mitochondrial function O 2 flux determined in mitochondria isolated from TA muscle of lean water-treated (LW), lean pioglitazone-treated (LP), diabetic water-treated (DW), and diabetic pioglitazone-treated (DP) ZDF rats, fueled with either TCA cycle substrates pyruvate plus malate (Complex I-respiration) (A) or succinate plus rotenone (Complex II-respiration) (B), or β-oxidation substrate palmitoyl- L-carnitine (C). The OXPHOS state was initiated by addition of an ADP-regenerating system, the LEAK state by blocking ATP synthesis with carboxyatractyloside (CAT), and the ETS state by addition of carbonyl cyanide 3-chlorophenyl hydrazone (CCCP). Data are represented as means ± SD (n=6 per group). For the respiratory capacity with palmitoyl-l-carnitine in the OXPHOS state the interaction between genotype and treatment was borderline significant (P=0.067) and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: * P<0.05 when compared with water-treated animals of the same genotype, ### P<0.001 when compared with lean animals of the same treatment group. Respiratory capacity with palmitoyl-lcarnitine in the ETS state was significantly higher in diabetic rats compared with lean rats, independent of treatment regimen: P<0.001 (two-way ANOVA). 106

108 Effect of pioglitazone on muscle mitochondrial function With the β-oxidation substrate palmitoyl-l-carnitine, respiratory capacity in both the OXPHOS and ETS state was higher in water-treated diabetic rats compared with water-treated lean rats (P<0.001; Figure 4C). In lean animals, pioglitazone treatment did not affect O 2 flux fueled by palmitoyl-l-carnitine in any of the respiratory states. However, in diabetic rats treatment with pioglitazone lowered palmitoyl-l-carnitine driven mitochondrial respiration in the OXPHOS state (P<0.05) to the level of lean controls. Markers of muscle mitochondrial biogenesis and content Protein expression level of PGC-1α in TA muscle was lower in water-treated diabetic rats compared with water-treated lean rats (P<0.001; Figure 5A). In lean animals, pioglitazone treatment did not affect PGC-1α expression, but in diabetic rats treatment with pioglitazone increased expression of PGC-1α (P<0.01), leading to a partial normalization compared with lean controls. However, mtdna copy number and citrate synthase activity did not differ between groups (Figure 5B and 5C), indicating that mitochondrial content did not differ between muscle of lean and diabetic rats and that it was not affected by pioglitazone treatment. PGC-1 a /GAPDH [-] LW LP DW DP PGC1α A B C GAPDH ### ** # LW LP DW DP mtdna copy no [-] LW LP DW DP 4 Figure 5. Markers of muscle mitochondrial biogenesis and content. Protein expression level of PGC-1α normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (A), relative copy number of mtdna (B), and citrate synthase (CS) activity (C), determined in whole TA muscle homogenate of lean water-treated (LW, n=6), lean pioglitazone-treated (LP, n=6), diabetic water-treated (DW, n=6) and diabetic pioglitazone-treated (DP, n=4 for PGC-1α and mtdna, and n=6 for CS activity) ZDF rats. Data are represented as means ± SD. For PGC-1α expression, the interaction between genotype and treatment was borderline significant (P=0.098) and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: ** P<0.01 when compared with water-treated animals of the same genotype, # P<0.05, ### P<0.001 when compared with lean animals of the same treatment group. 107

109 Chapter 4 Discussion The present study aimed to elucidate the effect of pioglitazone on in vivo mitochondrial function in muscle of both lean and diabetic ZDF rats. Using 31 P MRS, we showed that while treatment with pioglitazone did not affect in vivo muscle oxidative capacity in lean rats, it did improve oxidative capacity of muscle in diabetic rats and restored it to the level of the lean controls. The improvement of in vivo muscle oxidative capacity in diabetic rats was not accompanied by an increase in either ex vivo mitochondrial respiratory capacity or mitochondrial content, but rather was paralleled by an improvement in lipid homeostasis, i.e. decreases of plasma TG and muscle lipid and long-chain acylcarnitine content, and a normalization of β-oxidation capacity. In agreement with previous reports 12,24,42,50, we observed that plasma TG and muscle lipid and acylcarnitine content were elevated in diabetic rats compared with lean rats, demonstrating disturbed lipid homeostasis in the diabetic animals and supporting the reported association between ectopic lipid accumulation and insulin resistance Two weeks of treatment with pioglitazone did not affect plasma and muscle lipid levels in lean rats. In contrast, in diabetic rats pioglitazone lowered plasma TG levels and resulted in a partial normalization of IMCL and muscle long-chain acylcarnitine content compared with lean controls. The latter is consistent with the muscle lipid-lowering effect of pioglitazone as has been shown before in human subjects with insulin resistance and type 2 diabetes 9,27, supposedly as a result of redistribution of lipids from muscle into adipose tissue. In vivo muscle oxidative capacity is, apart from extramitochondrial factors such as vascular oxygen supply 51, dependent both on the amount of mitochondria present in skeletal muscle and their intrinsic function 52. In vivo 31 P MRS experiments showed that the PCr recovery rate constant (k PCr ), a measure of in vivo muscle oxidative capacity, was lower in diabetic rats compared with lean control animals. However, ex vivo mitochondrial respiratory capacity with TCA cycle substrates pyruvate and succinate, supporting Complex I- and II-dependent respiration, respectively, was similar between diabetic and lean animals, independent of the respiratory state. Moreover, mtdna copy number and citrate synthase activity were not different between muscle of lean and diabetic rats. These results imply that neither a lower mitochondrial content nor an impairment of their intrinsic function can account for the lower in vivo muscle oxidative capacity in diabetic rats. Instead, it suggests that the functioning of mitochondria in their natural cellular environment is impaired by factors that are not taken into account during ex vivo measurement of intrinsic mitochondrial function. We furthermore demonstrated that β-oxidation capacity was higher in muscle mitochondria of diabetic rats compared with lean rats. This finding confirms previous reports on increased capacity to oxidize FAs in order to cope with the high lipid loads in insulin-resistant states In addition, insulin-resistant muscle has been associated with the accumulation of incompletely metabolized FAs 24,53. Indeed, we showed that the content of long-chain acylcarnitines, which reflects the longchain acyl-coa content 54, was significantly higher in muscle of diabetic rats as compared with lean rats. The accumulation of acylcarnitines indicates an increased FA oxidation flux in diabetic muscle, which is not matched by an increase in energy demand. When FA oxidation flux outpaces the demand of the respiratory system this leads to increased formation of ROS 25, which in turn promotes mitochondrial uncoupling and lowers the efficiency of ATP synthesis 55. Moreover, FAs and their metabolites can also directly impair mitochondrial ATP production by mitochondrial uncoupling or by inhibiting enzymes involved in the process, such as inhibition of mitochondrial 108

110 Effect of pioglitazone on muscle mitochondrial function adenine nucleotide translocase by long-chain acyl-coa esters 56. Therefore, our data indicate that the mitochondrial functional impairment that we observed in vivo in muscle of diabetic rats is the result of mitochondrial lipid overload, likely due to an increased influx of FAs, rather than an intrinsic impairment in the pathway involved in mitochondrial ATP production, similar to our observation in long-term high-fat diet fed rats 22. In lean rats, pioglitazone treatment did not affect in vivo muscle oxidative capacity, nor did it lead to changes in mitochondrial content or ex vivo mitochondrial respiratory capacity with Complex I- and II-dependent substrates. In diabetic rats, treatment with pioglitazone increased in vivo muscle oxidative capacity and actually normalized it to the value of lean controls, indicating that the impairment in in vivo muscle oxidative capacity as observed in water-treated diabetic rats was completely alleviated by pioglitazone treatment. This was however not accompanied by increases in ex vivo mitochondrial respiratory capacity or muscle mitochondrial content, which implies that the improvement in in vivo muscle oxidative capacity in diabetic rats cannot be explained by effects of pioglitazone on mitochondrial intrinsic function and/or content. Previous studies on the effect of TZDs on mitochondrial function report contrasting data. Rosiglitazone has been reported to enhance expression of PGC-1α in skeletal muscle 34, but this was however not reflected in an increase of mitochondrial content 34,57. This is in agreement with the results we obtained with pioglitazone and is likely explained by the fact that the activity of PGC-1α is not only regulated at the transcriptional level, but also via several post-translational modifications, such as reversible acetylation, phosphorylation, and methylation 58. In contrast to our results with pioglitazone in diabetic rats, treatment of type 2 diabetes patients with rosiglitazone did not improve in vivo muscle oxidative capacity 59. Furthermore, while some studies have shown pioglitazone-induced impairments in Complex I and III activity and mitochondrial respiration, both after acute exposure in vitro and after regular treatment in vivo 29-31, others report enhanced expression of genes involved in mitochondrial function and increased mitochondrial respiration in muscle after in vivo treatment with and in vitro exposure to pioglitazone 32,33,38. 4 The improvement in in vivo muscle oxidative capacity in diabetic rats was paralleled by partially normalized levels of plasma TG and IMCL. In addition, pioglitazone treatment lowered ex vivo β-oxidation capacity in diabetic rats to the value of lean controls. The normalization of ex vivo β-oxidation capacity in diabetic muscle was paralleled by a decrease in muscle long-chain acylcarnitines, indicating that the reduced lipid supply upon pioglitazone treatment is accompanied by a decrease in FA oxidation flux, restoring the balance between energy supply and demand in diabetic muscle 25. These results corroborate our finding that the impairment of in vivo muscle oxidative capacity in diabetic rats is likely the result of lipid-induced mitochondrial uncoupling and/or inhibition and it shows that this effect can be completely alleviated by redirecting the lipids from the muscle into adipose tissue. 109

111 Chapter 4 In conclusion, we have shown that pioglitazone treatment restores in vivo muscle oxidative capacity in diabetic rats, which was not accompanied by improvements in intrinsic mitochondrial properties, but rather was paralleled by a reduction in muscle lipid content and FA oxidation capacity. These findings demonstrate that excessive accumulation of ectopic lipids can induce mitochondrial stress and thus hinder efficient ATP production, and that pioglitazone, by redistributing lipids from muscle into adipose tissue, effectively alleviates lipid-induced mitochondrial dysfunction in vivo. Acknowledgements We thank Leonie Niesen and David Veraart for their assistance in animal handling, and Dr. Koo Rijpkema for his advice on the statistical analysis. B.W. and J.J.P. are supported by a VIDI grant (project number ) from the Netherlands Organisation for Scientific Research (NWO). J.C. is supported by the NWO-funded Groningen Systems Biology Center for Energy Metabolism and Aging. 110

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114 Effect of pioglitazone on muscle mitochondrial function 49. Broder, I. & Srere, P.A. Immunochemical studies with citrate-condensing enzyme. Biochimica et Biophysica Acta (BBA) - Specialized Section on Enzymological Subjects 67, (1963). 50. Jonkers, R.A., van Loon, L.J., Nicolay, K. & Prompers, J.J. In vivo postprandial lipid partitioning in liver and skeletal muscle in prediabetic and diabetic rats. Diabetologia 56, (2013). 51. Kemp, G.J. Mitochondrial dysfunction in chronic ischemia and peripheral vascular disease. Mitochondrion 4, (2004). 52. Prompers, J.J., Wessels, B., Kemp, G.J. & Nicolay, K. MITOCHONDRIA: investigation of in vivo muscle mitochondrial function by 31 P magnetic resonance spectroscopy. Int. J. Biochem. Cell Biol. 50, (2014). 53. Noland, R.C., et al. Carnitine insufficiency caused by aging and overnutrition compromises mitochondrial performance and metabolic control. J. Biol. Chem. 284, (2009). 54. Brass, E.P. & Hoppel, C.L. Relationship between acid-soluble carnitine and coenzyme A pools in vivo. Biochem. J. 190, (1980). 55. Brand, M.D. Uncoupling to survive? The role of mitochondrial inefficiency in ageing. Exp. Gerontol. 35, (2000). 56. Lerner, E., Shug, A.L., Elson, C. & Shrago, E. Reversible inhibition of adenine nucleotide translocation by long chain fatty acyl coenzyme A esters in liver mitochondria of diabetic and hibernating animals. J. Biol. Chem. 247, (1972). 57. Pagel-Langenickel, I., et al. A discordance in rosiglitazone mediated insulin sensitization and skeletal muscle mitochondrial content/activity in Type 2 diabetes mellitus. Am J Physiol Heart Circ Physiol 293, H (2007). 58. Fernandez-Marcos, P.J. & Auwerx, J. Regulation of PGC-1alpha, a nodal regulator of mitochondrial biogenesis. Am. J. Clin. Nutr. 93, 884S-890 (2011). 59. Schrauwen-Hinderling, V.B., et al. The insulin-sensitizing effect of rosiglitazone in type 2 diabetes mellitus patients does not require improved in vivo muscle mitochondrial function. The Journal of clinical endocrinology and metabolism 93, (2008)

115 Metformin impairs mitochondrial function in skeletal muscle of both lean and diabetic rats in a dose-dependent manner Bart Wessels 1, Jolita Ciapaite 2, Nicole M.A. van den Broek 1, Klaas Nicolay 1 and Jeanine J. Prompers 1 1 Biomedical NMR, Department of Biomedical Engineering, Eindhoven University of Technology, the Netherlands, 2 Center for Liver, Digestive and Metabolic Diseases, Department of Pediatrics, University of Groningen, University Medical Center Groningen, Groningen, the Netherlands, Published in PLoS One 9(6):e (2014) doi: /journal.pone Epub 2014 Jun 20.

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118 Effect of metformin on muscle mitochondrial function Abstract Aim: Metformin is a widely prescribed drug for the treatment of type 2 diabetes. Previous studies have demonstrated in vitro that metformin specifically inhibits Complex I of the mitochondrial respiratory chain. This seems contraindicative since muscle mitochondrial dysfunction has been linked to the pathogenesis of type 2 diabetes. However, its significance for in vivo skeletal muscle mitochondrial function has yet to be elucidated. The aim of this study was to assess the effects of metformin on in vivo and ex vivo skeletal muscle mitochondrial function in a rat model of diabetes. Materials and Methods: Healthy (fa/+) and diabetic (fa/fa) Zucker diabetic fatty rats were treated by oral gavage with metformin dissolved in water (30, 100 or 300 mg/kg bodyweight/day) or water as a control for 2 weeks. After 2 weeks of treatment, muscle oxidative capacity was assessed in vivo using 31 P magnetic resonance spectroscopy and ex vivo by measuring oxygen consumption in isolated mitochondria using high-resolution respirometry. Results: Two weeks of treatment with metformin impaired in vivo muscle oxidative capacity in a dose-dependent manner, both in healthy and diabetic rats. Whereas a dosage of 30 mg/kg/ day had no significant effect, in vivo oxidative capacity was 21% and 48% lower after metformin treatment at 100 and 300 mg/kg/day, respectively, independent of genotype. High-resolution respirometry measurements demonstrated a similar dose-dependent effect of metformin on ex vivo mitochondrial function. Conclusion: Metformin compromises in vivo and ex vivo muscle oxidative capacity in Zucker diabetic fatty rats in a dose-dependent manner

119 Chapter 5 Introduction Metformin is the most commonly prescribed drug to treat type 2 diabetes and it has been in clinical use for decades. Metformin is a biguanide that lowers blood glucose levels primarily by improving insulin sensitivity in the liver, where it effectively inhibits gluconeogenesis 1, whereas it does not have marked hypoglycemic effects 2. Moreover, metformin enhances insulin sensitivity in skeletal muscle, thereby stimulating peripheral glucose utilization 3. A number of in vitro studies reported that metformin inhibits Complex I of the mitochondrial respiratory chain 3-9, thus limiting the respiratory capacity of the cell and possibly restricting ATP synthesis. The mechanism through which metformin acts on Complex I, however, is still not known. Some studies suggest that metformin binds directly to the mitochondrial membrane phospholipids, thereby altering physicochemical membrane properties 3,10. Others contradict this direct mechanism and postulate that an intact cell is required for metformin s inhibitory action on Complex I, involving an indirect pathway via the cell membrane 6,8. Treatment of patients with type 2 diabetes with a Complex I inhibitor seems contraindicative, since muscle mitochondrial dysfunction has been linked to the pathogenesis of this disease 11,12. Moreover, considering that regular exercise is recommended in most guidelines for the treatment of type 2 diabetes 13,14, it seems even more unfavorable to treat diabetes patients with a Complex I inhibitor, as it would attenuate their exercise capacity and consequently their ability to increase insulin sensitivity via exercise training. The latter has indeed been demonstrated by Sharoff et al. 15, who observed that the therapeutic effects of exercise training were absent in patients who were treated with metformin in conjunction with exercise therapy. Although the specific inhibitory action of metformin on Complex I has been shown using in vitro measurements, its significance for in vivo skeletal muscle mitochondrial function has yet to be elucidated. The aim of this study was to determine the effect of metformin on in vivo skeletal muscle oxidative capacity in a rat model of diabetes using phosphorous ( 31 P) magnetic resonance spectroscopy (MRS). Lean, healthy and obese, diabetic Zucker diabetic fatty (ZDF) rats were dosed orally for 2 weeks with metformin dissolved in water (30, 100 or 300 mg/kg body weight/ day) or water as a control. A dosage of 30 mg/kg/day is typically prescribed for diabetes patients, while, because of the lower bioavailability of metformin in rats compared with humans, ~ mg/kg/day metformin is needed to attain similar effects on glucose homeostasis in rats 3, P MRS measurements were complemented by ex vivo high resolution respirometry (HRR) measurements in isolated mitochondria to interpret the effects of metformin on in vivo oxidative capacity. We demonstrated that 2 weeks of treatment with metformin compromised in vivo and ex vivo muscle oxidative capacity in ZDF rats in a dose-dependent manner. 118

