protein thiol oxidation and muscle contractile …reversible thiol oxidation of muscle proteins as...

PROTEIN THIOL OXIDATION AND MUSCLE

CONTRACTILE PERFORMANCE

Isaac Lim Gui Jia (PhD)

This thesis is submitted as requirement for the

Doctor of Philosophy at the

University of Western Australia

Janurary 2012

i

EXECUTIVE SUMMARY

There is mounting evidence that reactive oxygen species (ROS) play an important role in

the development of muscle fatigue. Some of the proposed mechanisms involve the

reversible thiol oxidation of muscle proteins as well as ROS-induced protein carbonylation.

Although the functions of several muscle proteins have been reported to be affected by

these processes, it still remains to be determined whether the thiol oxidation levels of

these and other proteins increase with fatiguing muscle stimulation, and whether this is

an important mechanism of ROS-mediated muscle fatigue. It was the primary objective of

this thesis to address these issues using isolated extensor digitorum longus (EDL) muscles

from male Wistar rats, ARC(s) mice and Ins2Akita diabetic mice.

Using EDL muscle preparations from rats, this thesis provides evidence that reversible

protein thiol oxidation plays a more important role than protein carbonylation in muscle

fatigue. Firstly, total protein thiol oxidation level increases in response to fatiguing

stimulation, but not following sustained non-fatiguing stimulation. Secondly, unlike

protein carbonylation responses, the recovery of muscle contractile performance after

fatiguing stimulation is accompanied by a return of total protein thiol oxidation to pre-

stimulation level. Thirdly, both muscle fatigue and total protein thiol oxidation level are

more pronounced when fatiguing stimulation takes place in the presence of the thiol

oxidising agent, diamide. Fourthly, muscle contraction performed with the thiol reducing

agent, dithiotreitol (DTT), decreases the magnitude of both muscle fatigue and total

protein thiol oxidation level without affecting protein carbonylation level. The muscle

ii

proteins targeted by reversible protein thiol oxidation include the contractile proteins,

actin, myosin, troponin and tropomyosin.

One limitation with the aforementioned results is that they are based on experiments

performed at 25 °C. Since this temperature is markedly different from that of working

muscles, this raises the question of the importance of the thiol oxidation level of muscle

proteins contributing to fatigue at physiological muscle temperature. Using isolated

mouse EDL preparations incubated at 34 °C, this thesis shows that the thiol oxidation level

of muscle proteins increases in response to fatiguing stimulation, and that acute exposure

to DTT improves muscle contractile performance by as much as 87 %. Although this

suggests that reversible protein thiol oxidation has the potential to be a major contributor

of muscle fatigue at physiological temperature, the importance of this mechanism could

not be ascertained because total protein thiol oxidation level in resting muscles is higher

compared to in vivo muscles, suggesting DTT is beneficial, in part, because it normalises

muscle function rather than only opposing muscle fatigue.

One of the issues raised by these findings is the question of whether some of the medical

conditions associated with impaired muscle function also involve an increase in the thiol

oxidation level of muscle proteins. This was tested using isolated muscle preparations

from Ins2Akita mice. Despite the absence of difference in maximal specific force between

the muscles of non-diabetic mice and diabetic mice, the latter were less resistant to

fatigue. Interestingly, the finding that the total thiol oxidation level of muscle proteins

attained after fatiguing stimulation in muscles from non-diabetic mice is comparable to

iii

that of muscles from diabetic mice prior to contraction could be taken as evidence that

changes in protein thiol oxidation level may not play an important role in explaining the

increased fatigue response in diabetic mice. However, since the thiol oxidation level of

myosin is lower in pre-fatigue muscles from diabetic mice compared to fatigued muscles

from non-diabetic mice, this could explain why resistance to muscle fatigue is lower in

muscles from diabetic compared to non-diabetic mice despite comparable total protein

thiol oxidation level. Arguably, this interpretation holds as long as myosin responsiveness

to thiol oxidation plays a more important role in determining contractile force compared

with other contractile proteins such as actin, troponin and tropomyosin. Although further

work is required to support this interpretation, the findings with myosin highlight the

importance of examining the response of individual proteins in addition to that of total

proteins.

In conclusion, this thesis shows for the first time that the thiol oxidation level of muscle

proteins increases in response to fatiguing muscle stimulation, and identifies experimental

conditions where oxidative stress contributes to muscle fatigue through protein thiol

oxidation rather than oxidative protein damage. This thesis also suggests that reversible

protein thiol oxidation has the potential to be a major mechanism of ROS-mediated

muscle fatigue at physiological temperature. Finally, this thesis suggests that the increased

thiol oxidation level of some proteins may explain, at least in part, the impaired muscle

contractile performance associated with diseases such as diabetes.

iv

ACKNOWLEDGEMENTS

I would like to acknowledge and thank the following people, whose contributions have

been invaluable, enriching and defining in making my PhD possible.

To my supervisors:

Professor Paul Fournier. For your wisdom, passion and enthusiasm. Thank you for your

constant encouragement and support you gave throughout my PhD years. It has been a

pleasure working with you!

Associate Professor Peter Arthur. For your insight, devotion and dedication. Thank you

for your desire for perfection and quality throughout my PhD years. It definitely made

generation of high quality results possible!

Asscoiate Professor Anthony Bakker. For your knowledge, professionalism and patience.

Thank you for your time and guidance throughout my PhD years. It definitely made the

experimental work easier!

To Asscoiate Professor Chooi-May Lai.

Many thanks for your generous provision of the mice samples.

To my dear family.

For supporting me unconditionally throughout my entire time in Perth. Thank you for

making this wonderful experiences a reality.

To my friends.

For encouraging and cheering me on until the very end. Thank you for making my PhD

years so much enjoyable.

To my dearest, Narisa Yee.

Thank you for your endless support, understanding and listening. Most of all, thank you

for being there, always by my side!

v

TABLE OF CONTENTS

EXECUTIVE SUMMARY ………………………………………………………………………………… i

ACKNOWLEDGEMENTS …………………………………………………............................... iv

TABLE OF CONTENTS …………………………………………………………………………………… v

LISTS OF FIGURES …………..………………………………………….................................... ix

LIST OF TABLES ……………..…………………………………………..................................... xi

LIST OF ABBREVIATIONS ……………………………………………………………………………… xii

1 LITERATURE REVIEW

1.1 Introduction …………………………………………………………………………………..... 2

1.2 Damaging effect of ROS and the antioxidant defense system ………….. 3

1.3 Oxidative stress and biomarkers ………………………………………………………. 6

1.4 Exercise-induced oxidative stress …………………………………………………….. 8

1.5 Mechanisms of ROS production during exercise ………………………………. 9

1.5.1

1.5.2

1.5.3

1.5.4

1.5.5

Mitochondria and electron transport chain …………………………

Nicotinamide adenine dinucleotide phosphate oxidase ………

Phospholipase A2-dependent processes ……………………………..

Xanthine oxidase …………………………………………………………………

Summary …………………………………………………………………………….

10

11

12

13

13

1.6 Factors influencing ROS production during exercise ………………………… 15

1.6.1

1.6.2

1.6.3

Temperature and ROS production ………………………………………

Carbon dioxide and ROS production ……………………………………

Summary …………………………………………………………………………….

15

16

17

1.7 Role of ROS in unfatigued muscle …………………………………………………….. 18

1.8 Role of ROS as a mediator of muscle fatigue ……………………………………. 21

1.8.1

1.8.2

Animal studies …………………………………………………………………….

Human studies ……………………………………………………………………

22

24

vi

1.9 Mechanisms and sites of ROS action ……………………………………………..... 28

1.9.1 Protein thiol oxidation ……………………………………………………….. 28

1.9.2 Protein oxidative damage …………………………………………………… 32

1.10 Statement of problems and hypotheses …………………………………………… 34

2 PROTEIN THIOL OXIDATION AND MUSCLE CONTRACTILE

PERFORMANCE

2.1 Introduction …………………………………………………………………………………….. 40

2.2 Materials and methods ……………………………………………………………………. 44

2.2.1 Animals ………………………………………………………………………………. 44

2.2.2 Intact muscle experiments …………………………………………………. 44

2.2.3 Skinned fiber experiments …………………………………………………. 46

2.2.4 Assay of proteins thiol oxidation levels ………………………………. 47

2.2.5 Electrophoretic separation and assay of ‘2 Tag’-labelled

protein ………………………………………………………………………………..

48

2.2.6 Assay of protein carbonylation levels …………………………………. 49

2.2.7 Statistical analysis ………………………………………………………………. 50

2.3 Results ……………………………………………………………………………………………… 51

2.3.1

The effect of fatiguing and sustained stimulation on protein

oxidation …………………………………………………………………………….

