Regulation of energy metabolism in cultured skeletal muscle ...

Regulation of energy metabolism in

cultured skeletal muscle cells: Effects of

exercise, donor differences and perilipin 2

Studies in human and mouse myotubes

Jenny Lund

Dissertation for the degree of Philosophiae Doctor (Ph.D.)

Department of Pharmaceutical Biosciences

School of Pharmacy

Faculty of Mathematics and Natural Sciences

University of Oslo

2017

© Jenny Lund, 2017 Series of dissertations submitted to the Faculty of Mathematics and Natural Sciences, University of Oslo No. 1891 ISSN 1501-7710 All rights reserved. No part of this publication may be reproduced or transmitted, in any form or by any means, without permission. Cover: Hanne Baadsgaard Utigard. Print production: Reprosentralen, University of Oslo.

Contents

ACKNOWLEDGEMENTS ....................................................................................................... 1

LIST OF PUBLICATIONS ....................................................................................................... 2

ABBREVIATIONS ................................................................................................................... 4

ABSTRACT ............................................................................................................................... 6

INTRODUCTION ..................................................................................................................... 8

Energy metabolism in skeletal muscle ................................................................................... 8

Dynamics of skeletal muscle lipid pools .............................................................................. 12

Metabolic flexibility of skeletal muscle ............................................................................... 13

Skeletal muscle fiber types ................................................................................................... 15

Effects of exercise on energy metabolism in skeletal muscle .............................................. 16

Insulin resistance, obesity and type 2 diabetes ..................................................................... 20

AIMS........................................................................................................................................ 23

SUMMARY OF PAPERS ....................................................................................................... 24

METHODOLOGICAL CONSIDERATIONS ........................................................................ 32

Donor characteristics ............................................................................................................ 32

Study design of the in vivo exercise intervention ................................................................. 34

Cultured skeletal muscle cells as an in vitro model ............................................................. 35

Methods used to measure energy metabolism in cultured skeletal muscle cells ................. 38

Data analyses and statistics .................................................................................................. 42

DISCUSSION AND CONCLUSIONS ................................................................................... 43

Oxidative capacity ................................................................................................................ 43

Lipid storage and turnover ................................................................................................... 49

Fiber type transformations in skeletal muscles .................................................................... 50

Effects on insulin sensitivity ................................................................................................ 52

Final considerations.............................................................................................................. 53

REFERENCES ........................................................................................................................ 55

1

ACKNOWLEDGEMENTS

The work presented in this thesis was performed at Department of Pharmaceutical

Biosciences, School of Pharmacy, University of Oslo during the period of 2013-2017. I am

very grateful for getting the opportunity to perform this PhD, and would like to express my

gratitude towards the University and all the people that have made this possible for me.

First of all I would like to thank my supervisors Eili Kase, Arild Rustan, Hege Thoresen, and

Jørgen Jensen for your support, guidance and encouragement; special thanks to the three of

you I have seen and talked to almost every single day. You have made this a great experience.

Next, I would like to thank the rest of my colleagues Nataša Nikolić, Vigdis Aas, Nils

Gunnar Løvsletten, Hege Bakke, and Camilla Stensrud. Also, thanks to the rest of the

members of the muscle research group, past and present, and all the talented master students

that I have had the pleasure of being cosupervisor for throughout these years: Mari Brubak,

Siw Anette Helle, Nils Gunnar Løvsletten, Abel Mengeste, and Sevnur Turan. I also have to

thank the rest of the wonderful people at the department working in Gydas vei; this has been

a great working environment!

Last but not least I would like to thank my wonderful family and friends. There are several

that have been a great support, but I would like to thank six of you in particular. First and

foremost: Mom and dad, you have always been there for me with encouragement, questions

on how my work is going and lots of support. I am so thankful for all the opportunities you

have given me and I love you very much! Next, I would like to thank my friends. Maren, we

met our first day at University and we have been best of friends ever since! Kristine, we have

become great friends throughout these years and I am forever grateful! Kristin, you have

become our “honorary pharmacist” and I am so happy I got to know you! Solveig, we also

met our first day at University and even though you switched career path we have kept the

great friendship and now we are also going to be colleagues. I love you all and look forward

to plenty of more good times with you! Your friendship and support means the world to me.

Oslo, 2017

Jenny Lund

2

LIST OF PUBLICATIONS

Paper I

Lund J, Rustan AC, Løvsletten NG, Mudry JM, Langleite TM, Feng YZ, Stensrud C,

Brubak MG, Drevon CA, Birkeland KI, Kolnes KJ, Johansen EI, Tangen DS, Stadheim HK,

Gulseth HL, Krook A, Kase ET, Jensen J, and Thoresen GH.

Exercise in vivo marks human myotubes in vitro: Training-induced increase in lipid

metabolism.

PLOS ONE, 2017;12(4):e0175441.

Paper II

Lund J, Tangen DS, Wiig H, Stadheim HK, Helle SA, Birk JB, Rustan AC, Thoresen GH,

Wojtaszewski JFP, Kase ET, and Jensen J.

Glucose metabolism and metabolic flexibility in cultured skeletal muscle cells is related to

exercise status in young male subjects.

Submitted to Archives of Physiology and Biochemistry.

Paper III

Lund J, Helle SA, Kase ET, Li Y, Løvsletten NG, Stadheim HK, Jensen J, Thoresen GH,

and Rustan AC.

Higher fatty acid turnover and oxidation in cultured human skeletal muscle cells from trained

young male subjects.

Submitted to PLOS ONE.

Paper IV

Lund J, Aas V, Tingstad RH, Van Hees A, and Nikolić N.

Lactic acid is readily used as an energy source or stored as glycogen and intracellular lipids

in human myotubes.

Submitted to PLOS ONE.

3

Paper V

Feng YZ, Lund J, Li Y, Knabenes IK, Bakke SS, Kase ET, Lee YK, Kimmel AR, Thoresen

GH, Rustan AC, and Dalen KT.

Loss of perilipin 2 in cultured myotubes enhances lipolysis and shifts the metabolic energy

balance from glucose oxidation towards fatty acid oxidation.

Under revision before resubmission to Journal of Lipid Research.

Publications not included in this thesis:

Lund J, Stensrud C, Rajender, Bohov P, Thoresen GH, Berge RK, Wright M, Kamal A,

Rustan AC, Miller AD, Skorve J.

The molecular structure of thio-ether fatty acids influences PPAR-dependent regulation of

lipid metabolism.

Bioorganic & Medical Chemistry, 2016;24(6):1191-1203.

4

ABBREVIATIONS

ABHD5/CGI-58 abhydrolase domain containing 5/comparative gene identification-58

ACAD Acyl-CoA dehydrogenase

ACBP acyl-CoA-binding protein

ACC acetyl-CoA carboxylase

ACL ATP citrate lyase

ACOX acyl-CoA oxidase

ACSL acyl-CoA synthetase

AMPK AMP-activated protein kinase

ASM acid soluble metabolite

ATGL adipose triglyceride lipase

ATP adenosine triphosphate

BMI body mass index

CA cell-associated radioactivity

CE cholesteryl ester

CPT carnitine palmitoyltransferase

DAG diacylglycerol

DGAT diacylglycerol acyltransferase

FABPc/FABPpm cytoplasmic/plasma membrane-associated fatty acid binding protein

FAS fatty acid synthase

FAT/CD36 fatty acid translocase

FATP fatty acid transport protein

(F)FA (free) fatty acid

G0S2 G0/G1 switch 2

G-6-P glucose-6-phosphate

GLUT glucose transporter

GPR G-protein coupled receptor

G(Y)S glycogen synthase

H heart

HK hexokinase

HRmax maximal heart rate

HSL hormone-sensitive lipase

IMCL intramyocellular lipid

IMTG intramyocellular triacylglycerol

IRS insulin receptor substrate

KO knockout

LD lipid droplet

LDH lactate dehydrogenase

LMM linear mixed-model analysis

LPL lipoprotein lipase

M muscle

MAG monoacylglycerol

MCD malonyl-CoA decarboxylase

MCT monocarboxylate transporter

MEF myocyte enhancer factor

MGAT monoacylglycerol acyltransferase

MGL monoacylglycerol lipase

MHC myosin heavy chain

MYF5 myogenic factor 5

5

MYH myosin heavy chain, gene

MYOD myogenic differentiation protein

NAD+/NADH nicotinamide adenine dinucleotide

OA oleic acid

PA palmitic acid

PDC pyruvate dehydrogenase complex

PDHA1 pyruvate dehydrogenase alpha 1

PDK pyruvate dehydrogenase kinase

PHK phosphorylase kinase

PI3K phosphatidylinositol 3-kinase

PKA protein kinase A

PKB/Akt protein kinase B

PKC protein kinase C

PKM muscle pyruvate kinase

PL phospholipid

PLIN perilipin

PPAR peroxisome proliferator-activated receptor

P(PAR)GC peroxisome proliferator-activated receptor gamma coactivator

PYGM muscle-associated glycogen phosphorylase

Rac1 Ras-related C3 botulinum toxin substrate 1

RER respiratory exchange ratio

RM repetition maximum

SCD stearoyl-CoA desaturase

Ser serine

SLC2 solute carrier family 2

SMM skeletal muscle mass

SPA scintillation proximity assay

T2D type 2 diabetes

TAG triacylglycerol

TBC1D1 TBC1 domain family member 1

TBC1D4/AS160 TBC1 domain family member 4/Akt substrate of 160 kDa

TCA tricarboxylic acid

TFAM mitochondrial transcription factor A

Thr threonine

Tyr tyrosine

UCP uncoupling protein

VLDL very low-density lipoprotein

VO2max maximal oxygen uptake

WHO World Health Organization

WHR waist-to-hip ratio

WT wild-type

6

ABSTRACT

The prevalence of metabolic disorders such as overweight, obesity and type 2 diabetes (T2D)

has rapidly increased worldwide during the last decades, and physical activity has preventive

as well as therapeutic benefits for these conditions. Increasing evidence suggests that

dysregulations in lipid influx, storage and/or triacylglycerol (TAG) lipolysis have significant

impact on insulin sensitivity and glucose homeostasis in skeletal muscle. Furthermore, it has

been suggested that the insulin resistance in subjects that are overweight/obese and/or have

T2D is associated with lipid accumulation in their skeletal muscles. Much of the studies that

have been performed have been aimed towards the possibility of increasing lipid utilization

by exercise or pharmacological activation to avoid ectopic lipid accumulation in skeletal

muscle. The nuclear receptor peroxisome proliferator-activated receptor delta (PPAR) has

been shown to be an important regulator of skeletal muscle lipid metabolism. This thesis

aimed to study regulation of energy metabolism in cultured human skeletal muscle cells

isolated from biopsies from subjects with different metabolic profile and training status, and

we also studied effects of an in vivo exercise intervention on in vitro energy metabolism in

the cells. Plasma lactate concentrations increase rapidly during exercise, and was initially

thought of as a waste product; however, recently lactate was found to be a useable energy

source in skeletal muscle. Therefore, we aimed to study lactate metabolism in cultured human

myotubes at rest and to see if acute and chronic lactate exposure affected metabolism of

glucose and oleic acid (OA). The lipid droplet (LD)-associated protein perilipin 2 (PLIN2) is

one of several PPAR target genes, and to study the functional role of PLIN2 and LDs on

energy metabolism in skeletal muscle we also examined myotubes established from Plin2+/+

and Plin2-/-

mice.

The 12-week training intervention, consisting of combined endurance and strength training,

improved endurance, strength and insulin sensitivity in vivo, and reduced the participants’

body weight. Biopsy-derived cultured myotubes from these participants before and after the

exercise intervention showed exercise-induced increase in total cellular OA uptake, oxidation

and lipid accumulation, as well as increased fractional glucose oxidation (glucose oxidation

relative to glucose uptake). Most of these exercise-induced increases were significant in the

overweight group, whereas no changes in OA or glucose metabolism were observed in

myotubes from the normal weight subjects. On the other hand, when studying energy

metabolism in individuals with different inherent training status we observed higher

7

carbohydrate and fat oxidation in vivo in trained and intermediary trained subjects compared

to sedentary untrained subjects. Fiber type distribution did not differ between groups. In

myotubes established from the trained compared to untrained subjects we observed higher

fractional glucose oxidation, and those myotubes were also more sensitive towards the

suppressive action of acutely added OA to the cells. Furthermore, myotubes from trained

subjects had lower fatty acid (FA) accumulation, lower incorporation of OA into total lipids,

TAG, diacylglycerol and cholesteryl ester, higher TAG-related lipolysis and re-esterification,

and also higher FA complete oxidation (CO2) and β-oxidation compared to myotubes from

untrained subjects. When studying lactate metabolism in myotubes established from lean

healthy donors we observed that the cells expressed both of the monocarboxylate transporters,

MCT1 and MCT4, and we observed that lactic acid was a usable substrate for both glycogen

synthesis and incorporation into lipids. Acute addition of lactic acid inhibited glucose and OA

oxidation, whereas OA uptake increased. Pretreatment with lactic acid for 24 h did not affect

glucose or OA metabolism; however, when increasing the exposure time by replacing

glucose with lactic acid in the cell culture media during the whole proliferation and

differentiation period, glucose uptake and oxidation as well as OA oxidation were increased.

Ablation of Plin2 resulted in myotubes with reduced number of LDs, reduced accumulation

of TAG and higher lipolysis. Furthermore, ablation of Plin2 resulted in a metabolic shift in

energy metabolism from utilization of glucose towards FAs. Despite increased oxidative

capacity for FAs, the exercise intervention in vivo, high training status and ablation of Plin2

did not have any impact on insulin-stimulated responses.

The results presented in this thesis shows that exercise is able to induce changes in human

myotubes in vivo that are discernible in vitro and that cultured myotubes retain some the

phenotypic traits of their donors. Ablation of Plin2 shifted the cells from glucose to lipid

metabolism. Furthermore, the results suggest that prolonged exposure to lactate affect

metabolism of glucose and FAs. Also, for the first time we show that lactic acid is a usable

substrate for glycogen synthesis and it can be stored as intracellular lipids in myotubes. Thus,

lactate may be an important regulator of energy metabolism in human myotubes.

8

INTRODUCTION

Energy metabolism in skeletal muscle

Skeletal muscle constitutes about 40% of the body weight in non-obese, adult individuals. It

is the largest insulin-sensitive organ, accounting for more than 80% of insulin-stimulated

glucose disposal. Thus, skeletal muscle is quantitatively the most important site with regards

to insulin resistance [1-3]. Furthermore, skeletal muscle is the largest storage organ for

glycogen, having about 4-fold higher capacity than the liver [4]. At rest, skeletal muscle

accounts for approximately 30% of the resting metabolic rate [5]. During maximal exercise

this can be increased by up to 20-fold. Skeletal muscle is therefore the main contributor to

exercise-induced changes in whole-body energy metabolism [6]. The two main fuel sources

for skeletal muscle are carbohydrates and fatty acids (FAs) [7]. Lipid stores are very large,

and potentially inexhaustible. Carbohydrate stores are on the other hand limited, comprising

400-500 g of glycogen in skeletal muscle, 60-100 g of glycogen in the liver and 4-5 g of

glucose circulating in the blood in the resting situation [8]. With regards to lipid metabolism,

skeletal muscle is also the dominating organ [9, 10]; FA oxidation is the main metabolic

activity of skeletal muscle during fasting [9, 11]. As a consequence, factors regulating

skeletal muscle FA oxidation and mitochondrial function capacity will affect whole-body

energy homeostasis. Therefore, skeletal muscle is of particular interest in metabolic diseases

such as obesity and type 2 diabetes (T2D) due to the critical role that skeletal muscle plays in

glycemic control and metabolic homeostasis.

In skeletal muscle, glucose may be stored as glycogen, be oxidized to produce energy as

adenosine triphosphate (ATP) or act as a precursor for lipid synthesis. During rest, glucose

uptake across the plasma membrane is considered the rate-limiting step for glucose utilization

[12]. A family of transmembrane transport proteins named glucose transporters (GLUTs) is

responsible for this step. In human skeletal muscle cells the majority of the glucose uptake is

mediated by GLUT1 and GLUT4 [13-16]. GLUT1 appears to be the main facilitator of basal

glucose uptake [17], whereas GLUT4 is translocated during insulin-stimulation or contraction

from intracellular vesicles to the cell surface to mediate glucose uptake through different

signaling pathways [18-21]. Some of the factors that has been proposed involved in

regulation of this GLUT4-translocation in skeletal muscle cells are the Rab-GTPase-

activating proteins TBC1 domain family member 1 (TBC1D1) [22] and TBC1 domain family

member 4 (TBC1D4, also known as Akt substrate of 160 kDa (AS160)) [23], Rab8A and

Rab13 [24]. Binding of insulin to its receptor leads to activation by phosphorylation of

9

insulin receptor substrate 1 (IRS1), phosphatidylinositol 3-kinase (PI3K) and protein kinase B

(PKB/Akt) [25]. Thus, PKB/Akt is a principal insulin-regulated signal transductor of

GLUT4-translocation to the cell membrane in response to insulin [25, 26]. Once inside the

cell glucose is phosphorylated to glucose-6-phosphate (G-6-P) by hexokinase (HK) and goes

into glycolysis, generating pyruvate, ATP and nicotinamide adenine dinucleotide (NADH)

(Figure 1). Alternatively, G-6-P can be converted to glycogen by glycogen synthase (GS)

and stored [27]. However, skeletal muscle has as previously mentioned a limited ability to

store glycogen, and in this case or in the case of excessive energy supply most excess glucose

is converted to lipids through lipogenesis [28]. De novo lipogenesis does occur in skeletal

muscle, however to a low extent [29]. Pyruvate, either from plasma glucose or stored

glycogen, can enter mitochondrial oxidation via decarboxylation to acetyl-CoA mediated by

the pyruvate dehydrogenase complex (PDC) [30]. PDC is positioned in such a way that it

plays a central role in regulation of glucose metabolism as well as fuel selection in skeletal

muscle, and it is a crucial regulator of ATP levels and thus maintaining the cells’ energy

balance [31]. In skeletal muscle, pyruvate dehydrogenase kinase (PDK) 4 inhibits the activity

of PDC through phosphorylation. Thus, increased PDK4 levels are associated with reduced

PDC activity and thus reduced glucose oxidation [32].

FAs are delivered to skeletal muscle as free fatty acids (FFAs) bound to albumin, or derived

from triacylglycerol (TAG) in chylomicrons or very low-density lipoproteins (VLDLs) in

plasma, where the FAs are liberated by lipoprotein lipase (LPL) before they are taken up by

the cells [33]. FFAs enter the skeletal muscle cells through passive diffusion over the plasma

membrane or via transport proteins in the plasma membrane (Figure 1). The major transport

proteins regulating skeletal muscle FA uptake are fatty acid translocase (FAT/CD36), plasma

membrane-associated fatty acid-binding protein (FABPpm) and a family of fatty acid

transport proteins (FATP1-6) [34, 35], where FAT/CD36 and FATP4 are considered the

quantitatively most important transport proteins in skeletal muscle [36, 37]. Once inside the

cells, FAs are reversibly bound to the cytoplasmic FABP (FABPc), which protects against

lipotoxic accumulation of FFAs and shuttles FAs throughout the cellular compartments [38].

Mediated by acyl-CoA synthetase (ACSL), FAs are activated to FA-CoA (as acyl-CoA) [39].

Acyl-CoA-binding protein (ACBP) acts as an intracellular carrier of FA-CoA. Further, FA-

CoA can be oxidized in mitochondria for energy production as ATP, esterified to

monoacylglycerol and diacylglycerol (MAG and DAG, respectively), stored as TAG in lipid

droplets (LDs – discussed in more detail under “Dynamics of skeletal muscle lipid pools”,

10

pp. 12-13), incorporated into phospholipids (PLs) for use in cellular membranes, or

metabolized to lipid second messengers [40]. Yet, the FAs are mainly distributed between

mitochondrial oxidation, TAG synthesis and LD storage [35]. Conversion of FA-CoA to

MAG is catalyzed by monoacylglycerol acyltransferase (MGAT) and conversion of DAG to

TAG is catalyzed by diacylglycerol acyltransferase (DGAT) 1 and/or 2 [41, 42]. The fate of

the FAs is influenced by the concentration of the incoming FAs, what type of FAs it is, fiber

type of the muscle, energy requirements of the muscle, and the hormonal ambience [38].

After mitochondrial transport FA-CoA can be oxidized as acyl-L-carnitine by carnitine

palmitoyltransferase (CPT) 1 and 2, located on the outer and inner mitochondrial membrane,

respectively [43]. FAT/CD36 is also found in the mitochondrial membrane, and it has been

suggested that it works in cooperation with CPT1 [44, 45]. Once inside the mitochondrial

matrix FA-CoA is metabolized to acetyl-CoA through -oxidation. Thereafter, acetyl-CoA

from both -oxidation and glycolysis enters the tricarboxylic acid (TCA) cycle (Figure 1).

Previously, regulation of FA oxidation has been considered a trait of transport of FAs across

the mitochondrial membranes, especially by reduced malonyl-CoA inhibition of CPT1

derived from acetyl-CoA from the glycolytic pathways catalyzed by acetyl-CoA carboxylase

(ACC) 2 in oxidative tissues [43]. However, recent work has challenged this perception,

suggesting that the regulation of FA oxidation in skeletal muscle is a much more complicated

process, involving multiple regulatory sites such as FA transport across the cell membrane,

binding and transport of FAs in the cytoplasm, LD formation and degradation, FA transport

across the mitochondrial membrane, and potential regulations within the β-oxidation pathway,

TCA cycle and electron transport chain [46-48]. Malonyl-CoA decarboxylase (MCD)

catalyzes conversion of malonyl-CoA to acetyl-CoA [49]. Malonyl-CoA can be converted to

FAs by the action of FA synthase (FAS) [50] and FAs can be further elongated and

desaturated by elongases and stearoyl-CoA desaturases (SCDs), respectively [51].

11

Figure 1. Energy metabolism in skeletal muscle. Glucose is transported into cells by glucose

transporters (GLUT) and is stored as glycogen or utilized through glycolysis to yield pyruvate.