120 Effect of metformin on muscle mitochondrial function Research design and Methods Ethics statement All experimental procedures were reviewed and approved by the Animal Experimental Committee of Maastricht University (permit number: ). Surgery, MRS experiments and termination were performed under isoflurane (IsoFlo) anesthesia with additional pain relief using buprenorphine (Temgesic), and all efforts were made to minimize suffering. Animals Lean, non-diabetic fa/+ and obese, diabetic, fa/fa adult male ZDF rats (12 weeks of age) were purchased from Charles River Laboratories (Sulzfield, Germany). The animals were housed pairwise, in a controlled environment (20 and 50% relative humidity on a 12-h light-dark cycle) and given ad libitum access to water and specific standardized chow for ZDF rats (Purina Formula 5008, Bioservices, the Netherlands). For 15 days, animals were dosed with metformin (0, 30, 100 or 300 mg/kg body weight/day, n=6 per group) in 1 ml of water directly into the stomach by oral gavage. Dosing was performed once daily between 4 and 6 pm. At day 15, in vivo MRS experiments were performed on the animals between 8 am and 4 pm, i.e hours after the dosage of metformin on the previous day. Following the MRS measurements, animals were administered with the last dose of metformin (between 4 and 6 pm). The following day between 8 and 10 am, i.e hours after the last dosage of metformin, animals were sacrificed under anesthesia by incision of the vena cava. The terminal half-life of metformin after oral administration in rats has been determined to be ~3, 6 and 7 hours at doses of 50, 100 and 200 mg/kg, respectively 22, which implies that all in vivo and ex vivo experiments were performed under conditions in which plasma levels of metformin were less than 5% of the maximum plasma concentrations. One tibialis anterior (TA) muscle was used for isolation of mitochondria. The other TA was frozen in liquid nitrogen and stored at -80 C. 5 Plasma parameters After 2 weeks of treatment, a blood sample was taken between 12 and 2 pm (i.e. at least 18 hours after the previous dosage of metformin), after a 4-hour fast, for determination of post-therapy plasma glucose and insulin concentrations. Plasma glucose concentrations were determined using an automatic glucometer (Freestyle, Abbott, IL, USA). Plasma insulin concentrations were determined with an ultrasensitive rat insulin ELISA kit (Mercodia, Uppsala, Sweden). MRS measurements 31 P MRS measurements were performed using a horizontal 6.3-T MR scanner (Bruker, Ettlingen, Germany) with an ellipsoid (10/18 mm) 31 P surface coil. The animals were anaesthetized using isoflurane (2-3%) combined with medical air (0.6 L/min). 31 P MRS was applied to assess in vivo oxidative capacity of the TA muscle, as described previously 23. A fully relaxed spectrum (repetition time = 20 s, 32 averages) was recorded first, followed by a time series of spectra (repetition time = 5 s, 4 averages) obtained during a resting period of 3 min, 2 min of electrical stimulation and 15 min of recovery. Electrodes were implanted subcutaneously along the distal nerve trajectory of the N. peroneus communis to electrically stimulate the TA muscle. Pulses with a stimulation voltage of approximately 3 V were used to reach similar levels of phosphocreatine (PCr) depletion for the different animals. 119

121 Chapter 5 MRS data analysis MR spectra were fitted in the time domain using a nonlinear least squares algorithm (advanced method for accurate, robust, and efficient spectral fitting; AMARES) in the jmrui software package 24 as described previously 23. In short, spectral analysis of the 31 P MR spectra was done by fitting the PCr peak to Lorentzian and the inorganic phosphate (Pi) as well as the α-, β- and γ-atp peaks to Gaussian line shapes. Intracellular ph was calculated from the chemical shift difference between the Pi and PCr resonances 25. For the time series, the concentrations of PCr determined during recovery were fit to a mono-exponential function using Matlab (version , Mathworks, Natick, MA, USA) yielding a rate constant, k PCr, which is a measure of skeletal muscle mitochondrial oxidative capacity. For each rat, results from two time series with end-stimulation ph values higher than 6.9 were averaged 26. Determination of the relative mitochondrial DNA copy number The relative mitochondrial-dna copy number was measured as described previously 27. Briefly, genomic DNA was isolated from a 25 mg transversal slice of mid-belly TA using GenElute Mammalian Genomic DNA Miniprep Kit (Sigma-Aldrich, Zwijndrecht, The Netherlands). Mitochondrial DNA (mtdna) content relative to peroxisome proliferator-activated receptor-γ coactivator 1α (PGC-1α) gene was measured using real-time PCR as described in 28. High-resolution respirometry Skeletal muscle mitochondria were isolated from whole TA muscle through a differential centrifugation procedure as described elsewhere 27. Mitochondrial protein content was determined using a BCA protein assay kit (Pierce, Thermo Fisher Scientific Inc., Rockfort, IL, USA). Ex vivo mitochondrial function was evaluated by measuring oxygen consumption rates (O 2 flux) at 37 C using a 2-channel high-resolution Oroboros oxygraph-2k (Oroboros, Innsbruck, Austria) as described previously 27. O 2 flux was fueled either with 5 mm pyruvate plus 5 mm malate (Complex I respiration) or 5 mm succinate plus 1 µm rotenone (Complex II respiration). Maximal rates of oxygen consumption coupled to ATP synthesis, i.e. the OXPHOS state (classical state 3), was determined after addition of an ADP-regenerating system consisting of excess hexokinase (4.8 U/ml), glucose (12.5 mm) and ATP (1 mm). The resting state respiration, which compensates for proton leak, i.e. the LEAK state (classical state 4), was assessed after addition of 1.25 µm carboxyatractyloside (CAT). Finally, the maximal capacity of the electron transfer system (ETS), i.e. the ETS state (classical state U), was determined by uncoupling the ETS from ATP synthesis with the addition of 1 µm carbonyl cyanide 3-chlorophenyl hydrazone (CCCP) 29. The respiratory control ratio (RCR) was calculated as the ratio of OXPHOS to LEAK states. 120

122 Effect of metformin on muscle mitochondrial function High-resolution respirometry after in vitro incubation with metformin Isolated mitochondria from a cohort of water-treated lean and diabetic ZDF rats (n=5 per genotype) were incubated in assay medium supplemented with metformin (1 mm) for 5 minutes in the presence of pyruvate and malate or succinate plus rotenone (at 37 C), after which mitochondrial respiratory capacity was assessed in the OXPHOS state. Results were expressed relative to the oxygen consumption rates measured without incubation with metformin. For all HRR measurements, signals from the oxygen electrode were recorded at 0.5-s intervals and measurements were done in duplicate. Data acquisition and analysis was performed using Oxygraph-2k-Datlab software (Oroboros, Innsbruck, Austria). Statistical analysis Data are presented as means ± SD. Statistical significance of genotype and treatment effects were assessed by applying a two-way Analysis of Variance (ANOVA) in the IBM SPSS 20 statistical package (SPSS Inc., Chicago, IL, USA). In case of a significant effect of treatment, Bonferroni corrected post-hoc tests were carried out in order to identify differences between different treatment regimens. In case the interaction between genotype and treatment was significant or borderline significant (P<0.1), the differences were evaluated in more detail by separately analyzing the effects of genotype and treatment using Bonferroni-corrected two-sided unpaired t-tests. For determination of mitochondrial respiratory capacity changes after in vitro incubation of mitochondria with metformin, statistical analysis was done using a 2x2 mixed design ANOVA with one within-subjects factor (metformin incubation) and one between-subjects factor (genotype) in SPSS. The level of statistical significance was set at P<

123 Chapter 5 Results Animal characteristics Animal characteristics after 2 weeks of treatment are summarized in Table 1. Body weight was significantly higher in diabetic animals compared with lean animals (P<0.01), except for the watertreated groups (for which body weight also did not differ before start of treatment). Fasting plasma glucose (P<0.001) and insulin (P<0.01) were significantly higher in diabetic animals compared with lean animals, independent of treatment regimen. Two weeks of treatment with 30, 100 or 300 mg/kg/day metformin had no effect on body weight, fasting plasma glucose, or fasting plasma insulin in lean or diabetic animals. Table 1. Animal characteristics of lean and diabetic ZDF rats after 2 weeks of treatment with water or 30, 100 or 300 mg/kg body weight/day metformin (MET30, MET100 and MET300, respectively). Lean Diabetic Body weight (g) Fasting glucose (mm) Fasting insulin (pm) Water 366 ± ± ± 99 MET ± ± ± 63 MET ± ± ± 60 MET ± ± ± 42 Water 380 ± ± ± 50 MET ± 11 ## 13.2 ± ± 128 MET ± 15 ### 14.7 ± ± 166 MET ± 27 ### 15.1 ± ± 164 Data is represented as mean ± SD (n=6 per group). Fasting plasma glucose (ANOVA: P<0.001) and insulin (ANOVA: P<0.01) were significantly higher in diabetic animals compared with lean animals, independent of treatment regimen. For body weight, the interaction between genotype and treatment was significant and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: ## P<0.01, ### P<0.001 when compared with lean animals of the same treatment regimen. 122

124 Effect of metformin on muscle mitochondrial function In vivo muscle mitochondrial oxidative capacity 31 P MRS was applied to assess the effect of metformin treatment on in vivo mitochondrial oxidative capacity. Representative examples of 31 P MR spectra obtained from TA muscle at rest and after 2 minutes of electrical stimulation are shown in Figure 1A and 1B, respectively. PCr and Pi concentrations and intracellular ph measured in TA muscle at rest and after muscle stimulation are listed in Table 2. End-stimulation ph was significantly higher in diabetic animals compared with lean animals (P<0.01), independent of treatment regimen. However, the end-stimulation ph was higher than 7.0 for all animals and therefore did not influence PCr recovery kinetics. A monoexponential function was fitted through the PCr concentrations obtained during the recovery phase (Figure 1C), yielding the PCr recovery rate constant, k PCr, which is representative for muscle oxidative capacity in vivo. k PCr was 25% lower in diabetic rats compared with lean rats, independent of treatment regimen (P<0.001) (Figure 1D). Table 2. Metabolite concentrations and ph in TA muscle measured by 31 P MRS of lean and diabetic ZDF rats after 2 weeks of treatment with water or 30, 100 or 300 mg/kg body weight/day metformin (MET30, MET100 and MET300, respectively). REST END-STIMULATION ph (-) [PCr] (mm) [P i ] (mm) ph (-) [PCr] (mm) [P i ] (mm) PCr (%) Lean Water 7.17 ± ± ± ± ± ± ± 3.5 MET ± ± ± ± ± ± ± 3.3 Diabetic MET ± ± ± ± ± ± ± 4.7 MET ± ± ± ± ± ± ± Water 7.16 ± ± ± ± ± ± 2.2 # 63.7 ± 3.1 MET ± ± ± ± ± 1.2 ##, * 21.3 ± ± 3.4 ## MET ± ± ± ± ± ± 3.6 #, 65.6 ± 3.8 MET ± ± ± ± ± ± 2.6 # 62.2 ± 4.6 Data is represented as mean ± SD (n=6 per group). At rest, ph was significantly lower and [Pi] was significantly higher in diabetic animals compared with lean animals, independent of treatment regimen (ANOVA: P<0.05). At the end of stimulation, ph was significantly higher in diabetic animals compared with lean animals, independent of treatment regimen (ANOVA: P<0.01). For end-stimulation [PCr], [Pi] and PCr, the interaction between genotype and treatment was significant and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: * P<0.05 when compared with water-treated animals of the same genotype, # P<0.05, ## P<0.01 when compared with lean animals of the same treatment regimen, P<0.05, P<0.01 when compared with MET30-treated animals of the same genotype. 123

125 Chapter 5 Two weeks of treatment with metformin had a significant effect on in in vivo muscle oxidative capacity, independent of genotype (P<0.001). Post-hoc testing revealed that treatment of lean and diabetic rats with 30 mg/kg/day metformin did not affect in vivo muscle oxidative capacity when compared with water-treated controls. However, in rats treated with metformin at a dosage of 100 and 300 mg/kg/day, in vivo muscle oxidative capacity was 21% (P<0.001) and 47% (P<0.001) lower, respectively, when compared with water-treated animals. A B PCr ATP P i PCr ATP P i γ α β γ α β P chemical shift (ppm) 31 P chemical shift (ppm) C D 100 stimulation [PCr] (%) k PCr (min -1 ) * * * * Water MET30 MET100 MET Time (min) 0.0 Lean Diabetic Figure 1. In vivo oxidative capacity of tibialis anterior (TA) muscle, assessed by 31 P MRS Representative examples of 31 P MR spectra obtained during rest with 32 averages (A) and at the end of the electrical-stimulation protocol with 4 averages (B). (C) Representative examples of relative PCr concentrations during rest, muscle stimulation and recovery (time resolution = 20 s) for a water-treated diabetic rat (open symbols) and a diabetic rat treated with metformin at 300 mg/kg body weight/day (filled symbols). PCr concentrations are expressed as a percentage of the resting PCr concentration. Mono-exponential functions (dark lines) were fit to the recovery data and the PCr recovery rate constants were 0.63 and 0.21 min -1 for the water-treated and metformin-treated animal, respectively. (D) Rate constants of PCr recovery, k PCr, after electrical stimulation in TA muscle of lean and diabetic rats treated with water or 30, 100 or 300 mg/kg body weight/day metformin (MET30, MET100 and MET300 respectively). Data is represented as mean ± SD (n=6 per group). k PCr was significantly lower in diabetic rats compared with lean rats, independent of treatment regimen (ANOVA: P<0.001). In addition, treatment had a significant effect on k PCr, independent of genotype, and a pairwise analysis of differences is provided by Bonferroni-corrected post-hoc tests: * P<0.001 when compared with water-treated animals, P<0.001 when compared with MET30-treated animals, P<0.001 when compared with MET100-treated animals. 124

126 Effect of metformin on muscle mitochondrial function Mitochondrial content Skeletal muscle oxidative capacity is determined by intrinsic mitochondrial properties, as well as the number of mitochondria in the tissue. Relative mtdna copy number, which was used as an estimate of mitochondrial content, did not differ between lean and diabetic rats (Figure 2). Moreover, metformin treatment (300 mg/kg/day) did not affect relative mtdna copy number Relative mitochondrial copy no (-) Lean Diabetic Water MET300 Figure 2. Mitochondrial content Relative mitochondrial-dna copy number of lean and diabetic rats after 2 weeks of treatment with either water or metformin (300 mg/kg bodyweight/day). Data is represented as mean ± SD (n=6 per group). Ex vivo mitochondrial function In order to evaluate ex vivo intrinsic mitochondrial function after 2 weeks of oral treatment with metformin, HRR was used to measure O 2 flux in mitochondria isolated from TA muscle, using both Complex I- and Complex II-dependent substrates. OXPHOS LEAK ETS O 2 flux (nmol mg -1 min -1 ) * Lean * * * # Diabetic * * * Lean # Diabetic Lean * # # Diabetic Water MET30 MET100 MET300 Figure 3. Complex I-dependent respiration O 2 consumption rates determined in mitochondria isolated from TA muscle of lean and diabetic rats treated with water or 30, 100 or 300 mg/kg body weight/day metformin (MET30, MET100 and MET300, respectively) for 2 weeks, fueled by pyruvate plus malate (Complex I-dependent substrate). Respiratory capacity was determined in the OXPHOS state, when mitochondrial respiration is coupled to ATP synthesis; the LEAK-state, when the system is limited by ADP; and the ETS state, after uncoupling of the ETS from ATP synthesis. Data is represented as mean ± SD (n=6 per group). For the OXPHOS state, the interaction between genotype and treatment was borderline significant and for the LEAK and ETS state, the interaction between genotype and treatment was significant. A pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: * P<0.05 when compared with water-treated animals of the same genotype, P<0.05 when compared with MET30-treated animals of the same genotype, P<0.05 when compared with MET100-treated animals of the same genotype, # P<0.05 when compared with lean animals of the same treatment regimen. 125