51

2.3.2 The effect of recovery from fatiguing stimulation on protein

oxidation …………………………………………………………………………….

52

2.3.3 The effect of protein thiol modification on fatiguing

stimulation and protein oxidation .………………………………………

55

2.3.4

Evidence that oxidation of contractile proteins correlates

with the loss of contractile force …………………………………………

57

2.3.5 Thiol oxidation of contractile proteins and the loss of

contractile force ………………………………………………………………….

59

vii

2.4 Discussion ………………………………………………………………………………………… 65

3 PROTEIN THIOL OXIDATION AND MUSCLE CONTRACTILE

PERFORMANCE AT 34 °C

3.1 Introduction …………………………………………………………………………………….. 73


3.2.1 Animals ………………………………………………………………………………. 75

3.2.2 Intact muscle preparation …………………………….……………………. 75

3.2.3 Fatiguing stimulation protocol ……………………………………………. 76



protein ………………………………………………………………………………..

78


3.3 Results ……………………………………………………………………………………………… 80

3.3.1 The effect of fatiguing stimulation on protein thiol oxidation

at 34 °C ...................................................................................

80

3.3.2 The effect of the protein thiol reducing agent, DTT on

fatiguing stimulation and protein thiol oxidation at 34 °C ……

81


contractile force at 34 °C …………………………………………………….

84

3.4 Discussion ………………………………………………………………………………………… 88

4 PROTEIN THIOL OXIDATION AND MUSCLE CONTRACTILE FUNCTION

IN INS2AKITA DIABETIC MICE

4.1 Introduction …………………………………………………………………………………….. 95


4.2.1 Animals ………………………………………………………………………………. 97

viii

4.2.2 Intact muscle preparation …………………………….……………………. 97

4.2.3 Fatiguing stimulation protocol ……………………………………………. 98



protein ………………………………………………………………………………..

100


4.3 Results ……………………………………………………………………………………………… 102

4.3.1 Body mass, fasting plasma glucose level and fasting glycated

hemoglobin of non-diabetic mice and diabetic mice …………..

102

4.3.2 The relationship between protein thiol oxidation of non-

diabetic mice and diabetic mice on associated muscle

functions …………………………………………………………….………………

103


contractile force ………………………………………………………………….

104

4.3.4 The effect of protein thiol modification on muscle

contraction and protein thiol oxidation ………………………………

106

4.4 Discussion ………………………………………………………………………………………… 110

5 GENERAL DISCUSSION

5.1 General discussion ……………………………………………………………… 117

6 REFERENCES

6.1 References …………………………………………………………………………. 126

ix

LIST OF FIGURES

Figure 1.1 Potential sites for the production of ROS in skeletal muscle.. 9

Figure 1.2 Biphasic effects of ROS in skeletal muscle force production... 20

Figure 1.3 Reversible thiol oxidation of muscle proteins which

potentially cause muscle fatigue …………………………………………

29

Figure 2.1 The effect of fatiguing and sustained stimulation on protein

oxidation …………………………………………………………………………….

52

Figure 2.2 The effect of recovery from fatiguing stimulation on protein

oxidation …………………………………………………………………………….

54

Figure 2.3 The effect of protein thiol modification on fatiguing muscle

contraction and protein oxidation ………………………………………

56

Figure 2.4 The effect of protein thiol oxidation and oxidative damage

on contractile force of skinned muscle fibers ..………………….

59

Figure 2.5 The effect of fatiguing and sustained stimulation on thiol

oxidation of individual proteins …………………………………………..

60

Figure 2.6 The effect of protein thiol modification on thiol oxidation of

individual proteins ………………………………………………………………

62

Figure 2.7 Correlation of individual protein thiol oxidation and

contractile force ….………………………………………………………………

63

Figure 2.8 The effect of recovery from fatiguing stimulation on thiol

oxidation of individual proteins …………………………………………..

64

Figure 3.1 The effect of fatiguing stimulation on protein thiol oxidation

at 34 °C ……………………………………………………………………………….

81

Figure 3.2 The effect of protein thiol reducing agent, DTT on fatiguing

stimulation and protein thiol oxidation at 34 °C ………………….

83

Figure 3.3 The effect of fatiguing stimulation on thiol oxidation of

x

individual proteins at 34 °C ………………………………………………… 85

Figure 3.4 The effect of protein thiol reducing agent, DTT on thiol

oxidation of individual proteins at 34 °C ……………………………

87

Figure 4.1 Body mass, fasting plasma glucose level and fasting glycated

hemoglobin of non-diabetic mice (CON) and diabetic mice

(DM) ……………………………………………………………………………………

102

Figure 4.2 The relationship between protein thiol oxidation of non-

diabetic mice (CON) and diabetic mice (DM) on associated

muscle functions ..……………………………………………………………….

104

Figure 4.3 The effect of fatiguing stimulation on thiol oxidation of

individual proteins in non-diabetic mice (CON) and diabetic

mice (DM) …………………………………………………………………………..

106

Figure 4.4 The effect of protein thiol modification on single muscle

contraction and protein thiol oxidation ………………………………

108

Figure 4.5 The effect of protein thiol modification on thiol oxidation of

individual proteins ………………………………………………………………

109

xi

LIST OF TABLES

Table 1.1 Important physiologic enzymatic antioxidants ……………………. 4

Table 1.2 Important physiologic non-enzymatic antioxidants ……………. 5

Table 1.3 Indirect biomarkers of oxidative stress ………………………………. 7

Table 1.4 Animal studies showing that antioxidant pretreatment

inhibits fatigue …………………………………………………………………….

23

Table 1.5 Human studies showing that antioxidant pretreatment

inhibits fatigue …………………………………………………………………….

26

xii

LIST OF ABBREVIATIONS

(SR) Ca2+ Sarcoplasmic reticulum ATPase

·OH Hydroxyl radical

Ca2+ Calcium

CAT Catalase

CO2 Carbon dioxide

DNP Dinitrophenyl

DNPH Dinitrophenylhydrazine

DTT Dithiothreitol

ecSOD Extracellular superoxide dismutase

EDL Extensor digitorum longus

ESR Electronic spin resonance

FCS Flavonoids from corn silk

FLm FL-N-(2-aminoethyl) maleimide

FHA FeSO4, H2O2 and ascorbate

GPx Glutathione peroxidase

H2O2 Hydrogen peroxide

HbA1c Glycated haemoglobin levels

HbA1C Gycated hemoglobin

KRS Kreb’s mammalian ringer solution

MnSOD Manganese superoxide dismutase

xiii

NAC N-acetylcysteine

NADPH Nicotinamide adenine dinucleotide phosphate oxidase

NO Nitric oxide

O2·⁻ Superoxide anion

PLA2 Phospholipase A2

ROS Reactive oxygen species

SDS Sodium dodecyl sulfate

-SH Thiol moiety

SR Sarcoplasmic reticulum

T1DM Type 1 diabetes mellitus

TCEP Phosphine hydrochloride

TRm Texas red-C2-maleimide

T-tubule Transverse tubule

UCP3 Mitochondrial uncoupling protein 3

1

CHAPTER 1:

LITERATURE REVIEW

2

1.1 INTRODUCTION

Over half a century ago, the emerging technology of electron spin resonance (ESR)

spectroscopy provided the first data showing that skeletal muscle contains free radicals

(Commoner et al., 1954). The biological importance of this finding was unclear at the time.

It was not until the early 1980s that researchers identified the first link between muscle

function and free radical biology. Electron spin resonance was again used to show that

free radical content was elevated in isolated frog limb muscles stimulated to contract

repetitively (Koren et al., 1980). Shortly afterwards, Davies and colleagues (1982) reported

a 2- to 3-fold increase in the free radical content of skeletal muscles from rats run to

exhaustion. These discoveries stimulated widespread interest in the relationship between

exercise and oxidative stress.

A free radical is defined as any molecular species capable of independent existence while

containing one or more unpaired electrons (Aruoma, 1994). An unpaired electron is one

that occupies an atomic or molecular orbital by itself. Free radicals can be formed by the

loss of a single electron or by the gain of a single electron by a non-radical molecule. The

most important free radicals in the body are part of a family of molecules better known as

reactive oxygen species (ROS; Cheeseman & Slater, 1993). Reactive oxygen species are

chemically reactive molecules containing oxygen which include superoxide anion (O2·⁻),

hydroxyl radical (·OH) and hydrogen peroxide (H2O2).

3

1.2 DAMAGING EFFECT OF ROS AND THE ANTIOXIDANT DEFENSE SYSTEM

Reactive oxygen species have the capacity to damage proteins, cause DNA-strand

breakage and compromise the integrity of polyunsaturated membrane lipids, which in

turn can affect the homeostatic environment of the cell (Niess & Simon, 2007).