GLUT4 is translocated from intracellular vesicles to the cell surface through activation of the insulin

pathway. Lactate is taken up and extruded by the monocarboxylate transporters MCT1 and MCT4,

respectively. Once inside the cell, lactate dehydrogenases (LDHs) catalyze the interconversion of

pyruvate and NADH to lactate and NAD+; LDH1 and LDH5 are the two isoforms catalyzing the

conversions of lactate to pyruvate and pyruvate to lactate, respectively. Uptake of fatty acids (FAs) is

facilitated by different transport proteins (FAT, FATP and FABPpm). Intracellular FAs are bound to

cytoplasmic FA binding proteins (FABPc) and activated to FA-CoA by acyl-CoA synthetase (ACSL).

Acyl-CoA-binding protein (ACBP) acts as an intracellular carrier of FA-CoA. Under conditions of

excess energy supply FA-CoAs may be incorporated into complex lipids as diacylglycerol (DAG),

triacylglycerol (TAG) and phospholipids (PL), and assembled in lipid droplets (LDs) for storage via

the action of monoacylglycerol acyltransferase (MGAT) and diacylglycerol acyltransferase (DGAT),

respectively. Upon energy demand, TAG, DAG and monoacylglycerol (MAG) are hydrolyzed by

adipose triglyceride lipase (ATGL), hormone-sensitive lipase (HSL) and monoacylglycerol lipase

(MGL) to FAs. FA-CoAs from both exogenous and endogenous derived FAs are used as fuel and

transported into mitochondria as acyl-L-carnitine via carnitine palmitoyltransferase (CPT) 1 and 2,

and thereafter metabolized through β-oxidation, yielding acetyl-CoA which enters the tricarboxylic

acid (TCA) cycle. Pyruvate, either from plasma glucose, lactate or stored glycogen, can enter the

mitochondria via decarboxylation to acetyl-CoA through the action of pyruvate dehydrogenase

complex (PDC). Citrate that escapes from the TCA cycle can be converted to acetyl-CoA in the

cytosol by ATP citrate lyase (ACL), and thereafter to malonyl-CoA by acetyl-CoA carboxylase

(ACC). Malonyl-CoA decarboxylase (MCD) catalyzes the reverse reaction and converts malonyl-

CoA to acetyl-CoA. Malonyl-CoA can be converted to FAs by the action of FA synthase (FAS) and

FAs can be further elongated and desaturated by elongases and stearoyl-CoA desaturases (SCD),

respectively. Malonyl-CoA is a potent inhibitor of CPT1, and can therefore inhibit entry and oxidation

of FAs in the mitochondria. FAs are able to suppress glucose oxidation through inhibition of PDC by

pyruvate dehydrogenase kinase 4 (PDK4) and by acetyl-CoA, as well as inhibition of glycolytic

enzymes by cytosolic citrate.

12

Skeletal muscle is the major site of lactate production and removal in the body [52]. The

lactate G-protein-coupled receptor 81 (GPR81) is mainly expressed in adipocytes [53-55], but

it has also been found in skeletal muscle, liver and kidney [56] as well as in the brain [57].

Lactate transport is mediated by proton-linked monocarboxylate transporters (MCTs), where

MCT1 and MCT4 are the most important and well-described isoforms in skeletal muscle [58].

MCT1 has predominantly been found in oxidative muscle, whereas MCT4 does not appear to

correlate with fiber type [58-60]. It has been observed that MCT1 has higher affinity for

lactate than MCT4, and is therefore thought to be most central for lactate uptake, whereas

MCT4 is considered most central for lactate removal [61] (Figure 1). The molecular

mechanisms involved in MCT regulation are still unclear, but both transcriptional and post-

transcriptional mechanisms are involved. Lactate dehydrogenases (LDHs) are responsible for

the conversion of pyruvate and NADH to lactate and NAD+ [62]. The conversion of pyruvate

into lactate is a necessary step to maintain high glycolytic flux in cells [63]. The LDH

enzymes are active as homo- or heterotetramers composed of muscle (M) and heart (H)

protein subunits, which are encoded by distinct genes: LDHA and LDHB, respectively.

Therefore, the different possible combinations allow the existence of five isoforms of LDH:

LDH1-LDH5 [63]. LDH enzymes with high M-subunit content, i.e. LDH5 in particular

(containing four M-subunits), are abundant in glycolytic skeletal muscles, where they reduce

pyruvate to lactate [64] (Figure 1). LDH enzymes with high H-subunit content, i.e. LDH1 in

particular (containing four H-subunits), are mainly found in aerobic tissues, where they

convert lactate into pyruvate [64] (Figure 1).

Dynamics of skeletal muscle lipid pools

Approximately 50-60% of the FAs taken up by the skeletal muscle cells are stored as TAG in

the LDs [65], and is then usually referred to as intramyocellular lipids (IMCL) [66] or

intramyocellular triacylglycerol (IMTG) [65]. In addition to functioning as a fuel source for

mitochondria, LDs are dynamic cellular organelles involved in signaling and lipid shuttling.

Apart from TAG, LDs also contain DAG, cholesteryl ester (CE) and free cholesterol, and

they are surrounded by a monolayer of PLs and proteins [67, 68]. These LD-binding proteins

are called perilipins (PLINs), and they are thus important in LD biogenesis [69, 70]. It has

been characterized five PLINs in human skeletal muscle, with different tissue expression

patterns [71]. PLINs also differ in size, affinity to the LDs, stability when not bound to the

LDs, and transcriptional regulation. As the PLINs are positioned at the LD surface they

13

manage access of lipases to the lipids within the LD core. Thus, they regulate LD size and

turnover [72]. It has been shown that more than half of the surface area of LDs in human

muscle biopsies is covered by PLIN2 [73], and this PLIN is proposed to protect LDs from

lipolysis [74]. Furthermore, PLIN2 content has been shown to be higher in oxidative type I

muscle fibers compared to glycolytic type II muscle fibers [73]. As LDs in oxidative muscle

often are close to mitochondria they maintain coupling of lipid storage with lipid

consumption as fuel, which appears to be important for efficient energy utilization [68].

Upon energy demand, e.g. during exercise, the enzymatic degradation of the esterified neutral

lipids in the LD core into single lipid species such as FAs or glycerol depends on active

recruitment of lipases to the LD surface. Adipose triglyceride lipase (ATGL) is considered to

be the first step in catabolism of TAG [75]. This generates DAG, which is further degraded

by hormone-sensitive lipase (HSL) [76]. The final step is degradation of MAG to glycerol

and FFA by monoacylglycerol lipase (MGL), thus providing FAs that can undergo

mitochondrial oxidation (Figure 1). Other potentially important proteins in the breakdown

regulation of IMTG are abhydrolase domain containing 5 (ABHD5, also known as

comparative gene identification-58, CGI-58) [32] and G0/G1 switch 2 (G0S2) [77], which are

coactivator and inhibitor of ATGL, respectively. ATGL and ABHD5 are strongly associated

during contraction-induced muscle lipolysis and work together with PLINs to regulate

lipolysis [78]. Activity of HSL is mostly regulated by phosphorylation, such as

phosphorylation on serine 660 (Ser660) [32].

Metabolic flexibility of skeletal muscle

As described previously, skeletal muscle uses both carbohydrates and FAs as fuel. During the

fed state, increased availability of plasma glucose stimulates glucose oxidation and FA

synthesis, whereas FA oxidation increases both during fasting and sustained exercise [79, 80],

but shifts from FA to glucose metabolism when exercise intensity increases [7, 10]. The

molecular mechanisms for this regulation are suggested to involve L-carnitine as an exercise-

induced increase in glycolysis enhances the production of acetyl-CoA and eventually acetyl-

L-carnitine. This results in reduced availability of free L-carnitine, which is a substrate of

CPT1. As a consequence, FA entry into the mitochondria for β-oxidation is reduced [7, 10].

Substrate selection during exercise appears to also be affected by levels of malonyl-CoA,

although this factor may be more important in resting skeletal muscle [10]. Furthermore,

14

activation of β-adrenergic pathways in skeletal muscle is an important event in the early

phases of exercise, leading to activation of protein kinase A (PKA), which in turn activates

HSL, resulting in increased lipolysis [81].

The ability to switch between substrates for fuel, depending on substrate availability, exercise

intensity and physiological conditions, represent an important feature of healthy skeletal

muscle and is called metabolic flexibility [82, 83]. The inhibition of glucose oxidation by

FAs, referred to as “Randle cycle” [84], is mediated by inhibition of several glycolytic steps.

PDK4, the dominant isoform in skeletal muscle, inhibits PDC by phosphorylation and

thereby switch fuel source from glucose to FAs [31] (Figure 1). Excess production of citrate

from enhanced FA oxidation escapes from mitochondria and inhibits the rate-limiting

enzyme of glycolysis, 6-phosphofructo-1-kinase, leading to an increase in G-6-P, which

eventually inhibits HK and leads to reduced glucose uptake and oxidation [46, 85]. The

opposite situation, where glucose suppresses FA oxidation, is usually referred to as “reverse

Randle cycle” [86]. Citrate that escapes from glucose oxidation is transported back to the

cytosol where it is converted to acetyl-CoA by ATP citrate lyase (ACL), which in turn is

converted to malonyl-CoA by ACC. As described above (under “Energy metabolism in

skeletal muscle”, pp. 8-12), malonyl-CoA inhibits CPT1 and thereby entry and oxidation of

FAs in mitochondria [86, 87] (Figure 1). Thus, citrate signals both fed (high concentrations

of glucose) and fasted (high concentrations of FAs) states. Loss of ability to easily switch

between glucose and lipid oxidation is termed metabolic inflexibility [83], and is associated

with reduced lipid oxidation and thereby promotes lipid accumulation in skeletal muscle [88],

which may interfere with insulin signaling and function (discussed in more detail under

“Insulin resistance, obesity and type 2 diabetes”, pp. 20-22). Insulin resistance, obesity

and T2D are linked to reduced lipid oxidation during fasting and impaired postprandial

switch from lipid to glucose oxidation [89], and this inflexibility has also been observed in

individuals with impaired glucose tolerance [90], suggesting that inflexibility plays a role in

the early development of T2D. In fact, it has been observed that cultured skeletal muscle cells

(myotubes) established from subjects with T2D, as well as myotubes established from obese,

have reduced capacity to oxidize FAs compared to myotubes from lean subjects [91-93].

Furthermore, the fact that metabolic flexibility of substrate oxidation is preserved in cells

when grown in culture suggests that metabolic switching is an intrinsic property of skeletal

muscle [94].

15

Metabolic flexibility of myotubes in vitro is referred to as suppressibility, adaptability and

substrate-regulated flexibility [94, 95]. Suppressibility is defined as the ability of cells to

suppress FA oxidation by acute addition of glucose, adaptability is defined as the capacity of

cells to increase FA oxidation upon increased FA availability [94], and substrate-regulated

flexibility is defined as the ability to increase FA oxidation when changing from a “fed” (high

glucose, low FA) to a “fasted” (high FA, no glucose) condition [95]. In vitro suppressibility

has been shown to be inversely correlated with insulin sensitivity and metabolic flexibility in

vivo, whereas adaptability has been found to be positively correlated with the same

parameters [94]. Nevertheless, metabolic inflexibility may be due to both intrinsic and

extrinsic (induced) factors, and the molecular mechanisms underlying metabolic inflexibility

remains to be established.

Skeletal muscle fiber types

Skeletal muscles are composed of different fiber types, and they are all structurally,

functionally and metabolically different. The different phenotypes are classified by

contractile speed, either slow- or fast-twitch, based on the “time-to-peak tension” or “twitch”

characteristics, and the histochemical staining for myofibrillar (myosin) ATPase, where the

slow-twitch phenotype is type 1 and the fast-twitch phenotype is type 2 (highest ATPase

activity). Human skeletal muscle fibers mainly express three isoforms of myosin heavy chain

(MHC): MHC, MHC2A and MHC2X (with respective genes MYH7, MYH2 and MYH1)

[96]. MHC-expressing fibers are the slow, fatigue resistant and oxidative type I muscle

fibers, whereas the MHC2A-expressing fibers are the fast oxidative type IIa muscle fibers,

and the MHC2X-expressing muscle fibers are the fast glycolytic type IIX fibers. The slow-

twitch type I fibers are associated with higher mitochondrial content and GLUT4 protein

expression compared to the fast-twitch type II fibers [97-99]. Furthermore, a composition

consisting of more type I fibers have been associated with increased insulin responsiveness

[100]. Due to differences in abundance of the oxygen transporting protein myoglobin, the

fiber types are different color-wise, where type I fibers are more red in appearance, type 2X

are white and 2A has an intermediate color. This is in turn related to mitochondrial density

and the relative contribution of oxidative metabolism in the respective fibers. Muscle

phenotype is highly influenced by exercise, and muscle cells can change their fiber type and

enzymatic properties according to altered functional demands (reviewed by e.g. Gundersen

16

[101]). However, the mechanisms involved in muscle fiber type switching are complex and

not known in detail, but transcription factors such as myocyte enhancer factor 2 (MEF2) [102]

and its target gene peroxisome proliferator-activated receptor gamma coactivator 1 alpha

(PPARGC1A) [103] have been shown to be involved in control of the slow fiber type

program (discussed in more detail under “Effects of exercise on energy metabolism in

skeletal muscle”, pp. 16-19). Further, after studies on myostatin null mice [104], the growth

factor myostatin has been suggested to be a regulator of the fiber type composition in skeletal

muscles by regulating gene expression of MEF2 and myogenic differentiation 1 (MYOD1).

Effects of exercise on energy metabolism in skeletal muscle

Skeletal muscles are characterized by their ability to adapt and remodel in response to

contractile activity [105], and it allows the muscles to more efficiently utilize substrates for

ATP production and thus become more resistant to fatigue [105, 106]. Energy metabolism,

mitochondrial function, intracellular signaling, gene transcription, as well as contractile

proteins are all affected by contractile activity. Exercise-induced adaptations in energy

metabolism are reflected by changes in both mitochondrial content (both in size and number)

and function [107, 108] and improved oxidative capacity [105, 109-113]. Furthermore, an

acute bout of exercise improves glucose homeostasis by increasing skeletal muscle glucose

uptake, whereas regular exercise induces changes in expression of metabolic genes such as

those involved in mitochondrial activity, muscle fiber type characteristics and GLUT4

expression at protein levels [112]. The functional consequences of these alterations depends

on intensity, duration, frequency, and mode of exercise [105].

Glucose metabolism in skeletal muscle is strongly affected by contractile activity. Trained

muscle fibers import and use more glucose than untrained fibers [114, 115]. It has been

shown that contractile activity increases translocation of GLUT4 and glucose uptake [116],

but appears to involve a different signaling pathway than that activated by insulin (Figure 2).

The mechanism is unclear, but it is known that contraction activates AMP-activated protein

kinase (AMPK), which in turn activates glucose uptake. However, it has also been shown that

contractions could stimulate glucose uptake even in AMPK knockdown mice [117].

Furthermore, there have been indications that TBC1D1 and TBC1D4 might be regulated by

contractile activity through activation by AMPK, but not by PKB/Akt [22, 23]. The

Ca2+

/calmodulin-dependent kinase pathway is probably also involved in this process, and a

17

link between exercise-induced glucose uptake and muscle glycogen content has also been

proposed [118]. Furthermore, it has been suggested that Ras-related C3 botulinum toxin

substrate 1 (Rac1), a small GTPase and member of the Rho family, is a novel regulator of

contraction-induced glucose uptake in skeletal muscle [119]. Exercise-stimulated glucose

uptake by muscle occurs independently of insulin signal transduction [120-122]. In addition

to exerting acute effects on glucose uptake, exercise promotes a short-term increase in insulin

sensitivity after cessation of exercise [123-125].

At rest, about 70% of the FFAs released into the circulation during lipolysis are esterified

back to triacylglycerol instead of being oxidized as measured on whole body metabolism

using stable isotopes [126]. However, during exercise these FAs may be used to supply

energy to skeletal muscle [127]. Both lipid synthesis and oxidation have been shown to be

increased in exercising skeletal muscle [10, 128-131], which takes more of their required

energy from lipids and less from carbohydrates during submaximal work [87, 113, 131]. This

is accounted for by an increase in IMTG utilization in trained muscle [132, 133]. In skeletal

muscle it has previously been accepted that HSL is the principal enzyme responsible for

lipolysis of IMTG during exercise [134, 135], but recently ATGL has emerged as the major

regulator in lipolysis of IMTG during contractile activity [78, 136]. How FA oxidation is

regulated by contractile activity is not clear. Despite observed exercise-induced increase in

FA uptake [137, 138], studies examining effects of exercise on the FA transport proteins

(FAT/CD36 and FABPpm) have given conflicting results, probably due to differences in

exercise intensity and duration in the performed studies [139]. Nevertheless, rates of FA

oxidation may increase by 3- to 10-fold from resting values during exercise at mild to

moderate intensity (25-65% of maximal oxygen uptake (VO2max)) [7, 140]. As FA oxidation

in skeletal muscle is strongly regulated by the mitochondrial FA transport capacity through

translocation of FAT/CD36 to the plasma membrane [139], CPT1 activity and mitochondrial

oxidative capacity, these factors are often used as measures for exercise-induced changes.

Furthermore, several of the important enzymes regulating mitochondrial activity have been

shown to be upregulated after endurance training, including the β-oxidative enzymes short-

chain, medium-chain and very long-chain acyl-CoA dehydrogenases and the TCA cycle

enzyme citrate synthase [130, 141]. Similarly, exercise-induced increase in FA oxidation has

been reported to be accompanied by increased expression and activity of CPT1 [142, 143], as

well as decreased malonyl-CoA, which as previously stated is a potent inhibitor of CPT1

[127, 144, 145]. In fact, reduction of basal malonyl-CoA levels has been proposed to be one

18

of the main cellular mechanisms mediating exercise-induced increase in FA oxidation

(Figure 2).

Figure 2. Proposed signaling pathways for contraction-stimulated effects on metabolism in

skeletal muscle. Contraction leads to energy depletion (i.e. an elevated AMP/ATP ratio) and elevated

intracellular [Ca2+

], which in turn leads to activation of AMP-activated protein kinase (AMPK) and

calmodulin-dependent protein kinases (CaMK), respectively. Activated AMPK phosphorylates TBC1

domain family member 4 (TBC1D4, also known as Akt substrate of 160 kDa (AS160)), and TBC1

domain family member 1 (TBC1D1) at multiple phosphorylation sites and allows conversion of less

active GDP-loaded Rab to more active GTP-loaded Rab. The increased levels of active GTP-loaded

Rab then allows glucose transporter 4 (GLUT4) storage vesicles to translocate and fuse with the

plasma membrane. This translocation of GLUT4 is also mediated through the canonical insulin-

signaling pathway via activation of insulin receptor substrates (IRS), leading to phosphorylation of

Akt. Contraction also promotes expressions of GLUT4, carnitine palmitoyltransferase 1 (CPT1) and

pyruvate dehydrogenase kinase isozyme 4 (PDK4). Activated AMPK and/or CaMK promote

relocation of fatty acid transporter (FAT, also known as CD36) to the plasma membrane and the outer

mitochondrial membrane to increase fatty acid (FA) uptake and oxidation. Furthermore, contraction

leads to increased lipolysis of lipid droplets (LDs) by activation of adipose triglyceride lipase (ATGL)

and hormone-sensitive lipase (HSL). Prolonged FA and [Ca2+

] influx activates peroxisome

proliferator-activated receptor delta (PPARδ) and myocyte enhancer factor 2 (MEF2), respectively,

and thereby their target genes. AMPK and/or CaMK increase expressions of peroxisome proliferator-

activated receptor gamma coactivator 1 alpha (PGC1α) and nuclear respiratory factor 1 (NRF1),

which further orchestrates the enhancement of mitochondrial biogenesis and function. Furthermore,

PPARδ, MEF2 and PGC1α are all implicated in the oxidative fiber type program. Green arrows

represent activation, whereas yellow arrows represent processes that probably are not affected by

contraction.

19

Plasma lactate concentrations increase rapidly during exercise. Originally lactate was

considered a waste product that was transported to the liver, kidneys or other organs for

clearance [146]. However, it is now generally accepted that lactate is also taken up by the

muscles and oxidized [147]; in fact, the contribution of muscles to total body lactate

clearance is considerable during exercise [147]. For several types of exercise, and especially

sustained submaximal exercise, lactate is transiently released into the bloodstream before a

shift occurs, and active muscle starts to use lactate as an energy source instead of continuing

to produce more [148]. It has been observed that this situation mainly occurs in type I and

type IIa muscle fibers, where lactate predominantly is oxidized, whereas in type IIb fibers

lactate is mainly removed through glyconeogenesis [148]. When the blood lactate

concentration is above the resting value it makes an ideal concentration gradient for lactate

uptake [149]. However, other factors such as muscle metabolic rate, optimal levels of

intracellular and extracellular pH, adequate blood flow, and training status also determine the

rate of lactate consumption by working muscles [150].

Skeletal muscles’ plasticity in response to exercise requires changes in the expression pattern

of muscle-specific genes, and thus extends beyond the described metabolic adaptations.

Among the other reported responses to exercise is increased proportion of oxidative muscle

fibers after endurance exercise [151-153]. The nuclear receptor PPARδ and its coactivator

PGC1α appears to partially mediate some of the positive adaptations to exercise [154-161].

Furthermore, PPARα, PPARδ, PGC1α, and some of the known PPAR target genes, such as

PDK2 and PDK4, increase in the post-exercise period [162]. Therefore, these transcription

factors and the pathways they are involved in may represent some of the molecular substrates

for the effects of exercise in skeletal muscle.

All taken together, exercise leads to extensive adaptations in skeletal muscle, and regular

exercise plays a central role in prevention and also treatment of metabolic disorders such as

obesity and T2D by improvements in insulin sensitivity (discussed in more detail under

“Insulin resistance, obesity and type 2 diabetes”, pp. 20-22). A summary of the proposed

signaling pathways for contraction-stimulated effects on energy metabolism in skeletal

muscle are illustrated in Figure 2 above.