127 Chapter 5 Complex I Complex I-dependent respiratory capacity (driven by pyruvate plus malate) in the OXPHOS state was not different between lean and diabetic animals, except at the highest metformin dosage (300 mg/kg/day), for which OXPHOS respiratory capacity was lower in lean rats compared with diabetic rats (P<0.05) (Figure 3). Whereas 2 weeks of treatment with metformin at 30 mg/kg/ day did not affect Complex I-dependent respiratory capacity in the OXPHOS state, treatment at a dosage of 100 and 300 mg/kg/day lowered OXPHOS respiratory capacity compared with water treatment in both lean and diabetic animals (P<0.05). In lean animals, Complex I-dependent OXPHOS respiratory capacity was further reduced after metformin treatment at 300 mg/kg/day compared with 100 mg/kg/day (P<0.01), but this dose-dependent effect was not significant in diabetic animals. In lean animals, Complex I-dependent respiration in the LEAK state was lower after metformin treatment when compared with water treatment, for all metformin dosages (P<0.05) (Figure 3). As a consequence of the concomitant changes in OXPHOS and LEAK states in response to metformin treatment, the RCR s, which give an indication of the coupling efficiency between substrate oxidation and ATP synthesis, were not affected in lean animals (Table 3). In diabetic rats, the LEAK state was lower in the MET100 and MET300 groups compared with the MET30 group only (P<0.001) and the RCR was higher in the MET100 compared with MET30 group (P<0.05). In addition, the RCR was higher in water-treated diabetic rats compared with water-treated lean rats (P<0.05). In lean animals, treatment with metformin at a dosage of 300 mg/kg/day lowered Complex I-dependent respiratory capacity in the ETS state when compared to all other treatment regimens (P<0.01) (Figure 3). In contrast, metformin treatment had no significant effect on Complex I-dependent ETS respiratory capacity in diabetic rats. 126

128 Effect of metformin on muscle mitochondrial function Complex II Complex II-dependent respiratory capacity (driven by succinate plus rotenone) in the OXPHOS state was not different between lean and diabetic animals (Figure 4). Moreover, treatment with metformin had no effect on Complex II-dependent OXPHOS respiratory capacity, except for diabetic rats treated with 300 mg/kg/day metformin, for which OXPHOS respiratory capacity was lower than for diabetic rats treated with 100 mg/kg/day metformin (P<0.01). Complex II-dependent respiration in the LEAK state (Figure 4) and RCR s (Table 3) were not different between groups. 600 OXPHOS LEAK O 2 flux (nmol mg -1 min -1 ) Lean Diabetic Lean Diabetic Water MET30 MET100 MET300 Figure 4. Complex II-dependent respiration O 2 consumption rates determined in mitochondria isolated from TA muscle of lean and diabetic rats treated with water or 30, 100 or 300 mg/kg body weight/day metformin (MET30, MET100 and MET300 respectively) for 2 weeks, fueled by succinate plus rotenone (Complex II-dependent substrate). Respiratory capacity was determined in the OXPHOS state, when mitochondrial respiration is coupled to ATP synthesis; and the LEAK-state, when the system is limited by ADP. Data is represented as mean ± SD (n=6 per group). For the OXPHOS state, the interaction between genotype and treatment was significant and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: P<0.05 when compared with MET100-treated animals of the same genotype

129 Chapter 5 Table 3. Respiratory control ratios (RCR s) in mitochondria isolated from TA muscle of lean and diabetic rats treated with water or 30, 100 or 300 mg/kg body weight/day metformin (MET30, MET100 and MET300, respectively) for 2 weeks, fueled by pyruvate plus malate (Complex I-dependent substrate) and succinate plus rotenone (Complex II-dependent substrate). RCR Pyruvate (-) RCR Succinate (-) Lean Water 11.0 ± ± 0.4 MET ± ± 1.3 MET ± ± 0.8 MET ± ± 0.7 Diabetic Water 17.6 ± 4.7 # 4.2 ± 0.6 MET ± ± 1.0 MET ± ± 0.6 MET ± ± 0.7 Data is represented as mean ± SD (n=6 per group). For the RCR with pyruvate, the interaction between genotype and treatment was significant and a pairwise analysis of differences is provided by Bonferroni-corrected two-sided unpaired t-tests: # P<0.05 when compared with lean animals of the same treatment regimen, P<0.05 when compared with MET30-treated animals of the same genotype. Mitochondrial function after in vitro incubation with metformin In order to assess whether metformin would affect mitochondrial respiratory capacity in vitro, mitochondria were isolated from TA muscle excised from lean and diabetic rats, and incubated with 1 mm metformin for 5 min. Complex I- and Complex II-dependent OXPHOS respiratory capacity were then determined and normalized to OXPHOS respiratory capacity measured in the isolated mitochondria without addition of metformin (Figure 5). Complex I-dependent respiratory capacity in the OXPHOS state decreased 28% after in vitro incubation with metformin, independent of genotype (P<0.001). In contrast, incubation of isolated mitochondria with metformin did not affect Complex II-dependent respiratory capacity. O 2 flux (treated/control OXPHOS) * Lean * Diabetic pyruvate succinate Figure 5. Mitochondrial function after in vitro incubation with metformin O 2 flux measured in mitochondria isolated from TA muscle of lean and diabetic ZDF rats after 5 min of incubation with metformin (1 mm), normalized to O 2 flux measured in isolated mitochondria without addition of metformin. Respiratory capacity was determined in the OXPHOS state, when mitochondrial respiration is coupled to ATP synthesis, fueled with either pyruvate plus malate (Complex I respiration) or succinate plus rotenone (Complex II respiration). Data is represented as mean ± SD (n=6 per group). Incubation with metformin significantly lowered OXPHOS respiration fueled with pyruvate plus malate, independent of genotype (ANOVA: * P<0.001). Metformin did not affect Complex II respiration. 128

130 Effect of metformin on muscle mitochondrial function Discussion A number of in vitro studies have shown that metformin inhibits Complex I of the mitochondrial respiratory chain 3-9. However, the significance of this inhibition for in vivo skeletal muscle mitochondrial function has yet to be elucidated. The aim of this study was to clarify to which extent metformin affects in vivo and ex vivo skeletal muscle oxidative capacity. To this end we assessed the mitochondrial response to 2 weeks of treatment with metformin (0, 30, 100 or 300 mg/kg body weight/day) in a rat model of diabetes using 31 P MRS and HRR, respectively. We showed that 2 weeks of treatment with metformin impairs in vivo muscle oxidative capacity in a dose-dependent manner, both in healthy and in diabetic rats. Whereas a dosage of 30 mg/kg/ day had no significant effect, in vivo oxidative capacity was 21% and 48% lower after 2 weeks of metformin treatment at 100 and 300 mg/kg/day, respectively, independent of genotype. HRR measurements demonstrated a similar dose-dependent effect of metformin on ex vivo respiratory capacity with a Complex I-dependent substrate, whereas Complex II-dependent respiratory capacity was largely unaffected. In contrast to the current belief that metformin has only a mild effect on mitochondrial function 30, we observed that metformin may severely impair skeletal muscle oxidative capacity in vivo, depending on the dosage. Two weeks of metformin treatment at 300 mg/kg/day led to a 2-fold reduction in the rate of PCr recovery after muscle stimulation, both in lean and diabetic rats, which is comparable to the 40% lower PCr recovery rate found in sedentary individuals as compared with endurance athletes, who run a minimum of 30 miles per week 31. At 100 mg/kg/ day, metformin had a more moderate effect on in vivo muscle oxidative capacity, while at 30 mg/ kg/day no significant effect on PCr recovery was observed. Patients with type 2 diabetes typically receive an oral dose of metformin of approximately 30 mg/kg/day. It should be noted though that the bioavailability of metformin in the systemic circulation after oral treatment is lower in rats (F=30%) 22 compared with patients (F=56%) 32. Therefore treatment with ~ mg/kg/ day metformin in rats is considered to be more clinically relevant, also because therapeutic effects of metformin treatment in rats at that dosage are similar to the effects in patients treated with 30 mg/kg/day metformin 3, It thus seems likely that patients with type 2 diabetes, possibly already featuring some level of mitochondrial impairment, will be affected in daily life functioning or when performing exercise as a consequence of treatment with metformin. Interestingly, Braun et al. observed a small but significant (2.7%) decrease in whole-body peak aerobic capacity (peak VO 2 ) in healthy volunteers after 9-12 days of treatment with 2000 mg/day (which equals 30 mg/ kg body weight/day) metformin 30. Moreover, a study by Sharoff et al. demonstrated that exerciseinduced improvement of whole-body insulin sensitivity is lost in insulin-resistant individuals taking metformin 15. Their findings essentially imply that metformin treatment in these patients limits their ability to improve their peripheral insulin sensitivity through exercise. 5 In vivo skeletal muscle oxidative capacity is determined by intrinsic mitochondrial function as well as mitochondrial content. However, we did not find a difference in relative mtdna copy number or PGC-1α protein expression (not shown) between rats treated with metformin (300 mg/kg/ day) and water-treated controls, which implies that the observed reduction in in vivo skeletal muscle oxidative capacity after metformin treatment is not caused by a decrease in mitochondrial content. In fact, Suwa et al. 33 reported enhanced protein expression of PGC-1α and increased citrate synthase activity in Wistar rats after 2 weeks of treatment with metformin, suggesting a stimulation of mitochondrial biogenesis. However, the dosage regimen used in that study was twice as high as the highest dosage used in the present study, which might explain why we did 129

131 Chapter 5 not observe an effect on mitochondrial biogenesis. Our results are in agreement with other rodent studies showing that 2 or 4 weeks of metformin treatment at ~300 mg/kg/day does not lead to increased activity of citrate synthase 21, 34. In order to evaluate intrinsic mitochondrial function, we performed HRR measurements in isolated muscle mitochondria from rats treated with metformin using both Complex I- and Complex IIdependent substrates. Two weeks of treatment with metformin affected Complex I-dependent respiratory capacity in the OXPHOS state, similar to the dose-dependent effect observed for in vivo muscle oxidative capacity. At 300 mg/kg/day, Complex I-dependent respiratory capacity in the OXPHOS state was ~40% lower than in water-treated controls, which is comparable to the 48% reduction in in vivo muscle oxidative capacity. For lean rats, the effect of metformin on Complex I-dependent respiratory capacity in the OXPHOS state was similar to that in the ETS state, indicating that the effect of metformin is confined to the respiratory chain. Moreover, metformin did not increase respiration in the LEAK state. Therefore, it seems that the effect of metformin on in vivo muscle oxidative capacity can be fully explained by its inhibition of Complex I-dependent respiration. The inhibitory action of metformin on Complex I-dependent respiration has been demonstrated before in in vitro studies, in which isolated mitochondria from rat liver 3-5 and skeletal muscle 4, as well as permeabilized cells 3,6,7 were incubated with metformin. Reports on Complex I activity in cultured cells further support an inhibitory effect of metformin on Complex I 4,7. In contrast, other ex vivo animal studies reported no effects on the respiratory capacity of permeabilized muscle fibers obtained from the oxidative part of the gastrocnemius of obese Zucker rats after 4 weeks of treatment with metformin (320 mg/kg/day) 34 and the predominantly glycolytic TA of wild type mice after 2 weeks of treatment with metformin (300 mg/kg/day) 21. Likewise, it was shown that in permeabilized vastus lateralis muscle fibers of type 2 diabetes patients treated with metformin (2000±200 mg/day) Complex I-dependent respiratory capacity was not different compared with healthy control subjects, indicating that mitochondrial Complex I respiration is not inhibited by metformin 35. Surprisingly, in L6 muscle cell cultures 36 and in skeletal muscle of kinase dead AMPK mice 21 metformin even increased mitochondrial energy formation. The discrepancies across the literature could be caused by differences in species, dosing regimens, muscle fiber types, and the methods used to determine the effect of metformin on the mitochondria. However, when comparing our results with the ex vivo animal studies of Kane et al. 34 and Kristensen et al. 21, in which metformin did not affect mitochondrial respiration in either oxidative or glycolytic muscle from rats or mice after 2-4 weeks of metformin treatment at ~300mg/kg/day, it seems that all except methodological differences can be excluded. In the current study mitochondria were isolated from a whole TA muscle to allow comparison with the in vivo data, while Kane et al. and Kristensen et al. used permeabilized muscle fibers. It has recently been reported that the respiratory response in permeabilized fibers can be different from that of isolated mitochondria 37. In this study, Complex II-dependent OXPHOS respiratory capacity was largely unaffected by metformin treatment. This is in agreement with previous reports showing that metformin has no effect on Complex II-dependent respiratory capacity 4,6. Schäfer and Rieger postulated that metformin inhibits the activity of the oxidative phosphorylation enzymes by binding to the mitochondrial membrane phospholipids and modifying physicochemical membrane properties 10. Following this reasoning, it is not surprising that the activity of Complex I, the largest and most complex enzyme among the enzymes involved in the oxidative phosphorylation pathway, is impaired the most by metformin. Our observation that Complex II-dependent OXPHOS capacity in mitochondria from diabetic rats treated with 300 mg/kg/day metformin was lower than for 130

132 Effect of metformin on muscle mitochondrial function diabetic rats treated with 100 mg/kg/day metformin suggests that the activity of Complex II or of downstream electron transport chain complexes (i.e Complex III and/or IV) is impaired by a high dosage of metformin. This inhibitory effect could be partially caused by a progressively larger derangement of the inner mitochondrial membrane by a high concentration of metformin 10, impairing the activity of smaller ETC complexes. Possibly, mitochondria from skeletal muscle of diabetic animals are more sensitive to the toxic metformin effect due to other factors related to the diseased environment, since we do not observe inhibition of Complex II-dependent respiration in mitochondria from lean animals. However, our results on Complex I-dependent respiration do not support the notion that mitochondria from diabetic muscle are more sensitive to metformininduced membrane derangements. Although it is well established that metformin attenuates Complex I-dependent respiratory capacity, the mechanism through which metformin exerts its inhibitory action on Complex I is still subject of debate. A number of reports propose an indirect pathway, involving cell membrane events, via which metformin affects mitochondrial respiration 6,8. This is based on the observation that inhibition of Complex I is lost when metformin is added to mitochondria isolated from their cellular environment 6 or when metformin is micro-injected into the interior of an intact oocyte, suggesting membrane-mediated events are necessary for this effect to occur 6,8. Others, however, have shown that metformin does inhibit Complex I in mitochondria isolated from skeletal muscle and liver 3,4, thus contradicting the suggestion that an intact cell is needed for metformin to exert its effect on mitochondrial function. Early work of Schäfer and Rieger 10 showed that biguanides have an affinity to directly bind to mitochondrial membrane phospholipids, causing the accumulation of positive charge at the membrane surface, thereby rendering the electrostatic surface potential more positive. This will alter the physicochemical properties of the mitochondrial membrane, which may underlie the inhibition of metformin of Complex I and which supports a direct pathway for metformin to affect mitochondria. In order to determine whether metformin affects mitochondrial respiratory capacity via a direct or indirect pathway, we studied mitochondrial respiration after incubating isolated mitochondria with 1 mm metformin for 5 min. We observed a 28% inhibition of Complex I-dependent respiratory capacity, whereas Complex II-dependent respiratory capacity was unaffected. Our findings thus imply that metformin inhibits mitochondrial respiration through Complex I via a direct pathway. 5 Apart from the effects of metformin on muscle mitochondrial function, we observed that in vivo muscle oxidative capacity was 25% lower in diabetic rats compared with lean control animals, independent of treatment regimen. However, relative mtdna copy number and Complex I- and Complex II-dependent respiratory capacity were similar between diabetic and lean animals, which implies that neither a lower mitochondrial content nor an impairment of their ex vivo intrinsic function can account for the lower in vivo muscle oxidative capacity in diabetic rats. Instead, it suggests that in diabetic muscle the functioning of mitochondria in their natural cellular environment is impaired by factors that are not taken into account during the ex vivo measurements in isolated mitochondria, such as lipid-induced mitochondrial uncoupling 27. The beneficial effects of metformin on glucose homeostasis are well established both in patient and animal studies 19,34, However, in this study no changes in fasting plasma levels of glucose or insulin were observed in any of the animal groups after 2 weeks of treatment with metformin. It should be noted, though, that the therapy duration in our study was shorter than in the animal studies in which improved glucose tolerance was observed (typically 3 to 4 weeks) 34, which might explain why plasma parameters were unaffected in our study. 131