Fortunately, all cells are endowed with a system of antioxidant enzymes that degrades

ROS. The sarcoplasm contains CuZn-superoxide dismutase and glutathione peroxidase and

catalase (Lawler & Power, 1998). The mitochondrial matrix contains Mn-superoxide

dismutase and glutathione peroxidase. Superoxide dismutase is responsible for

dismutating O2·⁻ to form H2O2 and oxygen, while glutathione peroxidase is capable of

reducing H2O2 or organic hydroperoxides to water and alcohol respectively (Lawler &

Power, 1998). Catalase is able to catalyze the breakdown of H2O2 to form water and

oxygen.

Other thiol-based antioxidant enzyme systems, thioredoxin and thioredoxin reductase

(Nakamura, 2005) and the peroxiredoxins (Rhee et al., 2005), are also expressed in

skeletal muscle (Hamelin et al., 2007). Functionally, the thioredoxin system operates as a

major ubiquitous disulfide reductase responsible for maintaining proteins in their reduced

state (Nakamura, 2005), while peroxiredoxins removes H2O2 and reduces both

hydroperoxides and peroxynitrate with the use of electrons provided by a physiological

thiol similar to the thioredoxin system (Rhee et al., 2005). Table 1.1 provides a brief

overview of the important physiologic enzymatic antioxidants function of these molecules.

4

Table 1.1: Important physiologic enzymatic antioxidants

Enzymatic Antioxidants

Superoxide

dismutase

Glutathione

peroxidase

Catalase

Thioredoxin and

thioredoxin

reductase

Peroxiredoxins

Properties

Located in both mitochondria and cytosol; dismutates

O2·⁻ (Lawler & Power, 1998)

Located in both mitochondria and cytosol; removes

H2O2 and organic hydroperoxides (Lawler & Power,

1998)


H2O2 (Lawler & Power, 1998)

Located in both mitochondria and cytosol; thioredoxin

catalyse the reduction of protein disulfide bonds, and

thioredoxin active-site cysteines are regenerated by

thioredoxin reductase (Nakamura, 2005)


H2O2 and organic hydroperoxides (Rhee et al., 2005)

The function of antioxidant enzymes is complimented by lipid-soluble and water-soluble

non-enzymatic antioxidants. Vitamin E, carotenes, and ubiquinol are lipid soluble and

localised in cell membranes (Janero, 1991; Powers et al., 2004). Lipoate and glutathione

are water soluble and widely distributed within the cytosol (Packer et al., 1995; Powers et

al., 2004). Glutathione is the most abundant non-protein thiol, present at near millimolar

concentrations, and is a primary determinant of the reducing environment within cells

5

(Ferreira & Reid, 2008). In the context of muscle fatigue, glutathione is among the most

important non-enzymatic antioxidants. These antioxidants protect the cell against ROS

toxicity in several ways. These include conversion of ROS into less-active molecules (i.e.,

scavenging) and prevention of the transformation of less reactive ROS into more

damaging forms (i.e., prevention of hydrogen peroxide transforming to the damaging

hydroxyl radical). Table 1.2 provides a brief overview of the important physiologic non-

enzymatic antioxidants function of these molecules.

Table 1.2: Important physiologic non-enzymatic antioxidants

Enzymatic Antioxidants

Vitamin E

Glutathione

alpha-Lipoic acid

Carotenoids

Ubiquinones

Properties

Lipid soluble phenolic compound; major chain breaking

antioxidant found in cell membranes (Janero, 1991)

Nonprotein thiol in cells; serves multiple roles in the

cellular antioxidant defense (Powers et al., 2004)

Endogenous thiol; effective as an antioxidant and in

recycling vitamin C; may also be an effective glutathione

substitute (Packer et al., 1995)

Lipid soluble antioxidants located primarily in membranes

of tissues (Powers et al., 2004)

Lipid soluble quinone derivatives; reduced forms are

efficient antioxidants (Powers et al., 2004)

6

1.3 OXIDATIVE STRESS AND BIOMARKERS

The term oxidative stress was first defined in 1985 as “a disturbance in the pro-oxidant-

antioxidant balance in favor of the former” (Sies & Cadenas, 1985). Because of the

complexity associated with the assessment of cellular redox balance, Jones (2006)

proposed that this term should be redefined as “a disruption of redox signaling and

control”. More recently, Sies and Jones (2007) defined oxidative stress as “an imbalance

between oxidants and antioxidants in favor of the oxidants, leading to a disruption of

redox signalling and control and/or molecular damage”.

A common approach to assess oxidative stress in biological systems involves the

measurement of changes in the level of redox-sensitive molecules that respond to

oxidative stress. In general, reliable markers of oxidative stress possess the following

qualities, which are: 1) chemically unique and detectable, 2) increase or decrease during

periods of oxidative stress, 3) possess relatively long half-lives, and 4) are not influenced

by other cellular processes (Halliwell & Gutteridge, 2007).

There are many molecules that possess one or more of these attributes, and techniques

are available to measure these biomarkers (Powers & Jackson, 2008). During periods of

oxidative stress, pro-oxidants may overwhelm the antioxidant defenses in cells and

damage cellular constituents. The oxidative stress in biological systems is characterised by

the increase in the formation of radicals and other oxidants. However, due to the

difficulties in measuring ROS production directly, most studies have used indirect

biomarkers to demonstrate oxidative stress. These indirect biomarkers of oxidative stress

7

typically fall into one of three categories 1) decrease in small-molecular-weight and/or

lipid-soluble antioxidants, 2) oxidative damage to cellular components (i.e., lipids,

proteins, and/or DNA), 3) disturbance in cellular redox balance (Table 1.3).

Table 1.3: Indirect biomarkers of oxidative stress

1. Antioxidants

Glutathione

Ascorbate

Alpha-tocopherol

Total antioxidant

capacity

2. Oxidation products

Protein carbonyls

Isoprostanes

Nitrotyrosine

8-OH-dG

4-hydroxy-nonenal

Malondialdehyde

3. Antioxidant/Pro-oxidant balance

Reduced glutathione to

oxidised glutathione

ratio

Cysteine redox state

Thiol/disulfide state

There are many biomarkers to quantify oxidative stress. Unfortunately, each category of

biomarkers of oxidative stress biomarkers has limitations and the development of a single

and ideal biomarker has proven to be a difficult task. Hence, it appears that no one

biomarker best assesses oxidative stress and that in most cases the measurement of

multiple biomarkers is required to confirm the presence of oxidative stress in tissues

(Halliwell & Gutteridge, 2007).

8

1.4 EXERCISE-INDUCED OXIDATIVE STRESS

There is compelling evidence that exercise can result in oxidative stress. These studies

reveal that skeletal muscle continually generates ROS (O’Neill et al., 1996; Kolbeck et al.,

1997; Stofan et al., 2000; Zuo et al., 2000; Pattwell et al., 2001; Pattwell et al., 2004).

Although some studies did not show an increase in oxidative stress with exercise (Viguie et

al., 1993; Sacheck et al., 2003; Ramel et al., 2004), with a decrease even reported by some

(Lovlin et al., 1987; Groussard et al., 2003), a large number of studies have reported

significant increases in ROS level associated with muscle contraction and exercise (Kawai

et al., 1994; Laaksonen et al., 1999; Mastaloudis et al., 2001; McAnulty et al., 2003;

Mastaloudis et al., 2004; Vasilaki et al., 2006; Gomez-Cabrera et al., 2010; Sahlin et al.,

2010).

There may be several reasons why some investigators have failed to observe signs of

exercise-induced oxidative stress. First, the use of different test subjects might have

influenced the findings of different studies. There is also a lack of information on factors

such as training status, age and sex which could potentially play a role in modulating

oxidative stress (Vollaard et al., 2005). Furthermore, research has adopted a large range of

exercise intensities. Only exercise of sufficient intensity or duration appears to lead to a

large enough increase in free radical production (Poulsen et al., 1996; Quindry et al.,

2003), and the sensitivity of the different markers of oxidative stress is such that some

markers of oxidative stress are sensitive enough to be affected by exercise, whereas

others are not (Niess et al., 1996; Liu et al., 2000).

9

1.5 MECHANISMS OF ROS PRODUCTION DURING EXERCISE

The mechanisms whereby exercise increases the production of ROS are contentious.

Although various mechanisms have been proposed, their relative contributions have yet

to be established. Various potential sites for ROS production exist in skeletal muscle and

are summarised and discussed below (Figure 1.1).