20

Insulin resistance, obesity and type 2 diabetes

Insulin resistance develops when the cells of the body becomes less sensitive and eventually

resistant towards insulin. When the cells no longer are able to absorb adequate amounts of

glucose, manifested as a decrease in glucose uptake and muscle glycogen synthesis [163,

164], it leads to hyperglycemia, affecting the β-cells to produce more insulin which in turn

leads to hyperinsulinemia. When the β-cells no longer are able to produce enough insulin,

hyperglycemia and eventually T2D will become a fact.

One of the biggest risk factors for developing insulin resistance is being overweight or obese,

and it has been shown that both visceral adiposity [165, 166] and subcutaneous adiposity

[166] are correlates of insulin resistance; however, visceral adiposity is more strongly

associated with insulin resistance than subcutaneous adiposity [166]. This association is

thought to be explained by the fact that abdominal fat is resistant towards the anti-lipolytic

effect of insulin, leading to exaggerated release of FAs and increased levels of plasma FFAs

[167], causing lipotoxicity and further reduction of insulin sensitivity, which in turn increases

gluconeogenesis in the liver, inhibits insulin-mediated glucose uptake in skeletal muscle and

elevates serum glucose concentrations. Obesity may also in itself lead to insulin resistance if

the adipocytes reaches a certain size and no longer are able to store more fat, leading to

ectopic lipid accumulation in liver and skeletal muscle [163, 168, 169]. Furthermore, visceral

adipose tissue is prone to inflammation and inflammatory cytokine production, contributing

to a chronic low-grade inflammation [170]. It has become more evident that adipose tissue

secretes several bioactive peptides known as adipokines, and it has been suggested that these

play an important role in the crosstalk between adipose tissue and skeletal muscle [168, 171-

173]. High rates of TAG degradation (lipolysis) and release of FFAs into the circulation are

typical features of a dysfunctional adipose tissue [174]. Thus, imbalance in the secretion of

the pro- and anti-inflammatory adipokines caused by increased TAG accumulation might

contribute to induction and/or promotion of insulin resistance in skeletal muscle [171, 175,

176]. Furthermore, visceral adipose tissue is also associated with intrahepatic TAG content,

and it has been reported that intrahepatic TAG content might be an even better predictor of

metabolic disorders than visceral adiposity [177].

Overweight and obesity are defined either by body mass index (BMI), waist circumference or

waist-to-hip ratio (WHR). The World Health Organization (WHO) has classified BMI

between 18.50 kg/m2 and 24.99 kg/m

2 as normal body weight, BMI ≥ 25 kg/m

2 as overweight

and BMI ≥ 30 kg/m2 as obese [178]. However, several studies indicate that measurement of

21

waist circumference or waist-to-hip ratio, which reflect visceral (abdominal) fat, may be more

suitable for classification of overweight and obesity [179, 180]. For Caucasian European

females/males, a waist circumference and WHR are considered above normal if they are

higher than 80/94 cm and 0.85/0.90, respectively [181, 182]. WHO reported in 2008 that 1.4

billion adults were overweight, and of these 500 million were obese. In most cases,

overweight and obesity is caused by an imbalance between energy intake and energy

expenditure, although genetics and chronic stress also are known contributors [183-185].

More recently evidence for the influence of gut microbiota on metabolic processes and

contribution to low-grade inflammation and obesity has emerged [186-189].

Overweight and obesity are strongly associated with insulin resistance and T2D [190, 191]; in

fact the majority of subjects diagnosed with T2D are classified as overweight or obese [192].

It is established that a family history of T2D markedly increases the risk of developing the

disease, particularly in the first-degree relatives [193-195]. However, genome-wide

association studies have revealed that only 10% of the estimated heritability of T2D can be

explained [196]. Other lifestyle factors than body weight are thus involved in the

development of T2D, including physical inactivity and consumption of a high-fat diet [197-

199]. However, overweight and obesity is preventable, and T2D can be delayed, prevented

and treated by lifestyle interventions such as healthy diet, regular physical activity and weight

loss, as well as by pharmacological treatment [197, 198, 200-202]. Still, the prevalence of

T2D has rapidly increased worldwide during the last decades. In 2015 it was estimated that

415 million people had diabetes, of which most had T2D. By the year of 2040 the number of

people with diabetes is projected to reach 642 million worldwide [203].

The hallmarks of T2D on cellular level are insulin resistance in liver, adipose tissue and

skeletal muscle, increased lipolysis from adipose tissue, increased glucose production in liver,

and increased pancreatic β-cell dysfunction [196]. In skeletal muscle of diabetics,

dysfunctional insulin-stimulated GLUT4 translocation has been associated with increased

lipid accumulation or disruptions in FA metabolism, which may include altered FA uptake,

TAG synthesis and breakdown (lipolysis), FA oxidation, or any combination of these [204-

208]. It has been shown that IMTG content and insulin sensitivity are inversely correlated

[205], but not for endurance-trained athletes [10, 66, 209, 210]. Exercise is known to increase

accumulation of IMTG, but also to improve insulin sensitivity, whereas increased IMTG

content is strongly associated with insulin resistance for individuals with obesity and/or T2D.

This phenomenon has been known as “the athlete’s paradox” [66, 211, 212]. Furthermore, it

22

has been shown that a decrease in IMTG after diet-induced weight loss correlates with

improved insulin sensitivity [213, 214]. Recently it has become more evident that

accumulation of other intracellular lipid intermediates than TAG, like long-chain acyl-CoA,

DAGs and ceramides, are more harmful, having a negative effect on the activation of the

insulin-signaling cascade in skeletal muscle cells [215-217]. Ceramides may inhibit

serine/threonine (Ser/Thr) phosphorylation of PKB/Akt [218], and skeletal muscle ceramide

levels have been reported to be increased in obese [219], insulin resistant [220] and insulin

resistant obese [151] individuals. Activation of protein kinase C (PKC) may disturb GLUT4

translocation by Ser phosphorylation of IRS1 [88, 221]. Furthermore, it has been shown that

lipid-induced insulin resistance is associated with increased intramuscular DAG content [207],

and Bergman et al. showed that membrane, but not cytosolic DAG was associated with PKCε

activation [222]. As mitochondria are the main cellular site devoted to FA oxidation it has

been proposed that impaired mitochondrial function leads to accumulation of IMTG and lipid

intermediates in skeletal muscle [195]. However, this has been argued as the IMTG

accumulation may precede the development of mitochondrial dysfunction and/or that insulin

resistance may arise when mitochondrial function is unaffected or even improved [223, 224].

It has been shown that PLIN2 gene expression is lower in insulin resistant obese subjects

compared to obese controls [220], whereas higher PLIN2 protein content has been found in

skeletal muscle of insulin resistant subjects that have undergone weight loss or used

pharmacological treatment to increase muscle insulin sensitivity [225]. This suggests that

PLIN2 might play a role in decreasing intramuscular lipotoxicity by promoting lipid storage.

Moreover, improvements in insulin sensitivity following endurance [226] or resistance

training [227] are linked to an increase in content of PLIN2 and PLIN5, whereas similar

muscular PLIN2 protein content has been observed between obese non-diabetics and obese

diabetics and it correlated negatively with insulin-stimulated glucose uptake [228]. Overall

this indicates that a high expression level of LD-associated proteins might be preferable, and

further insight into how PLIN2 regulates LDs in skeletal muscle is necessary.

23

AIMS

The overall aim of the present thesis was to study regulation of glucose and lipid metabolism

in cultured skeletal muscle cells related to different metabolic status of the donors and

myotubes; myotubes from young sedentary vs. young trained subjects, myotubes from older

normal weight vs. older overweight subjects before and after 12 weeks of exercise, myotubes

from healthy subjects cultured with lactate, and myotubes from wild-type (WT) vs. Plin2

knockout (KO) mice. More specifically, the objectives of the different studies were:

1) Study the effects of 12 weeks extensive endurance and strength training in vivo on

energy metabolism in cultured human myotubes in vitro from sedentary normal

weight (BMI < 25 kg/m2) and sedentary overweight men (BMI ≥ 25 kg/m

2) in the age

of 40-62 years (paper I).

2) Investigate the effect of training status in vivo on glucose metabolism in myotubes

from sedentary untrained (VO2max < 45 ml/kg/min) and trained (VO2max > 60

ml/kg/min) subjects in the age of 21-38 years (paper II).

3) Explore fatty acid metabolism in myotubes established from trained (VO2max > 60

ml/kg/min) and sedentary untrained (VO2max < 45 ml/kg/min) younger subjects

(paper III).

4) Investigate lactate metabolism in cultured human myotubes and examine if lactate

exposure could affect metabolism of oleic acid and glucose (paper IV).

5) Study lipid storage capacity and turnover, as well as lipid and glucose metabolism and

muscle fiber type characteristics in myotubes from mice lacking Plin2 vs. WT mice

myotubes (paper IV).

24

SUMMARY OF PAPERS

Paper I: Exercise in vivo marks human myotubes in vitro: Training-induced increase in lipid

metabolism

The 12-week training intervention improved endurance, strength and insulin sensitivity in

vivo, and reduced the participants’ body weight and BMI. Biopsy-derived cultured human

myotubes after exercise showed increased total cellular oleic acid (OA) uptake (30%),

oxidation (46%) and lipid accumulation (34%), as well as increased fractional glucose

oxidation (14%) compared to cultures established prior to the exercise intervention. Most of

these exercise-induced changes were significant in the overweight group, whereas the normal

weight group showed no change in OA or glucose metabolism.

In conclusion, 12 weeks of combined endurance and strength training promoted lipid and

glucose metabolism in biopsy-derived cultured human myotubes, showing that training in

vivo are able to induce changes in human myotubes that are discernible in vitro.

The findings from the work in paper I are summarized in Table 1.

25

Table 1. Effects of 12 weeks of combined endurance and strength training in vivo on energy

metabolism in cultured myotubes established from normal weight and overweight subjects. An

increase or decrease in energy metabolism in vitro after participating in an in vivo exercise

intervention is indicated with ↑ and ↓, respectively. - indicates no exercise-induced differences. N/A

indicates that it was not evaluated due to too small sample size. Abbreviations: CPT1A, carnitine

palmitoyltransferase 1A; CYC1, cytochrome c1; FA, fatty acid; IRS1, insulin receptor substrate 1;

pAkt/Akt, phosphorylation of Akt/total Akt; pAMPKα/AMPKα, phosphorylation of AMP-activated

protein kinase α/total AMP-activated protein kinase α; PDK4, pyruvate dehydrogenase kinase 4;

PPARGC1A, peroxisome proliferator-activated receptor γ coactivator 1α; pTBC1D4/TBC1D4,

phosphorylation of TBC1 domain family member 4/total TBC1 domain family member 4; TFAM,

mitochondrial transcription factor A.

Normal weight Overweight Both groups combined

Lipid accumulation - ↑ ↑

FA uptake - - ↑

FA oxidation - ↑ ↑

FA fractional oxidation - ↑ ↑

Glucose uptake - - -

Glycogen synthesis - - -

Glucose oxidation - - -

Glucose fractional oxidation - ↑ ↑

PPARGC1A

(gene & methylation) - - -

PDK4 (gene & methylation) - - -

CPT1A (gene) - - -

CYC1 (gene) - - -

TFAM (methylation) N/A N/A -

ATP synthase (protein) - - -

pAMPKα/AMPKα - - -

pAkt/Akt - - -

pTBC1D4/TBC1D4 - - -

IRS1 (gene) ↓ - ↓

IRS1 (methylation) N/A N/A ↑

IRS1 (protein) - - -

26

Paper II: Glucose metabolism and metabolic flexibility in cultured skeletal muscle cells is

related to exercise status in young male subjects

Trained and intermediary trained subjects had higher maximal blood lactate levels than

untrained subjects after an incremental test, and skeletal muscle biopsies from the trained

group contained significantly more glycogen compared to biopsies from the intermediary

trained and untrained groups. Fiber type distribution in biopsies and myotubes was similar

between the groups. In cultured myotubes an increased glucose uptake was observed in cells

from trained subjects compared to cells from untrained. Fractional glucose oxidation was also

enhanced in trained myotubes, which were also more sensitive to the suppressive action of

acutely added OA to the cells.

In conclusion, differentiated skeletal muscle cells established from trained subjects with

increased capacity for energy production retained some of their phenotypes in vitro with

respect to enhanced glucose metabolism and metabolic flexibility.

Paper III: Higher fatty acid turnover and oxidation in cultured human skeletal muscle cells

from trained young male subjects.

Myotubes from trained subjects had lower FA accumulation, lower incorporation of OA into

total lipids, TAG, DAG, and CE, higher TAG-related lipolysis and re-esterification, and

higher FA complete oxidation and β-oxidation compared to myotubes from untrained

subjects. There were no significant differences in mRNA or protein expression between cells

from the two groups, but mRNA expression of CPT1B correlated positively with maximal fat

oxidation in vivo.

To conclude, myotubes established from trained subjects have increased FA turnover and

oxidation compared to myotubes from untrained subjects. Whether these properties in the

satellite cells are inherent from birth or acquired through lifestyle remains unknown.

The findings from the work in papers II and III are summarized in Table 2.

27

Table 2. Energy metabolism in vivo, ex vivo and in vitro in trained subjects compared to

untrained subjects. Higher or lower energy metabolism in trained compared to untrained subjects is

indicated with ↑ and ↓, respectively, whereas - indicates no differences. N/A indicates that it was not

evaluated. Abbreviations: ATGL, adipose triglyceride lipase; CD36, fatty acid translocase; CPT1,

carnitine palmitoyltransferase 1; CYC1, cytochrome c1; FA, fatty acid; GLUT4, glucose transporter 4;

GS, glycogen synthase; HKII, hexokinase II; HSL, hormone-sensitive lipase; MHC/MYH, myosin

heavy chain on protein and gene level, respectively; OXPHOS, oxidative phosphorylation; pAkt/Akt,

phosphorylation of Akt/total Akt; PDK4, pyruvate dehydrogenase kinase 4; PLIN, perilipin; PPARD,

peroxisome proliferator-activated receptor ; PPARGC1A, peroxisome proliferator-activated receptor

γ coactivator 1α; pTBC1D4/TBC1D4, phosphorylation of TBC1 domain family member 4/total TBC1

domain family member 4; TAG, triacylglycerol.

In vivo Ex vivo In vitro

Glucose uptake N/A N/A ↑

Glycogen synthesis/content N/A - -

Glucose/carbohydrate oxidation ↑ N/A -

Glucose fractional oxidation N/A N/A ↑

Suppression of glucose oxidation by FA N/A N/A ↑

Lipid accumulation & distribution N/A N/A ↓

TAG-related lipolysis & re-esterification N/A N/A ↑

FA oxidation ↑ N/A ↑

β-oxidation N/A N/A ↑

Fiber type I/MHCI/MYH7 & fiber type II/MHCIIa/MYH2 N/A - -

GLUT4 (gene) N/A N/A -

GLUT4 (protein) N/A - N/A

GS (protein) N/A - N/A

HKII (protein) N/A - N/A

Akt (protein) N/A - -

OXPHOS complexes (protein) & CYC1 (gene) N/A - -

PDK4 (gene & protein) N/A N/A -

PLIN2 & PLIN3 (gene & protein) N/A N/A -

ATGL & HSL (protein) N/A N/A -

pAkt/Akt & pTBC1D4/TBC1D4 N/A N/A -

CD36 (gene) N/A N/A -

PPARD & PPARGC1A (gene) N/A N/A -

CPT1A & CPT1B (gene) N/A N/A -

28

Paper IV: Lactic acid is readily used as an energy source or stored as glycogen and

intracellular lipids in human myotubes.

Myotubes expressed both lactate transporters, MCT1 and MCT4, and lactic acid was found to

be a substrate for both glycogen synthesis and lipid formation. Pyruvate and palmitic acid

(PA) inhibited lactic acid oxidation, whilst glucose and α-cyano-4-hydroxycinnamic acid (a

specific inhibitor of monocarboxylic acid transporters) inhibited lactic acid uptake. Acute

addition of lactic acid inhibited glucose and OA oxidation, whereas lactic acid increased OA

uptake. 24 h incubation with lactic acid did not affect glucose or OA metabolism. However,

when increasing the exposure time to lactic acid by replacing glucose with lactic acid during

the whole proliferation and differentiation period, glucose and OA oxidation as well as

glucose uptake were increased.

In conclusion, prolonged exposure to lactic acid increased glucose metabolism and OA

oxidation. Furthermore, lactic acid was found to be a substrate for glycogen synthesis, and it

was incorporated into lipids. Thus, lactic acid may be an important regulator of energy

metabolism in human skeletal muscle cells.

The findings from the work in paper IV are summarized in Table 3.

29

Table 3. Energy metabolism in myotubes after lactic acid treatment. Increased or decreased

energy metabolism in myotubes after treatment with lactic acid is indicated with ↑ and ↓, respectively,

whereas - indicates no differences. N/A indicates that it was not assessed. Abbreviations: FA, fatty

acid.

Basal 4-5 mM

lactic acid

10 mM

lactic acid

SLC16A1 (gene) Expressed - -

MCT1 (protein) Expressed - -

SLC16A4 (gene) Expressed ↓ -

MCT4 (protein) Possible - -

Glycogen synthesis

from [14

C]lactic acid Possible N/A N/A

Lipid distribution

from [14

C]lactic acid Possible N/A N/A

Acute exposure

(4 h)

Glucose uptake N/A - -

Glucose oxidation N/A - ↓

Fractional glucose oxidation N/A - -

FA uptake N/A ↑ -

FA oxidation N/A - ↓

Fractional FA oxidation N/A ↓ ↓

Chronic exposure

(24 h)

Glucose uptake N/A - -

GLUT4 (protein) N/A - -

Glucose oxidation N/A - -

FA uptake N/A - -

FA oxidation N/A - -

MHCI & MHCIIa (protein) N/A - -

Chronic exposure

(~14 days)

Glucose uptake N/A ↑ N/A

Glucose oxidation N/A ↑ N/A

FA uptake N/A - N/A

FA oxidation N/A ↑ N/A

Suppressibility - N/A N/A

Adaptability - N/A N/A

30

Paper V: Loss of perilipin 2 in cultured myotubes enhances lipolysis and shifts the metabolic

energy balance from glucose oxidation towards fatty acid oxidation.

Myotubes established from Plin2-/-

mice contained reduced content of LDs and accumulated

less OA in TAG and DAG, due to elevated LD hydrolysis compared to corresponding

Plin2+/+

myotubes. The reduced ability to store TAG in LDs in Plin2-/-

myotubes altered

energy metabolism from utilization of glucose towards that of FAs. Plin2-/-

myotubes were

characterized by higher oxidation of OA, lower glycogen synthesis and reduced glucose

oxidation compared to Plin2+/+

myotubes. In accord with these metabolic changes, ablation

of a functional Plin2 protein resulted in higher mRNA expression of Pparα and Ppargc1α,

transcription factors that stimulate expression of genes important for FA oxidation, the FA

transporter Cd36, the mitochondrial FA transporter Cpt2, and uncoupling proteins Ucp2 and

Ucp3. mRNA expressions of Cpt1b, a known facilitator of FA transport into mitochondria,

the peroxisomal acyl-CoA oxidase 2 (Acox2) and the basal glucose transporter Slc2a1 (Glut1)

were lower in Plin2-/-

compared to Plin2+/+

myotubes. On the other hand, mRNA expressions

of the other basal glucose transporter Slc2a4 (Glut4), the hexokinase Hk1, important for

activation of glucose, and the glycogen synthase, Gys1, were unaltered. In contrast, it was

observed a lower expression in mRNA expression of Pygm, involved in pathways leading to

glycogen degradation. Further, mRNA expression of genes involved in mobilization of

pyruvate for the TCA cycle, pyruvate kinase (Pkm) and pyruvate dehydrogenase (Pdha1),

were lower in Plin2-/-

myotubes compared to Plin2+/+

myotubes. mRNA expression of Pdk4,

a negative regulator of Pdha1 and key enzyme for switching fuel source from glucose

towards FAs, was higher in Plin2-/-

compared to Plin2+/+

myotubes. Loss of Plin2 had no

impact on insulin-stimulated Akt phosphorylation.

In conclusion, the results suggest that Plin2 is essential for balancing the pool of skeletal

muscle LDs to avoid an uncontrolled hydrolysis of the intracellular TAG pool. The

consequences of an increased release of FAs due to lack of Plin2 may therefore impact

skeletal muscle energy metabolism.

The findings from the work in paper V are summarized in Table 4.

31

Table 4. Energy metabolism in Plin2-/-

myotubes compared to Plin2+/+

myotubes. Higher or lower

energy metabolism in Plin2-/-

compared to Plin2+/+

myotubes is indicated with ↑ and ↓, respectively,

whereas - indicates no differences. Abbreviations: Acad, acyl-CoA dehydrogenase family; Acox,

acyl-CoA oxidase; CA, cell-associated radioactivity; Cd36, fatty acid translocase; Cpt, carnitine

palmitoyl transferase; FA, fatty acid; Fabp3, fatty acid binding protein 3; Gys1, glycogen synthase 1;

Hk1, hexokinase 1; LDs, lipid droplets; Pdha1, pyruvate hydrogenase α1; Pdk4, pyruvate

dehydrogenase kinase 4; Phk, phosphorylase kinase subunits; Pkm, muscle pyruvate kinase; Plin,

perilipin; Ppar, peroxisome proliferator-activated receptor; Ppargc1, peroxisome proliferator-activated

receptor γ coactivator 1; Pygm, muscle-associated glycogen phosphorylase; Slc2a, solute carrier

family 2; TAG, triacylglycerol; Ucp, uncoupling protein.