133 Chapter 5 There are indications that the inhibition of Complex I contributes to metformin s therapeutic efficacy. It is well-known that metformin lowers blood glucose levels primarily by lowering glucose production in the liver, which is an ATP-dependent process. Therefore it is possible that the reduction of mitochondrial oxidative capacity underlies the mechanism through which metformin suppresses glucose release from the liver 39. Moreover, there are several reports indicating that metformin promotes glucose uptake in peripheral tissues, thus contributing to its antihyperglycemic efficacy. This could be conciliated with its action on mitochondria by the energy charge hypothesis postulated by Brunmair et al. 4. This hypothesis states that agents that interfere with Complex I-dependent cellular respiration affect enzymes like AMP-dependent protein kinase and hence induce a metabolic response, such as increased glucose uptake and glycolysis, to compensate for decreased ATP synthesis rates 3,4. In conclusion, we demonstrated that 2 weeks of treatment with metformin compromised in vivo and ex vivo muscle oxidative capacity in ZDF rats in a dose-dependent manner. Moreover, our finding that also in vitro incubation of isolated mitochondria with metformin lowers Complex I-dependent respiratory capacity supports the hypothesis that metformin inhibits Complex I via a direct pathway. Acknowledgements We thank Leonie Niesen and David Veraart for their assistance in animal handling. 132

134 Effect of metformin on muscle mitochondrial function References 1. Hundal, R.S., et al. Mechanism by which metformin reduces glucose production in type 2 diabetes. Diabetes 49, (2000). 2. Argaud, D., Roth, H., Wiernsperger, N. & Leverve, X.M. Metformin decreases gluconeogenesis by enhancing the pyruvate kinase flux in isolated rat hepatocytes. European journal of biochemistry / FEBS 213, (1993). 3. Owen, M.R., Doran, E. & Halestrap, A.P. Evidence that metformin exerts its anti-diabetic effects through inhibition of complex 1 of the mitochondrial respiratory chain. Biochem J 348 Pt 3, (2000). 4. Brunmair, B., et al. Thiazolidinediones, like metformin, inhibit respiratory complex I: a common mechanism contributing to their antidiabetic actions? Diabetes 53, (2004). 5. Carvalho, C., et al. Metformin promotes isolated rat liver mitochondria impairment. Mol Cell Biochem 308, (2008). 6. El-Mir, M.Y., et al. Dimethylbiguanide inhibits cell respiration via an indirect effect targeted on the respiratory chain complex I. J Biol Chem 275, (2000). 7. Guigas, B., et al. Metformin inhibits mitochondrial permeability transition and cell death: a pharmacological in vitro study. Biochem J 382, (2004). 8. Detaille, D., Guigas, B., Leverve, X., Wiernsperger, N. & Devos, P. Obligatory role of membrane events in the regulatory effect of metformin on the respiratory chain function. Biochem Pharmacol 63, (2002). 9. Palenickova, E., Cahova, M., Drahota, Z., Kazdova, L. & Kalous, M. Inhibitory effect of metformin on oxidation of NADH-dependent substrates in rat liver homogenate. Physiological research / Academia Scientiarum Bohemoslovaca 60, (2011). 10. Schafer, G. & Rieger, E. Interaction of biguanides with mitochondrial and synthetic membranes. Effects on ion conductance of mitochondrial membranes and electrical properties of phospholipid bilayers. European journal of biochemistry / FEBS 46, (1974). 11. Szendroedi, J., Phielix, E. & Roden, M. The role of mitochondria in insulin resistance and type 2 diabetes mellitus. Nature reviews. Endocrinology 8, (2012). 12. Morino, K., Petersen, K.F. & Shulman, G.I. Molecular mechanisms of insulin resistance in humans and their potential links with mitochondrial dysfunction. Diabetes 55 Suppl 2, S9-S15 (2006). 13. ADA. Standards of Medical Care in Diabetes Diabetes care 36, S11-S66 (2013). 14. Praet, S.F. & van Loon, L.J. Optimizing the therapeutic benefits of exercise in Type 2 diabetes. J Appl Physiol 103, (2007). 15. Sharoff, C.G., et al. Combining short-term metformin treatment and one bout of exercise does not increase insulin action in insulin-resistant individuals. Am J Physiol Endocrinol Metab 298, E (2010). 16. Penicaud, L., Hitier, Y., Ferre, P. & Girard, J. Hypoglycaemic effect of metformin in genetically obese (fa/fa) rats results from an increased utilization of blood glucose by intestine. Biochem J 262, (1989). 17. Wilcock, C. & Bailey, C.J. Sites of metformin-stimulated glucose metabolism. Biochem Pharmacol 39, (1990). 18. Rouru, J., et al. Effects of metformin treatment on glucose transporter proteins in subcellular fractions of skeletal muscle in (fa/fa) Zucker rats. Br J Pharmacol 115, (1995). 19. Smith, A.C., et al. Metformin and exercise reduce muscle FAT/CD36 and lipid accumulation and blunt the progression of high-fat diet-induced hyperglycemia. Am J Physiol Endocrinol Metab 293, E (2007). 20. Bailey, C.J. & Puah, J.A. Effect of metformin on glucose metabolism in mouse soleus muscle. Diabete & metabolisme 12, (1986). 21. Kristensen, J.M., Larsen, S., Helge, J.W., Dela, F. & Wojtaszewski, J.F. Two weeks of metformin treatment enhances mitochondrial respiration in skeletal muscle of AMPK kinase dead but not wild type mice. PloS one 8, e53533 (2013). 22. Choi, Y.H., Kim, S.G. & Lee, M.G. Dose-independent pharmacokinetics of metformin in rats: Hepatic and gastrointestinal first-pass effects. Journal of pharmaceutical sciences 95, (2006). 23. De Feyter, H.M., et al. Increased intramyocellular lipid content but normal skeletal muscle mitochondrial oxidative capacity throughout the pathogenesis of type 2 diabetes. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 22, (2008)

135 Chapter Vanhamme, L., van den Boogaart, A. & Van Huffel, S. Improved method for accurate and efficient quantification of MRS data with use of prior knowledge. J Magn Reson 129, (1997). 25. Taylor, D.J., Bore, P.J., Styles, P., Gadian, D.G. & Radda, G.K. Bioenergetics of intact human muscle. A 31 P nuclear magnetic resonance study. Molecular biology & medicine 1, (1983). 26. van den Broek, N.M., De Feyter, H.M., de Graaf, L., Nicolay, K. & Prompers, J.J. Intersubject differences in the effect of acidosis on phosphocreatine recovery kinetics in muscle after exercise are due to differences in proton efflux rates. American journal of physiology. Cell physiology 293, C (2007). 27. van den Broek, N.M., et al. Increased mitochondrial content rescues in vivo muscle oxidative capacity in longterm high-fat-diet-fed rats. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 24, (2010). 28. Ciapaite, J., et al. Functioning of oxidative phosphorylation in liver mitochondria of high-fat diet fed rats. Biochim Biophys Acta 1772, (2007). 29. Gnaiger, E. Capacity of oxidative phosphorylation in human skeletal muscle: new perspectives of mitochondrial physiology. Int J Biochem Cell Biol 41, (2009). 30. Braun, B., et al. Impact of metformin on peak aerobic capacity. Appl Physiol Nutr Metab 33, (2008). 31. Larsen, R.G., Callahan, D.M., Foulis, S.A. & Kent-Braun, J.A. In vivo oxidative capacity varies with muscle and training status in young adults. J Appl Physiol 107, (2009). 32. Graham, G.G., et al. Clinical pharmacokinetics of metformin. Clinical pharmacokinetics 50, (2011). 33. Suwa, M., Egashira, T., Nakano, H., Sasaki, H. & Kumagai, S. Metformin increases the PGC-1alpha protein and oxidative enzyme activities possibly via AMPK phosphorylation in skeletal muscle in vivo. J Appl Physiol 101, (2006). 34. Kane, D.A., et al. Metformin selectively attenuates mitochondrial H 2 O 2 emission without affecting respiratory capacity in skeletal muscle of obese rats. Free radical biology & medicine 49, (2010). 35. Larsen, S., et al. Metformin-treated patients with type 2 diabetes have normal mitochondrial complex I respiration. Diabetologia 55, (2012). 36. Vytla, V.S. & Ochs, R.S. Metformin increases mitochondrial energy formation in L6 muscle cell cultures. J Biol Chem 288, (2013). 37. Picard, M., et al. Mitochondrial structure and function are disrupted by standard isolation methods. PloS one 6, e18317 (2011). 38. Klip, A. & Leiter, L.A. Cellular mechanism of action of metformin. Diabetes care 13, (1990). 39. Leverve, X.M., et al. Mitochondrial metabolism and type-2 diabetes: a specific target of metformin. Diabetes Metab 29, 6S88-94 (2003). 40. Rossetti, L., et al. Effect of metformin treatment on insulin action in diabetic rats: in vivo and in vitro correlations. Metabolism: clinical and experimental 39, (1990). 41. Inzucchi, S.E., et al. Efficacy and metabolic effects of metformin and troglitazone in type II diabetes mellitus. N Engl J Med 338, (1998). 134

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137 Metformin treatment impairs in vivo skeletal muscle oxidative capacity as well as contractile function in diabetic rats Bart Wessels 1, Joep P. Schmitz 1, Robert W. Wiseman 2, Klaas Nicolay 1 and Jeanine J. Prompers 1 1 Biomedical NMR, Department of Biomedical Engineering, Eindhoven University of Technology, the Netherlands, 2 Department of Physiology, Michigan State University, East Lansing, MI, USA

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140 Metformin affecting muscle contractile function Abstract Aim: We previously demonstrated that metformin impairs in vivo muscle oxidative capacity in both healthy and diabetic rats in a dose-dependent manner. In this study we investigated the effect of metformin treatment on skeletal muscle energy metabolism and muscle contractile function in a rat model of diabetes. Materials and Methods: 31 P magnetic resonance spectroscopy was performed on 14-week old lean fa/+ (LN), diabetic fa/fa (Db) and metformin-treated (300 mg/kg/day for 14 days) diabetic fa/fa (DbM) Zucker Diabetic Fatty rats (n=8 per group) for the in vivo assessment of muscle oxidative capacity and intracellular ph changes during muscle stimulation. Muscle contractile properties were evaluated by determining the peak force, time-tension integral (TTI) and muscle relaxation time from force measurements with a home-built force transducer. Results: Muscle oxidative capacity was 21% lower in Db and 54% lower in DbM rats compared with the LN animals, and it was 42% lower in DbM rats compared with Db rats (P<0.001). Intracellular proton accumulation during 3 min of muscle stimulation was higher in both Db (P<0.05) and DbM (P<0.001) rats compared with LN animals. Reductions in peak force and TTI during the stimulation protocol did not differ among groups, indicating that muscle fatigue was not affected in Db and DbM rats compared with LN animals. However, we found that the muscle relaxation time was elongated in Db and DbM rats compared with LN animals (P<0.001) and in DbM rats compared with Db rats (P<0.001). Conclusion: Metformin treatment in diabetic rats increases the rate of intracellular muscle acidosis and delays muscle relaxation during muscle contractions. These findings demonstrate that treatment with metformin impairs skeletal muscle function, which suggests that metformin could lower the ability of patients with type 2 diabetes to perform exercise

141 Chapter 6 Introduction Type 2 diabetes is a metabolic disorder that is reaching epidemic proportions worldwide and the rapid rise in the number of type 2 diabetes patients has been associated with the increasing prevalence of obesity 1. One of the underlying mechanisms implicated in the etiology of insulin resistance and type 2 diabetes is skeletal muscle mitochondrial dysfunction 2-4. However, whether the latter is truly a cause or rather a consequence of insulin resistance is under debate 5-7. The primary role of mitochondria is to generate ATP through oxidative phosphorylation, which is the main source of ATP under aerobic conditions. An alternative route for ATP production is through glycolysis, which also functions during anaerobic conditions, since this pathway does not require oxygen. Although glycolysis is mostly considered to be an emergency back-up of cellular energy homeostasis, glycolysis and oxidative phosphorylation are actually complementary processes, and their relative contributions to ATP delivery depend on the timescale and level of energy demands 8. Mitochondrial oxidative phosphorylation is highly efficient and provides a steady rate of ATP supply. The glycolytic pathway, on the other hand, is less efficient, but produces ATP at high conversion rates and can be activated within milliseconds in order to meet high and/or highly fluctuating energy demands 9. Metformin is the most commonly prescribed drug to treat type 2 diabetes. Interestingly, it has been shown that metformin inhibits Complex I of the mitochondrial respiratory chain We previously demonstrated that metformin impairs in vivo muscle oxidative capacity in both healthy and diabetic rats in a dose-dependent manner 17. However, in type 2 diabetes patients the only serious side-effect of metformin that has been reported is lactic acidosis 18. Possibly, lactic acidosis during metformin treatment is due to an increase in glycolysis flux as an adaptive response to compensate for impaired mitochondrial oxidative phosphorylation capacity to fulfill the energy demand of the cell. Considering the benefits of regular physical activity in the treatment of diabetes patients 19-21, it seems unfavorable to treat diabetes patients with a drug that limits their oxidative ATP production. Metformin could attenuate their exercise capacity and consequently their ability to increase insulin sensitivity via exercise training, as was observed by Sharoff et al. 22. In addition to impairments in muscle energy metabolism, type 2 diabetes has been associated with muscle contractile dysfunction, as demonstrated by delayed myofibrillar relaxation 23,24. An increase in muscle relaxation time can be related to a decrease in the rate of uptake of Ca 2+ by the sarcoplasmic reticulum 25 and a lower dissociation rate of myofibrillar cross-bridges 26, which are both dependent on the availability of ATP 27. Safwat et al. found that the increased delay of muscle contractile relaxation in diabetic rats was associated with a reduced expression and activity of sarco- and endoplasmic reticulum Ca 2+ ATPase (SERCA) 24. Interestingly, it has been suggested that AMPK activators, including metformin, attenuate endoplasmic reticulum stress by inhibiting SERCA oxidation and maintaining SERCA activity 28. The aim of this study was to determine the effect of metformin on skeletal muscle energy metabolism and contractile function in a rat model of diabetes. Healthy, lean and obese, diabetic Zucker Diabetic Fatty (ZDF) rats were used in this study. One group of diabetic rats was orally dosed with metformin (300 mg/kg/day) for 2 weeks. Muscle intracellular ph dynamics and oxidative capacity were measured in vivo using 31 P magnetic resonance spectroscopy (MRS) during and after electrically induced muscle contractions, respectively. A home-built force transducer was used to perform force measurements during muscle stimulation in the MR scanner. 140