Figure 1.1. Potential sites for the production of ROS in skeletal muscle (Powers et al., 2010). Abbreviations

used: SR/T-tubule, sarcoplasmic reticulum/transverse tubule; PLA2, phospholipase A2; ecSOD, extracellular

superoxide dismutase; MnSOD, manganese superoxide dismutase; GPx, glutathione peroxidase; CAT,

catalase.

10

1.5.1 Mitochondria and electron transport chain

Until recently, it was generally accepted that the most important source of ROS, both at

rest and during exercise, involved an ‘electron leak’ from the electron transport chain in

the mitochondrial inner-membrane (Vollaard et al., 2005). Early reports suggested that 2-

5 % of the total oxygen consumed by mitochondria might undergo one electron reduction

with the generation of ROS (Boveris & Chance, 1973; Loschen et al., 1974). Since then,

studies have identified the major site(s) of ROS generation within mitochondria, with most

data now indicating that complexes I and III of the electron transport chain are the main

sites of mitochondrial ROS production (Barja, 1999; Muller et al., 2004). In complex I, the

main site of electron leakage to oxygen appears to be the iron-sulfur clusters, whereas in

complex III, it appears to be the Q10 semiquinone (Muller et al., 2004). In complex III, the

ROS is released from both sides of the inner mitochondrial membrane (Muller et al.,

2004).

Until recently, it was assumed that the increased ROS generation that occurs during

contractile activity is directly related to the elevated oxygen consumption which arises

with increased mitochondrial activity. This implies a 50- or 100- fold increase in ROS

generation by skeletal muscle during aerobic contractions (Kanter, 1994; Urso & Clarkson,

2003). St-Pierre and colleagues (2002) reassessed the rate of production of ROS by

mitochondria and indicated that the upper estimate of the proportion of the electron flow

that gives rise to ROS may be as low as ∼0.15 %. The low rate of ROS production may

include a role for uncoupling proteins (specifically mitochondrial uncoupling protein 3 in

11

skeletal muscle) as regulators of mitochondrial production of ROS acting to protect

mitochondria against oxidative damage (Brand et al., 2004; Brand & Esteves, 2005). In

addition, studies reveal that mitochondria produce more ROS during state 4 (basal)

respiration as compared with state 3 (maximal ADP-stimulated respiration) (Adhihetty et

al., 2005; Kozlov et al., 2005). This is significant as skeletal muscle mitochondria are

predominantly in state 3 during aerobic contractile activity which in turn limits their

generation of ROS during contractions (Adhihetty et al., 2005; Kozlov et al., 2005; Kavazis

et al., 2009). Collectively, these findings suggest that mitochondria may not be the primary

source of ROS production in skeletal muscle during exercise and further studies are

required to fully elucidate the role that mitochondria play in contraction-induced

production of ROS in skeletal muscle (Jackson et al., 2007).

1.5.2. Nicotinamide adenine dinucleotide phosphate oxidase

Nicotinamide adenine dinucleotide phosphate (NADPH) oxidase is another potential

source of ROS production during exercise. This enzyme generates superoxide by

transferring electrons from NADPH to molecular oxygen (Muller et al., 2004).

Nicotinamide adenine dinucleotide phosphate oxidase exists in several cellular locations in

muscle fibers (sarcoplasmic reticulum, transverse tubules, and sarcolemma) and has been

postulated to contribute to exercise induced ROS production in skeletal muscle (Powers et

al., 2010). The generation of ROS by NADPH oxidase has been shown at the sarcoplasmic

reticulum (SR) sites of both cardiac (Cherednichenko et al., 2004) and skeletal muscle (Xia

et al., 2003). The transverse tubules of skeletal muscle also contain NADPH oxidase whose

12

activity is increased by depolarisation (Espinosa et al., 2006; Hidalgo et al., 2006). This

enzyme contains some of the classical subunits found in the NADPH oxidase of phagocytic

cells, and releases superoxide to the cytosol of skeletal muscle cells (Jackson, 2008).

NADPH oxidase complex is also expressed at the plasma membrane level in skeletal

muscle (Javesghani et al., 2002). However, whether this complex releases ROS

predominantly to the plasma membrane cannot be ascertained (Javesghani et al., 2002).

Unfortunately, although NADPH oxidase is a source of ROS, the role this enzymes plays in

ROS production during exercise still remains to be determined.

1.5.3. Phospholipase A2-dependent processes

Phospholipase A2 (PLA2) is an enzyme that also has the potential to mediate ROS

production during exercise. This enzyme cleaves membrane phospholipids to release

arachidonic acid, which is a substrate for ROS-generating enzyme systems such as the

lipoxygenases (Zuo et al., 2004). Further, activation of PLA2 can activate NADPH oxidases

(Zhao et al., 2002), and increased PLA2 activity has been reported to stimulate ROS

generation in both muscle mitochondria (Nethery et al., 2000) and cytosol, (Gong et al.,

2006).

Both calcium-dependent and independent forms of PLA2 are reported to play a role in

muscle ROS generation (Power & Jackson, 2008). In this regard, the calcium-independent

enzymes has been proposed to modulate cytosolic oxidant activity in skeletal muscle cells

(Gong et al., 2006), whereas the calcium-dependent isoform located within mitochondria

has been reported to stimulate mitochondrial ROS generation during contractile activity

13

(Nethery et al., 1999). Gong and colleagues (2006) hypothesised that the calcium-

independent PLA2 is a major determinant of ROS activity under resting conditions,

whereas during contractions, heat stress, or other processes elevating intracellular

calcium, the calcium-dependent PLA2 is activated to promote ROS production. However,

the relative contribution of these enzymes to ROS production during exercise is still an

unresolved issue.

1.5.4. Xanthine oxidase

Xanthine oxidase is another enzyme with the capacity to generate ROS. This enzyme

oxidises hypoxanthine to produce xanthine and generate superoxide radicals (Halliwell &

Gutteridge, 2007). Several studies suggest that activation of xanthine oxidase in rat

skeletal muscle is an important source of contraction-induced ROS production (Judge &

Dodd, 2004; Gomez-Cabrera et al., 2005; Wang et al., 2009; Gomez-Cabrera et al., 2010).

However, while rat skeletal muscles contain significant levels of xanthine oxidase (Judge &

Dodd, 2004), human skeletal muscle cells appear to possess low amounts of xanthine

dehydrogenase or oxidase (Hellsten et al., 1996). Due to the lack of data in humans,

additional research is required to determine the role that xanthine oxidase plays in

exercise-induced ROS production.

1.5.5. Summary

Despite the initial indications that mitochondria are the predominant site for ROS

generation during contractile activity, a number of other potential cellular sites have been

14

identified. It is still unclear if all of these multiple sites contribute to the increase in ROS

levels during contractions or whether one site predominates. It is possible that the

multiple sites of generation are active in differing situations and that the effects of the

ROS generated are relatively localised and important for disparate functions.

15

1.6 FACTORS INFLUENCING ROS PRODUCTION DURING EXERCISE

Temperature and carbon dioxide (CO2) have been identified as factors with the potential

to influence ROS production during exercise.

1.6.1. Temperature and ROS production

Skeletal muscles are exposed to increased temperatures during intense exercise,

particularly in high environmental temperatures. There have been many studies showing

the importance of increased temperature on the production of ROS (Zuo et al., 2000;

Arbogast & Reid, 2004; Moopanar & Allen 2005; Edwards et al., 2007; van der Poel et al.,

2007). In general, as the temperature increases above 37 °C, the performance of isolated

preparations of skeletal muscle markedly decreases, and there is strong evidence

supporting the generation of ROS as a potential cause for the poor longevity of in vitro

skeletal muscle at physiological temperatures (37- 43 °C, Zuo et al., 2000; Edwards et al.,

2007; van der Poel et al., 2002; van der Poel & Stephenson, 2002; Arbogast & Reid, 2004;

Moopanar & Allen, 2005, 2006; van der Poel et al., 2007).

The effect of temperature on ROS production is likely influenced by intracellular sources.

Mitochondrial oxygen consumption (Brooks et al., 1971) and the enzymatic activities of

oxidoreductases, e.g., NADPH oxidase (Morgan et al., 2003) are strongly increased by a

rise in temperature. Antioxidant enzyme activities are also temperature sensitive,

affecting the pathways that buffer muscle-derived oxidants. Therefore, the changes in

total oxidant activity likely reflect changes in the net balance of these variables.