Plin2-/-

vs. Plin2+/+

myotubes

Plin2 (gene & protein) Not detected

Plin3 (gene & protein) ↓

LDs per nucleus ↓

Lipid accumulation & distribution ↓

TAG content ↓

Lipolysis ↑

CA of FA ↓

FA oxidation & β-oxidation ↑

Glycogen synthesis ↓

CA of glucose ↓

Glucose oxidation ↓

Ppara (gene) ↑

Pparg (gene) ↓

Ppard (gene) -

Ppargc1a (gene) ↑

Ppargc1b (gene) -

Cd36 (gene) ↑

Fabp3 (gene) -

Cpt1b (gene) ↓

Cpt2 (gene) ↑

Ucp2 & Ucp3 (genes) ↑

Acadm, Acadl & Acadvl (genes) -

Acox1 (gene) -

Acox2 (gene) ↓

Slc2a1 (gene) ↓

Slc2a4 (gene) -

Hk1 (gene) -

Gys1 (gene) -

Phka1 & Phkb (genes) -

Phkg1 (gene) ↑

Pygm (gene) ↓

Pkm & Pdha1 (genes) ↓

Pdk4 (gene) ↑

Akt (protein) ↓

Insulin-stimulated glycogen synthesis -

Insulin stimulated pAkt/Akt -

32

METHODOLOGICAL CONSIDERATIONS

Donor characteristics

Cultured myotubes used in this thesis were established from biopsies from adult donors of

different ages (21-58 years). Subjects in papers I-III had different degrees of metabolic

status and/or exercise status. All subjects were male in papers I-III, whilst in paper IV both

female and male donors were included. The biopsies were obtained after informed written

consent and approval by the Regional Committee for Medical and Health Research Ethics:

reference number 2011/882 for the study in paper I, reference number 2011/2207 for the

study in papers II-III and reference number S-04133 for the study in paper IV. A summary

of selected donor characteristics across the four studies with human myotubes included in this

thesis is given in Table 5 below.

Table 5. Summary of selected donor characteristics across the studies. Donor characteristics for

the seven donor groups used in the present thesis: normal weight before and after the exercise

intervention, overweight before and after the exercise intervention, untrained, trained, and lean.

Values are presented as means ± SEM. BMI, body mass index; f, fasting; n, number of subjects; S,

serum; TAG, triacylglycerol; VO2max, maximal oxygen uptake.

Donor

group

Paper n Age

(years)

BMI

(kg/m2)

fS-

insulin

(pmol/l)

fS-C-

peptide

(pmol/l)

fS-TAG

(mmol/l)

VO2max

(ml/kg/min)

Normal

weight

before

I

7

48.0

± 2.8

23.3

± 0.7

34.8

± 4.6

568.8

± 25.6

2.1

± 0.7

42.5

± 0.9

Normal

weight after

7 23.2

± 0.7

38.8

± 6.6

680.3

± 55.1

1.4

± 0.2

47.1

± 2.2

Overweight

before

11

51.9

± 1.8

29.4

± 0.7

51.2

± 5.9

818.6

± 73.8

1.9

± 0.2

37.1

± 1.5

Overweight

after

11 28.7

± 0.7

69.7

± 9.4

918.1

± 70.3

2.2

± 0.3

42.3

± 1.8

Untrained

II&III

6 26.8

± 2.8

28.0

± 2.5

119.5

± 35.6

636.8

± 110

1.2

± 0.3

41.1

± 1.6

Trained 6 24.3

± 0.7

23.9

± 0.9

49.5

± 10.6

314.3

± 10.3

1.4

± 0.3

64. 9

± 1.7

Healthy IV 11 40.3

± 3.7

23.6

± 0.8 - - - -

33

The use of BMI as a measure of obesity is as previously mentioned much disputed, but due to

its simplicity and validation in multiple epidemiological studies it is widely used. The main

limitations of BMI measurements are that it does not differentiate body fat from lean mass

and central (visceral) fat from peripheral (subcutaneous) fat. Thus, subjects who have a large

muscle mass, e.g. athletes, may be classified as overweight or even obese if BMI alone is

used as the diagnostic criterion. On the other hand, individuals with low lean body mass but

high body fat content may still have BMI within the normal range. Furthermore, individuals

with normal BMI and high body fat percentage also show a high degree of metabolic

dysregulation, known as normal weight obesity [178]. Therefore, other criteria such as waist

circumference or WHR should be included when diagnosing obesity.

It is known that there are differences with regards to energy metabolism between sexes in

vivo [229]. Women have higher expression levels of genes involved in lipid storage and

turnover, FA uptake, insulin sensitivity, and higher level of muscle fiber type 1 than men

[230-235]. On the other hand, men have higher whole-body resting energy expenditure and

skeletal muscle mass (SMM) [236]. This difference does not appear to be reflected in

cultured myotubes, unless incubated in presence of sex hormones [237], as studies have

shown that the metabolism of glucose and PA were similar in myotubes from male and

female donors [238]. The same studies showed similar mRNA expression levels of several

genes important in regulation of glucose and lipid metabolism. Thus, in the studies where

donor differences were of interest (papers I-III) only male donors were used. In paper IV

where we studied effect of lactic acid exposure the influence of using cultured myotubes from

both female and male donors were considered irrelevant.

Another factor that affects several metabolic processes in skeletal muscle in vivo is age. With

increasing age it has been observed an increase in impaired insulin sensitivity and obesity

[239-241], increased IMTG content [242], increased proportion of fast muscle fibers [243],

reduced SMM [244], reduced PPAR content of skeletal muscle [245], reduced

mitochondrial content [242, 246] and function [239], and it has also been associated with

damaged mitochondrial DNA [247-249]. However, it is thought that it is the combination of

obesity and inactivity and not age per se that are more important in age-related declines in

insulin sensitivity [240]. Furthermore, several of these age-related effects may be prevented

by exercise [241, 250], and it has been seen that an exercise-induced increase in muscle

mitochondria is maintained at least up to 70 years of age [251-253]. In this thesis, all muscle

biopsy donors were below 59 years old.

34

Study design of the in vivo exercise intervention

The research performed in paper I was approved, as part of a larger project: Skeletal Muscles,

Myokines and Glucose Metabolism (MyoGlu) [254]. The study (Clinical Trial Registration:

NCT01803568), a non-randomized parallel group study, was approved by the regional ethics

committee, and adhered to the Declaration of Helsinki. All participants performed two whole-

body strength training sessions and two spinning bike interval sessions under close

supervision weekly, representing in total 4 h of intensive exercise. The endurance training

sessions included one session of 7 min intervals at 85% of maximum heart rate (HRmax) and

one session of 2 min intervals at >90% of HRmax per week. Between the intervals the

participants were allowed to rest or cycle with a light load; time between intervals were 3 min

and 2 min for the 7 min and 2 min interval sessions, respectively. The number of 7 min

intervals progressed from 3 to 4 after the second week of training, and from 4 to 5 after six

weeks of training. Number of 2 min intervals progressed from 6 to 7 after the second week,

and from 7 to 10 after six weeks. The strength training sessions included a 10 min warm-up

and three sets of each of the exercises: leg curl, leg press, chest press, shoulder press, cable

pull-down, seated rowing, abdominal crunches, and back extensions. During the first three

weeks of training, a load that could be performed with 12 as repetition maximum (12-RM)

was used. From week four, sets of 10-RM were used, and after eight weeks sets of 8-RM

were used. Weight loads were continuously adjusted to ensure maximum resistance for the

requested number of repetitions. Abdominal crunches and back extensions were performed

with 12-20 repetitions. Before and after the exercise intervention several clinical parameters

were measured, including anthropometry, insulin sensitivity by euglycemic-hyperinsulinemic

clamp, body composition by bioelectric impedance analysis, maximum strength, and VO2max

by bicycle ergometry. Hyperinsulinemia during clamp suppresses endogenous (hepatic)

glucose production [255, 256]. However, the degree of suppression was not possible to

evaluate as hepatic insulin sensitivity was not directly measured. As glucose infusion rate

primarily is a quantification of peripheral insulin sensitivity, measurements of endogenous

glucose production, e.g. using glucose tracers, would have strengthened the study. The

muscle biopsies used for isolation of satellite cells were taken 2 h after an acute bicycle test.

For measurement of maximal strength the participants performed leg press, chest press and

cable pull-down with a 1-RM for each of the three exercises. To reduce learning effects 1-

RM was tested twice in the beginning of the study, where one strength training session was

performed before the first test and an additional training was performed before the second test.

35

After aerobic warm-up for 10 min the strength test started with 10 repetitions at a load below

50% of expected 1-RM, then 6 repetitions below 70%, three repetitions below 80%, one

repetition below 90%, and one repetition at a load below 95% of expected 1-RM. Thereafter,

resistance was gradually increased to reach 1-RM; subjects were allowed two attempts at this

load. Encouragement was given and the subjects were allowed 3 min rest between the

attempts.

The endurance tests were performed with a Monark Ergomedic 839E cycle and oxygen

consumption was measured with an Oxycon Pro. Expiration air was collected with a V2-

mask [257]. Capillary fingertip blood samples were taken for analyses of glucose and lactate.

Endurance sessions and tests were performed with Polar heart rate monitors, and heart rates

were analyzed with Polar ProTrainer 5 software. Subjects reported their perceived exertion

during training sessions and tests according to Borg’s scale [258, 259]. After a standardized

warm-up, the VO2max test started at a workload similar to the final load of an incremental test

in which the relationship between work (watt) and oxygen uptake was established. Subjects

cycled for 1 min and then the workload was increased by 15 watts every 30 seconds until

exhaustion. The test was concluded once oxygen uptake reached plateau (i.e. oxygen

consumption increased less than 0.5 ml/kg/min over a 30 watt increase in workload),

respiratory exchange ratio (RER) values > 1.1 and blood lactate > 7 mM. Before the exercise

intervention VO2max was tested twice (more than 3 days apart), but after the 12 weeks of

training only a single test was performed. The subjects abstained from training two days

before the VO2max tests. HRmax was defined during the VO2max test.

Cultured skeletal muscle cells as an in vitro model

The human muscle cells used in this thesis was obtained from musculus vastus lateralis

(papers I-III) or musculus obliquus internus abdominis (paper IV). The isolation of satellite

cells from all biopsies was performed at the same location and by the same trained

researchers. Multinucleated myotubes were established by activation and proliferation of

satellite cells based on the method of Henry et al. [260] and modified according to Gaster et

al. [261, 262]. During differentiation of activated satellite cells (myoblasts) into myotubes the

expression pattern of key proteins for both glucose [263] and lipid metabolism [264]

increases, and this pattern express a higher resemblance to adult skeletal muscle than the

pattern of myoblasts. Thus, myotubes are preferred in experiments [265]. Moreover, cultured

36

human myotubes have the most relevant genetic background for the study of metabolic

pathways and processes as compared to rodent cell cultures [266].

To be able to study the role of PLIN2 in skeletal muscle, muscle myoblast cultures were

established form the hind leg containing musculus gastrocnemius and musculus soleus from

Plin2+/+

and Plin2-/-

mice (paper V). We disrupted the Plin2 gene using standard

homologous recombination in embryonic stem cells. The use of KO mice enables us to study

the role of one specific protein at a time; however, silencing of a gene may elicit

compensatory cellular responses by up-regulation of genes sharing overlapping functions.

However, as we show in paper V, complete loss of Plin2 was not compensated by other

related Plin genes. Full KO of the gene is superior to silencing with siRNA, as some

expression of the target gene will be retained after silencing with siRNA. When comparing

metabolic studies between different species, it is important to consider the differences

existing between the species with regards to metabolic regulation, as nicely reviewed in [267].

For instance, mice have a basal metabolic rate that is approximately 7.5-fold higher than that

of humans. Furthermore, it appears that glucose tolerance in mice is more closely related to

hepatic insulin action rather than skeletal muscle insulin action.

Cultured human skeletal muscle cells are generally characterized by low mitochondrial

capacity and having fuel preference for carbohydrates over lipids [266]. The GLUT1:GLUT4

ratio is higher in cultured myotubes than in adult skeletal muscle [263, 268], resulting in

lower insulin responsiveness on glucose transport. Typically, insulin increases glucose uptake

by 40-50% in myotubes [263, 269]. However, despite the reduced insulin-responsiveness the

mechanisms involved in glucose uptake in vivo are conserved in vitro [263].

Another aspect to consider when comparing myotubes obtained from different sources of

muscle satellite cells is heterogeneity with respect to muscle fiber types. However, it has been

shown that human satellite cells isolated from either fast or slow muscle fibers form

myotubes in vitro which coexpress both fast and slow fibers independently of the fiber type

from which they were established [270]. Furthermore, it has been reported that myotubes

express fast fiber type regardless of donor muscle having mixed fiber type expression in vivo

[262]. Further, murine satellite cells isolated from various muscles have been shown to be

uniform regardless of muscle origin, and the dominant fiber type is the intermediate fiber

type, i.e. MHCIIa [271]. In our cells we have observed a significant amount of slow fiber

type after 8 days of differentiation [266]. In paper II we for the first time performed a

37

quantitative comparison between MHC expression in myotubes and the skeletal muscle

biopsies of which they originated from. The three isoforms expressed in human skeletal

muscle biopsies (MHCI, MHCIIa and MHCIIx) were expressed in the myotubes, but in

addition at least three other bands were visible. These may possibly represent developmental

forms of myosin as previously described by others [262, 270], highlighting the more limited

ability of cultured myotubes to fully differentiate to a similar degree as skeletal muscle in

vivo. We did not attempt to determine which isoform of MHCs that were present; however,

we have from raw data of a microarray mRNA expression analysis (reported in [272]), using

the same donors as in paper IV, observed substantial mRNA expression of both the

embryonic myosin MYH3 and the neonatal myosin MYH8 (developmental myosins, reviewed

by Schiaffino et al. [273]) as shown in Figure 3 (unpublished data). All taken together, these

findings demonstrate that myotubes differ from donor muscle with respect to MHC

expression. However, we have previously shown that cell content of MHCI is increased in

electrically pulse stimulated myotubes [274], showing plasticity potential of muscle cells.

Figure 3. mRNA expression of different myosin heavy chains in cultured human myotubes. Cells

established from healthy donors were cultured and differentiated into myotubes. Total RNA was

isolated and gene expression of myosin heavy chains (MYHs) assessed by microarray (data reported to

Gene Expression Omnibus, GSE40789). Data are presented as means ± SEM (n = 4).

Several characteristics of the in vivo phenotype are conserved in culture. For example, the

diabetic phenotype is conserved in myotubes [275, 276]. The ability to switch between lipid

and glucose oxidation also appears to be an intrinsic characteristic, as it is conserved in vitro

[94, 277]. The precise mechanisms by which skeletal muscle cells are able to retain the in

vivo characteristics are not known. However, a combination of genetic and epigenetic

mechanisms are likely to be involved, as reviewed in [266]. For instance, epigenetic

38

regulation of skeletal muscle stem cells and skeletal muscle differentiation, exercise, diet, and

a family history of T2D have all been described to influence DNA methylation and/or histone

modifications in human skeletal muscle [278, 279], and these are traits that might follow the

isolated satellite cells into their corresponding cultured myotubes.

Even though several traits seems to be intrinsic and conserved in vitro it has to be mentioned

that it has also been shown that the ability of the myoblasts to fuse and differentiate into

myotubes, as well as metabolic processes such as glucose uptake, glycogen synthesis, glucose

and FA oxidation, gradually can become impaired with increased passaging of the muscle

cells [280]. Therefore, all experiments performed in this thesis were on cells from passage

numbers that exerted normal responses, and passage number remained constant for each

experiment and within each donor, e.g. passage number remained constant for cells from

same donor before and after the exercise intervention in paper I.

Taken together, although some limitations, the cell model used in this thesis appears to be

valuable for studying skeletal muscle metabolism including conditions such as exercise

responses, effects of overweight/obesity and insulin resistance. Having the most relevant

genetic background, along with morphological, metabolic and biochemical properties of adult

skeletal muscle, differentiated primary human myotubes derived from satellite cells represent

the best available alternative system to intact human skeletal muscle that can be used to study

human diseases [260, 262, 266]. Moreover, as this system of cells is not immortalized it

allows investigation of the innate characteristics of the donors of which they were obtained

from. Since the extracellular environment can be precisely monitored this system is clearly

valuable for the purpose of research under controlled experimental conditions.

Methods used to measure energy metabolism in cultured skeletal muscle cells

Metabolic processes in this thesis were described by combining functional studies using

radiolabeled substrates with gene expression analyses using qPCR, protein expression

analyses using immunoblotting, DNA-methylation by pyrosequencing after bisulfite

treatment, as well as lipid composition analyses on isolated LDs, staining of LDs and nuclei

followed by live imaging or staining in fixated cells followed by imaging.

Scintillation proximity assay (SPA) [281] was used to study both real-time substrate

accumulation and lipolysis (papers II, III and V). Lipid accumulation of radiolabeled OA

39

was monitored over 24 h with measurements at several time points, and thereafter media was

changed to a media not containing radiolabeled OA to measure efflux of accumulated OA as

a measure of lipolysis (papers III and V). This corresponded with incline in radiolabeled

OA released to the cultivation media, measured as remaining radioactivity in the cells. The

assumption for this method is that most of the TAG used for lipolysis accumulates in LDs.

An inhibitor of ACSL, triacsin C, was used to inhibit FA re-esterification and oxidation

(paper III), and it would therefore reflect lipolysis more selectively (total lipolysis). Re-

esterification was calculated as the difference in lipolysis with and without triacsin C present

[282-284]. Previous studies have in cultured human myotubes shown that TAG resynthesis

[32, 285] and FA oxidation [283] are efficiently blocked by triacsin C, while lipolysis is

increased [283]. Glucose accumulation was also measured with SPA (paper II), but over 6 h.

The substrate oxidation assay [281] was used to study complete oxidation (CO2 production),

cell-associated radioactivity (CA) and metabolic flexibility. In addition, radioactivity released

to the culture media, acid soluble metabolites (ASM), from β-oxidation of OA was measured

(papers III and V). Glycogen synthesis assay was used to study incorporation of

radiolabeled glucose (papers I, II and V) and lactic acid (paper IV) into glycogen, and also

to measure an eventual insulin response as glucose uptake is a more uncertain method with

regards to this. Thin layer chromatography was used to study incorporation of radiolabeled

OA (papers III and V) and lactic acid (paper IV) into intracellular lipids and lipid classes.

In paper III we used pharmacological activation of PPAR with the selective and highly

potent agonist GW501516, to explore mitochondrial fuel utilization in myotubes from two

groups with different training status, i.e. high or low VO2max. In paper V we also used

various compounds to modify lipid turnover from the IMTG pool in myotubes, an inhibitor of

HSL/ATGL (CAY10499) [286] and an inhibitor of ATGL (Atglistatin) [287]. In paper IV

we treated the myotubes with various concentrations of lactic acid, from 2 mM to 20 mM

depending on the experiment. Human lactate concentrations may reach 10-20 mM in the

circulation [288, 289]. In the study behind papers II and III we measured blood lactate

concentrations during an incremental test as shown in Figure 4 below. We observed lactate

values were in the range ~1-5.5 mM during performance at 50-80% of maximal exercise

intensity, further supporting the chosen concentrations for lactic acid treatment in paper IV.

40

Figure 4. Blood lactate at different exercise intensities in subjects with differences in inherent

training status. Blood lactate concentrations were measured in capillary blood at several exercise

intensities during an incremental test using an YSI 1500 Sport Lactate Analyzer. Data are presented as

means ± SEM (n = 6 in untrained group, n = 11 in intermediary trained group and n = 6 in trained

group).

qPCR is useful for investigation of a limited number of anticipated regulated genes. DNA

methylation by pyrosequencing was assessed after bisulfite conversion in paper I. In skeletal

muscle, DNA methylation has been shown to respond to and be modulated by multiple

stimuli, including exercise, diet and metabolic diseases such as diabetes [279, 290-292]. For

selected genes, these studies have suggested a possible role of DNA methylation in muscle

cell metabolism by drawing positive or negative correlations with mRNA expression or

protein abundance. Sequencing techniques do not discriminate between a cytosine and a

methylated cytosine. Bisulfite treatment of DNA samples transforms unmethylated cytosines

into uracil, whereas methylated cytosines remain unchanged. Subsequent PCR amplifies the

target region while tyrosine (Tyr) is incorporated instead of uracils, which also represents

unmethylated cytosines of the original sequence. A challenge is to ensure complete bisulfite

conversion without DNA breakdown. In this thesis the EpiTect Fast Bisulfite Conversion Kit

from QIAGEN was used for this purpose. Pyrosequencing is a method used for de novo

sequencing and, when combined with bisulfite treatment, to determine methylation status of

short regions of DNA. The major challenge that was encountered was to design primers that

would successfully amplify and sequence our target region. Cautions in the design were CpG

density and repetitive motifs, because if the density of CpG was high then the primers would

overlap with CpGs and the binding would be biased or in worst case not effective. In the case

of repetitive motifs, the primer might then have bound at several locations, resulting in

41

amplification of several products. To ensure that good primers had been designed,

electrophoresis was performed before pyrosequencing.

Immunoblotting was applied to study expression and phosphorylation of relevant proteins,

and thus address changes in post-translational modifications. However, immunoblotting is

highly dependent on the quality of the antibodies used, and is a semi-quantitative method.

When studying phosphorylation by immunoblotting it is important to choose the right

phosphorylation site with regards to what the outcome measure is, e.g. effects of exercise or

insulin. The works presented in this thesis have collaterally studied phosphorylation of four

different proteins: Akt/PKB at Ser473, TBC1D4 at Thr642, AMPKα at Thr172, and IRS1 at

Tyr612. TBC1D4 was first discovered while screening for novel substrates of the Ser/Thr

kinase Akt2, which phosphorylates TBC1D4 at Thr642 and Ser588 [23, 293] after insulin

stimulation [294], which in turn is involved in translocation of GLUT4 to the plasma

membrane in adipose and muscle cells [293]. The phosphorylation site we chose for TBC1D4

in papers I and II was therefore Thr642. Akt is a Ser/Thr kinase which is activated by a

variety of growth factors in a PI3K-dependent manner through translocation to the plasma

membrane and phosphorylation at two regulatory sites, namely Thr308 in the activation loop

and Ser473 in the hydrophobic C-terminal regulatory domain [295, 296]. Therefore, an

antibody directed to one of these phosphorylation sites, Ser473, was chosen for

immunoblotting in papers I, II and V. IRS1 is a substrate of the insulin receptor tyrosine

kinase which upon tyrosine phosphorylation functions as a docking molecule to engage and

activate SH2 domain-containing proteins, e.g. PI3K [297]. Phosphorylation of Tyr612 is

mediated by the PKC pathway [298], and was chosen as the phosphorylation site in paper I.