142 Metformin affecting muscle contractile function Materials and methods Animals Lean, healthy fa/+ and obese, diabetic fa/fa adult male ZDF rats (14 weeks of age) were purchased from Charles River Laboratories (Sulzfield, Germany) and given ad libitum access to water and standardized chow for ZDF rats (Purina Formula 5008, Bioservices, the Netherlands). The animals were housed in pairs in a controlled environment (20 C and 50% relative humidity), on a 12-h lightdark cycle. The animals were divided into three experimental groups: A group of untreated lean rats (LN; n = 8), a group of untreated diabetic animals (Db; n = 8), and a group of diabetic animals treated with metformin (DbM; n = 8). The DbM rats were dosed daily with metformin (300 mg/kg body weight, n = 8) in 1 ml of water directly into the stomach by oral gavage for 14 days. After 14 days of treatment, in vivo MRS experiments were performed and after the in vivo measurements, the animals were killed under anesthesia by incision of the vena cava. All experimental procedures were reviewed and approved by the Animal Experimental Committee of Maastricht University. 31 P Magnetic Resonance Spectroscopy All MRS measurements were performed on a 6.3-T horizontal Bruker MR scanner (Bruker, Ettlingen, Germany). The animals were anaesthetized using isoflurane (2-3%) combined with medical air (0.6 L/min) and their body temperature was maintained at 36 ± 1 C using heating pads. A pressure sensor registering thorax movement (Rapid Biomedical, Rimpar, Germany) was used to monitor respiration. 31 P MRS was performed with an ellipsoid (10/18 mm) 31 P surface coil, as described previously 17,29. An adiabatic excitation pulse with a flip angle of 90 was used to obtain 31 P MR spectra of the tibialis anterior (TA) muscle. A fully relaxed spectrum (repetition time TR=20 s, 32 averages) was recorded first, followed by a time series of spectra (TR=5 s, 4 averages) consisting of a resting period of 3 min, 3 min of electrical stimulation, and min of recovery. The TA muscle was stimulated via subcutaneously implanted electrodes positioned along the distal nerve trajectory of the N. peroneus communis. Stimulation pulses were applied every second (during the relaxation delay of the MRS experiment) with a stimulation pulse train with a length of 100 ms and a frequency of 80 Hz. The voltage of the stimulation pulses (3-5 V) was optimized at the start of the protocol to reach maximal contractile force, as measured by a home-built force transducer, in order to obtain similar workloads for the different animals P MR spectra were fitted in the time domain using a nonlinear least squares algorithm (advanced method for accurate, robust, and efficient spectral fitting; AMARES) in the jmrui software package 30 as described previously 17,29. In short, spectral analysis of the 31 P MR spectra was done by fitting the phosphocreatine (PCr) peak to Lorentzian and the inorganic phosphate (P i ) as well as the α-, β- and γ-atp peaks to Gaussian line shapes. Absolute concentrations of the phosphorylated metabolites were calculated after correction for partial saturation with the assumption that the ATP concentration is 8.2 mm at rest 31. Intracellular ph was calculated from the chemical shift difference between the P i and PCr resonances (δ; in parts per million or ppm) using Equation 1 32 : δ 3.27 ph=6.75+log (1) 5.63 δ For the determination of intracellular ph during recovery, 3 spectra were averaged (time resolution is 1 min) to improve the signal-to-noise ratio for the P i peak. Intracellular proton concentration, [H + ], 141

143 Chapter 6 was derived from the intracellular ph according to: [H + ] = 10 -ph. Intracellular proton accumulation during the 3 min stimulation protocol, [H + ], was calculated from the difference of [H + ] at the end of stimulation and [H + ] before the start of stimulation. The initial PCr consumption at the start of muscle stimulation, [PCr], was calculated as the decrease in PCr concentration during the first minute of the stimulation protocol 33. The concentrations of PCr determined during recovery were fit to a mono-exponential function (Figure 1D) using Matlab (version R2010b, Mathworks, Natick, MA, USA) yielding a rate constant, k PCr, which is a measure of in vivo skeletal muscle mitochondrial oxidative capacity. Force measurements During the muscle stimulation protocol and the acquisition of the 31 P MR spectra, the force produced by the dorsal flexor muscles was measured using a home-built, MR compatible force transducer. Data acquisition was performed at a sampling rate of 2000 Hz with LabView software (National Instruments, Woerden, The Netherlands). Before the start of the MRS experiment, the optimal voltage for the muscle stimulation was determined by inducing single contractions at increasing voltages until no further increase in force production was detected. The force produced during the muscle stimulation protocol was normalized relative to the maximal contractile force at the start of the stimulation protocol. The time-tension integral (TTI) was calculated using Matlab 2010b (Mathworks, USA) as: tend 1 TTI= Force dt (2) T tot t start where t start and t end represent the start and end times of a single contraction, respectively, Force is the force produced during muscle contractions normalized to the initial maximal contractile force, and T tot is the total period of contraction and relaxation (defined as the time period between the first point after the force reached >10% of peak force and the first point after the force declined to <10% of peak force post-contraction) 34. The muscle relaxation time was determined as the time period between the end of muscle contraction/start of muscle relaxation (defined as the first point after the force dropped to <90% of peak force) and the end of muscle relaxation (defined as the first point after the force declined to <10% of peak force). Statistical analysis Data are presented as means ± SD. The listed n values represent the number of animals used for a particular experiment. Statistical significance of the differences was assessed by applying a one-way analysis of variance (ANOVA) using Bonferroni corrected post-hoc analyses in the SPSS 20.0 statistical package (SPSS Inc., Chicago, IL, USA). The level of statistical significance was set at P<

144 Metformin affecting muscle contractile function Results 31 P Magnetic Resonance Spectroscopy 31 P MRS was applied to assess in vivo muscle energy metabolism during and after contractions. In Figure 1A a representative 31 P MR spectrum obtained from TA muscle of a lean rat at rest is depicted. Figure 1B shows the concentrations of phosphorous metabolites obtained from the 31 P MR spectra acquired before, during and after the muscle stimulation protocol. During muscle stimulation, ATP levels are maintained at resting-state values, while the PCr concentration drops and the P i concentration rises. During the recovery phase after electrical stimulation has ended, PCr and P i levels return to resting-state values. An example of the time course of intracellular ph dynamics during muscle stimulation and recovery is depicted in Figure 1C. Group averages for the PCr and P i concentrations and intracellular ph measured in TA muscle at rest and at the end of muscle stimulation are listed in Table 1. Table 1 Metabolite concentrations and ph measured using 31 P MRS in TA muscle of lean ZDF rats (LN), diabetic ZDF rats (Db), and diabetic ZDF rats treated with metformin (DbM). LN Db DbM Rest parameters ph 7.16 ± ± ± 0.06 [PCr] (mm) 34.6 ± ± ± 2.2 [P i ] (mm) 1.8 ± ± ± 0.5 End-Stimulation parameters ph 6.55 ± ± 0.02** 6.33 ± 0.10*** [PCr] (mm) 10.0 ± ± 1.4* 9.4 ± 1.3 [P i ] (mm) 24.7 ± ± ± 5.2 Data are represented as means ± SD (n=8 per group). * P<0.05, ** P<0.01, *** P<0.001 when compared with LN rats, P <0.01 when compared with Db rats. At rest, concentrations of PCr and P i and intracellular ph did not differ between groups. At the end of the stimulation protocol, the PCr concentration was slightly higher in diabetic rats (Db) compared with lean rats (LN; P<0.05) and compared with diabetic metformin-treated rats (DbM; P<0.01)

145 Chapter 6 A B C D Figure 1. In vivo 31 P MRS in TA muscle Representative example of 31 P MRS in tibialis anterior (TA) muscle of a lean ZDF rat. (A) 31 P MR spectrum measured at rest (36 averages). (B) PCr, P i and ATP concentrations determined from 31 P MR spectra acquired during 3 min of rest, 3 min muscle stimulation and 15 min recovery (time resolution is 20 s). (C) Intracellular ph calculated from the chemical shift difference between the P i and PCr resonances during muscle stimulation and recovery. During recovery 3 spectra were averaged (time resolution is 1 minute) to improve the signal-to-noise ratio for the P i peak. (D) PCr concentrations during recovery fit with a mono-exponential function (solid line), yielding the rate constant of PCr recovery, k PCr, which was 0.52 min -1 for this specific example. However, no differences were detected in [PCr] during the first minute of muscle stimulation (Figure 2A), which indicates that ATP consumption at the start of the muscle stimulation protocol was similar among groups. Intracellular muscle ph dropped more quickly in Db and DbM rats during muscle stimulation compared with the LN animals (Figure 2B). End-stimulation ph was lower in both Db (P<0.01) and DbM rats (P<0.001) with respect to LN animals (Table 1). Furthermore, the end-stimulation ph in DbM rats tended to be lower than in Db rats (P=0.08). Intracellular proton accumulation during the 3 min of muscle stimulation, [H + ], was higher in both Db (P<0.05) and DbM rats (P<0.001) compared with LN animals (Figure 2C). 144

146 Metformin affecting muscle contractile function A B LN Db DbM [PCr] (mm) ph (-) LN Db DbM ** *** Time (sec) C 0.4 *** D 0.6 [H + ] (mm) * k PCr (min -1 ) * *** LN Db DbM 0.0 LN Db DbM 6 Figure 2. Results from 31 P MRS measurements of TA muscle Results from 31 P MRS measurements in tibialis anterior (TA) muscle of lean ZDF rats (LN), diabetic ZDF rats (Db), and diabetic ZDF rats treated with metformin (DbM). Data are represented as means ± SD (n=8 per group). (A) PCr consumption during the first minute of the stimulation protocol, [PCr]. (B) Time course of intracellular ph during muscle stimulation (error bars are omitted for better visibility of the means). (C) Intracellular proton accumulation during the 3 min stimulation protocol, [H + ]. (D) PCr recovery rate constant, k PCr. * P<0.05, ** P<0.01, *** P<0.001 when compared with LN rats, P<0.001 when compared with Db rats. A mono-exponential function was fitted through the PCr concentrations obtained during the recovery phase (Figure 1D), yielding the PCr recovery rate constant, k PCr, which is representative for muscle oxidative capacity in vivo. k PCr was 21% lower in Db rats (P<0.05) and 54% lower in DbM rats (P<0.001) compared with LN animals (Figure 2D). Moreover, k PCr was 42% lower in DbM rats compared with Db rats (P<0.001). 145

147 Chapter 6 Figure 3. Force measurements (A) Result of a typical force measurement in a lean ZDF rat, obtained during the 3 min stimulation protocol inside the MR scanner. Force was measured with a home-built force transducer at a sample rate of 2000 Hz, and was expressed relative to the maximum contractile force determined before the start of the muscle stimulation protocol. (B) Examples of a single contraction curves (close-up of a 1 sec time interval) towards the end of the stimulation protocol of a lean (LN), a diabetic (Db) and a diabetic metformin-treated (DbM) ZDF rat. Force measurements A representative example of a force measurement during the 3 min muscle stimulation protocol in a lean rat is depicted in Figure 3A. Figure 3B shows a close-up of the force measurement during a single contraction towards the end of the stimulation protocol for one animal of each experimental group. The relative reduction in peak force generation and TTI after 3 min of muscle stimulation did not significantly differ between groups, indicating that muscle fatigue was similar in all groups (Figures 4A and 4B). At the end of the muscle stimulation protocol, the muscle relaxation time was, however, elongated in both Db and DbM rats compared with LN animals (P<0.001; Figure 4C) and in DbM rats compared with Db rats (P<0.001). The slower relaxation of the muscles of Db and DbM rats towards the end of the stimulation protocol can also be appreciated from Figure 3B. Muscle relaxation time at the end of the stimulation protocol was significantly correlated with k PCr (r=-0.81, P<0.001; Figure 4D). 146

148 Metformin affecting muscle contractile function A Peak force (relative to max force) C Relaxation time (sec) 1.4 LN Db DbM contraction no (-) LN Db DbM *** *** B TTI (-) D Relaxation time (sec) LN Db DbM contraction no (-) P < r = LN Db DbM contraction no (-) k PCr (min -1 ) Figure 4. Results of force measurements Results from the force measurements during the muscle stimulation protocol in lean ZDF rats (LN), diabetic ZDF rats (Db), and diabetic ZDF rats treated with metformin (DbM). Data are represented as means ± SD (n=8 per group). (A) Peak force normalized relative to the maximal contractile force at the start of the stimulation protocol. (B) Time-tension integral (TTI). (C) Muscle relaxation time. (D) Correlation plot between k PCr and the muscle relaxation time at the end of the stimulation protocol

149 Chapter 6 Discussion The aim of this study was to determine the effect of metformin on skeletal muscle energy metabolism and contractile function in diabetic rats. In vivo muscle oxidative capacity was lower in diabetic rats compared with lean rats, and was further impaired in diabetic rats upon 2 weeks of treatment with metformin. The decreases in in vivo muscle oxidative capacity in diabetic rats and diabetic metformin-treated rats were associated with increases in intracellular proton accumulation during muscle stimulation, indicating a larger reliance on glycolytic ATP production. Muscle fatigue, as determined from reductions in peak force and TTI, was not affected by diabetes or metformin treatment. However, at the end of the muscle stimulation protocol, muscle relaxation was significantly longer in diabetic rats compared with lean rats, and was further prolonged upon metformin treatment in diabetic rats. In this study we found that in vivo muscle oxidative capacity was 21% lower in Db rats and 54% lower in DbM rats compared with the LN animals. These results are in agreement with findings of our previous study on the effects of metformin on in vivo muscle oxidative capacity 17. However, although the relative reductions in k PCr in Db and DbM rats with respect to LN animals are similar, the absolute values for k PCr are approximately 10-20% lower in this study compared with our previous report. The lower values for k PCr can be attributed to the lower end-stimulation ph that was observed in the present study, which has previously been shown to affect the rate of PCr resynthesis 35. We demonstrated that muscle ph dropped more quickly in untreated and metformin-treated diabetic rats during muscle stimulation, which was also reflected by the elevated proton accumulation after muscle stimulation in these animals compared with lean rats The faster intracellular acidosis and the elevated intracellular accumulation of protons indicate that in Db and DbM rats glycolytic activity during muscle stimulation is higher than in LN animals. Muscle fatigue, as determined from reductions in peak force and TTI, was similar among animals, irrespective of genotype or treatment. Earlier studies did report impairments in peak twitch tension and peak tetanic tension of diabetic rats 24,36,37. This reduction in force production was thought to be the result of the lower muscle mass of diabetic animals 37, but we corrected for differences in muscle mass by normalizing for the maximal contractile force at the start of the stimulation protocol. The force measurements further revealed that, towards the end of the muscle stimulation protocol, the muscle relaxation time was elongated in diabetic rats compared with lean rats. Moreover, treatment with metformin aggravated this delay in muscle relaxation in diabetic animals compared with the untreated diabetic animals. These findings confirm earlier reports of a delay in muscle relaxation in diabetic muscle 23,24. The time it takes for muscle to relax after contraction depends on the rate of myofibrillar cross-bridge cycling 26 and the uptake of Ca 2+ by the sarcoplasmic reticulum 25. The faster intracellular acidosis and elevated intracellular proton accumulation during muscle stimulationcould therefore explain the longer muscle relaxation times found in Db and DbM rats, since intracellular acidification is known to inhibit SERCA activity 38,39. However, although intracellular acidification has been associated with delayed muscle relaxation, it is not thought to be the primary cause for a decrease in the relaxation rate 40,41. Instead, Dawson et al. found that the muscle relaxation rate was linearly dependent on the free-energy change for ATP hydrolysis, which in turn may be related to the rate of Ca 2+ uptake by SERCA or the rate of myofibrillar cross-bridge cycling 40. Another possible explanation for differences in muscle relaxation times is related to the fact that the delivery of ATP is compartmentalized within cells and that some ATP-dependent cell functions primarily derive energy from mitochondrial ATP, while other cell functions are driven by glycolytic ATP 42. Mitochondria are spatially closely associated with 148

150 Metformin affecting muscle contractile function myofibrils and the sarcoplasmic reticulum membranes, and appear to be the preferred provider of ATP needed for sarcoplasmic calcium uptake 43,44. In addition, besides providing ATP for Ca 2+ uptake by the sarcoplasmic reticulum, mitochondria themselves play a role in Ca 2+ buffering, which has been shown to affect the relaxation rate after muscle contractions 45, dependent on muscle fiber type 46. All of these mechanisms do point to a causal link between the rate of mitochondrial ATP synthesis and the rate of muscle relaxation, which is corroborated by the strong correlation we found between k PCr and the muscle relaxation time (Figure 4D). Safwat et al. related the delay in muscle relaxation in diabetic rats to decreased SERCA expression and activity 24. Interestingly, AMPK activators, like metformin, are reported to reduce sarcoplasmic reticulum stress and recover SERCA activity 28. However, in this study we found that diabetic animals treated with metformin presented with a prolonged relaxation time compared with untreated diabetic animals. This finding indicates that metformin does not improve muscle contractile function as suggested previously 24, but instead exacerbates muscle contractile dysfunction in diabetic muscle. In conclusion, metformin treatment in diabetic rats increases the rate of intracellular muscle acidosis and delays muscle relaxation during muscle contractions. These findings demonstrate that treatment with metformin impairs skeletal muscle function, which suggests that metformin could lower the ability of patients with type 2 diabetes to perform exercise. Furthermore the correlation between muscle relaxation time and muscle oxidative capacity supports the hypothesis that delayed relaxation in diabetic muscle is caused by impaired mitochondrial ATP synthesis