16

1.6.2. Carbon dioxide and ROS production

During exercise, skeletal muscles are exposed to elevated CO2 levels (Ranatunga et al.,

1987). There is evidence suggesting that CO2 influences redox homeostasis (Stofan et al.,

2000; Arbogast & Reid, 2004). Carbon dioxide and bicarbonate react directly with nitric

Oxide (NO) derivatives (Vesela & Wilhelm, 2002) and undergo electron exchange reactions

to form carbonate radicals (Liochev & Fridovich, 2001). Dissolved CO2 also promotes

acidosis in biological systems, stimulating cellular production of ROS and NO (Carbonell et

al., 2002). Studies in skeletal muscle about the net effect of CO2-mediated reactions on

ROS generation are conflicted as increased CO2 is reported to both promote oxidative

stress when measurement is taken outside the cell (Stofan et al., 2000) and protect

against oxidative stress when measurement is taken within the cell (Arbogast & Reid,

2004).

This apparent conflict can be explained based on compartmentalisation of the two assays

used in these studies (Arbogast & Reid, 2004). The reaction between CO2, NO and other

oxidants forms redox-active intermediates, such as peroxynitrite and carbonate radicals,

that are more unstable, have shorter half-lives, and diffuse shorter distances than the

parent molecules. This tends to localise nitrosative and oxidative reactions near the sites

of production, which are likely to be within the cell. Thus reactions with intracellular

targets become more likely, enhancing the ROS signal measured within the cell (Arbogast

& Reid, 2004), and diffusion of ROS out of cells becomes less likely, lessening the signal

measured outside the cell (Stofan et al., 2000).

17

1.6.3. Summary

In conclusion, oxidative stress during exercise is strongly influenced by temperature.

Although indications that CO2 could mediate in ROS production, it is still uncertain if the

presence of CO2 promotes or protects against oxidative stress. The limited data imply that

additional research is required to determine the role of CO2 in ROS production during

exercise.

18

1.7 ROLE OF ROS IN UNFATIGUED MUSCLE

It is well established that ROS have an important influence on force production in

unfatigued skeletal muscle. Low levels of ROS in skeletal muscle during basal conditions

are required for normal force production (Reid, 2001; Supinski et al., 2007). This has been

shown in studies using fiber bundles from rat diaphragm (Reid et al., 1993) and single fiber

preparations from limb muscles of frogs (Oba et al., 1996) or mice (Andrade et al., 1998).

When incubated with antioxidants such as catalase and dithiothreitol (DTT), these muscle

preparations experience a decrease in early twitch force, while the initial response to H2O2

exposure is a rise in developed force in rat diaphragm fiber bundles (Reid et al., 1993) and

in single fibers from limb muscles of frogs (Oba et al., 1996) or mice (Andrade et al, 1998).

The force–frequency relationship is also depressed by ROS depletion, shifting the curve

rightward in a dose-dependent manner. Furthermore, maximal tetanic force under low

ROS conditions is depressed by up to 50 %, a loss that is spontaneously reversed when

antioxidants are removed (Reid et al., 1993; Oba et al., 1996; Andrade et al., 1998). In

contrast, exposure to a superoxide generating system stimulates a leftward shift of the

force–frequency curve that is associated with an increase in twitch and submaximal

tetanic forces, but with no change in maximal tetanic force (Lawler et al., 1997).

As muscular activity increases ROS levels, the redox balance is driven to an oxidised state,

which also depresses force. Studies using single fibers from limb muscles of frogs (Oba et

al., 1996) or mice (Andrade et al., 1998) show that prolonged exposure to an oxidant such

as H2O2 depresses force production. Hence, the force–frequency relationship is depressed

19

by prolonged exposure with oxidant, shifting the curve rightward in a dose-dependent

manner (Oba et al., 1996; Andrade et al., 1998). This deleterious effect of H2O2 is

completely reversible by DTT (Oba et al., 1996; Andrade et al., 1998). It follows from these

observations that there is an intermediate cellular redox state that optimises force

production in skeletal muscle. This is evident when low-levels of ROS increase force in

unfatigued muscle (Reid et al., 1993; Oba et al., 1996; Andrade et al., 1998).

The aforementioned findings supported the view of Reid and colleagues (1993) who

proposed a theoretical model to explain the relationship between muscle redox balance

and isometric force production (Figure 2). Their model predicts that the muscle redox

state is a physiologically regulated variable that maintains equilibrium by matching the

rates of ROS production with the cellular antioxidant capacity. This model predicts that an

optimal cellular redox state exists whereby conditions are ideal for muscle force

production and that a deviation from the most favourable redox balance leads to a decline

in force production (Figure 1.2).

20

Figure 1.2. Biphasic effects of ROS in skeletal muscle force production (Reid et al., 1993). Point 1 illustrates

the force generated by muscle in basal state (i.e., no antioxidants or oxidants added). Point 2 illustrates the

force produced by unfatigued skeletal muscle exposed to low levels of oxidants; this represents the optimal

redox state for force production. Point 3 illustrates the deleterious effects of excessive ROS on skeletal

muscle force.

21

1.8 ROLE OF ROS AS A MEDIATOR OF MUSCLE FATIGUE

Muscle fatigue is defined as the reversible decrease in muscle force or power that occurs

with muscle contraction (Allen et al., 2008). Muscle fatigue plays an important protective

mechanism against ATP depletion and associated risk of rigor mortis state. Fatigue can

result from many sites along the complex pathways starting in the cortex and leading to

excitation of lower motor neurons in the spinal cord to the neuromuscular junction of the

muscle. Fatigue resulting from processes inside the spinal cord and above is defined as

central fatigue, whereas fatigue resulting from processes in the peripheral nerve,

neuromuscular junction, and muscle is defined as peripheral fatigue (Allen et al., 2008).

Peripheral fatigue can result from different type of exercise intensities. Exercise that

requires intense, near maximal muscle contraction results in the rapid onset of fatigue

that is likely caused by metabolic stresses (Allen & Westerblad, 2001). Force declines

rapidly under these conditions and recovers within seconds-to-minutes after exercise

stops. In contrast, fatigue resulting from less-intense, submaximal contractions is believed

to result, in part, from metabolic stresses such as exercise-mediated depletion of muscle

glycogen stores (Ortenblad et al., 2011), reduced availability of blood glucose due to a fall

in hepatic glycogen level (Callow et al., 1986), increased inorganic phosphate levels

(Westerblad et al., 2002), hyperthermia and dehydration (Nybo, 2008) to list a few. This is

the type of fatigue where oxidative stress appears to be important and will be explored in

more detail.

22

1.8.1. Animal studies

As discussed previously, there is ample evidence that ROS production increases in working

muscle and that strenuous exercise leads to oxidative stress (Kawai et al., 1994; Laaksonen

et al., 1999; Mastaloudis et al., 2001; McAnulty et al., 2003; Mastaloudis et al., 2004;

Vasilaki et al., 2006; Gomez-Cabrera et al., 2010; Sahlin et al., 2010). Many animal studies

implicate ROS as a contributor to contraction-induced muscle fatigue (Table 1.4). Several

studies using pharmacologic antioxidants have established a cause-and-effect link

between increased ROS production and fatigue of exercising respiratory and limb muscles.

For instance, ROS-selective antioxidant enzymes and other antioxidant drugs have been

shown to slow mechanical losses during electrically stimulated fatigue of muscle

preparations in vitro (Shindoh et al., 1990; Reid et al., 1992a; Diaz et al., 1994; Khawli &

Reid, 1994; Moonpanar & Allen, 2005; Ferreira et al., 2009) and in in vivo experiments

(Novelli et al., 1990; Davis et al., 2009; Yu et al., 2010; Agten et al., 2011; Hu & Deng,

2011). It is important to note, however, that antioxidant efficacy depends on the fatigue

protocol. Fatigue is delayed in the presence of antioxidants during contractile protocols

that use submaximal activation patterns, but not when contraction is at maximal or near-

maximal intensity (Reid et al., 1992; Matuszczak et al., 2005).