However, IRS1 is also phosphorylated on Ser residues, and modulation of IRS1 Ser

phosphorylation in cells has been observed after treatment with a variety of substances,

including insulin [299, 300], thus phosphorylation at a Ser motif, such as Ser312 and Ser636

[301, 302], would also have been relevant. AMPK is a heterotrimeric complex composed of a

catalytic α subunit and regulatory β and γ subunits, each of which is encoded by two or three

distinct genes [303]. Phosphorylation of Thr172 is required for activation of AMPK [304-

306]; however, AMPKα is also phosphorylated at Thr258 and Ser485 (for α1, Ser491 for α2),

but the upstream kinase and the biological significance of phosphorylation at these sites has

not yet been elucidated [307], thus we chose to study phosphorylation at Thr172 in paper I.

Staining of LDs and nuclei in fixated cells followed by imaging (paper III and V) or

staining in cultured cells followed by live imaging (paper V) were other methods that were

42

used to get a better understanding of the cellular changes. The cells were incubated with

fluorescent substances that are known to diffuse through the plasma membrane and bind to

intracellular organelles. After excitation of the fluorophore the emitted light can be quantified.

In this thesis Hoechst 33258 and Bodipy493/503 were used. Hoechst 33258 binds to double

stranded DNA, and is used for a quantitative measure of DNA, whereas Bodipy 493/502 is a

lipophilic dye that accumulates in non-polar neutral lipids and is quite specific for LDs [308].

Images were taken randomly and acquisition and analysis were according to a standardized

protocol to avoid bias from the operator.

Data analyses and statistics

Most of the statistical methods used in this thesis were non-parametric, i.e. Mann-Whitney

test between different groups or Wilcoxon matched-pairs signed rank test within groups, as

the number of observations were too few to assume normal distribution or as testing of

normal distribution showed otherwise. Where tests confirmed normality, unpaired t test was

used between groups and paired t test was used within groups. Furthermore, linear mixed-

model analysis (LMM) was used, e.g. for time-dependent experiments, to compare the

differences between groups with between-donor variation and simultaneously compare

differences between groups with between-group variation. The LMM analysis includes all

observations and at the same time takes into account that not all observations are independent.

For correlation studies, Spearman correlation analyses were performed. Spearman’s test of

correlation is a non-parametric measure of statistical dependence between two variables. It

assesses how well the relationship between two variables can be described using a monotonic

function. A perfect correlation of +1 or -1 occurs when each of the variables is a perfect

monotone function of the other, and it is Spearman’s correlation coefficient (, most often

denominated as r) that defines the correlation’s strength. All values are reported as means ±

SEM, unless stated otherwise. The value n represents the number of different donors used,

each with at least duplicate samples. Differences were considered significant when p < 0.05.

43

DISCUSSION AND CONCLUSIONS

In this thesis, energy metabolism in cultured human skeletal muscle cells established from

lean, normal weight, overweight, sedentary untrained, and trained subjects was studied, as

well as in cultured skeletal muscle cells from mice expressing or lacking the Plin2 gene. In

paper I, energy metabolism was studied in myotubes established from normal weight and

overweight subjects before and after 12 weeks of combined endurance and strength training

in vivo. In paper II, effects of inherent training status on glucose metabolism and fiber type

distribution were studied in vivo, ex vivo and in vitro, whereas in paper III FA metabolism

was studied in cultured myotubes established from the same untrained and trained subjects as

included in paper II. In paper IV lactate metabolism and the impact of lactate on glucose

and FA metabolism was studied in cultured myotubes established from healthy donors.

Finally, in paper V the functional role of Plin2 on lipid turnover and energy metabolism as

well as muscle fiber remodeling was investigated in myotubes from mice.

Oxidative capacity

Cultured human myotubes are characterized by their low mitochondrial oxidative potential.

Three of the papers presented in this thesis using human myotubes (papers I-III) showed

increased mitochondrial oxidative capacity of OA and somewhat of glucose, using different

experimental approaches. Impairments in skeletal muscle glucose and FA oxidation have

been reported in association with obesity and T2D in the resting state [87, 93, 309]; thus, it is

important to determine whether interventions, either exercise or pharmacological, can reverse

these impairments.

For oxidative use, carbohydrates (derived from circulating plasma glucose or intramuscular

glycogen) can only enter the mitochondria via the decarboxylation of pyruvate to acetyl-CoA,

which is a reaction catalyzed by PDC. Therefore, PDC is positioned in such a way that it

plays one of the most essential roles in regulation of glucose metabolism, as well as fuel

selection in skeletal muscle, and it is also imperative for maintaining the cells’ energy

balance by regulation of ATP levels [31]. In skeletal muscle, PDK4 inhibits the activity of

PDC by phosphorylation, and thus increased PDK4 expression is associated with reduced

PDC activity and thereby reduced glucose oxidation [32]. Therefore, PDK4 plays an

important part in switching the oxidation towards FAs [310].

44

Skeletal muscle FA oxidation reaches its maximum at moderate intensities between 55% and

65% of VO2max, above which a gradual decrease in FA oxidation is observed [10]. This was

confirmed in paper II, where we observed peak fat oxidation in vivo for all participants,

trained, intermediary trained and untrained, at 55% exercise intensity with increased fat

oxidation with better training status. In contrast, prolonged exercise, at low to moderate

intensities, is associated with a steady increase in utilization of long-chain FAs [10].

In paper I we show for the first time effects on both lipid and glucose metabolism in vitro

after an in vivo exercise intervention, i.e. certain impacts of exercise in vivo are retained in

myotubes established from satellite cells from the same donors. Furthermore, we observed

that these exercise-induced changes were predominant in the overweight group. Thus,

cultured, passaged myoblasts established from satellite cells (i.e. progenitor cells) and

differentiated into myotubes, may be used as a model system for studying mechanisms

related to exercise and metabolic diseases. This was first indicated by Bourlier et al. [311],

which observed enhanced glucose metabolism in vivo that also was conserved in vitro after

an 8-week aerobic exercise intervention. However, they concluded that a longer exercise

intervention combining aerobic and anaerobic exercise was warranted to conclude whether

lipid metabolism in fact was not affected [311]. As we show in paper I both glucose and FA

metabolism were affected when increasing duration of the exercise intervention and

combining endurance and strength training. Bourlier et al. also suggested that epigenetic

modifications should be studied [311]. Therefore, in paper I we also examined whether

epigenetic modifications could be involved in the observed exercise-induced changes in

energy metabolism in the myotubes. In skeletal muscle, DNA methylation has been shown to

respond to and be modulated by several stimuli, including exercise [279, 291, 292]. We

investigated selected targets, PDK4, PPARGC1A, PPARD, mitochondrial transcription factor

A (TFAM), and IRS1, and found a significant hypermethylation of IRS1 overall for the three

CpGs examined, and a reduced IRS1 mRNA expression (paper I). No change in protein level

of IRS1 was observed (paper I). Further, one of the eight CpGs within the TFAM sequence

was hypomethylated, but no other CpGs were differentially regulated (paper I). Subtle

differences in DNA methylation could be explained by factors extrinsic to skeletal muscle or

changes in the cellular composition of skeletal muscle rather than changes in the epigenome

of the myonuclei. Furthermore, we only explored the DNA methylation in cultured myotubes,

and differences in the DNA methylation between cells in vivo and in vitro have been reported

45

[312]. Further, other epigenetic mechanisms may be involved, e.g. histone modifications, but

this was not studied.

Plasma lactate values increase rapidly during exercise [146]. Animal studies have shown that

oxidation accounts for approximately 75% of lactate removal during exercise, with the

remainder being used for gluconeogenesis in the liver and kidney [313]. Elevated blood

lactate levels have been shown to downregulate the use of glucose and FFAs as energy

substrates [313]. In paper IV we observed increased uptake and oxidation of glucose in

cultured human myotubes after prolonged chronic exposure (approximately 14 days) with

lactic acid. We also observed increased OA oxidation after the same treatment (paper IV).

Pharmacological activation with a PPARδ agonist and exercise training are proposed to

synergistically increase oxidative myofibers in adult mice [314]. PDK4 is important for FA

oxidation and a known target gene for PPAR [315-317], as was confirmed in paper III and

shown in Figure 5 (unpublished data).

Figure 5. PPAR activation by GW501516 increased mRNA expression of PDK4. Myotubes

established from trained and untrained subjects were treated for 96 h with 10 nM of the PPAR

agonist GW501516 or 0.1% DMSO (as control). Total RNA was isolated and mRNA expression of

PDK4 assessed by qPCR. Data are presented as means ± SEM (n = 6 in both groups). *Statistically

significant vs. Control (p < 0.05). Abbreviations: DMSO, dimethyl sulfoxide; PDK4, pyruvate

dehydrogenase kinase 4; PPAR, peroxisome proliferator-activated receptor delta; RPLP0, ribosomal

phosphoprotein lateral stalk subunit P0.

Results from studies in vivo indicate that PDK4 transcription and PDK4 mRNA levels are

increased in human skeletal muscle both during prolonged exercise and during recovery from

exercise [318-320], in response to fasting [321, 322] and to high-fat diets [323, 324].

However, it has been shown that factors important for FA oxidation such as CPT1B and

46

PDK4 are not further induced with exercise compared to PPARδ activation alone [314],

which is consistent with the findings in paper III that myotubes from subjects with higher

training status did not experience a higher PPAR agonist-induced mRNA expression of

CD36 (paper III) or PDK4 (Figure 5) compared to myotubes from subjects with lower

training status.

In both rodent and human skeletal muscle, PPARD mRNA is expressed to a greater extent

than PPARA, whereas mRNA expression of PPARG is very low [325, 326], as also observed

in cultured mice myotubes in paper V. They are transcription factors known to be activated

by FAs [327, 328]. When comparing myotubes from Plin2-/-

with Plin2+/+

mice we observed

similar mRNA levels of Ppard, the predominant subtype in skeletal muscle, but higher levels

of Ppara, which is abundantly expressed in tissues with high FA catabolism including

skeletal muscle [329-331], and lower levels of Pparg, which primarily is expressed in white

adipose tissue and stimulates expression of genes promoting lipid storage [332] (paper V).

Similar differences in PPARα expression have been observed in liver of Plin2+/+

and Plin2-/-

mice [333], whereas the opposite expression pattern has been found in muscle electroporated

to overexpress Plin2 [334]. Ablation of Plin2 in mice also resulted in a metabolic shift in

energy metabolism from utilization of glucose towards utilization of FAs, i.e. a higher FA

oxidation and lower glucose metabolism was observed in Plin2-/-

myotubes (paper V). In line

with this, mRNA expressions of the fatty acid transporter Cd36 as well as Ppargc1a, a master

regulator of mitochondrial biogenesis [103] and known to facilitate FA oxidation [158, 264],

the mitochondrial FA transporter Cpt2, and the uncoupling proteins (Ucp) 2 and 3 were

higher in Plin2-/-

compared to Plin2+/+

myotubes. Genes encoding for mitochondrial acyl-

CoA dehydrogenases (Acadm, Acadl and Acadvl) as well as peroxisomal acyl-CoA oxidase 1

(Acox1) were unaltered, whereas mRNA expression of Cpt1b, known to facilitate FA

transport into mitochondria [45], and Acox2 were lower in Plin2-/-

than Plin2+/+

myotubes

(paper V). With regards to genes involved in glucose metabolism, mRNA expression of

solute carrier family 2 member 1 (Slc2a1, encoding for Glut1) was lower in Plin2-/-

than

Plin2+/+

myotubes, whereas mRNA expressions of Slc2a4 (encoding for Glut4), Hk1,

important for activation of glucose, glycogen synthase 1 (Gys1), and genes encoding for two

of the subunits in phosphorylase kinase (Phka1 and Phkb) were similar between the two WT

and KO myotubes (paper V). Expression of a third phosphorylase kinase subunit (Phkg1)

was elevated, whereas lower mRNA expressions of muscle-associated glycogen

phosphorylase (Pygm), involved in the pathways leading to glycogen degradation, pyruvate

47

kinase (Pkm) and pyruvate dehydrogenase alpha 1 (Pdha1), both involved in mobilization of

pyruvate for the TCA cycle, were observed in Plin2-/-

compared to Plin2+/+

myotubes (paper

V). Also, mRNA expression of Pdk4, a negative regulator of Pdha1, was higher in Plin2-/-

than Plin2+/+

myotubes (paper V). Altogether, although these transcriptional changes in the

Plin2-/-

myotubes do not fully overlap with transcriptional changes reported for mice with

overexpression of Ppara in muscle [335], repression of genes important for glucose uptake

(Slc2a1), degradation of glycogen (Pygm) and glycolysis (Pkm, Phda1 and Pdk4) may

explain the reduced glucose oxidation in the Plin2-/-

myotubes (paper V).

As described above, PDC is positioned in such a way that it has one of the most central roles

in regulation of glucose metabolism as well as fuel selection in skeletal muscle [31]. Both

pharmacological activation of PPAR by GW501516 (paper III and Figure 5) and ablation

of Plin2 (paper V) resulted in an increase in mRNA expression of PDK4. However, no

difference in protein expression of PDK4 was observed (paper II). As PDK4 is known to

inhibit PDC, and thereby reduce glucose oxidation [31], we indicate in paper V that PDK4 is

an important enzyme for switching the fuel source from glucose towards FAs. However, in

papers II and III we observed both increased fractional glucose oxidation and FA oxidation

in myotubes from trained compared to myotubes from untrained. Others have also observed

that PDK4 expression is increased after PPAR activation [315-317, 336]. Furthermore,

PDK4 inhibition of glucose oxidation in response to elevated plasma FA availability has been

observed both during fasting [321, 322] and high-fat diet [337]. At these conditions the

availability of carbohydrates is low, so PDK4 contributes to glucose preservation by

preventing its oxidation. Therefore, by regulation of PDK4, PPAR is suggested to play a

role in utilization of FAs under physiological conditions. On the other hand, trained muscle

fibers have been shown to use more glucose than untrained fibers [114, 115]. This was

confirmed in myotubes from trained subjects compared to myotubes from untrained subjects

in paper II, and with enhanced oxidation of carbohydrates after exercise in obese subjects

[311], in line with what we observed in paper I in cells from overweight subjects compared

to cells from normal weight subjects after an exercise intervention in vivo. Increased glucose

oxidation after exercise has also been shown in subjects with T2D [338], and our group have

also shown that electric pulse stimulation is able to increase the oxidative capacity of glucose

in human myotubes isolated from lean, healthy subjects and severely obese subjects with or

without T2D [339].

48

To summarize the results in papers I-V, there are several common traits between the studies,

as shown in Figure 6 below, which may further elucidate the positive effects of exercise as

prevention and treatment of metabolic diseases. 12 weeks of exercise in vivo (paper I) as

well as high inherent exercise status (papers II and III) and chronic lactate treatment (~14

days, paper IV) increased glucose and FA oxidation in the cells. Although healthy donors

without any exercise stimuli were used to study lactate metabolism, chronic treatment with

lactate throughout the entire proliferation and differentiation period can perhaps be somewhat

related to, although more extreme, the lactic acid build-up during prolonged exercise. Indeed,

we did observe an inhibitory effect of lactic acid on glucose and oleic acid metabolism

(paper IV). KO of Plin2 (paper V) also increased the oxidative capacity of the cells

compared to WT cells.

Figure 6. Comparison of glucose and lipid/fatty acid (FA) metabolism across the five studies. An

increase or decrease is indicated with ↑ and ↓, respectively. - indicates no differences. If no symbol is

given it indicates that it was not assessed. Dark green represents results from paper I, light green

represents results from papers II and III, purple represents results from paper IV, whereas orange

represents results from paper V. The processes have been explained previously (Figure 1, p. 11).

49

Lipid storage and turnover

LDs are dynamic organelles resulting from the balance between storage and breakdown of

TAG by lipases to generate FAs available for oxidation in the mitochondria [340]. Exercise

has been found to increase lipid storage in skeletal muscle [152, 153, 341], but as we show in

paper III myotubes from trained subjects had in fact lower fatty acid accumulation and

incorporation into complex lipids compared to myotubes from untrained subjects.

High levels of IMTG have been shown to correlate with insulin resistance [205, 342-344].

However, it has also been observed that insulin resistance can occur independently of

changes in IMTG content, and thereby dissociating IMTG concentrations from insulin

resistance [345]. Studies using cultured human myotubes have shown increased storage of

neutral lipids together with improved glucose metabolism and metabolic switching of the

cells [95, 346, 347], which suggests that to enhance partitioning of excess FAs towards

storage is beneficial to prevent insulin resistance by limiting accumulation of lipotoxic

intermediates [348-351]. It has been hypothesized that insulin resistance develops due to

lower lipid accumulation and higher lipolysis without an increase in FA oxidation [352],

processes which may lead to increased accumulation of lipotoxic intermediates that could

interfere with insulin signaling [217, 222, 350, 353]. In accordance, as we show in paper III,

myotubes from trained subjects had lower lipid accumulation and higher TAG-related

lipolysis and re-esterification than myotubes from untrained subjects, but myotubes from

trained also had higher FA oxidation than myotubes from untrained. We did not measure

lipolysis in paper I; however, lipid accumulation and FA oxidation were measured.

Myotubes from overweight participants isolated before the exercise intervention showed

lower lipid accumulation and no differences in FA oxidation compared to myotubes from

normal weight participants (paper I). However, after the exercise intervention myotubes

from overweight had higher lipid accumulation and FA oxidation compared to myotubes

from normal weight (paper I). Moreover, there were no exercise-induced differences in gene

expressions of either PLIN2 or CD36 (paper I).

Using Plin2-/-

myotubes we demonstrated that Plin2 plays an essential role in skeletal muscle

lipid storage and turnover (paper V). Similar to earlier studies based on partial ablation of

Plin2 [334, 354], we showed that complete removal of Plin2 in myotubes (paper V)

generated cells with reduced number of LDs and with less accumulated TAG. We also

showed that loss of Plin2 did not affect the FA uptake rate across the plasma membrane or

the amount of FAs incorporated into LDs, but merely increased degradation of the TAG

50

deposited within LDs by interfering with lipolysis (paper V). In support of this, earlier

studies have shown that overexpression of PLIN2 in embryonic kidney cells from hamster

limited the interaction of lipases with LDs [355] and increased lipolysis in mouse hepatocytes

with combined PLIN2 and PLIN3 knockdown [354]. Furthermore, as we show in paper V,

inhibitors against ATGL and HSL only had an effect on LD accumulation in Plin2-/-

myotubes, suggesting that ATGL and HSL gained more access to LDs or increased in activity

in cells lacking Plin2. In support of this, it has been observed higher TAG accumulation and

lower lipolysis in skeletal muscle of ATGL-/-

mice [356], whereas HSL-/-

mice has been

shown to contain higher levels of DAG [357]. In line with a functional role of PLIN2 as TAG

protector, Plin2-/-

myotubes exposed to OA accumulated less DAG (paper V), a finding that

contradicts the previous report of increased incorporation of PA into DAG in PLIN2

knockdown C2C12 cells [334]. As it has been shown a functional redundancy among some

PLINs [354], the incomplete removal of the PLIN2 protein in PLIN2 knockdown cells [334]

compared to complete removal of PLIN2 in our Plin2-/-

cells (paper V) likely contributes to

these discrepancies. Furthermore, the different types of labeled FAs used may also contribute,

as PA is accumulated to a lower extent into LDs than OA in myotubes [283]. Interestingly, in

paper IV we show for the first time in cultured human myotubes that lactic acid is able to be

stored as lipids. Chen et al. has shown that lactate can be metabolized to lipids in cultured

HeLa cells and H460 human lung cancer cells [358], and Jin et al. has shown that lactic acid

can be converted to glycerol in rat muscle [359].

Fiber type transformations in skeletal muscles

It has been suggested that there is an association between insulin sensitivity and the amount

of oxidative type I fibers, as lower expression of type I fibers has been observed in muscle

biopsies from subjects with insulin resistance and T2D when compared to healthy subjects

[360, 361]. However, it has also been shown that there were no differences in fiber type

distribution between muscle biopsies from lean non-diabetic and obese diabetic subjects

[362]. Furthermore, myotubes in culture differ from donor muscle as human satellite cells

form myotubes in vitro independently of the fiber type from which they originated [262, 270,

271]. In paper I we did not observe any differences in protein expressions of MHCI

(representing type I muscle fibers) or MHCIIa (representing type II muscle fibers) between

the groups or as a result of the exercise intervention. We did not observe any differences in

protein expression of these MHCs in paper IV either after 24 h incubation with lactic acid. In

51

paper II we performed for the first time a quantitative comparison between MHC expression

in myotubes and the skeletal muscle biopsies from which they originated. The three isoforms

expressed in human skeletal muscle biopsies (MHCI, MHCIIa and MHCIIx) were expressed

in the myotubes, but in addition at least three other bands were visible (discussed in more

detail in “Cultured skeletal muscle cells as an in vitro model”, pp. 36-37). In paper V,

ablation of Plin2 in mice resulted in myotubes that were more oxidative and less glycolytic.