151 Chapter 6 References 1. Sullivan, P.W., Morrato, E.H., Ghushchyan, V., Wyatt, H.R. & Hill, J.O. Obesity, inactivity, and the prevalence of diabetes and diabetes-related cardiovascular comorbidities in the U.S., Diabetes Care 28, (2005). 2. Lowell, B.B. & Shulman, G.I. Mitochondrial dysfunction and type 2 diabetes. Science 307, (2005). 3. Morino, K., Petersen, K.F. & Shulman, G.I. Molecular mechanisms of insulin resistance in humans and their potential links with mitochondrial dysfunction. Diabetes 55 Suppl 2, S9-S15 (2006). 4. Szendroedi, J., Phielix, E. & Roden, M. The role of mitochondria in insulin resistance and type 2 diabetes mellitus. Nat Rev Endocrinol 8, (2012). 5. Turner, N. & Heilbronn, L.K. Is mitochondrial dysfunction a cause of insulin resistance? Trends Endocrinol Metab 19, (2008). 6. Dumas, J.F., Simard, G., Flamment, M., Ducluzeau, P.H. & Ritz, P. Is skeletal muscle mitochondrial dysfunction a cause or an indirect consequence of insulin resistance in humans? Diabetes Metab 35, (2009). 7. Patti, M.E. & Corvera, S. The role of mitochondria in the pathogenesis of type 2 diabetes. Endocr Rev 31, (2010). 8. Epstein, T., Xu, L., Gillies, R.J. & Gatenby, R.A. Separation of metabolic supply and demand: aerobic glycolysis as a normal physiological response to fluctuating energetic demands in the membrane. Cancer Metab 2, 7 (2014). 9. Pfeiffer, T., Schuster, S. & Bonhoeffer, S. Cooperation and competition in the evolution of ATP-producing pathways. Science 292, (2001). 10. Brunmair, B., et al. Thiazolidinediones, like metformin, inhibit respiratory complex I: a common mechanism contributing to their antidiabetic actions? Diabetes 53, (2004). 11. Owen, M.R., Doran, E. & Halestrap, A.P. Evidence that metformin exerts its anti-diabetic effects through inhibition of complex 1 of the mitochondrial respiratory chain. Biochem J 348 Pt 3, (2000). 12. Carvalho, C., et al. Metformin promotes isolated rat liver mitochondria impairment. Mol Cell Biochem 308, (2008). 13. El-Mir, M.Y., et al. Dimethylbiguanide inhibits cell respiration via an indirect effect targeted on the respiratory chain complex I. J Biol Chem 275, (2000). 14. Guigas, B., et al. Metformin inhibits mitochondrial permeability transition and cell death: a pharmacological in vitro study. Biochem J 382, (2004). 15. Detaille, D., Guigas, B., Leverve, X., Wiernsperger, N. & Devos, P. Obligatory role of membrane events in the regulatory effect of metformin on the respiratory chain function. Biochem Pharmacol 63, (2002). 16. Palenickova, E., Cahova, M., Drahota, Z., Kazdova, L. & Kalous, M. Inhibitory effect of metformin on oxidation of NADH-dependent substrates in rat liver homogenate. Physiol Res 60, (2011). 17. Wessels, B., Ciapaite, J., van den Broek, N.M., Nicolay, K. & Prompers, J.J. Metformin impairs mitochondrial function in skeletal muscle of both lean and diabetic rats in a dose-dependent manner. PLoS One 9, e (2014). 18. Howlett, H.C. & Bailey, C.J. A risk-benefit assessment of metformin in type 2 diabetes mellitus. Drug Saf 20, (1999). 19. Winnick, J.J., et al. Short-term aerobic exercise training in obese humans with type 2 diabetes mellitus improves whole-body insulin sensitivity through gains in peripheral, not hepatic insulin sensitivity. J Clin Endocrinol Metab 93, (2008). 20. Nojima, H., et al. Effect of aerobic exercise training on oxidative stress in patients with type 2 diabetes mellitus. Metabolism 57, (2008). 21. De Feyter, H.M., et al. Exercise training improves glycemic control in long-standing insulin-treated type 2 diabetic patients. Diabetes Care 30, (2007). 22. Sharoff, C.G., et al. Combining short-term metformin treatment and one bout of exercise does not increase insulin action in insulin-resistant individuals. Am J Physiol Endocrinol Metab 298, E (2010). 23. Cotter, M.A., Cameron, N.E., Robertson, S. & Ewing, I. Polyol pathway-related skeletal muscle contractile and morphological abnormalities in diabetic rats. Exp Physiol 78, (1993). 150

152 Metformin affecting muscle contractile function 24. Safwat, Y., Yassin, N., Gamal El Din, M. & Kassem, L. Modulation of skeletal muscle performance and SERCA by exercise and adiponectin gene therapy in insulin-resistant rat. DNA Cell Biol 32, (2013). 25. Gollnick, P.D., Korge, P., Karpakka, J. & Saltin, B. Elongation of skeletal muscle relaxation during exercise is linked to reduced calcium uptake by the sarcoplasmic reticulum in man. Acta Physiol Scand 142, (1991). 26. Crow, M.T. & Kushmerick, M.J. Correlated reduction of velocity of shortening and the rate of energy utilization in mouse fast-twitch muscle during a continuous tetanus. J Gen Physiol 82, (1983). 27. Russell, D.M., et al. The effect of fasting and hypocaloric diets on the functional and metabolic characteristics of rat gastrocnemius muscle. Clin Sci (Lond) 67, (1984). 28. Dong, Y., et al. Activation of AMP-activated protein kinase inhibits oxidized LDL-triggered endoplasmic reticulum stress in vivo. Diabetes 59, (2010). 29. De Feyter, H.M., et al. Increased intramyocellular lipid content but normal skeletal muscle mitochondrial oxidative capacity throughout the pathogenesis of type 2 diabetes. FASEB J 22, (2008). 30. Vanhamme, L., van den Boogaart, A. & Van Huffel, S. Improved method for accurate and efficient quantification of MRS data with use of prior knowledge. J. Magn. Reson. 129, (1997). 31. Taylor, D.J., et al. Energetics of human muscle: exercise-induced ATP depletion. Magn Reson Med 3, (1986). 32. Taylor, D.J., Bore, P.J., Styles, P., Gadian, D.G. & Radda, G.K. Bioenergetics of intact human muscle. A 31P nuclear magnetic resonance study. Mol Biol Med 1, (1983). 33. Giannesini, B., et al. Citrulline malate supplementation increases muscle efficiency in rat skeletal muscle. Eur J Pharmacol 667, (2011). 34. Klawitter, P.F. & Clanton, T.L. Tension-time index, fatigue, and energetics in isolated rat diaphragm: a new experimental model. J Appl Physiol (1985) 96, (2004). 35. van den Broek, N.M., De Feyter, H.M., de Graaf, L., Nicolay, K. & Prompers, J.J. Intersubject differences in the effect of acidosis on phosphocreatine recovery kinetics in muscle after exercise are due to differences in proton efflux rates. Am J Physiol Cell Physiol 293, C (2007). 36. Paulus, S.F. & Grossie, J. Skeletal muscle in alloxan diabetes. A comparison of isometric contractions in fast and slow muscle. Diabetes 32, (1983). 37. Chonkar, A., Hopkin, R., Adeghate, E. & Singh, J. Contraction and cation contents of skeletal soleus and EDL muscles in age-matched control and diabetic rats. Ann N Y Acad Sci 1084, (2006). 38. Ganitkevich, V. Clearance of large Ca 2+ loads in a single smooth muscle cell: examination of the role of mitochondrial Ca 2+ uptake and intracellular ph. Cell Calcium 25, (1999). 39. Missiaen, L., et al. Ca 2+ extrusion across plasma membrane and Ca2+ uptake by intracellular stores. Pharmacol Ther 50, (1991). 40. Dawson, M.J., Gadian, D.G. & Wilkie, D.R. Mechanical relaxation rate and metabolism studied in fatiguing muscle by phosphorus nuclear magnetic resonance. J Physiol 299, (1980). 41. Edwards, R.H., Hill, D.K. & Jones, D.A. Metabolic changes associated with the slowing of relaxation in fatigued mouse muscle. J Physiol 251, (1975). 42. Saks, V.A., Khuchua, Z.A., Vasilyeva, E.V., Belikova, O. & Kuznetsov, A.V. Metabolic compartmentation and substrate channelling in muscle cells. Role of coupled creatine kinases in in vivo regulation of cellular respiration- -a synthesis. Mol Cell Biochem , (1994). 43. Rossi, A.M., Eppenberger, H.M., Volpe, P., Cotrufo, R. & Wallimann, T. Muscle-type MM creatine kinase is specifically bound to sarcoplasmic reticulum and can support Ca 2+ uptake and regulate local ATP/ADP ratios. J Biol Chem 265, (1990). 44. Kaasik, A., et al. Energetic crosstalk between organelles: architectural integration of energy production and utilization. Circ Res 89, (2001). 45. Gillis, J.M. Inhibition of mitochondrial calcium uptake slows down relaxation in mitochondria-rich skeletal muscles. J Muscle Res Cell Motil 18, (1997). 46. Lannergren, J., Westerblad, H. & Bruton, J.D. Changes in mitochondrial Ca 2+ detected with Rhod-2 in single frog and mouse skeletal muscle fibres during and after repeated tetanic contractions. J Muscle Res Cell Motil 22, (2001)

153 Summarizing Discussion

154 7

155 Chapter 7 154

156 Summarizing Discussion Mitochondrial function as a target for therapy in type 2 diabetes Summary The prevalence of type 2 diabetes (T2D) is growing at an alarming rate worldwide, which is associated with the fact that more and more people are becoming overweight or obese as a result of excessive caloric intake and/or lack of physical activity. Because skeletal muscle is the primary site for postprandial glucose disposal, the impairment of insulin-dependent glucose uptake in skeletal muscle is considered an early step in the development of T2D 1. It is well established that accretion of muscle intracellular lipids contributes to the onset of T2D 2-6. A lower capacity of mitochondria to oxidize fatty acids (FA) has been suggested to underlie this lipid spillover; however reports are contradictory regarding the cause-and-effect relationship between mitochondrial dysfunction and the accumulation of ectopic lipids. It is obvious though that ectopic lipid accumulation and reduced mitochondrial oxidative capacity are entangled in a vicious cycle that underlies the progression of muscle insulin resistance and T2D 7,8. A better understanding of the interaction between lipid accumulation and functioning of mitochondria in skeletal muscle is therefore invaluable for the development of effective treatment strategies for T2D. The aim of this thesis was to assess the pathogenic roles of lipid accumulation and mitochondrial dysfunction in diabetic muscle and to determine the effects of anti-diabetic drug therapies on muscle lipid content and mitochondrial function. The main factors contributing to in vivo mitochondrial function in skeletal muscle naturally include the number of mitochondria per tissue volume and their intrinsic capacity for ATP synthesis, but also extramitochondrial factors such as the availability of substrates and O 2 affect in vivo muscle oxidative capacity 9,10. Since O 2 availability is a primary determinant of mitochondrial ATP production, we determined in Chapter 2 whether O 2 availability is a limiting factor for in vivo mitochondrial function in lean rats, as determined by 31 P MRS measurement of PCr recovery. Furthermore we studied whether an impairment in O 2 availability might play a role in muscle mitochondrial dysfunction in diabetic rats. We demonstrated that muscle stimulation induced a larger decrease in muscle blood oxygenation in anesthetized lean and diabetic rats under normoxic breathing conditions as compared with hyperoxic breathing conditions. PCr recovery was slower in diabetic rats compared with lean rats; however, it was unaffected by supplemental O 2 in both groups of animals, showing that under normoxic conditions muscle oxidative capacity in both lean and diabetic rats is limited by mitochondrial capacity rather than O 2 availability. These findings validate that in vivo assessment of PCr recovery using 31 P MRS is a representative measure of muscle oxidative capacity in our studies in anesthetized rats. Moreover, our findings demonstrate that O 2 availability does not play a role in the impairment of in vivo muscle oxidative capacity in diabetic rats. 7 Next, we used a combination of in vivo and ex vivo read-outs of mitochondrial function to evaluate the effects of intervention strategies that aim to reduce the accumulation of lipids and lipid intermediates in insulin-resistant and diabetic muscle. Long-chain FA are a rich source of reducing equivalents, i.e. NADH and FADH 2, that drive the electron transport chain (ETC). When energy intake exceeds energy usage, fatty acid oxidation flux will outpace the demand of the respiratory system. The resultant increased ratio of NADH/NAD will exert negative (reducing) pressure on the ETC. This in turn promotes the generation of reactive oxygen species (ROS) and mitochondrial uncoupling, which may lead to compromised mitochondrial ATP production. 155

157 Chapter 7 Carnitine supplementation has been proposed to promote the export of intermediate products of FA oxidation out of the mitochondria, and thus relieve the reducing pressure put on the ETC. In Chapter 3 we presented the effects of L-carnitine supplementation on lipid homeostasis, muscle mitochondrial function and insulin sensitivity in insulin-resistant Wistar rats. Carnitine supplementation alleviated carnitine insufficiency induced by high-fat diet feeding; however this did not lead to improvements in muscle lipid status, in vivo muscle mitochondrial function or insulin resistance. Treatment of diabetic rats with pioglitazone, however, improved plasma and muscle lipid homeostasis and restored in vivo muscle mitochondrial function to the level of lean controls, as was described in Chapter 4. The beneficial effects of pioglitazone on in vivo muscle oxidative capacity in diabetic rats were not accompanied by an increase in mitochondrial content or an improvement in ex vivo intrinsic functioning of the mitochondria. Pioglitazone is a peroxisome proliferator-activated receptor-γ (PPAR-γ) agonist and is known to induce a relocation of lipids from ectopic sites, such as skeletal muscle, into adipose tissue. Our findings indicate that the impairment of in vivo muscle oxidative capacity in diabetic rats results from mitochondrial lipid overload and that pioglitazone restores in vivo mitochondrial function in diabetic muscle by alleviating the overflow of FA into mitochondria. Finally, we studied the effects of metformin on in vivo muscle mitochondrial and contractile function. Metformin is the most commonly prescribed drug to treat T2D and was demonstrated to inhibit Complex I of the mitochondrial respiratory chain in vitro In Chapter 5 we investigated the significance of this inhibition for in vivo muscle mitochondrial function. We demonstrated that metformin compromises in vivo muscle oxidative capacity in a dose-dependent manner in both lean and diabetic rats. Moreover, high-resolution respirometry measurements showed a similar dose-dependent effect of metformin on ex vivo mitochondrial function, indicating that the reductions in in vivo muscle mitochondrial function are directly related to the inhibitory effects of metformin on Complex I. In addition, treatment with metformin increased the rate of muscle intracellular acidosis during muscle stimulation in diabetic rats, as was described in Chapter 6. Aside from the impairment in cellular energy homeostasis, metformin treatment also exacerbated skeletal muscle contractile dysfunction in diabetic rats, as demonstrated by prolonged myofibrillar relaxation after contraction. The metformin-induced impairments in in vivo muscle mitochondrial and contractile function could explain why therapeutic benefits of exercise training are lost in patients on metformin therapy

158 Summarizing Discussion Discussion Throughout this thesis, the combination of in vivo PCr recovery measurements with a number of ex vivo read-outs was proven crucial to be able to distinguish between intra- and extramitochondrial factors affecting the capacity for oxidative ATP synthesis in vivo. The PCr recovery rate constant measured in skeletal muscle using 31 P MRS yields an in vivo measure of the muscle oxidative capacity. On the other hand high-resolution respirometry provides an ex vivo measure of the intrinsic respiratory capacity per mitochondrion for a range of different substrates (unlimited by the supply of substrates and O 2, as those are present in excess), while citrase synthase activity and mtdna copy number are measures of the number of mitochondria per tissue volume (mitochondrial content). Additionally, the supply of substrates and O 2 to the muscle, removal of potentially toxic metabolites, and the degree of coupling of the ETS to ATP synthesis are factors that can influence the capacity for ATP synthesis by mitochondria in vivo 15 (Figure 1). O 2 intrinsic mitochondrial capacity Sub Mito content X ROS OXPHOS FAO LEAK ATP Figure 1. Schematic representation of the network contributing to in vivo mitochondrial function O 2 and substrates (Sub) are required for mitochondria to generate energy in the form of ATP during oxidative metabolism (OXPHOS). Oxidation of FA substrates (FAO) is a source of reactive oxygen species (ROS) and promotes mitochondrial uncoupling (by increasing proton leak; LEAK), which could lead to compromised mitochondrial ATP production. O 2 and substrate supply, as well as the amount of mitochondria (Mito content), their intrinsic function, and their coupling to ATP synthesis (together termed intrinsic mitochondrial capacity) combined, determine muscle oxidative capacity in vivo. 7 When evaluating the effect of metformin on in vivo muscle oxidative capacity, assessment of mitochondrial respiration ex vivo revealed that the observed impairment of muscle oxidative capacity originates from metformin s specific inhibitory effect on Complex I. The additive value of combined ex vivo and in vivo measures of mitochondrial function was furthermore emphasized by the distinctive effects of pioglitazone in diabetic rats. While pioglitazone restored in vivo muscle oxidative capacity in diabetic rats to the level of healthy controls, this was not accompanied by increases in mitochondrial content or ex vivo mitochondrial respiratory capacity. Surprisingly, pioglitazone treatment actually decreased ex vivo β-oxidation capacity in isolated mitochondria of diabetic rats. However, pioglitazone, most likely as a result of the relocation of lipids from muscle into adipose tissue, also reduced the content of muscle lipids and lipid intermediates, which indicates that the improvement in in vivo muscle oxidative capacity is probably explained 157