23

Table 1.4 Animal studies showing that antioxidant pretreatment inhibits fatigue

Study

Treatment

Performance Indicator

Performance

increases (%)

Shindoh et al

(1990)

150 mg/kg N-

acetylcysteine

Fatigue for 20 min at 20 Hz using

electrically stimulated

diaphragm muscle (In vitro)

21

Reid et al

(1992a)

5 x 102 U/ml

Superoxide dismutase,

1.8 x 104 U/ml

catalase




20

Diaz et al (1994) 4 mg/ml N-

acetylcysteine




35

Khawli & Reid

(1994)

10 mM N-

acetylcysetine

Fatigue for 10 min at 30-40 Hz

using electrically stimulated


Significant

Moopanar &

Allen (2005)

5 mM Tiron Fatigue for 2 min at 100 Hz using

electrically stimulated flexor

digitorum brevis (In vitro)

40

Ferreira et al

(2009)

10 mM L-2-

oxothiazolidine-4-

carboxylate (OTC)




13-16

Novelli et al

(1990)

6.5 mM Vitamin C and

spin trappers

Time to exhaustion using

swimming mices (In vivo)

50, 32-86

24

Davis et al (2009) 12.5-25 mg/kg

quercetin × 10 days

Maximal endurance exercise

performance using running

mices (In vivo)

36-37

Yu et al (2010) 500-2000 mg/kg

flavonoids × 10 days

Maximal endurance exercise

performance using swimming

rats (In vivo)

8-62

Agten et al

(2011)

150 mg/kg N-

acetylcysteine

Maximal diaphragm force

production using breathing mice

(In vivo)

Significant

Hu & Deng

(2011)

100-400 mg/kg

flavonoids daily for 28

days

Time to exhaustion using

running mice (In vivo)

42-72

1.8.2. Human studies

Several human trials have reported that nutritional antioxidants such as vitamins C and E,

beta-carotene, and linoleic acid do not improve exercise performance and are not

effective ergogenic aids (Snider et al., 1992; Alessio, 1993; Rokitzki et al., 1994; Goldfarb,

1999; Avery et al., 2003; Bryant et al., 2003; Jackson et al., 2004; Powers et al., 2004;

Gaeini et al., 2006). However, the antioxidant N-Acetylcysteine (NAC) has been shown to

be effective in preventing muscle fatigue in humans (Table 1.5). N-Acetylcysteine, a

reduced thiol donor that supports glutathione resynthesis (Ruffmann & Wendel, 1991),

not only inhibits muscle fatigue of isolated muscle preparations (Shindoh et al., 1990; Diaz

et al., 1994; Khawli & Reid, 1994), but also delays muscle fatigue in human experiments

25

involving electrically stimulated fatigue of limb muscle (Reid et al., 1994), breathing

against an inspiratory resistive load (Travaline et al., 1997), cycling exercise (Medved et al.,

2004; McKenna et al., 2006; Kelly et al., 2009; Bailey et al., 2011), repeated knee

extensions (Koechlin et al., 2004) and repetitive handgrip exercise (Matuszczak et al.,

2005). The NAC-mediated improvements in performance range from 15 % (Reid et al.,

1994; Matuszczak et al., 2005) to 69 % (Bailey et al., 2011). Nonetheless, it is noteworthy

that NAC does not appear to retard muscle fatigue during exercise at near-maximal

intensities in human studies where ROS effects will be minimal (Medved et al., 2003;

Matuszczak et al., 2005).

Recently, it has been reported that supplementation with the flavonoid antioxidant,

quercetin, may also delay muscle fatigue and improve human exercise performance (Table

1.5; Davis et al., 2010; Nieman et al., 2010). Flavonoids from corn silk (FCS) have been

investigated and confirmed to possess various pharmacological activities such as

antihypertensive, anti-infectious, anti-oxidative and anti-diabetic (Li & Yu, 2009; Liu et al.,

2011). In recent studies, FCS has been shown to be a potent scavenger of hydroxyl and

peroxyl radicals (Ren et al., 2005; Ebrahimzadeh et al., 2008; Hu et al., 2009). It has also

been reported that FCS had anti-fatigue activity (Hu et al., 2010). Overall, although there is

compelling evidence that antioxidant treatment can delay fatigue in healthy humans, it is

clear that the efficacy depends upon the chemical properties of the antioxidant.

26

Table 1.5 Human studies showing that antioxidant pretreatment inhibits fatigue

Study

Treatment

Performance Indicator

Performance

increases (%)

Reid et al (1994)

150 mg/kg/hr NAC for

60 min pre-exercise

Repetitive electrical stimulation

of tibialis anterior for 30 min at

10 Hz

15

Travaline et al

(1997)


60 min pre-exercise

Time to exhaustion from

breathing against inspiratory

resistor

50

Koechlin et al

(2004)

1800 mg/day NAC × 4

days then 600 mg

NAC pre-exercise


repetitive knee extensions

5

Medved et al

(2004)


15 min then 25

mg/kg/hr NAC pre-

exercise and during

exercise

Time to exhaustion from cycling 26

Matuszczak et al

(2005)


45 min pre-exercise


repetitive isometric handgrip

15

Mckenna et al

(2006)


15 min then 25

mg/kg/hr NAC pre-

exercise and during

exercise


27

Kelly et al (2009) 1800 mg NAC 45 min

pre-exercise

Breathing against inspiratory

resistor for 30 min

14

Davis et al (2010)

2 × 500 mg/day

quercetin × 7 days


Nieman et al

(2010)

1000 mg/day

quercetin × 14 days

Treadmill running for 12 min

3

Bailey et al

(2011)


15 min then 25

mg/kg/hr NAC pre-

exercise and during

exercise

Time to exhaustion from cycling 24-69

28

1.9 MECHANISMS AND SITES OF ROS ACTION

The intracellular mechanism(s) by which ROS contribute to muscle fatigue are unclear.

Two possible mechanisms involving the effect of ROS on these muscle proteins involved in

the control and activation of muscle force production during the excitation-contraction

coupling are described in the next sections. During contraction, calcium (Ca2+) ions are

released from the sarcoplasmic reticulum. These Ca2+ ions then binds to Troponin-C, and

this induces a conformational change in the regulatory complex exposing the actin

molecule for binding to the myosin ATPase located on the myosin head. These changes

result in the actin and myosin filaments sliding past each other thereby shortening the

sarcomere length resulting in a contraction.

1.9.1. Protein thiol oxidation

One mechanism whereby ROS might play a role in muscle fatigue involves the reversible

oxidation of protein thiols. The thiol moiety (-SH) of the amino acid cysteine can undergo

reversible, covalent reactions with muscle-derived oxidants to form, for instance, disulfide

bonds (Eaton, 2006). Thiol oxidation can alter protein function by interfering with the

enzyme active sites or by altering protein structure (Ferreira & Reid, 2008). Numerous

proteins involved in muscle contraction can undergo reversible thiol oxidation, and this

can potentially cause muscle fatigue as summarised in the following diagram (Figure 1.3).

For example, Ca2+ handling proteins, such as the SR Ca2+ ATPase and the SR Ca2+ release

channel have received attention because their activity can be affected by the oxidation of

their thiol groups (Scherer & Deamer 1986; Abramanson & Salama, 1989; Viner et al.,

http://www.cvphysiology.com/Cardiac%20Function/CF020.htm



29

1996; Anzai et al., 2000; Pessah & Feng, 2000; Gutierrez-Martin et al., 2004; Aracena-Parks

et al., 2006; Zissimopoulos et al., 2007).

Figure 1.3. Reversible thiol oxidation of muscle proteins which potentially cause muscle fatigue.

Thiol oxidation of contractile proteins could also contribute to muscle fatigue. Exposure of

actin to ROS, leads to the oxidation of cysteine 374 results in structural alterations of the

actin molecule (Dalle-Donne et al., 2003b), disrupting the interactions between actin and

actin binding proteins (Milanzi et al., 2000). Thiol oxidation of myosin and actin decreases

Ca2+-activated force and its Ca2+ sensitivity through impaired Ca2+ regulation of the actin–

K+

Na+

2 K+

3 Na+

K+Na+

P

P

PP

ATP

ATP

ATP + P

ATP + P

Mito

S.R.

tropomyosin

troponin

ATP

ATP + P

myosin

GP

PFK

T.T.

ACh

K+

Na+

actinCa2++

Ca2++

+

ROS

30

myosin cycle (Hertelendi et al., 2008). The oxidation of myosin causes a marked decrease

in myosin ATPase activity (Root & Reisler, 1992; Prochniewicz et al., 2008a). This is a

consequence of cysteine 707 oxidation in the myosin head which results in a decreased

fraction of myosin heads that undergo the force-generating transition during the

actomyosin ATPase cycle (Tiago et al., 2006). The effects of oxidation on actin in the

actomyosin complex are predominantly determined by oxidation-induced changes in

myosin heads, and changes in weak-to-strong structural transitions in actin and myosin

are coupled to each other and are associated with oxidative inhibition of muscle

contractile force (Prochniewicz et al., 2008a; Pizarro & Ogut, 2009).

In addition to actin and myosin, thiol oxidation of troponin (Putkey et al., 1993; Sousa et

la., 2006; Pinto et al., 2008; Pinto et al., 2011) and tropomyosin (Williams & Swenson,

1982) also affects muscle contractile function. Oxidation of cysteine 98 on troponin

reduces the binding affinity of troponin for actin, which destabilises the troponin-actin

relationship (Pinto et al., 2011) and results in a loss of force production (Sousa et la., 2006;

Pinto et al., 2008). For tropomyosin, a disulfide bridge can be formed on cysteine 190 in

response to oxidation, which results in reduced binding affinity of tropomyosin to actin,

which is reported to affect the ATPase activity of actomyosin (Williams & Swenson, 1982).