Further, loss of Plin2 (paper V) also caused higher gene expression of oxidative type I fiber

marker (Myh7) and lower expressions of the glycolytic fiber type markers (Myh1, Myh2 and

Myh4) together with higher gene expressions of Mef2c and the myogenic markers Myod1 and

myogenic factor 5 (Myf5, Figure 7 (unpublished data)). We also observed higher mRNA

expression of Ppargc1a (paper V). Underpinning these results, MEF2 [363-365] and

PPARGC1A [103] have been shown to regulate fiber type switching from glycolytic type II

fibers to oxidative type I fibers. In fact, muscle specific KO of MEF2C resulted in decreased

proportion of oxidative fibers, whilst overexpression of MEF2C increased the proportion of

oxidative fibers [102]. Furthermore, we have previously shown that overexpression of

PPARGC1A in human myotubes resulted in enhanced lipid oxidative capacity and decreased

gene expression of MYH2 [158]. Both MYOD and MYF5 are important transcription factors

that activates many downstream genes to initiate muscle cell differentiation to multinucleated

myotubes [366]. Additionally, MYOD is proposed to be implicated in the fast-fiber formation

[367]. However, other studies have shown that neither KO of the MYOD gene in mice [368]

nor increased expression of MYOD after exercise in rat [369] were involved in fiber type

transformations. How lacking Plin2 can influence skeletal muscle differentiation/fiber type

remodeling remains unclear, but it is likely that increased flux of FAs may contribute to the

observed metabolic shift in energy metabolism from utilization of glucose towards FAs, and

to the reduced expression of glycolytic and increased expression of the oxidative muscle fiber

type markers in Plin2-/-

myotubes (paper V and Figure 7).

52

Figure 7. mRNA expression of fiber type markers in Plin2+/+

and Plin2-/-

myotubes. Differentiated

Plin2+/+

and Plin2-/-

myotubes was analyzed by qPCR. (A) Expression of myosin heavy chains (Myhs)

linked to muscle fiber type: Myh1 (type IIx fibers), Myh2 (type IIa fibers), Myh4 (type IIb fibers), and

Myh7 (type I fibers). (B) Expression of myogenic regulatory factors: Myocyte enhancer factor 2c

(Mef2c), myogenic differentiation 1 (Myod1) and myogenic factor 5 (Myf5). Data are presented as

means ± SEM (n = 3) normalized to expression of the housekeeping control TATA box binding

protein (Tbp).

Effects on insulin sensitivity

Disturbances in energy metabolism of skeletal muscle are associated with metabolic diseases

related to insulin resistance [370, 371]. In paper I we found a significantly increased glucose

infusion rate in vivo after the exercise intervention in both normal weight and overweight

subjects, indicating increased insulin sensitivity. However, we did not observe any exercise-

induced changes in in vitro insulin response, i.e. insulin-stimulated Akt phosphorylation,

TBC1D4 phosphorylation or glycogen synthesis. This may possibly be explained by

suboptimal experimental conditions in vitro, e.g. treatment with an insulin concentration

leading to maximal insulin stimulation; however, we have previously been able to detect

donor-specific differences in insulin-response with the same experimental setup as in paper I

[284, 352]. Thus, the lack of exercise-induced changes in insulin response in vitro may also

be a result of the underlying study in vivo where the difference in insulin sensitivity was quite

small, though significant, between the normal weight group and overweight group, and that

this difference was too small to be able to detect in vitro. Others have shown higher exercise-

induced changes in insulin sensitivity in subjects who were more insulin-resistant at baseline

[372, 373], and even more evident in obese men compared to lean men [373], which also has

been observed after electric pulse stimulation of myotubes in vitro [339]. We were not able to

replicate this in paper I, where most of the subjects in the overweight group also were

dysglycemic, but with no greater insulin-stimulated responses in their myotubes after the in

53

vivo exercise intervention. However, we do not rule out the possibility that interventions with

an increased physical activity level may be effective to improve insulin sensitivity in

myotubes from population groups with insulin resistance or are prone to have insulin

resistance.

We did not observe any differences in in vitro insulin response between myotubes from

trained and untrained subjects (paper II), and these two donor groups also had quite similar

insulin sensitivity in vivo. As previously stated, it has been hypothesized that insulin

resistance is a result of lower lipid accumulation and higher lipolysis without increased FA

oxidation [352], processes which may lead to higher accumulation of lipotoxic intermediates

that could interfere with insulin signaling. This is in accordance with what we showed in

paper III where myotubes from trained subjects also had lower lipid accumulation and

higher lipolysis, but additionally they showed higher FA oxidation.

Ablation of Plin2 (paper V) did not affect insulin sensitivity in myotubes measured as an

effect on insulin-stimulated responses despite of improvement in lipid oxidative capacity.

This suggests that insulin resistance in skeletal muscle is not solely dependent on increased

lipid accumulation. Although a complete loss of Plin2 in myotubes did not affect insulin-

stimulated responses, it generated cells with reduced expression of total Akt protein that may

contribute to the reduced glucose metabolism observed in Plin2-/-

cells (paper V). However,

the three different isoforms of Akt (Akt1-3) have been shown to have distinct roles, where

Akt2 is specifically involved in the maintenance of glucose homeostasis [122, 374], and

therefore it remains to clarify which isoform(s) of the Akt proteins is reduced in our Plin2-/-

muscle cells.

Final considerations

The myotubes used in this thesis were established from donors of different origin, age and

physical activity level. Metabolism of OA and glucose in the cells was increased in human

myotubes as a result of an in vivo exercise intervention (paper I) and higher in myotubes

from trained compared to untrained donors (papers II and III). We have also, to the best of

our knowledge, shown for the first time that myotubes from trained subjects were more

flexible with regards to exploiting the available fuel source as seen by more effective

inhibition of glucose oxidation by FAs in myotubes from trained subjects compared to

myotubes from untrained subjects (paper II). This may possibly be explained by altered

54

access for FAs to mitochondria. Furthermore, myotubes from trained subjects had a higher

FA turnover and oxidation compared to myotubes from untrained subjects, indicating a

higher preference for FA metabolism with higher training status (paper III). In summary

(papers II and III), myotubes from trained subjects appear to have different metabolic

properties than myotubes from untrained, but whether these differences are inherent from

birth or acquired through e.g. lifestyle remains unknown. Lactate exposure affected

metabolism of two other major substrates for energy metabolism, namely glucose and OA, in

cultured human myotubes, and the impact of lactate exposure appeared to be related to the

exposure time (paper IV). This may further be related to the lactic acid build-up during

strenuous exercise, although more extreme. Thus, we suggest that lactate may have an

important role in regulation of energy metabolism on a cellular level (paper IV). Oxidative

capacity was also increased in myotubes established from mice lacking Plin2 (paper V), and

the increase was associated with a shift from glucose utilization towards FA utilization. We

also concluded that Plin2 was essential for balancing the pool of skeletal muscle LDs to

avoid an uncontrolled hydrolysis of the intracellular TAG pool (paper V). Interestingly, in

paper IV we showed for the first time in cultured human myotubes that lactic acid was

incorporated into lipids and glycogen. Increased physical activity (paper I), having high

training status (papers II and III) or ablation of Plin2 (paper V) did not affect insulin-

stimulated responses in cultured myotubes. Finally, we conclude that cultured, passaged

myotubes are valuable as an in vitro model system for studying mechanisms related to

exercise and metabolic diseases.

55

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I

RESEARCH ARTICLE

Exercise in vivomarks humanmyotubes invitro: Training-induced increase in lipidmetabolism

Jenny Lund1 , Arild C. Rustan1 , Nils G. L vsletten1 , Jonathan M. Mudry2, TorgrimM. Langleite3, Yuan Z. Feng1, Camilla Stensrud1, Mari G. Brubak1, Christian A. Drevon3,K re I. Birkeland4, Kristoffer J. Kolnes5, Egil I. Johansen5, Daniel S. Tangen5, HansK. Stadheim5, Hanne L. Gulseth4, Anna Krook2, Eili T. Kase1, J rgen Jensen5, G.Hege Thoresen1,6

1 Department of Pharmaceutical Biosciences, School of Pharmacy, University of Oslo, Oslo, Norway,2 Integrative Physiology, Department of Physiology and Pharmacology, Karolinska Institutet, Stockholm,Sweden, 3 Department of Nutrition, Institute of Basic Medical Sciences, University of Oslo, Oslo, Norway,4 Department of Endocrinology, Morbid Obesity and Preventive Medicine, Oslo, University Hospital andInstitute of Clinical Medicine, University of Oslo, Oslo, Norway, 5 Department of Physical Performance,Norwegian School of Sport Sciences, Oslo, Norway, 6 Department of Pharmacology, Institute of ClinicalMedicine, University of Oslo, Oslo, Norway

These authors contributed equally to this work.* [email protected]

Abstract

Background and aimsPhysical activity has preventive as well as therapeutic benefits for overweight subjects. In

this study we aimed to examine effects of in vivo exercise on in vitrometabolic adaptations

by studying energy metabolism in cultured myotubes isolated from biopsies taken before

and after 12 weeks of extensive endurance and strength training, from healthy sedentary

normal weight and overweight men.

MethodsHealthy sedentary men, aged 40–62 years, with normal weight (body mass index (BMI) < 25kg/m2) or overweight (BMI� 25 kg/m2) were included. Fatty acid and glucose metabolism

were studied in myotubes using [14C]oleic acid and [14C]glucose, respectively. Gene and

protein expressions, as well as DNAmethylation were measured for selected genes.

ResultsThe 12-week training intervention improved endurance, strength and insulin sensitivity in

vivo, and reduced the participants’ body weight. Biopsy-derived cultured human myotubes

after exercise showed increased total cellular oleic acid uptake (30%), oxidation (46%) and

lipid accumulation (34%), as well as increased fractional glucose oxidation (14%) compared

to cultures established prior to exercise. Most of these exercise-induced increases were sig-

nificant in the overweight group, whereas the normal weight group showed no change in

oleic acid or glucose metabolism.

PLOSONE | https://doi.org/10.1371/journal.pone.0175441 April 12, 2017 1 / 24

a1111111111a1111111111a1111111111a1111111111a1111111111

OPENACCESS

Citation: Lund J, Rustan AC, L vsletten NG, MudryJM, Langleite TM, Feng YZ, et al. (2017) Exercise invivomarks humanmyotubes in vitro: Training-induced increase in lipid metabolism. PLoS ONE 12(4): e0175441. https://doi.org/10.1371/journal.pone.0175441

Editor:Makoto Kanzaki, Tohoku University, JAPAN

Received:November 7, 2016

Accepted:March 27, 2017

Published: April 12, 2017

Copyright: 2017 Lund et al. This is an openaccess article distributed under the terms of theCreative Commons Attribution License, whichpermits unrestricted use, distribution, andreproduction in any medium, provided the originalauthor and source are credited.

Data Availability Statement: All relevant data arewithin the paper and its Supporting Informationfiles.

Funding: This work was funded by research grantsfrom the University of Oslo, Karolinska Institutet,the Norwegian Diabetes Association, Throne HolstFoundation of Nutrition Research, AktieselskabetFreia Chocolade Fabriks Medical Foundation,Norwegian PhD School of Pharmacy, South-Eastern Norway Regional Health Authority,Swedish Diabetes Association, Swedish Researchcouncil, Anders Jahres Foundation, and EU-

Conclusions12 weeks of combined endurance and strength training promoted increased lipid and glu-

cose metabolism in biopsy-derived cultured human myotubes, showing that training in vivo

are able to induce changes in human myotubes that are discernible in vitro.

IntroductionPhysical activity has preventive as well as therapeutic benefits for metabolic diseases associated

with insulin resistance such as obesity and type 2 diabetes mellitus (T2D) [1, 2]. In addition to

increased physical activity, dietary changes and weight loss are important lifestyle changes for

prevention as well as treatment of T2D [2], as increased body mass index (BMI) is strongly

associated with the prevalence of metabolic diseases [3, 4], and most type 2 diabetics are over-

weight or obese [5]. Physical activity is known to improve insulin sensitivity and glucose

homeostasis and to increase fatty acid oxidation in skeletal muscle [6–8], as well as to reduce

blood pressure and beneficially influence plasma lipoproteins [9].

Skeletal muscle is the largest glucose-consuming organ in the body and accounts for more

than 80% of the insulin-stimulated glucose disposal [10]. Skeletal muscle is also the primary

site for insulin resistance [11]. Also with regard to fatty acid metabolism, skeletal muscle is

quantitatively the most dominant tissue during exercise [7]. Satellite cells [12] are dormant

cells in mature skeletal muscle in vivo, but are activated in response to stress, e.g. muscle

growth [13], and may be activated in culture to proliferating myoblasts and differentiated into

multinucleated myotubes. Epigenetic changes such as DNA methylation of key regulatory

genes has been proposed as one of several molecular mechanisms to explain the beneficial

effects of lifestyle changes, as both diet and exercise can influence DNAmethylation [14, 15].

Several studies indicate that cultured myotubes retain the in vivo characteristics (see e.g. [11,16–20]), and although the precise mechanisms are not known, epigenetic changes may be

involved (discussed in [21]). Thus, cultured human myotubes may represent an ex vivomodel

system for intact human skeletal muscle [19].

Most studies on the effect of exercise on metabolic diseases have been performed in vivo[22, 23] or directly on muscle biopsies [24, 25]. However, a study on obese donors revealed

that enhanced glucose metabolism noted in vivo following 8 weeks aerobic exercise, was pre-served in cultured primary myotubes [16]. To further explore the effects of in vivo exercise onin vitrometabolic adaptations, we studied different aspects of energy metabolism in cultured

myotubes established from biopsies from healthy sedentary normal weight and overweight

men. Biopsies were obtained before and after 12 weeks of physical training, consisting of both

endurance and strength exercises.

Materials andmethods

MaterialsMaterials are reported in Table 1.

Ethics statementThe biopsies were obtained after informed written consent and approval by the Regional Com-

mittee for Medical and Health Research Ethics North, Tromsø, Norway (reference number:

2011/882). The research performed in this study was approved, as part of a larger project:

Changedmetabolism in myotubes from overweight post-training

PLOSONE | https://doi.org/10.1371/journal.pone.0175441 April 12, 2017 2 / 24

financed FP7 project (NutriTech grant agreementno.: 289511). JJ is a Visiting Professor atDepartment of Nutrition, Exercise and Sports,University of Copenhagen, supported by TheDanish Diabetes Academy and Novo NordiskFoundation.

Competing interests: The authors report noconflicts of interests.

Abbreviations: AMPK, AMP-activated proteinkinase; ANGPTL4, angiopoietin-like 4; BMI, bodymass index; CA, cell-associated radioactivity;CD36, fatty acid translocase; CPT1A, carnitinepalmitoyltransferase 1A; CYC1, cytochrome c-1;GAPDH, glyceraldehyde 3-phosphatedehydrogenase; GIR, glucose infusion rate; IRS1,insulin receptor substrate 1; MHC, myosin heavychain; OXPHOS, oxidative phosphorylation; PDK4,pyruvate dehydrogenase kinase, isoenzyme 4;PPARGC1A, peroxisome proliferator-activatedreceptor gamma, coactivator 1 alpha; PLIN2,perilipin 2; PPARD, peroxisome proliferator-activated receptor delta; RPLP0, large ribosomalprotein P0; T2D, type 2 diabetes mellitus; TBC1D4,TBC1 domain family member 4; TFAM,transcription factor A, mitochondrial; VO2max,maximal oxygen uptake.

Table 1. List of materials and respective producers.

Material Producer

Nunc™ Cell Culture Treated Flasks with Filter Caps ThermoFisher Scientific (Roskilde,Denmark)Nunc™ 96-MicroWell™ plates

Pierce™ BCA Protein Assay Kit

SuperSignal West Femto Maximum Sensitivity Substrate

O´GeneRuler 100 bp DNA ladder

Antibody against phosphorylated IRS1 at Tyr612 (#44-816G)

Primers for TaqMan PCR

DMEM-Glutamax™ low glucose with sodium pyruvate Gibco Invitrogen (Gibco, LifeTechnologies, Paisley, UK)FBS

Trypsin-EDTA

Penicillin-streptomycin (10000 IE/ml)

Amphotericin B

DPBS (without Mg2+ and Ca2+)

Ultroser G Pall (Cergy-Saint-Christophe, France)

Insulin (Actrapid® Penfill® 100 IE/ml) Novo Nordisk (Bagsvaerd, Denmark)

Trypan blue 0.4% solution Sigma-Aldrich (St. Louis, MO, US)

DMSO

L-glutamine

BSA (essentially fatty-acid free)

L-carnitine

D-glucose

Oleic acid (OA, 18:1, n-9)

HEPES

Glycogen

β-mercaptoethanol

Primers for PyroMark PCR and pyrosequencing

96-well Corning® CellBIND® tissue culture plates Corning (Schiphol-Rijk, theNetherlands)

VWR®Grade 703 Blotting Paper VWR (Poole, UK)

[1-14C]oleic acid (2.083 GBq/mmol) PerkinElmer NEN® (Boston, MA, US)

D-[14C(U)]glucose (9.25 GBq/mmol)

OptiPhase Supermix PerkinElmer (Shelton, CT, US)

96-well Isoplate®

Unifilter®-96 GF/B

TopSeal®-A transparent film

MultiScreen® HTS hydrophobic filter plates with high-proteinbinding Immobilon-P membrane

Millipore (Billerica, MA, US)

GelRed™ Nucleic Acid Gel Stain 10000X in water Biotium (Hayward, CA, US)

Clarity™Western ECL Substrate BioRad (Copenhagen, Denmark)

Tris/glycine buffer

Tris/glycine/SDS buffer

SDS

Tween 20

Bromophenol blue

Goat Anti-Rabbit IgG (H+L)-HRP Conjugate (#170–6515)

Goat Anti-Mouse IgG (H+L)-HRP Conjugate (#170–6516)

Mini-Protean® TGX™ gels (4–20%)

Bio-Rad Protein Assay Dye Reagent Concentrate

(Continued )

Changedmetabolism in myotubes from overweight post-training

PLOSONE | https://doi.org/10.1371/journal.pone.0175441 April 12, 2017 3 / 24

Skeletal Muscles, Myokines and Glucose Metabolism (MyoGlu) [26]. The study adhered to the

Declaration of Helsinki, and it was registered with the US National Library of Medicine Clini-

cal Trials registry (NCT01803568).

Donor characteristicsThe biopsies were obtained from 18 volunteer men before and after participating in a 12-week

exercise intervention program at the Norwegian School of Sports Sciences, Oslo, Norway. The

biopsies were taken 2 hours after an acute exercise test [26]. To take part in the study the par-

ticipants had to be sedentary men (not regularly exercising more than once a week), 40 to 62

years old, non-smokers and of Nordic ethnicity. Blood samples were analyzed at Oslo Univer-

sity Hospital during clamp measurements or at Furst Laboratories (Oslo, Norway). Prior to a

euglycemic hyperinsulinemic clamp, body composition by bioelectric impedance analysis was

performed with Tanita Body Composition Analyzer BC-418 MA. Both the clamp and bioim-

pedance measurements were performed under strict criteria, e.g. fasting from the night before,

no alcohol or exercise the last 48 hours and empty bladder before bioimpedance analysis.

Table 1. (Continued)

Material Producer

Glycerol Merck (Darmstadt, Germany)

Tris-HCl

Amersham™ Protran™ Premium 0.45 μmNC NitrocelluloseBlotting Membrane

Amersham™ (GE Healthcare,Esbjerg, Denmark)

Antibodies against human total and phosphorylated Akt at Ser473(#9272 and #9271S, respectively)

Cell Signaling Technology Inc.(Beverly, MA, US)

Antibodies against total and phosphorylated TBC1D4 at Thr642(#2670 and #4288, respectively)

Antibodies against total and phosphorylated AMPKα at Thr172(#2531 and #2532, respectively)

Antibody against total IRS1 (#3407)

Antibody against MHCIIa (#3403S)

Antibody against α-tubulin (#2144)

Antibody against MHCI (#MAB1628) Millipore (Temecula, CA, US)

Antibodies against human total OXPHOS (#110411) Abcam (Cambridge, UK)

RNeasy Mini Kit QIAGEN (Venlo, the Netherlands)

DNeasy Blood & Tissue Kit

EpiTect Fast DNA Bisulfite Conversion Kit

PyroMark® PCR Kit

PyroMark®Q24 Advanced CpG Reagents

PyroMark®Q24 Plate

PyroMark®Wash Buffer

PyroMark® Denaturation Buffer

PyroMark®Q24 Cartridge

Streptavidin Sepharose® High Performance beads GE Healthcare Life Sciences (LittleChalfont, UK)

TaqMan reverse transcription kit reagents Applied Biosystems (Warrington, UK)

MicroAmp®Optical 96-well Reaction Plate

MicroAmp®Optical Adhesive Film

High-Capacity cDNA Reverse Transcription Kit

Power SYBR®Green PCRMaster Mix

https://doi.org/10.1371/journal.pone.0175441.t001

Changedmetabolism in myotubes from overweight post-training

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The group was further divided in two groups, normal weight and overweight, i.e. below and

above the World Health Organization’s lower limit for overweight (BMI 25 kg/m2), respec-

tively, for all analyses except glycogen synthesis and DNAmethylation experiments where

only a subset of the donors were examined (n< 3 in the normal weight group).

Exercise trainingThe training program was performed at the Norwegian School of Sport Sciences. Each partici-

pant exercised 4 times per week for 12 weeks, both endurance sessions twice weekly and

strength training sessions twice weekly. Endurance sessions consisted of interval-based

cycling, and strength training sessions consisted of 3 sets of 8 exercises (leg press, arm press,

chest press, cable pull-down, leg curls, crunches, seated rowing, and a back exercise). All ses-

sions were supervised by one instructor for two participants. Each session, whether endurance

or strength training, lasted about 60 min, excluding 10–20 min aerobic warm-up. The endur-

ance exercise was performed with two different intervals; one of the sessions was performed at

7 min intervals, whereas the other session was performed at 2 min intervals. Compliance to the

exercise intervention was equally good in the two BMI groups [26].

Maximal strength was tested before and after the exercise intervention in maximal leg

press, cable pull-down, and breast press, whereas endurance capacity before and after the exer-

cise intervention was evaluated as maximal oxygen uptake (VO2max) after 45 min cycling at

70% of estimated VO2max. Each participant followed a standardized warm-up before testing.

Dietary intakes were registered by a food frequency questionnaire [27] before and after the

exercise intervention. There was no significant change in intake of energy-providing nutrients

during the study [28].