159 Chapter 7 by the alleviation of lipid-induced mitochondrial uncoupling and/or inhibition. We noted a similar discrepancy between in vivo and ex vivo assessments of mitochondrial function in one of our earlier studies 16. These findings clearly demonstrate that, by combining in vivo and ex vivo measurements, we can distinguish between intrinsic impairments in the oxidative phosphorylation pathway and other, possibly extramitochondrial, factors affecting mitochondrial ATP synthesis. In addition, different aspects of the study design, including age and nutritional state of the subjects/ animals and treatment regimen in case of a drug study, could affect the outcome, for example due to adaptive responses to mitochondrial impairments in order to restore muscle oxidative capacity, known as mitochondrial retrograde signaling 17. Therefore, by examining only a subset of parameters involved in lipid homeostasis and muscle mitochondrial function, one might overlook vital information regarding the context of the findings. This could explain contradictory reports found in literature on changes in mitochondrial function, e.g. in the pathophysiology of insulin resistance and T2D, or due to treatment with anti-diabetic drugs. Our approach of combining both in vivo and ex vivo readouts of mitochondrial function is invaluable for mechanistic studies toward the pathogenic role of mitochondrial function in T2D, but also in other diseases such as chronic heart failure. The main goal of the research presented in this thesis was to determine the respective roles of lipid accumulation and mitochondrial dysfunction in the pathology of type 2 diabetes and the effects of therapy aimed at improving insulin sensitivity. The findings regarding pioglitazone treatment in diabetic rats reaffirm that mitochondrial dysfunction is likely not a primary cause of insulin resistance and T2D Rather, impaired mitochondrial oxidative capacity seems the result of increased mitochondrial influx of FA, causing reducing pressure on the respiratory chain 7,8 and a consequent increase in ROS generation, which prompts mitochondrial uncoupling in order to relieve mitochondrial stress 22. Normalizing muscle free carnitine levels in high-fat diet fed rats with carnitine supplementation did not improve muscle lipid status, in vivo mitochondrial function or insulin sensitivity, suggesting that promoting the efflux of FA intermediates out of the mitochondria is less effective than reducing mitochondrial FA influx, as with pioglitazone treatment. Moreover, metformin treatment at dosages that severely impaired muscle oxidative capacity did not compromise glucose homeostasis in lean or diabetic rats. These findings comply with the notion that mitochondrial dysfunction in T2D is not the primary cause of the disease. The results described in this thesis suggest that interventions aimed at treating insulin resistance and T2D should focus on reducing mitochondrial lipid influx in metabolic tissues, such as skeletal muscle, to alleviate lipid-induced mitochondrial dysfunction and restore insulin sensitivity. Future perspectives We demonstrated that the anti-diabetic drug pioglitazone lowers muscle lipid content in obese, diabetic animals and further showed its direct relation with muscle mitochondrial function. Even with the small sample sizes used in our studies, we observed strong correlations between plasma insulin levels, muscle lipid content and mitochondrial function, confirming a close association between these markers of T2D. Future studies to further investigate the cellular mechanisms underlying T2D could employ different dosage regimens and/or treatment periods of thiazolidinediones, such as pioglitazone, for a more detailed characterization of the interrelation between insulin sensitivity, muscle lipid accumulation and mitochondrial dysfunction. Furthermore, to examine the transition from insulin resistance to over type 2 diabetes, healthy rats on a high-fat diet, instead of diabetic rats, could be used as a model of pre-diabetes. 158

160 Summarizing Discussion Complementing in vivo and ex vivo read-outs of mitochondrial function with in vivo measurement of O 2 availability in skeletal muscle, for example with the non-invasive BOLD technique, could prove beneficial in research also toward other pathologies featuring impaired O 2 availability, such as chronic heart failure or peripheral arterial disease. Patients with chronic heart failure are characterized by an impairment in central hemodynamics, which results in inadequate tissue oxygenation. Additionally, chronic heart failure patients present with prolonged muscle PCr recovery after submaximal exercise Restrictions in tissue oxygenation and/or mitochondrial capacity can cause chronic heart failure patients to experience early fatigue and exercise intolerance. Combining 31 P MRS with BOLD imaging allows for non-invasive simultaneous assessment of muscle energetics and tissue oxygenation, and may provide a better understanding of the pathophysiological basis of the functional impairments in these patients, leading to improved therapeutic approaches. A downside of 31 P MRS is that this modality is not available on most clinical MR scanners and therefore is not widely applied for clinical purposes. Chemical exchange saturation transfer (CEST) is a technique that enhances contrast to make MRI sensitive to the concentrations of metabolites and their chemical environment. Since creatine shows a concentration-dependent sensitivity to CEST, recently a CEST-based technique was developed that allows monitoring of muscle creatine levels with high spatial resolution in humans 26. Moreover, it was demonstrated that this method can be used to measure dynamic changes in creatine in muscle in vivo at 3T 27. This technique would make the evaluation of muscle energy metabolism in diseases such as T2D, chronic heart failure, or chronic pulmonary disease more clinically available. The storage and breakdown of glycogen play an important role in glucose metabolism and is an important energy source in skeletal muscle during exercise. Muscle glycogen was shown to be reduced in T2D 28,29 and using 13 C MRS it is possible to evaluate glycogen levels in vivo at high-field C MRS can also be used to determine the rate of glycolysis flux in vivo using hyperpolarized 13 C-labeled glucose 31. This latter technique could possibly be used to gain more quantitative information on the effect of metformin on glycolysis flux in muscle during exercise. Another possible approach to attain information on glycogen dynamics is based on CEST, and this so-called glycocest 32 method would make it possible to study the storage and spatial distribution of glycogen on clinically available scanners. However the validity of this approach and its practical feasibility in a clinical setting needs further testing. Finally, another valuable tool to study the interrelation of lipid homeostasis and muscle mitochondrial function would be the application of computational models to help interpret or predict changes in muscle metabolism. Throughout this thesis it has been pointed out that interpreting experimental data on mitochondrial function is not straightforward, because factors such as mitochondrial content and intrinsic function, O 2 delivery, substrate selection and retrograde signaling can all affect the outcome. An elaborate computational model of the metabolic system 33 can therefore be helpful to determine cause-and-effect relations across multiple levels of the metabolic pathway, ranging from gene expression to effective ATP synthesis in a whole muscle. In addition, crucial parameters of the metabolic system could be identified with the use of computational methods and guide the design of hypothesis-driven experimental studies

161 Chapter 7 References 1. Cline, G.W., et al. Impaired glucose transport as a cause of decreased insulin-stimulated muscle glycogen synthesis in type 2 diabetes. N Engl J Med 341, (1999). 2. Goodpaster, B.H., He, J., Watkins, S. & Kelley, D.E. Skeletal muscle lipid content and insulin resistance: evidence for a paradox in endurance-trained athletes. J Clin Endocrinol Metab 86, (2001). 3. Kuhlmann, J., et al. Intramyocellular lipid and insulin resistance: a longitudinal in vivo 1 H-spectroscopic study in Zucker diabetic fatty rats. Diabetes 52, (2003). 4. Krssak, M., et al. Intramyocellular lipid concentrations are correlated with insulin sensitivity in humans: a 1 H NMR spectroscopy study. Diabetologia 42, (1999). 5. Perseghin, G., et al. Intramyocellular triglyceride content is a determinant of in vivo insulin resistance in humans: a 1 H-13C nuclear magnetic resonance spectroscopy assessment in offspring of type 2 diabetic parents. Diabetes 48, (1999). 6. van Loon, L.J., et al. Intramyocellular lipid content in type 2 diabetes patients compared with overweight sedentary men and highly trained endurance athletes. Am J Physiol Endocrinol Metab 287, E (2004). 7. Patti, M.E. & Corvera, S. The role of mitochondria in the pathogenesis of type 2 diabetes. Endocrine reviews 31, (2010). 8. Muoio, D.M. & Neufer, P.D. Lipid-induced mitochondrial stress and insulin action in muscle. Cell Metab 15, (2012). 9. Kemp, G.J. Mitochondrial dysfunction in chronic ischemia and peripheral vascular disease. Mitochondrion 4, (2004). 10. Prompers, J.J., Wessels, B., Kemp, G.J. & Nicolay, K. MITOCHONDRIA: investigation of in vivo muscle mitochondrial function by 31 P magnetic resonance spectroscopy. Int J Biochem Cell Biol 50, (2014). 11. Brunmair, B., et al. Thiazolidinediones, like metformin, inhibit respiratory complex I: a common mechanism contributing to their antidiabetic actions? Diabetes 53, (2004). 12. Owen, M.R., Doran, E. & Halestrap, A.P. Evidence that metformin exerts its anti-diabetic effects through inhibition of complex 1 of the mitochondrial respiratory chain. Biochem J 348 Pt 3, (2000). 13. Guigas, B., et al. Metformin inhibits mitochondrial permeability transition and cell death: a pharmacological in vitro study. Biochem J 382, (2004). 14. Sharoff, C.G., et al. Combining short-term metformin treatment and one bout of exercise does not increase insulin action in insulin-resistant individuals. Am J Physiol Endocrinol Metab 298, E (2010). 15. Kemp, G.J., Ahmad, R.E., Nicolay, K. & Prompers, J.J. Quantification of skeletal muscle mitochondrial function by 31 P magnetic resonance spectroscopy techniques: a quantitative review. Acta Physiol (Oxf) (2014). 16. van den Broek, N.M., et al. Increased mitochondrial content rescues in vivo muscle oxidative capacity in longterm high-fat-diet-fed rats. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 24, (2010). 17. Butow, R.A. & Avadhani, N.G. Mitochondrial signaling: the retrograde response. Mol Cell 14, 1-15 (2004). 18. Sparks, L.M., et al. A high-fat diet coordinately downregulates genes required for mitochondrial oxidative phosphorylation in skeletal muscle. Diabetes 54, (2005). 19. Kelley, D.E. & Simoneau, J.A. Impaired free fatty acid utilization by skeletal muscle in non-insulin-dependent diabetes mellitus. J Clin Invest 94, (1994). 20. Simoneau, J.A., Colberg, S.R., Thaete, F.L. & Kelley, D.E. Skeletal muscle glycolytic and oxidative enzyme capacities are determinants of insulin sensitivity and muscle composition in obese women. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 9, (1995). 21. Lowell, B.B. & Shulman, G.I. Mitochondrial dysfunction and type 2 diabetes. Science 307, (2005). 22. Brand, M.D. Uncoupling to survive? The role of mitochondrial inefficiency in ageing. Experimental gerontology 35, (2000). 23. Kemps, H.M., et al. Skeletal muscle metabolic recovery following submaximal exercise in chronic heart failure is limited more by O(2) delivery than O(2) utilization. Clin Sci (Lond) 118, (2010). 24. Toussaint, J.F., et al. Local relation between oxidative metabolism and perfusion in leg muscles of patients with heart failure studied by magnetic resonance imaging and spectroscopy. J Heart Lung Transplant 17, (1998). 160

162 Summarizing Discussion 25. Hanada, A., et al. Dissociation between muscle metabolism and oxygen kinetics during recovery from exercise in patients with chronic heart failure. Heart 83, (2000). 26. Kogan, F., et al. Method for high-resolution imaging of creatine in vivo using chemical exchange saturation transfer. Magnetic resonance in medicine : official journal of the Society of Magnetic Resonance in Medicine / Society of Magnetic Resonance in Medicine 71, (2014). 27. Kogan, F., et al. In vivo chemical exchange saturation transfer imaging of creatine (CrCEST) in skeletal muscle at 3T. Journal of magnetic resonance imaging : JMRI 40, (2014). 28. He, J. & Kelley, D.E. Muscle glycogen content in type 2 diabetes mellitus. Am J Physiol Endocrinol Metab 287, E (2004). 29. Befroy, D.E., et al. Direct assessment of hepatic mitochondrial oxidative and anaplerotic fluxes in humans using dynamic 13C magnetic resonance spectroscopy. Nature medicine 20, (2014). 30. Heinicke, K., et al. Reproducibility and Absolute Quantification of Muscle Glycogen in Patients with Glycogen Storage Disease by 13C NMR Spectroscopy at 7 Tesla. PloS one 9, e (2014). 31. Rodrigues, T.B., et al. Magnetic resonance imaging of tumor glycolysis using hyperpolarized 13C-labeled glucose. Nature medicine 20, (2014). 32. van Zijl, P.C., Jones, C.K., Ren, J., Malloy, C.R. & Sherry, A.D. MRI detection of glycogen in vivo by using chemical exchange saturation transfer imaging (glycocest). Proc Natl Acad Sci U S A 104, (2007). 33. Schmitz, J.P., Vanlier, J., van Riel, N.A. & Jeneson, J.A. Computational modeling of mitochondrial energy transduction. Crit Rev Biomed Eng 39, (2011)

163 Dankwoord Zoals eenieder die voor mij afscheid genomen heeft van de Biomedical NMR groep, is dit een afscheid dat ook mij zwaar valt. De afgelopen jaren heb ik hier met veel plezier gewerkt en ik heb erg veel waardering op de manier waarop iedereen altijd voor elkaar klaarstaat. Daarnaast zorgen de vele borrels, groepsactiveiten, de weekendjes Ardennen en natuurlijk niet te vergeten de wereldberoemde meerkamp ervoor dat de werksfeer altijd heel positief is. Ik wil iedereen, past and present, daarom hartelijk bedanken voor een geweldige tijd! Hiernaast wil ik nog een aantal mensen in het bijzonder bedanken voor de bijdrage die zij geleverd hebben aan het tot stand komen van dit proefschrift. Allereerst Klaas, het boegbeeld en het hart van deze vakgroep. Naast uw aanstekelijke enthousiasme voor alles wat MR-gerelateerd is, zet uw open en directe persoonlijkheid de toon qua sfeer binnen de vakgroep. De groepsweekenden waren een duidelijk voorbeeld hiervan en hier greep u vervolgens de kans om ons op de mountainbike hard in het stof te laten bijten met beide handen aan. De afgelopen paar jaar zijn zeker voor u niet makkelijk geweest maar uw positieve instelling is volgens mij de reden dat de sfeer en de saamhorigheid binnen de Biomedical NMR groep zo goed is gebleven. Jeanine, jouw betrokkenheid bij het wel en wee van ons SPEM-groepje, binnen- en buiten de werkvloer, is ongeëvenaard en zonder jou was dit werk nooit tot een goed einde gekomen. Jouw deur staat altijd open voor een snelle vraag, een korte blik op nieuwe resultaten of een leuke anecdote. Je enthousiasme en plezier op en buiten de werkvloer maakt dat het voor ons altijd een plezier is naar het werk te komen. Ik heb de afgelopen jaren met veel plezier met je samengewerkt en ben tevens gegroeid als onderzoeker. Het is verder ook mooi om te zien hoe je fanatisme buiten je werk als onderzoekster ook zichtbaar aanwezig is tijdens activiteiten buiten het werk zoals het korfbaltoernooi, de opdrachten tijdens het groepsweekend en het acteren voor alle filmpjes. Ook doe je hard je best om op de hoogte te blijven van hoe het gaat met oud-collega s zoals Geralda, Nicole en Ewelina. Ik hoop dat je dit doorzet en dat er nog veel oud-bnmr dineetjes volgen. Richard en Nicole, jullie twee waren mijn vangnet in de eerste maanden van mijn promotie. Jullie hebben me bijgebracht wat er allemaal komt kijken bij het doen en laten van dierenstudies en, ondanks mijn soms wat gebrekkige planning in het begin, stonden jullie altijd à la minute voor me klaar. Dit alles deden jullie vol goede humor en terwijl dit jullie van eigen onderzoek of het schrijven van je eigen boekje afhield. Dit heb ik altijd erg gewaardeerd en ik heb geprobeerd dit voorbeeld te volgen en om op mijn beurt zo klaar te staan voor mijn (jongere) collega s. Desiree, my sweet roomie. I know you can read Dutch perfectly well but somehow it still feels weird talking Dutch to you. We have shared an office for 5+ years and I have to say that it was a pleasure from the start. It was very gezellig and I think it was in both our interest that we have similar interior design ideas with regard to our desks... Since we started and finished around the same time we could also be there for each other, with me helping you calm down a bit during thesis stress and you getting me a bit more stressed when I needed to