As mentioned above, numerous proteins involved in muscle contraction can undergo

reversible thiol oxidation. However, the myofilaments may be a more physiologically

important target for ROS during muscle fatigue compared to alternation of SR function, as

there are indications that intracellular Ca2+ handling proteins are less sensitive to the

31

effects of increased ROS exposure compared to the contractile proteins. In these

experiments, the oxidant H2O2 was found to decrease myofilament Ca2+ sensitivity in

single fast twitch fibers, without affecting SR Ca2+ release, and these changes were largely

reversible by exposure to the antioxidant DTT, indicating that thiol group modification was

playing a major role in these effects (Andrade et al., 1998; Lamb & Posterino, 2003;

Posterino et al., 2003). Endogenous ROS-induced during muscle fatigue were also shown

to decrease myofilament Ca2+ sensitivity in single fast twitch fibers, without altering SR

Ca2+ handling, and again these effects could be inhibited by DTT exposure (Moopanar &

Allen, 2006).

One major limitation with all of the aforementioned studies examining the effect of ROS

on muscle proteins is that these studies have been performed in vitro with isolated

proteins and most have used unphysiological levels of oxidants that may not reflect

physiologically relevant conditions. To date, no study has examined the effect of fatiguing

contraction on the thiol oxidation levels of muscle proteins, thus leaving unanswered the

identity of the protein targeted by exercise-mediated oxidative stress. This gap in our

knowledge is primarily due to the difficulty of investigating changes in the thiol oxidation

level of muscle proteins due to the poor sensitivity of the assay techniques available.

However, a newly developed assay, referred to as the ‘2 TAG’ protein thiol method

(Armstrong et al., 2011), has been shown to be sensitive enough to examine the thiol

oxidation levels of muscle proteins, and, as discussed later, this method will be adopted in

this thesis to identify the protein targeted by exercise-mediated oxidative stress.

32

1.9.2. Protein oxidative damage

Another possible intracellular mechanism by which ROS could contribute to muscle fatigue

involves protein dysfunction caused by oxidative damage. Protein oxidative damage is

generally considered to be irreversible and occurs when proteins directly react with ROS,

leading to the formation of protein derivatives or peptide fragments containing carbonyl

groups, such as aldehydes and ketones (Barrerio & Hussain, 2010). Increased protein

oxidative damage (as measured by the carbonyl assay) has been observed during exercise

and muscle contraction (Reznick et al., 1992; Saxton et al., 1994; Radák et al., 1998;

Barrerio et al., 2006; Pinho et al., 2006; Matsunaga et al., 2008; Veskoukis et al., 2008;

Dubersteina et al., 2009; Agten et al., 2011). An improvement in contractile force is also

observed with decreased protein carbonylation following the administration of

antioxidants (Reznick et al., 1992; Barrerio et al., 2006; Agten et al., 2011).

Numerous proteins involved either directly, or indirectly, in muscle contraction can

undergo irreversible oxidative damage, and potentially cause muscle fatigue. For example,

oxidation of the SR Ca2+-ATPase can inhibit SR Ca2+-pumping function, with an increase in

protein carbonylation levels being associated with a decrease in Ca2+-ATPase activity

(Matsunaga et al., 2008). In addition, exposure of actin to ROS causes a rapid increase in

the carbonylation level of actin which results in strong inhibition of actin polymerization

and a complete disruption of actin-filament organization (Dalle-Donne et al., 2001;

Fedorova et al., 2010). Mysoin is also targeted by ROS-induced carbonylation in skeletal

and cardiac muscles. The increase in the carbonylation of myosin is associated with

33

impaired contractile performance in cardiac muscle (Coirault et al., 2007; Shao et al.,

2010) and respiratory muscle (Yamada et al., 2007). Finally, carbonylation of tropomyosin

can also contribute to contractile dysfunction (Canton et al., 2006).

Although protein oxidative damage is generally considered to be irreversible (Barrerio &

Hussain, 2010) and for this reason not considered to be a mediator of muscle fatigue, this

view has recently been challenged by the observation in skeletal muscle and in cell

cultures of pulmonary smooth muscle cells that a rise in protein carbonylation level in

response to oxidative stress can be short lived (less than 30-60 min; Matsunaga et al.,

2008; Wong et al., 2008, 2010), with the decarbonylation of proteins occurring

independently of proteosomal activity (Wong et al., 2008), but via a mechanism which

remains to be elucidated (Wong et al., 2008). These observations raise the question of

whether reversible carbonylation could also be involved in promoting muscle fatigue. If

protein decarbonylation post-fatiguing muscle contraction were to occur in parallel with

recovery from fatigue, this could be taken as evidence that protein carbonylation may not

be only a marker of muscle damage, but also a mediator of muscle fatigue. In this regard,

it is noteworthy that the work of Matsunaga and colleagues (2008) shows that the

recovery of Ca2+ ATPase activity after exercise parallels that of the changes in its

carbonylation rather than thiol oxidation level.

Although it is possible that a short-lived transient rise in protein carbonylation might

contribute to fatigue, one major limitation with current literature is that the relative

importance of this mechanism compared to reversible protein thiol oxidation has never

34

been examined before. Since the possible involvement of reversible protein carbonylation

as a mediator of muscle fatigue still remains to be assessed, one of the objectives of this

thesis is to address these issues.

1.10 STATEMENT OF PROBLEM AND HYPOTHESES

As discussed above, it still remains to be determined whether the thiol oxidation level of

muscle proteins increases with fatiguing muscle contraction. For this reason, the first

objective of this thesis was to establish the extent to which the rise in protein thiol

oxidation levels correlates with muscle fatigue induced in response to a submaximal

muscle contraction protocol.

Given the importance of controlling for muscle damage and the possibility that short-lived

transient rise in protein carbonylation might also mediate fatigue, the second objective of

this thesis was to examine the response of protein carbonylation to fatiguing muscle

contraction.

Since temperature plays an important role in the production of ROS, the third objective of

this thesis was to examine the role of protein thiol oxidation on musce fatigue of isolated

skeletal muscle preparation at 34 °C.

Finally, as a means to explore whether muscle contractile dysfunction associated with a

number of medical conditions involve the reversible thiol oxidation of muscle proteins, the

last objective of this thesis was to provide evidence that muscle contractile dysfunction in

35

Ins2Akita diabetic mice is associated with an increase in the thiol oxidation level of muscle

proteins.

Using the aforementioned newly developed ‘2 TAG’ protein thiol assay method

(Armstrong et al., 2011), this thesis proposes to test the following hypotheses:

Chapter 2

The fatigue resulting from the isometric contraction of rat EDL muscle in vitro is

mediated by an increase in protein thiol oxidation rather than an increase in

protein carbonylation.

The fatigue resulting from the isometric contraction of rat EDL muscle in vitro is

associated with an increase in the thiol oxidation levels of myosin, actin, troponin

and tropomyosin.

Chapter 3

The fatigue resulting from the isometric contraction of rat EDL muscle at 34 °C in

vitro is associated with a rise in protein thiol oxidation.

The normalisation of the thiol oxidation of muscle proteins opposes the fatigue

resulting from the isometric contraction of rat EDL muscle at 34 °C

Chapter 4

Protein thiol oxidation is increased before and after isometric fatigue of in vitro

EDL muscles obtained from Akita diabetic mice

36

Increased protein thiol oxidation in the EDL muscles of diabetic mice is associated

with a reduced muscle contractile performance.