Culturing of human myotubesMultinucleated human myotubes were established by activation and proliferation of satellite

cells isolated frommusculus vastus lateralis from 7 sedentary normal weight men and from

11 sedentary overweight men. This was based on the method of Henry et al. [29] and modi-

fied according to Gaster et al. [30, 31]. For proliferation of myoblasts a DMEM-Glutamax™(5.5 mmol/l glucose) medium supplemented with 2% FBS and 2% Ultroser G were used. At

approximately 80% confluence the medium was changed to DMEM-Glutamax™ (5.5 mmol/l

glucose) supplemented with 2% FBS and 25 pmol/l insulin to initiate differentiation into

multinucleated myotubes. The cells were allowed to differentiate for 7 days; no difference in

cell differentiation could be detected based on protein expressions of MHCI and MHCIIa

(S1 Fig), and by visual examination in the microscope. During the culturing process the

muscle cells were incubated in a humidified 5% CO2 atmosphere at 37˚C, and medium was

changed every 2–3 days. Experiments were performed on cells from passage number 2 to 4.

For each experiment and within each donor, i.e. before and after exercise, the passage num-

ber remained constant. Isolation of satellite cells from all biopsies was performed at the same

location and by the same trained researchers. Skeletal muscle cultures have previously been

checked for the adipocyte marker fatty acid binding protein (FABP) 4 to ensure a homoge-

nous skeletal muscle cell-population. All cell cultures were visually checked for fibroblast

content throughout proliferation.

Fatty acid and glucose metabolismSkeletal muscle cells (7000 cells/well) were cultured on 96-well CellBIND1 microplates.

[1-14C]oleic acid (18.5 kBq/ml), 20, 100 or 400 μmol/l, or D-[14C(U)]glucose (21.46 kBq/ml),

200 μmol/l, were given during 4 h CO2 trapping as previously described [32]. In brief, a

Changedmetabolism in myotubes from overweight post-training

PLOSONE | https://doi.org/10.1371/journal.pone.0175441 April 12, 2017 5 / 24

96-well UniFilter1-96 GF/B microplate was mounted on top of the CellBIND1 plate and

CO2 production was measured in DPBS medium with 10 mmol/l HEPES and 1 mmol/l L-

carnitine adjusted to pH 7.2–7.3. CO2 production and cell-associated (CA) radioactivity

were assessed using a 2450 MicroBeta2 scintillation counter (PerkinElmer). The sum of14CO2 and remaining CA radioactivity was taken as a measurement of total cellular uptake

of substrate: CO2+CA. Fractional complete oxidation was calculated as: CO2

CO2þCA. Fractional

oxidation gives a picture of what proportion of the substrate taken up that is oxidized and

may or may not correlate to oxidation calculated per amount protein (or cells), depending

on the regulation of the different processes: uptake and oxidation. Thus, an increased frac-

tional oxidation indicates that substrate oxidation is increased relative to the substrate

uptake. Protein levels in the lysate were measured by the Bio-Rad protein assay using a VIC-

TOR™ X4 Multilabel Plate Reader (PerkinElmer).

Determination of lipid accumulationTo study whether an alteration of the radiolabeled oleic acid occurs and if it is incorporated

into complex lipids within the myotubes, lipid filtration was performed. Lysate from the fatty

acid oxidation assays were filtrated through hydrophobic MultiScreen1 HTS filter plates. The

total amount of complex lipids in the cell lysates was determined by liquid scintillation. Lipid

filtration has previously been evaluated against thin layer chromatography and found equal in

describing levels of total complex lipids in a cell lysate [33].

Glycogen synthesisMyotubes were exposed to serum-free DMEM supplemented with [14C(U)]glucose (18.5 kBq/

ml, 0.17 mmol/l) and 0.5 mmol/l unlabeled glucose, in presence or absence of 100 nmol/l

insulin (Actrapid1 Penfill 100 IE/ml) for 3 h to measure glycogen synthesis. In preliminary

unpublished studies, we have seen a defective insulin-stimulated glycogen synthesis at all

concentrations of insulin, ranging from 1 nmol/l to 100 nmol/l. Thus, we decided to use 100

nmol/l insulin to reach maximal insulin stimulation in all experiments. The cells were washed

twice with PBS and harvested in 1 mol/l KOH. Protein content was determined by use of the

Pierce BCA Protein Assay Kit, before 20 mg/ml glycogen and more KOH (final concentration

4 mol/l) were added to the samples. Then, [14C(U)]glucose incorporated into glycogen was

measured as previously described [34].

ImmunoblottingMyotubes were incubated with or without 100 nmol/l insulin for 15 min before the cells were

harvested in Laemmli buffer (0.5 mol/l Tris-HCl, 10% SDS, 20% glycerol, 10% -mercap-

toethanol, and 5% bromophenol blue). The proteins were electrophoretically separated on

4–20%Mini-Protean1 TGX™ gels with Tris/glycine buffer (pH 8.3) followed by blotting to

nitrocellulose membrane and incubation with antibodies for total Akt kinase and Akt phos-

phorylated at Ser473, total insulin receptor substrate (IRS) 1 and IRS1 phosphorylated at

Tyr612, total TBC1 domain family member 4 (TBC1D4, also known as Akt substrate of 160

kDa, AS160) and TBC1D4 phosphorylated at Thr642, total AMP-activated protein kinase

(AMPK) and AMPK phosphorylated at Thr172, MHCI, MHCIIa, total oxidative phosphoryla-

tion (OXPHOS) complexes, and -tubulin. Immunoreactive bands were visualized with

enhanced chemiluminescence (Chemidoc XRS, BioRad, Copenhagen, Denmark) and quanti-

fied with Image Lab (version 4.0) software. Myotubes from 10 donors were used for the

pTBC1D4/total TBC1D4, MHCI, MHCIIa, and OXPHOS analyses, whereas myotubes from 9

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donors were used for the pAkt/total Akt, pIRS1/total IRS1 and pAMPK /total AMPK analy-

ses. All samples were derived at the same time and processed in parallel. Expression levels were

normalized to one sample used as loading control. Expressions of MHCI, MHCIIa, OXPHOS

complex V, and total IRS1 were further normalized to the endogenous control -tubulin.

RNA isolation and analysis of gene expression by qPCRTotal RNA was isolated from myotubes using RNeasy Mini Kit according to the supplier´s

protocol. RNA was reversely transcribed with a High-Capacity cDNA Reverse Transcription

Kit and TaqMan Reverse Transcription Reagents using a PerkinElmer 2720 Thermal Cycler

(25˚C for 10 min, 37˚C for 80 min, 85˚C for 5 min). Primers were designed using Primer

Express1 (Applied Biosystems). qPCR was performed using a StepOnePlus Real-Time PCR

system (Applied Biosystems). Target genes were quantified in duplicates carried out in a 25 μlreaction volume according to the supplier´s protocol. All assays were run for 44 cycles (95˚C

for 15 s followed by 60˚C for 60 s). Expression levels were normalized to the average of the

housekeeping gene GAPDH (acc.no. NM002046). The housekeeping gene large ribosomal pro-

tein P0 (RPLP0, acc.no. M17885) was also analyzed; there were no differences between nor-

malizing for GAPDH or RPLP0. The following forward and reverse primers were used at

concentration of 30 μmol/l, GAPDH; RPLP0; pyruvate dehydrogenase kinase, isoenzyme 4

(PDK4, acc.no. BC040239); angiopoietin-like 4 (ANGPTL4, acc.no. NM139314); carnitine pal-

mitoyltransferase 1A (CPT1A, acc.no. L39211); perilipin 2 (PLIN2, acc.no. NM001122); fatty

acid translocase (CD36, acc.no. L06850); cytochrome c-1 (CYC1, acc.no. NM001916); peroxi-

some proliferator-activated receptor gamma, coactivator 1 alpha (PPARGC1A, acc.no.NM013261.3); peroxisome proliferator-activated receptor delta (PPARD, acc.no. BC002715);IRS1 (acc.no. NM_005544.2).

DNAmethylation measurementgDNA was extracted from myotubes using DNeasy Blood & Tissue Kit. A concentration of

�20 ng/μl was used. The gDNA was bisulfite treated using EpiTect Fast DNA Bisulfite Kit.

Forward, reverse and sequencing primers for PDK4, PPARGC1A, PPARD, mitochondrial

transcription factor A (TFAM), and IRS1were designed using PyroMark AssayDesign 2.0

(QIAGEN, Venlo, the Netherlands). We tested 3 CpGs in the promoter region of PKD4(chr7:95,226,252–95,226,322), 2 CpGs in the promoter of PPARGC1A (chr4:23,891,715–

23,891,726), 4 CpGs in the promoter of PPARD (chr6:35,309,819–35,309,931), 8 CpGs in the

promoter of TFAM (chr10:60,144,788–60,144,828), and 3 CpGs in the first exon of IRS1(chr2:227,661,201–227,661,293). For each primer-set, bisulfite-treated DNA was amplified by

PCR using PyroMark PCR Kit and MyCycler Thermal Cycler (BioRad, Copenhagen, Den-

mark). The reaction was visualized by gel electrophoresis to check if it was the right product

according to the size and if it was well amplified with no secondary product. The reaction was

optimized if necessary. DNAmethylation for each region of interest was measured by pyrose-

quencing using QIAGEN PyroMark Q24.

Presentation of data and statisticsData are presented as means ± SEM. The value n represents the number of different donors;

each in vitro experiment with at least duplicate observations. For immunoblotting, results for

normal weight group before exercise was set to 100%, and for experiments with insulin-stimu-

lation, basal before exercise was set to 100%. Statistical analyses were performed using Graph-

Pad Prism 6.0c for Mac (GraphPad Software, Inc., La Jolla, CA, US) or SPSS version 22 (IBM1

SPSS1 Statistics for Macintosh, Armonk, NY, US). Linear mixed-model analysis was used to

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compare differences between conditions with within-donor variation and simultaneously

compare differences between groups with between-donor variation. The linear mixed-model

analysis includes all observations in the statistical analyses and takes into account that not all

observations are independent. Paired t test was used within groups, whereas unpaired t test

with equal standard deviation was used to evaluate effects between groups. Correlation studies

were performed with Pearson’s test and are presented as Pearson’s correlation coefficient (r).

A p-value< 0.05 was considered significant.

Results

Donor characteristicsDonor characteristics pre- and post-training are presented in Table 2. After 12 weeks of exer-

cise both normal weight and overweight donor groups significantly increased maximal

strength and insulin sensitivity measured as the glucose infusion rate (GIR). Only the normal

weight group significantly reduced percentage body fat (overweight: p = 0.07) after the exercise

intervention, whereas only the overweight group significantly increased VO2max (normal

weight: p = 0.053) and reduced body weight and BMI. Visceral fat area also tended to be

smaller after the exercise intervention in the overweight group (p = 0.07).

As expected, there were significant differences between the normal weight group and

the overweight group both pre- and post-training for body weight, BMI, waist-hip ratio,

Table 2. Clinical and biochemical variables in normal weight (BMI 25 kg/m2) and overweight men (BMI� 25 kg/m2) at baseline (pre-training) andafter 12 weeks of extensive endurance and strength training (post-training).

Pre-training alldonors

Post-training alldonors

Pre-training normalweight

Post-training normalweight

Pre-trainingoverweight

Post-trainingoverweight

n 18 18 (17†) 7 7 (6†) 11 11

Age, y 50.4 ± 1.6 - 48.0 ± 2.8 - 51.9 ± 1.8 -

Body weight, kg 88.6 ± 3.2 87.1 ± 3.0* 78.4 ± 3.2 78.1 ± 3.3 95.1 ± 3.6# 92.8 ± 3.5*#

BMI, kg/m2 27.0 ± 0.9 26.6 ± 0.8* 23.3 ± 0.7 23.3 ± 0.6 29.4 ± 0.7# 28.7 ± 0.7*#

Waist-hip ratio 0.92 ± 0.01 0.91 ± 0.01 0.88 ± 0.01 0.88 ± 0.01 0.95 ± 0.01# 0.94 ± 0.01#

Fat mass, % 23.2 ± 1.2 22.1 ± 1.2* 18.0 ± 1.0 16.8 ± 0.8* 26.5 ± 0.9# 25.4 ± 1.0#

Visceral fat area,cm2

138.0 ± 12.5 118.4 ± 9.7 101.6 ± 15.0 90.5 ± 5.9 161.1 ± 14.4# 136.1 ± 13.0#

Fasting glucose,mmol/l

5.6 ± 0.1 5.7 ± 0.1* 5.3 ± 0.2 5.5 ± 0.2 5.7 ± 0.1 5.9 ± 0.1

GIR, mg/kg/min 6.0 ± 0.6 7.8 ± 0.8* 7.7 ± 0.7 9.4 ± 0.9* 5.0 ± 0.7# 6.7 ± 1.0*

VO2max, ml/kg/min

39.2 ± 1.1 44.1 ± 1.5* 42.5 ± 0.9 47.1 ± 2.2 37.1 ± 1.5# 42.3 ± 1.8*

Chest pressmax,kg

67.5 ± 3.6 77.6 ± 4.0*† 61.8 ± 4.7 71.7 ± 6.4*† 71.1 ± 5.0 80.9 ± 5.1*

Cable pull-downmax, kg

72.5 ± 3.3 82.4 ± 3.0*† 68.9 ± 3.7 78.3 ± 3.4*† 74.8 ± 4.9 84.5 ± 4.4*

Leg pressmax, kg 224.9 ± 10.4 249.3 ± 11.7* 192.9 ± 14.6 209.3 ± 13.9* 245.2 ± 10.5# 274.8 ± 11.7*#

Glucose infusion rate (GIR) measurements were performed with euglycemic hyperinsulinemic clamp analysis; visceral fat area and skeletal muscle mass

were based on bioelectrical impedance analysis with Tanita. Values are given as means ± SEM (n = 7 in the normal weight group and n = 11 in the

overweight group).

*Statistically significant vs. pre-training (p < 0.05, paired t test).#Statistically significant vs. normal weight (p < 0.05, unpaired t test with equal SD).†Missing data from one normal weight participant for the post-exercise tests in the two arm exercises (chest press and cable pull-down) due to an arm injury.

BMI, body mass index; VO2max, maximal oxygen uptake.

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percentage body fat, visceral fat area, and maximal strength in leg press (Table 2). GIR and

VO2max only differed pre-training between the groups.

Increased fatty acid and glucose metabolism in cultured humanmyotubes after 12 weeks of exerciseFatty acid metabolism in myotubes obtained from biopsies before and after 12 weeks of exer-

cise is presented in Fig 1. Results for all participants combined (n = 18) are shown in Fig 1A–

1D, and separated by BMI in Fig 1E–1H. The overall statistically significant exercise-induced

increase in total cellular oleic acid uptake was 30%, in oleic acid oxidation 46%, in fractional

oxidation 45%, and in lipid accumulation of oleic acid 34% (Fig 1D). When study group was

Fig 1. Effects of 12 weeks of exercise onmyotube fatty acid metabolism. Satellite cells isolated from biopsies fromm. vastus lateralis before andafter 12 weeks of exercise were cultured and differentiated to myotubes. Oxidation, cell-associated (CA) radioactivity and lipid accumulation of [14C]oleic acid were measured, and total cellular uptake (CO2+CA), oxidation (CO2), fractional oxidation ( CO2

CO2þCA), and lipid accumulation were determined.(A) Lipid accumulation presented as cpm/μg protein. Values are presented as means ± SEM for all participants combined (n = 18). (B)Oleic acidoxidation and total cellular uptake presented as nmol/mg protein. Values are presented as means ± SEM for all participants combined (n = 18). (C)Fractional oleic acid oxidation. Values are presented as means ± SEM for all participants combined (n = 18). (D) Fatty acid metabolism relative tobefore exercise. Values are presented as means ± SEM for all participants combined (n = 18). (E) Lipid accumulation presented as cpm/μg protein instudy group when separated by BMI. Values are presented as means ± SEM (n = 7 in the normal weight group and n = 11 in the overweight group). (F)Oleic acid oxidation and total cellular uptake presented as nmol/mg protein in study group when separated by BMI. Values are presented asmeans ± SEM (n = 7 in the normal weight group and n = 11 in the overweight group). (G) Fractional oleic acid oxidation in absolute values in studygroup when separated by BMI. Values are presented as means ± SEM (n = 7 in the normal weight group and n = 11 in the overweight group). (H) Fattyacid metabolism relative to before exercise in study group separated by BMI. Values are presented as means ± SEM (n = 7 in the normal weight groupand n = 11 in the overweight group). *Statistically significant vs. before exercise (p < 0.05, linear mixed-model analysis, SPSS). Statistically significantvs. normal weight group after exercise (p < 0.05, linear mixed-model analysis, SPSS). $Statistically significant vs. normal weight group (p < 0.05, linearmixed-model analysis, SPSS).

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separated by BMI, myotubes from the overweight group showed exercise-induced increase in

oleic acid oxidation, fractional oxidation and lipid accumulation by 71%, 70%, and 51%,

respectively, after exercise (Fig 1H). Total cellular oleic acid uptake also tended to be increased

after the exercise intervention in the overweight group (p = 0.08, Fig 1H). There were no statis-

tically significant exercise-induced changes in oleic acid metabolism in myotubes from the

normal weight group (Fig 1H). In myotubes established before exercise, lipid accumulation

was lower in the overweight group compared to the normal weight group (Fig 1E). Pre-train-

ing lipid accumulation correlated significantly positively with GIR (r = 0.47, and p = 0.05) and

negatively with fasting glucose (r = -0.53 and p = 0.03), suggesting a relationship between lipid

accumulation and insulin sensitivity (data not shown).

Glucose metabolism in myotubes obtained from biopsies before and after 12 weeks of exer-

cise is presented in Fig 2. Results for all participants combined (n = 18) are shown in Fig 2A–

2C, and separated by BMI in Fig 2D–2F. We observed a 14% exercise-induced increase in frac-

tional oxidation of glucose, but no exercise-induced effect on total cellular glucose uptake or

oxidation for all participants (Fig 2C). When study group was separated by BMI, a significant

exercise-induced increase in fractional glucose oxidation was observed in myotubes from the

overweight group (Fig 2F), while total cellular glucose uptake and oxidation tended to be

higher in the normal weight group compared to the overweight group after exercise (p = 0.07

and p = 0.06, respectively, Fig 2F). Furthermore, we found a significant correlation between

exercise-induced improvement in maximal leg press and exercise-induced increase in glucose

oxidation after exercise (Fig 2G, full line, r = 0.52, and p = 0.03), indicating a relationship

between in vivo and in vitro findings that is not visible when only comparing before and after

exercise. This correlation was also significant for the overweight group (Fig 2G, stapled line,

r = 0.68, and p = 0.02). In myotubes established before exercise, oxidation and uptake of glu-

cose were increased in the overweight group compared to the normal weight group (Fig 2D).

No changes in AMPK phosphorylation in cultured human myotubes after12 weeks of exerciseAMPK plays an important role in cellular energy homeostasis, acting as a sensor of AMP/ATP

or ADP/ATP ratios and thus cell energy level [35, 36]. To study whether AMPK could be a

part of the observed exercise-induced changes on energy metabolism in vitro cultured myo-

tubes was assessed by AMPK (Thr172) phosphorylation (Fig 3). No changes in pAMPK /

total AMPK ratio (Fig 3B) were observed in cells after exercise, nor between the two BMI

groups (Fig 3C).

No changes in mitochondria-related genes and proteins in culturedhuman myotubes after 12 weeks of exerciseTo study possible exercise-induced changes in oxidative capacity in the mitochondria we stud-

ied genes and proteins related to mitochondria (Fig 4). PPARGC1A codes for the master regu-

lator of mitochondrial biogenesis PGC-1 [37–39], PDK4, CPT1A and CYC1 are genes codingfor proteins involved in metabolism in mitochondria [40–43], while TFAM codes for a mito-

chondrial transcription factor [44]. There were no significant exercise-induced changes in

PPARGC1A, PDK4 (p = 0.08), CPT1A, or CYC1 for all participants combined (Fig 4A), nor

when separated by BMI (Fig 4B). However, we observed a significant correlation between exer-

cise-induced reduction in visceral fat area in vivo and increased mRNA expression of PDK4 inthe myotubes (Fig 4C, full line, p = 0.02, r = -0.54). This correlation was also significant for the

overweight group (Fig 4C, stapled line, p = 0.04, r = -0.63). We also monitored DNAmethyla-

tion of PPARGC1A, PDK4 and TFAM genes in myotubes from a small subset of donors (n = 6,

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Fig 2. Effects of 12 weeks of exercise onmyotube glucosemetabolism. Satellite cells isolated from biopsies fromm. vastus lateralis before andafter 12 weeks of exercise were cultured and differentiated to myotubes. Oxidation and cell-associated (CA) radioactivity of [14C]glucose weremeasured, and total cellular uptake (CO2+CA), oxidation (CO2), and fractional oxidation ( CO2

CO2þCA) were determined. (A)Glucose oxidation and totalcellular uptake presented as nmol/mg protein. Values are presented as means ± SEM for all participants combined (n = 18). (B) Fractional glucoseoxidation. Values are presented as means ± SEM for all participants combined (n = 18). (C)Glucose metabolism relative to before exercise. Values arepresented as means ± SEM for all participants combined (n = 18). *Statistically significant vs. before exercise (p < 0.05, linear mixed-model analysis,SPSS). (D)Glucose oxidation and total cellular uptake presented as nmol/mg protein in study group when separated by BMI. Values are presented asmeans ± SEM (n = 7 in the normal weight group and n = 11 in the overweight group). (E) Fractional glucose oxidation in absolute values in study groupwhen separated by BMI. Values are presented as means ± SEM (n = 7 in the normal weight group and n = 11 in the overweight group). (F)Glucosemetabolism relative to before exercise in study group when separated by BMI. Values are presented as means ± SEM (n = 7 in the normal weight

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combination of both donor groups) before and after exercise (Fig 4D). Overall, there were no

differences in CpG methylation within the regions we tested in PPARGC1A, PDK4 or TFAM.