164 Sharon, mijn roomie en buurvrouw sinds de verhuizing naar de HTC. Volgens mij staan wij in de gangen vd HTC bekend als de battle om de verwarming (24 graden is gewoon VEEL te warm!), maar het was altijd fijn de dag te beginnen met een kort verslag van de laatste ontwikkelingen op tv. Verder, net als met de andere SPEM-ers, ben ik blij dat we altijd voor elkaar klaar stonden of het nu in het lab was of op kantoor. It is hard to describe how much of a help you have been to accomplishing the work in this thesis Jolita. You helped me getting started on the famous oxygraph and have been our personal encyclopedia regarding everything mitochondria related. Even when you started working in Groningen you found time to do additional experiments for our article submissions. Thanks a lot for all the effort and fast results! I would also like to take this opportunity to express my gratitude to dr. Carlier, prof.dr. Wanders, prof.dr. Tack, dr. de Galan for their willingness to participate in my PhD committee. Geralda, of liever G, ik denk dat ik dankzij jou de Biomedical NMR groep ontdekt hebt. Jij was een van de pioniers hier en door de vele malen dat ik hier op de koffie of lunch kwam heb ik de vakgroep alvast leren kennen. Voor iedereen die een of meerdere keren geroepen heeft, Wie is toch die Geralda waar altijd over gepraat wordt?, de 26 e is je kans om kennis te maken met deze legende. Bedankt voor al je support de afgelopen jaren. In feite hebben wij onze interesse voor MR spectroscopie min of meer samen ontdekt Ot. Wij waren, tijdens het MDP project, twee van de meer fanatiekere deelnemers wat ons uiteindelijk die prachtige prent in de 1.5 T heeft opgeleverd. Verder was het spotvrije kantoor van jou en Bastiaan voor mij en Desiree altijd een ideaal streven dat we helaas nooit werkelijkheid hebben kunnen maken. Joep, bedankt voor al je hulp met de krachtmetingen en discussie over de resultaten. Het was weer even wennen toen ik weer zelf de beroemde knop van de spierstimulatie moest bedienen. Ik hoop verder dat de koffie bij DSM je niet teleurgesteld heeft na die weken dat je op de koffie uit onze automaat teerde. Igor, wij hebben heel wat late avonden gedeeld op de HTC onder het genot van de beste maaltijden die er op Thuisbezorgd te vinden zijn en meteen getest of de beamer geschikt is om op full HD een film te kijken (dat is zo). Jij bent verder de drijfveer dat Tom en ik steevast de wanden van Monk blijven bedwingen. Tom jij bent altijd een positieve noot binnen de groep en tijdens je optredens (zolang er maar geen accordeon bij is...). Miranda, jij was de laatste aanwinst van ons SPEM-groepje maar je kwam binnen als een wervelstorm. Binnen de groep greep je met veel enthousiasme alles aan wat je op je weg tegenkwam en sleurde je ons mee van de sportschool tot de volgende 90s party. Jij hebt echt in no-time je stempel op onze vakgroep gezet. Als je zoveel achter een computer zit als wij dan moet je af en toe gebruik maken van de sportfaciliteiten die de HTC biedt. Luc, jij was degene die wees op het bestaan van deze mogelijkheid op de HTC maar je had vervolgens zelf wat aandringen nodig voor je overtuigd was. Marloes was hierin, samen met Miranda, een drijvende kracht die onvermoeibaar bleek 163

165 tijdens bootcamp, het spinnen of het aanmoedigen van het Nederlands elftal. Blijkbaar heeft ze maar 1 achilleshiel en dat schijnt de Long-Island ice tea te zijn. Dat feestje heb ik helaas gemist. Larry, bedankt voor al je snelle technische hulp bij alle kleine rampjes die we soms op dagelijkse basis lijken te hebben. Wanneer je problemen hebt met je media center, die het volgens de geruchten nu toch echt doet, kan ik misschien jou eens uit de brand helpen. David en Leonie bedankt voor alle hulp bij mijn experimenten. De opzet van mijn studie zorgde ervoor dat jullie soms s ochtends vroeg, s avonds laat of zelfs in de weekenden paraat moesten staan om mij een handje te helpen. Caren, jij bent mij ook een aantal keer, tijdens die lange periodes van oraal doseren, in het weekend te hulp geschoten zodat ik een keer in bed kon blijven liggen. Maar als ik het goed heb ben je mij wel nog een aantal borrels schuldig... Heel erg bedankt voor al jullie flexibiliteit en hulp! Gustav, bedankt voor alle last-minute hulp bij problemen met de scanner of het opzetten van nieuwe sequenties en de gezelligheid op weg naar de strip of tijdens een van de vele uitjes. De volgende keer laat ik me wel overhalen bij het optreden van jouw band te zijn want uit de vele berichten die ik ontving schijnt dat een ongekend spektakel te zijn. Floortje, zoals we zojuist gemerkt hebben tijdens het onverwachte bezoek van de goedheiligman, ben jij er altijd om weer iets ludieks, ontspannends, of bloedserieus (de Meerkamp), voor ons te organiseren. Als we iets niet weten, er iets geregeld moet worden, of iemands veters gestrikt moeten worden, dan weten we je te vinden. Tessa, Bram en Leonie, we zijn maar kort kamergenootjes geweest maar dat hebben we goed gemaakt tijdens camping disco s, karaoke ontwijkend gedrag, trips naar New York en andere uitjes. Rik, ik ben blij dat jij de naam van borrel-initiator van mij hebt overgenomen! Stefanie, ik hoop dat Igor jou beter in de gaten houdt bij jouw promotie dan bij het feestje van Ot. Wolter, ik hoop dat je nieuwe collega s je humor net zo waarderen als wij dat deden. Siem, ik wacht op de dag dat Gordon Ramsey bij jou in de leer komt. Jules, onze strijder in the move part II. Bada, my karaoke skills were sub-par but I enjoyed your cuisine nonetheless. Nils Y.O.L.O., Renske, Maaike!, Pedro, Valentina, Bastiaan, Katrien, Esther, Pedro, Steffie, Holger, Sin Yuin, Jo, Laura, René, alle andere collega s en studenten die ik gemist heb, bedankt voor alle plezier dat ik gehad heb tijdens mijn tijd hier! Hareld, Ruud en Victor, ik heb met veel plezier met jullie samengewerkt aan jullie onderzoek omtrendt inspanningsintolerantie in patiënten met hartfalen, zowel tijdens mijn afstuderen als erna. Naast al mijn collega s heb ik natuurlijk ook veel steun gehad aan familie en vrienden tijdens mijn studie en promotie. Soms ter ontspanning of een begripvol oor of soms om je lekker af te kunnen reageren. Mijn echtgenote uit de Verwerstraat, Iloontje, waar wij jarenlang samen aten en relaxten na een dag studie of werk. Mijn systeem in de keuken laat tegenwoordig duidelijk te wensen over sinds ik mijn lieve huisgenootje niet verderop in de gang heb wonen. De koppen koffie met slap geouwehoer en een bordspelletje laat op de avond met Aloys zijn altijd erg gezellig. Hopelijk kunnen we dat nog regelmatig doen. Big Bart, ik ken niemand die een pilsje op het terras met een portie bittergarnituur zo waardeert als jij. Ik stel voor dat we die snel weer gaan eens gaan nemen (ik zal niks tegen Ingvild zeggen...). Glaudie, Jeff, TJ en Su, wij zijn al bevriend 164

166 sinds mijn tijd in V-town. Volgens mij is het heel speciaal dat we na al die tijd nog steeds hecht contact hebben en ieder jaar een spectaculair Boys Weekend hebben. J en TJ, ik verwacht jullie op de training woensdag! De Legends van Tantalus zoals ze bekend staan bij, nou ja eigenlijk alleen bij de Legends van Tantalus, vragen zich al geruime tijd af of dit zogenaamde boekje waarvoor ik trainingen en soms wedstrijden heb moeten afzeggen wel echt bestaat. Ik hoop dat dit bewijs in jullie hand voldoende is. Naast sportieve ontspanning hoort er af en toe ook een glaasje gehesen te worden, zoals dat in de Slappe Klets genoemd wordt. Steef, Jochem, Dave, Bobski, Chris, Patje, Lambooij, Lex en iedereen uit de Slappe Klets, bedankt voor de gezelligheid en moge Hinterglemm 2015 weer een groot succes worden. Ten slotte, Pap, Mam, Pim, Marion, Rik, Lieske, Peter, Cathalijne, Benjamin en David, zonder jullie steun, geduld en af en toe een duw in mijn rug was ik nooit zover gekomen! Thuis is het credo altijd geweest dat alles op tafel mag komen en uitgesproken kan worden. Ongeacht wat er aan de hand is, we kunnen thuis altijd aankloppen en het hele gezin staat voor elkaar klaar als het nodig is en dat is erg fijn om te beseffen. En voor alle anderen die een rol van betekenis gespeeld hebben tijdens mijn promotie en ik misschien vergeten ben expliciet te noemen: Hartstikke bedankt voor de geweldige tijd! 165

167 List of Publications Published in international Journal with impact factor 1. Wessels B, Ciapaite J, van den Broek NM, Houten SM, Nicolay K, Prompers JJ. Pioglitazone treatment restores in vivo muscle oxidative capacity in a rat model of diabetes. Diabetes Obes Metab Wessels B, Ciapaite J, van den Broek NM, Nicolay K, Prompers JJ. Metformin impairs mitochondrial function in skeletal muscle of both lean and diabetic rats in a dose-dependent manner. PLoS One. 2014; 9: e Abdurrachim D, Ciapaite J, Wessels B, Nabben M, Luiken JJ, Nicolay K, Prompers JJ. Cardiac diastolic dysfunction in high-fat diet fed mice is associated with lipotoxicity without impairment of cardiac energetics in vivo. Biochim Biophys Acta Prompers JJ, Wessels B, Kemp GJ, Nicolay K. MITOCHONDRIA: investigation of in vivo muscle mitochondrial function by 31P magnetic resonance spectroscopy. Int J Biochem Cell Biol 2014;50: Schmitz JP, Groenendaal W, Wessels B, Wiseman RW, Hilbers PA, Nicolay K, Prompers JJ, Jeneson JA, van Riel NA. Combined in vivo and in silico investigations of activation of glycolysis in contracting skeletal muscle. Am J Physiol Cell Physiol Jan 15;304(2):C doi: /ajpcell Kemps HM, Schep G, Zonderland ML, Thijssen EJ, De Vries WR, Wessels B, Doevendans PA, Wijn PF. Are oxygen uptake kinetics in chronic heart failure limited by oxygen delivery or oxygen utilization? Int J Cardiol Jul 9;142(2): doi: /j.ijcard Kemps HM, Prompers JJ, Wessels B, De Vries WR, Zonderland ML, Thijssen EJ, Nicolay K, Schep G, Doevendans PA. Skeletal muscle metabolic recovery following submaximal exercise in chronic heart failure is limited more by O(2) delivery than O(2) utilization. Clin Sci (Lond) Oct 26;118(3): Scientific articles in preparation 1. Wessels B, Ciapaite J, van den Broek NM, Houten SM, Wanders RJA, Nicolay K, Prompers JJ. Carnitine insufficiency in high-fat diet fed rats does not contribute to lipid-induced impairment of skeletal muscle mitochondrial function in vivo. 2. Wessels B, De Feyter HM, Nicolay K, Prompers JJ. In vivo assessment of muscle oxidative capacity using 31 P-MRS is not dependent on oxygen delivery in healthy and diabetic rats.

168 Conference proceedings (first author only) 1. Wessels B, Ciapaite J, van den Broek NM, Nicolay K, Prompers JJ. Metformin severely impairs in vivo muscle oxidative capacity in ZDF rats. European Bioenergetics Conference (EBEC), Freiburg, Germany, September Wessels B, Nicolay K, Prompers JJ. Oxygen delivery does not limit mitochondrial function in skeletal muscle of healthy and diabetic rats in vivo. 5 th Scientific meeting of International Society of Magnetic Resonance in Medicine (ISMRM) Benelux, Rotterdam, Nederland, January Wessels B, Schmitz JPJ, Nicolay K, Prompers JJ. Mitochondrial dysfunction in diabetic rats enhances glycolytic flux in skeletal muscle during stimulation. 4 th Scientific meeting of ISMRM Benelux, Leuven, Belgium, January Wessels B, Ciapaite J, Nicolay K, Prompers JJ. Pioglitazone treatment improves muscle oxidative capacity and lowers muscle lipid content in diabetic rats.71 st Scientific Sessions of American Diabetes Association, San Diego, CA, June Wessels B, Ciapaite J, van den Broek NM, Nicolay K, Prompers JJ. Metformin severely impairs in vivo muscle oxidative capacity in ZDF rats. Annual meeting and Exhibition of International Society of Magnetic Resonance in Medicine (ISMRM), Montreal, Canada, May Wessels B, Ciapaite J, Nicolay K, Prompers JJ. In vivo assessment of the effects of pioglitazone on muscle oxidative capacity and intramyocellular lipid content in diabetic rats using 31 P and 1 H MRS. ISMRM, Montreal, Canada, May Wessels B, Ciapaite J, Nicolay K, Prompers JJ. Pioglitazone treatment improves muscle oxidative capacity and lowers muscle lipid content in diabetic rats. Nederlandse Vereniging voor Diabetes Onderzoek (NVDO), Oosterbeek, The Netherlands, November Wessels B, Ciapaite J, van den Broek NM, Nicolay K, Prompers JJ. Metformin severely impairs in vivo muscle oxidative capacity in ZDF rats. NVDO, Oosterbeek, The Netherlands, November Wessels B, Ciapaite J, Nicolay K, Prompers JJ. Pioglitazone increases insulin sensitivity by improved mitochondrial function and IMCL. 3 rd Scientific meeting of ISMRM Benelux, Bovendonk, The Netherlands, January Wessels B, Ciapaite J, van den Broek NM, Nicolay K, Prompers JJ. Metformin severely impairs in vivo muscle oxidative capacity in ZDF rats. Nederlandse organisatie voor Wetenschappelijk Onderzoek (NWO), Veldhoven, The Netherlands, December Wessels B, Ciapaite J, van den Broek NM, Nicolay K, Prompers JJ. Metformin severely impairs in vivo muscle oxidative capacity in ZDF rats. 2 nd Scientific meeting of ISMRM Benelux, Utrecht, The Netherlands, January 2010.

169 About the Author Bart Wessels was born on August 19 th, 1977, in Eindhoven, the Netherlands. He grew up in Valkenswaard and finished secondary school (VWO) in 1996 at Were Di college, Valkenswaard. The same year, he started studying chemical engineering at the Eindhoven University of Technology. He switched to biomedical engineering the next year and obtained his Master of Science degree in 2008, at the Eindhoven University of Technology. During his studies he did an externship at Philips Research using mechanical testing of hair as a diagnostic tool for type 2 diabetes. His master thesis was on 31 P magnetic resonance spectroscopy combined with nearinfrared spectroscopy to assess exercise intolerance in patients with chronic heart failure. This Master s project was performed in the Biomedical NMR group of Prof. Dr. Klaas Nicolay in collaboration with the Máxima Medical Center Veldhoven. He started his PhD project in the Biomedical NMR group under supervision of Dr. Jeanine J. Prompers and Prof. Dr. Klaas Nicolay at Eindhoven University of Technology. The results of his PhD project, on the role of mitochondria in drug therapy against type 2 diabetes, are discussed in this thesis. At the end of his PhD candidacy Bart worked for 6 months on a joint project between AstraZeneca, Sweden and the Biomedical NMR group. The focus of this project was on validating the therapeutic efficacy of novel agents against muscle disease, using 31 P magnetic resonance spectroscopy. 168