However, 1 out of 8 CpGs tested in the TFAM-promoter was hypomethylated after exercise

compared to before exercise (34% decrease, data not shown). Furthermore, we measured

group and n = 11 in the overweight group). *Statistically significant vs. before exercise (p < 0.05, linear mixed-model analysis, SPSS). $Statisticallysignificant vs. normal weight group (p < 0.05, linear mixed-model analysis, SPSS). (G) Pearson’s test of correlation between exercise-induced changesin leg press and glucose oxidation in myotubes. Δ = after exercise–before exercise. Full line represents the regression line for all donors (n = 18,Pearson’s correlation coefficient, r = 0.52, and p = 0.03), whereas stapled line represents the regression line for the overweight group (n = 11,Pearson’s correlation coefficient, r = 0.68, and p = 0.02).

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Fig 3. Effects of 12 weeks of exercise onmyotube AMPK phosphorylation. Satellite cells isolated frombiopsies fromm. vastus lateralis before and after 12 weeks of exercise were cultured and differentiated tomyotubes. (A-C) AMPKα phosphorylation by immunoblotting. Protein was isolated and total AMPKα andpAMPKα expressions assessed by immunoblotting. A, one representative immunoblot. Bands selected fromone membrane have been spliced together to show only relevant samples, as indicated by lines separatingthe spliced blots. B, quantified immunoblots for participants combined (n = 9) relative to before exercise. C,quantified immunoblots for study group when separated by BMI relative to normal weight before exercise(n = 5 in the normal weight group and n = 4 in the overweight group). Values are presented as means ± SEM.All samples were derived at the same time and processed in parallel.

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protein expression of the mitochondrial oxidative phosphorylation (OXPHOS) complexes

(Fig 4E–4G), detected with an antibody cocktail recognizing complex I subunit NDUFB8,

complex II subunit 30 kDa, complex III subunit Core 2, complex IV subunit II, and ATP

synthase subunit alpha. Only complex V was quantifiable across the membranes. No clear

exercise-induced changes were observed for participants combined (Fig 4F), nor when sepa-

rated by BMI (Fig 4G).

No change in genes related to lipid metabolism after 12 weeks ofexercise in cultured human myotubesSome genes related to lipid metabolism were also examined to further probe mechanisms

behind the exercise-induced metabolic changes observed in vitro. mRNA of PLIN2, involvedin coating of lipid droplets and thus lipid accumulation [45, 46], was not significantly different

after the exercise intervention for all participants (Fig 5A) or when the study group was sepa-

rated by BMI (Fig 5B). Neither was mRNA of CD36, an important transporter of fatty acids

across the plasma membrane [47, 48] (Fig 5A and 5B). We have previously shown that

Fig 4. Effects of 12 weeks of exercise onmitochondria-related genes and proteins. Satellite cells isolated from biopsies fromm. vastus lateralisbefore and after 12 weeks of exercise were cultured and differentiated to myotubes. (A)mRNA expression of PPARGC1A, PDK4,CPT1A, andCYC1after exercise relative to before exercise. mRNAwas isolated and expression assessed by qPCR. All values were corrected for the housekeepingcontrolGAPDH, and presented as means ± SEM for all participants combined (n = 18). (B)mRNA expression of PPARGC1A, PDK4,CPT1A, andCYC1 after exercise relative to before exercise in study group when separated by BMI. mRNA was isolated and expression assessed by qPCR. Allvalues were corrected for the housekeeping controlGAPDH, and presented as means ± SEM (n = 7 in the normal weight group and n = 11 in theoverweight group). (C) Pearson’s test of correlation was performed between exercise-induced changes in visceral fat area and mRNA expression ofPDK4 in myotubes. Δ = after exercise–before exercise. Full line represents the regression line for all donors (n = 18, Pearson’s correlation coefficient, r= -0.54, and p = 0.02), whereas stapled line represents the regression line for the overweight group (n = 11, Pearson’s correlation coefficient, r = -0.63,and p = 0.04). (D)DNAmethylation of PPARGC1A, PDK4 and TFAM after exercise relative to before exercise. gDNA was isolated and bisulfitetreated, and methylation assessed by immunoblotting. Values are presented as means ± SEM (n = 6). (E-G)OXPHOS complexes by immunoblotting.Protein was isolated and OXPHOS complexes assessed by immunoblotting. E, one representative immunoblot. F, quantified immunoblots of complexV for participants combined. All values were corrected for the housekeeping control α-tubulin, and presented as means ± SEM (n = 10). G, quantifiedimmunoblots of complex V in study group when separated by BMI. All values were corrected for the housekeeping control α-tubulin, and presented asmeans ± SEM (n = 5 in each group).

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Fig 5. Effects of 12 weeks of exercise onmyotube expression of lipid metabolism associated genes.Satellite cells isolated from biopsies fromm. vastus lateralis before and after 12 weeks of exercise werecultured and differentiated to myotubes. mRNAwas isolated and expression assessed by qPCR. (A)mRNAexpression after exercise relative to before exercise for all participants combined. All values were correctedfor the housekeeping controlGAPDH, and presented as means ± SEM (n = 18). (B)mRNA expression afterexercise relative to before exercise for study group when separated by BMI. All values were corrected for the

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activation of PPAR increased lipid oxidation in human skeletal muscle cells [49]. Gene

expression of PPARD or the PPAR-target gene ANGPTL4 [50–52] also showed no exercise-induced changes (Fig 5A), nor when study group was separated by BMI (Fig 5B). We also

monitored DNAmethylation of PPARD in the small subset of donors (n = 6, combination of

both donor groups) before and after exercise, but no differences in CpG methylation within

the region we tested were observed (data not shown).

No changes in insulin response in cultured human myotubes after 12weeks of exerciseBoth donor groups experienced increased GIR after exercise (Table 2). To examine whether

the improved insulin sensitivity in vivowas mirrored in vitro in the myotubes, the response to

100 nmol/l insulin was assessed by measurement of Akt (Ser473) phosphorylation, TBC1D4

(Thr642) phosphorylation, IRS1 (Tyr612) phosphorylation, and glycogen synthesis (Fig 6). No

housekeeping controlGAPDH, and presented as means ± SEM (n = 7 in the normal weight group and n = 11in the overweight group).

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Fig 6. Effects of 12 weeks of exercise onmyotube Akt phosphorylation, TBC1D4 phosphorylation and glycogen synthesis with or without100 nmol/l insulin. Satellite cells isolated from biopsies fromm. vastus lateralis before and after 12 weeks of exercise were cultured and differentiatedto myotubes. (A-C) Akt phosphorylation by immunoblotting. Protein was isolated and total Akt and pAkt expressions assessed by immunoblotting. A,one representative immunoblot. B, quantified immunoblots relative to basal before exercise for participants combined. Values are presented asmeans ± SEM (n = 9). C, quantified immunoblots relative to basal before exercise for study group when separated by BMI (n = 4 in the normal weightgroup and n = 5 in the overweight group). (A, D and E) TBC1D4 phosphorylation by immunoblotting. Protein was isolated and total TBC1D4 andpTBC1D4 expressions assessed by immunoblotting. A, one representative immunoblot. D, quantified immunoblots relative to basal before exercise forparticipants combined. Values are presented as means ± SEM (n = 10). E, quantified immunoblots relative to basal before exercise for study groupwhen separated by BMI (n = 5 in both groups). All samples were derived at the same time and processed in parallel. (F)Glycogen synthesis relative tobasal before exercise. Values are presented as means ± SEM (n = 5). Absolute values (range) representing 100%: Basal glycogen synthesis 3.9–15.4nmol/mg protein. #Statistically significant vs. basal before exercise (p < 0.05, paired t test).

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changes in the basal level of pAkt/total Akt ratio or pTBC1D4/total TBC1D4 ratio were

observed in cells after exercise. As expected, insulin significantly increased the pAkt/total Akt

ratio in myotubes from both groups before and after exercise (Fig 6A and 6B), whereas there

were no significant effect of insulin on pTBC1D4/total TBC1D4 ratio (Fig 6A and 6D). When

the study group was separated by BMI, no significant differences in basal or insulin-stimulated

levels of pAkt/total Akt ratio or pTBC1D4/total TBC1D4 ratio were observed (Fig 6C and 6E,

respectively). No changes in the basal level or insulin-stimulated levels of pIRS1/total IRS1

ratio were observed (data not shown). Furthermore, no changes in the basal level of glycogen

synthesis were observed in myotubes, and insulin significantly increased glycogen synthesis by

about 1.5-fold both before and after exercise (Fig 6F). Thus, there was no exercise-effect on

insulin-stimulated Akt phosphorylation, TBC1D4 phosphorylation or glycogen synthesis.

Decreased IRS1mRNA expression and increased DNAmethylationwithin first exon region of IRS1 after 12 weeks of exercise in culturedhuman myotubesTo further study the insulin signaling pathway, we also measured mRNA expression, DNA

methylation and protein expression of IRS1 (Fig 7). We found that the mRNA expression of

IRS1was significantly decreased by 31% after exercise (n = 8, Fig 7A), which was only signifi-

cant in myotubes from the normal weight group upon separation by BMI (n = 3 in the normal

weight group and n = 5 in the overweight group, Fig 7B). Furthermore, DNA methylation of 1

out of 3 CpGs tested within the first exon region of IRS1was significantly increased by 23%

(n = 6, Fig 7C). There were no exercise-induced changes in protein expression of IRS1 detected

with immunoblotting (n = 9, Fig 7E), nor when study group was separated by BMI (n = 5 in

the normal weight group and n = 4 in the overweight group, Fig 7F).

DiscussionWe show that 12 weeks of exercise alters metabolism and gene expression of cultured human

myotubes. Fatty acid metabolism and fractional glucose oxidation were significantly increased

in myotubes established from skeletal muscle isolated from sedentary men after 12 weeks of

exercise. These exercise-induced metabolic changes in fatty acid metabolism in myotubes were

more predominant in cells from overweight subjects. Moreover, we observed a significant

exercise-induced decrease in mRNA expression of IRS1, as well as DNA hypermethylation in

the first exon of IRS1, however not detectable on protein level.

Bourlier et al. showed that cultured myotubes retained the exercise-trained phenotype invitro concerning some aspects of glucose metabolism [16]. Their study involved 8 weeks of aer-

obic exercise intervention and included only obese individuals [16]. In the present study we

examined a broader group of subjects including normal weight and overweight men, a longer

exercise intervention as well as a combination of aerobic and anaerobic exercise, to observe

and possibly explain differences in energy metabolism in cultured myotubes in vitro after thein vivo exercise intervention, and also to explore whether BMI of the subjects affected the

results.

As expected, the exercise intervention significantly increased VO2max (overweight group),

chest press, cable pull-down, and leg press capacity. The exercise intervention also improved

the metabolic health, with a significant increase in GIR, as well as a small, but significant

reduction in BMI. VO2max was not significantly increased in the normal weight group

(p = 0.053) even though they complied to the exercise intervention equally well [26]. The mean

increase was variable between the participants, and combined with the smaller sample size it

may explain the lack of statistical difference.

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With the combination of aerobic and anaerobic exercises and longer intervention we have

several interesting findings with regard to fatty acid metabolism in myotubes established from

biopsies taken before and after 12 weeks of exercise. We observed a significantly increased

oleic acid oxidation, fractional oxidation and lipid accumulation in the cells, statistically signif-

icant only in the overweight group (except total cellular oleic acid uptake).

In our study there are no data on lipid utilization in vivo or ex vivo to directly compare

with in vitro data. However, from the same clinical study muscle lipid content, measured by

Fig 7. Effects of 12 weeks of exercise onmyotube IRS1 gene expression and IRS1 first exon DNAmethylation. (A) IRS1mRNAexpression after exercise relative to before exercise for participants combined. mRNAwas isolated and expression assessed by qPCR. Allvalues were corrected for the housekeeping controlGAPDH, and presented as means ± SEM (n = 8). *Statistically significant vs. beforeexercise (p < 0.05, paired t test). (B) IRS1mRNA expression after exercise relative to before exercise for study group when separated by BMI.mRNA was isolated and expression assessed by qPCR. All values were corrected for the housekeeping controlGAPDH, and presented asmeans ± SEM (n = 3 in the normal weight group and n = 5 in the overweight group). *Statistically significant vs. before exercise (p < 0.05,paired t test). (C) IRS1 first exon DNAmethylation after exercise relative to before exercise. gDNA was isolated and bisulfite treated, andmethylation was assessed by pyrosequencing. Values are presented as means ± SEM (n = 6). *Statistically significant vs. before exercise(p < 0.05, paired t test). (D-F) IRS1 total protein expression. Protein was isolated and total IRS1 expression assessed by immunoblotting. D,one representative immunoblot. Bands selected from one membrane have been spliced together to show only relevant samples, as indicatedby lines separating the spliced blots. E, quantified immunoblots relative to before exercise for participants combined. All values were correctedfor the housekeeping control α-tubulin, and presented as means ± SEM (n = 9). G, quantified immunoblots relative to before exercise for studygroup when separated by BMI. All values were corrected for the housekeeping control α-tubulin, and presented as means ± SEM (n = 5 in thenormal weight group and n = 4 in the overweight group). All samples were derived at the same time and processed in parallel.

https://doi.org/10.1371/journal.pone.0175441.g007

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magnetic resonance spectrometry in vivo and electron microscopy ex vivo, was found to be

significantly reduced after the exercise intervention [26, 28], in line with an increase in lipid

metabolism in vitro.An exercise-induced increase in lipid oxidation in cultured myotubes is also in accordance

with findings from others in skeletal muscle in vivo during and after combined types of exer-

cise [7, 53]. A study by Ramos-Jimenez et al. [8] showed that lipid oxidation was increased in

endurance trained men (athletes trained at a competitive level) compared to untrained men, as

measured by lower respiratory exchange ratio. Increased lipid oxidation after exercise is also

in line with observations from an in vitromodel (electrical pulse stimulation) of myotube exer-

cise [54, 55]. Bourlier et al. did not observe exercise-induced differences in lipid metabolism

in cultured myotubes, however, they hypothesized that longer exercise interventions and/or

interventions including different types of exercise might lead to functional changes in lipid

metabolism [16].

Bourlier et al. [16] reported increased glucose metabolism in myotubes from obese subjects

after an 8-week aerobic exercise intervention. In our study we observed increased fractional

oxidation of glucose, statistically significant only in the overweight group, as well as a signifi-

cant correlation between exercise-induced increased maximal leg press capacity and increased

oxidation of glucose in the cells, indicating a relationship between glucose oxidation and exer-

cise outcome. However, the effects of exercise on glucose metabolism were less pronounced in

our study than described by Bourlier et al. [16], possibly explained by different donor groups

and exercise programs. Increased storage of glycogen is a well-reported physiologic response

to exercise as a mean to increase endurance capacity during submaximal exercise [11, 56], and

Bourlier et al. also reported increased basal glycogen synthesis in myotubes cultured from sat-

ellite cells after exercise in vivo [16]. However, this was not observed in this study, possibly

caused by different study conditions.

In this study we have compared myotubes from normal weight and overweight subjects. In

pre-training myotubes we found increased oxidation and uptake of glucose and lower lipid

accumulation in the overweight group compared to the normal weight group, as well as a pos-

sible association between lipid accumulation in vitro and insulin sensitivity in vivo. Severalprevious studies show no significant donor-related differences i basal glucose oxidation in

myotubes [54, 57–59], however Gaster [17] observed increased glucose oxidation in myotubes

from obese patients with T2D compared to myotubes from lean donors. It was suggested that

under certain conditions metabolism of myotubes from diabetic donors relies more on glucose

oxidation than myotubes from lean donors [17]. We have previously reported lower lipid accu-

mulation in myotubes from obese subjects with T2D compared to myotubes from obese non-

diabetic donors, explained by a reduced capacity for lipid accumulation and increased lipolysis

[60]. Our overweight donors are not diabetic, however this donor group had reduced pre-

training insulin sensitivity and myotubes from this group may resemble cells from T2D donors

in some ways. The donor-dependent differences in glucose metabolism and lipid accumula-

tion found in pre-training myotubes were evened out after exercise, in line with the increased

response to exercise in myotubes from the overweight group.

Satellite cells are usually dormant in vivo until they are challenged with growth or injury

[13], e.g. exercise. We observed changes in energy metabolism in skeletal muscle cells following

exercise intervention, and aimed to determine whether gene or protein expression were coin-

cident with the observed changes in energy metabolism.

Despite the increased fatty acid oxidation, we did not observe any significant exercise-

induced differences in phosphorylation of AMPK , and no changes in mRNA expression

levels of mitochondria-related genes or genes related to fatty acid metabolism. However,

there was a significant correlation between reduced visceral fat area in vivo and higher

Changedmetabolism in myotubes from overweight post-training

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mRNA expression of PDK4 in vitro. PDK4 is involved in phosphorylation and inactivation of

the pyruvate dehydrogenase complex (PDC). Increased expression of PDK4 inhibits PDCand reduces glucose oxidation, which makes PDK4 a major regulatory metabolic enzyme in

skeletal muscle as it is involved in switching from carbohydrate to lipid utilization [41, 61,

62]. Bourlier et al. [16] found a reduced PDK4mRNA expression after exercise in cultured

myotubes, in line with the increased glucose oxidation [16], while we previously have found

that increased lipid oxidation of cultured human myotubes in vitro simultaneously also

increased PDK4 expression [49, 55, 63]. Thus, the correlation between reduced visceral fat

area and increased PDK4 expression may indicate a relationship between lipid metabolism invivo and in vitro.

DNA methylation has been proposed as a molecular mechanism for exercise-mediated

changes in metabolic health [15] and has been associated with transcriptional silencing [64],

possibly by blocking the promoter that activating transcription factors normally bind. In

our study, DNA methylation of the mitochondrial genes TFAM and PDK4 were not changedin myotubes after exercise. This is in contrast to findings ex vivo after acute exercise. Barrèset al. [65] showed that acute exercise increased mRNA expression of PDK4 and PPARGC1Ain skeletal muscle, and that changes in methylation was part of the explanation. However,

we found both hypermethylation of IRS1 and reduction of IRS1mRNA expression in cul-

tured myotubes after 12 weeks of training, whereas protein expression apparently was

unchanged. The functional significance of these findings is unknown and not easy to

explain. Protein expression of IRS1 has previously been shown to be both increased [66]

and decreased [67] in human skeletal muscle after exercise. We have recently shown

enhanced tyrosine phosphorylation of IRS1, concomitant with increased glucose metabo-

lism in cultured myotubes obtained from donors before and after gastric by-pass surgery

[68]. Our study indicates that exercise-induced changes in promoter methylation may be

retained in satellite cells and during transition of these precursor cells to myoblasts and

finally to myotubes, however, at present we cannot explain a possible link between this and

the metabolic changes observed.

Disturbances in energy metabolism of skeletal muscle are associated with metabolic dis-

eases related to insulin resistance [69, 70]. In vivo we found a significant increased GIR after

training i both donor groups, indicating increased insulin sensitivity, while no exercise-

induced changes in in vitro insulin response (i.e. insulin-stimulated Akt phosphorylation,

TBC1D4 phosphorylation or glycogen synthesis) were observed. This could be explained by

sub-optimal experimental conditions (i.e. a maximal insulin stimulation), though we have

previously been able to detect donor-specific differences in insulin-response with the same

experimental setup [20, 60]. We hypothesize therefore that the lack of these effects are a

result of the underlying study in vivo where the two donor groups were quite similar with

regard to insulin sensitivity, and that the difference were too small to be able to detect invitro.

In conclusion, our data show that a combination of aerobic and anaerobic exercise mediates

changes in fatty acid and glucose metabolism in skeletal muscle cells. Thus, certain impacts of

exercise in vivo are retained in myotubes established from satellite cells, and our findings may

indicate that cultured, passaged myoblasts established from these progenitor cells and differen-

tiated into myotubes, can be used as a model system for studying mechanisms related to exer-

cise and metabolic diseases. Furthermore, we observed that the exercise-induced changes were

predominant in the overweight group. Future studies are required to explore whether epige-

netic or other changes can explain this relationship further, and to get a deeper insight into

molecular mechanisms behind changes in energy metabolism in myotubes after an exercise

intervention.

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Supporting informationS1 Fig. No differences in protein expression of differentiation markers. Satellite cells iso-

lated from biopsies fromm. vastus lateralis before and after 12 weeks of exercise were culturedand differentiated to myotubes. (A, B)MHCI expression by immunoblotting. Protein was iso-

lated and MHCI expression assessed by immunoblotting. A, one representative immunoblot.

Bands selected from one membrane have been spliced together to show only relevant samples,

as indicated by lines separating the spliced blots. B, quantified immunoblots for study group

when separated by BMI relative to normal weight before exercise (n = 5 in both groups). (A,

C)MHCIIa expression by immunoblotting. Protein was isolated and MHCIIa expression

assessed by immunoblotting. A, one representative immunoblot. Bands selected from one

membrane have been spliced together to show only relevant samples, as indicated by lines sep-

arating the spliced blots. C, quantified immunoblots for study group when separated by BMI

relative to normal weight before exercise (n = 5 in both groups). All values were corrected for

the housekeeping control -tubulin. Values are presented as means ± SEM. All samples were

derived at the same time and processed in parallel.

(TIF)

AcknowledgmentsThe authors thank the scientific staff at both Oslo and Stockholm groups for scientific

discussions.

Author Contributions

Conceptualization: JL ACR TML CAD KIB KJK EIJ DST HKS HLG JJ GHT.

Formal analysis: JL.

Funding acquisition: JL ACR CAD KIB AK JJ GHT.

Investigation: JL NGL JMM TML YZF CS MGB KJK EIJ DST HKS HLG.

Methodology: JL ACR AK ETK GHT.

Project administration: JL.

Resources: ACR KIB HLG AK JJ GHT.

Supervision: JL ACR ETK GHT.

Validation: JL ACR ETK GHT.

Visualization: JL.

Writing – original draft: JL.

Writing – review & editing: JL ACR JMM CAD KIB HLG AK ETK JJ GHT.

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