37

CHAPTER 2:

PROTEIN THIOL OXIDATION AND

MUSCLE CONTRACTILE PERFORMANCE

38

ABSTRACT

The aim of this study was to examine whether protein thiol oxidation or protein damage

(measured by protein carbonylation) resulting from oxidative stress could contribute to

muscle fatigue. Extensor digitorum longus (EDL) muscles from male Wistar rats (Rattus

norvegicus) incubated in Kreb’s ringer bicarbonate solution were subjected to a fatiguing

stimulation protocol which consisted of 10 min submaximal tetanic contractions of 500 ms

duration at 15 s intervals at a stimulation frequency of 50 Hz. These muscles were

compared to either control muscles in a rested state or muscles stimulated to contract

while exposed to either a protein thiol oxidising agent, diamide, or a protein thiol reducing

agent, dithiothreitol (DTT). In addition, some EDL muscles were subjected twice to the

fatiguing stimulation protocol mentioned above with a 60 min rest between the two

fatiguing stimulation. In response to fatiguing stimulation, contractile force fell by 50.8 ±

3.1 % (p<0.05) and this was accompanied by a 34.2 ± 7.2 % (p<0.05) increase in mean total

protein thiol oxidation level. Mean actin, myosin, troponin, tropomyosin thiol oxidation

levels were 22.4 ± 2.7 % (p<0.05), 55.5 ± 9.2 % (p<0.05), 34.0 ± 4.5 % (p<0.05), 29.2 ± 5.7

% (p<0.05) higher, respectively. The fall in contractile force following fatiguing stimulation

in muscles treated with DTT or diamide were 19.7 ± 2.7 % (p<0.05) lesser and 19.0 ± 6.5 %

(p<0.05) greater, respectively, compared to untreated stimulated muscles. Mean total

protein thiol oxidation levels were 24.8 ± 8.9 % (p<0.05) lower and 42.8 ± 9.6 % (p<0.05)

higher in response to DTT and diamide treatments, respectively. Mean actin, myosin,

troponin, tropomyosin thiol oxidation levels were 19.9 ± 2.5 % (p<0.05), 23.0 ± 2.9 %

(p<0.05), 22.4 ± 3.9 % (p<0.05), 22.5 ± 3.4 % (p<0.05) lower and 42.7 ± 7.0 % (p<0.05), 25.3

39

± 6.3 % (p<0.05), 34.0 ± 6.5 % (p<0.05), 26.0 ± 6.9 % (p<0.05) higher in response to DTT

and diamide treatments, respectively. Mean protein carbonylation was not significantly

different in response to diamide and DTT treatments. In response to the 60 min rest,

contractile force recovered to 92.1 ± 1.4 % (p<0.05), and this was accompanied with a full

recovery in mean total protein thiol oxidation level, while mean protein carbonylation

level remained elevated. The recovery in contractile force was associated with recovery in

the mean thiol oxidation levels of the actin, myosin, troponin and tropomyosin. However,

only actin and myosin was associated with a full recovery. In conclusion, this study

corroborates for the first time the view that oxidative stress contributes to muscle fatigue

through protein thiol oxidation, and that oxidative protein damage does not necessarily

contribute to muscle fatigue. These findings also raise the interesting possibility that the

thiol oxidation of multiple proteins directly, or indirectly, associated with contractile force,

contribute to ROS-mediated muscle fatigue.

Keywords: Reactive oxygen species, protein thiols, dithiothreitol, muscle fatigue

40

2.1 INTRODUCTION

The onset of muscle fatigue, defined as a reversible loss of muscle performance, limits

locomotive performance not only of individuals engaged in competitive sporting events or

recreational pursuits (Allen et al., 2008), but also of older individuals and patients with

chronic diseases such as heart failure (Flynn et al., 2009), chronic fatigue syndrome (Myhill

et al., 2009), and chronic obstructive pulmonary disease (Baghai-Ravary et al., 2009).

Among the many factors postulated to play a role in muscle fatigue, reactive oxygen

species (ROS) have been identified as potentially important contributors (Ferreira et al.,

2009; Kelly et al., 2009; Reardon & Allen, 2009a; Reardon & Allen, 2009b; Agten et al.,

2011). This is suggested by the marked increase in ROS production that occurs during

endurance exercise (Mastaloudis et al., 2001; McAnulty et al., 2003; Mastaloudis et al.,

2004; Vasilaki et al., 2006; Gomez-Cabrera et al., 2010; Sahlin et al., 2010) and from the

observations that pharmacologic antioxidants slow the rate of force decline in electrically

stimulated muscle preparations in vitro (Shindoh et al., 1990; Reid et al., 1992a; Reid et

al., 1992b; Diaz et al., 1994; Khawli & Reid, 1994; Moopanar & Allen, 2006; Ferreira et al.,

2009) and improve exercise performance in in vivo experiments (Novelli et al., 1990; Davis

et al., 2009; Yu et al., 2010; Agten et al., 2011; Hu & Deng, 2011). Similarly, ingestion of

some antioxidants in human participants has been shown in most, but not all studies

(Jackson et al., 2004; Powers et al., 2004; Matuszczak et al., 2005) to delay muscle fatigue

associated with a broad range of experimental protocols (Reid et al., 1994; Traveline et al.,

1997; Koechlin et al., 2004; Medved et al., 2004; Matuszczak et al., 2005; Mckenna et al.,

2006; Kelly et al., 2009; Davis et al., 2010; Nieman et al; 2010; Bailey et al., 2011).

41

One mechanism whereby ROS might contribute to muscle fatigue involves reversible

protein thiol oxidation. The thiol moiety (-SH) of the cysteine residues of proteins can

undergo reversible covalent reactions with muscle-derived oxidants to form disulfide

bonds (Eaton, 2006). Since muscle fatigue is also reversible, this mechanism is generally

considered to explain how ROS contribute to fatigue (Ferreira & Reid, 2008). In support of

this mechanism, numerous muscle proteins have been shown to undergo reversible thiol

oxidation when incubated in vitro with ROS. These include, for instance, calcium handling

proteins, such as the sarcoplasmic reticulum (SR) Ca2+ ATPase and the SR Ca2+ release

channel (Scherer & Deamer 1986; Abramanson & Salama, 1989; Viner et al., 1996; Anzai

et al., 2000; Pessah & Feng, 2000; Gutierrez-Martin et al., 2004; Aracena-Parks et al.,

2006; Zissimopoulos et al., 2007). Moreover, several contractile proteins are susceptible

to thiol oxidation and thus could also contribute to muscle fatigue. For example, thiol

oxidation of actin could disrupt the organisation of the actin filaments, leading to

reduction in the force generating ability of the sarcomere (Milzani et al., 2000). The

oxidation of myosin has been shown to inhibit myosin ATPase activity (Tiago et al., 2006).

The oxidation of the thiol groups of troponin also alters its function (Putkey et al., 1993).

Finally, the oxidation of tropomyosin can affect the ATPase activity of actomyosin

(Williams & Swenson, 1982). It is important to stress, however, that the aforementioned

effects of thiol oxidation on individual muscle proteins have been based mainly on studies

performed on purified proteins, and it still remains to be determined whether the thiol

oxidation of individual muscle proteins increases with muscle fatigue.

42

During muscle contraction, ROS also have the capacity to impair contraction by causing

oxidative damage to proteins through prcoesses such as protein carbonylation. Protein

carbonylation is reported to be an unavoidable physiological consequence of aerobic

activity (Dalle-Donne et al., 2003a), with increased oxidative protein damage observed

during exercise and repeated muscle contraction (Reznick et al., 1992; Saxton et al., 1994;

Radák et al., 1998; Barrerio et al., 2006; Pinho et al., 2006; Matsunaga et al., 2008;

Veskoukis et al., 2008; Dubersteina et al., 2009; Agten et al., 2011). Nevertheless, protein

carbonylation associated with protein damage is not generally considered to be

contributing substantially to muscle fatigue because muscle fatigue is a reversible process

and protein carbonylation is considered to be irreversible (Dalle-Donne et al., 2003a). This

view, however, has recently been challenged by the observation in skeletal muscles and in

cell cultures of pulmonary smooth muscle cell that the rise in protein carbonylation level

in response to oxidative stress can be short lived (less than 30-60 min; Matsunaga et al.,

2008; Wong et al., 2008, 2010). This decarbonylation of proteins can occur independently

of proteosomal activity (Wong et al., 2008), via a mechanism which remains to be

elucidated (Wong et al., 2008). These observations raise the question of whether

decarbonylation could also occur following muscle fatigue. In this regard, it is noteworthy

that the work of Matsunaga and colleagues (2008) shows that the recovery of Ca2+ ATPase

activity after exercise parallels that of the changes in its carbonylation level. Moreover,

many other proteins experience an increase in their carbonylation levels in response to

muscle contraction, such as actin (Dalle-Donne et al., 2001; Dalle-Donne et al., 2007;

Fedorova et al., 2010), myosin (Coirault et al., 2007; Shao et al., 2010), and tropomyosin

43

(Canton et al., 2006). Decreased protein carbonylation levels have also been linked to

improved contractile force following the administration of antioxidants (Reznick et al.,

1992; Barrerio et al., 2006; Agten et al., 2011).

Since it still remains to be determined whether the thiol oxidation of muscle proteins

increases with fatiguing muscle stimulation, the primary objective of this study was to

provide information about whether protein thiol oxidation could contribute to ROS-

mediated muscle fatigue. Also, given the possibility that a short-lived transient rise in

protein carbonylation might also contribute to fatigue, the next objective was to examine

the response of protein carbonylation to fatiguing muscle stimulation. Using the isolated

extensor digitorum longus (EDL) muscle preparation as the experimental model, this study

provides new evidence that protein thiol oxidation contributes, in part, to the ROS-

meidated muscle fatigue associated with submaximal muscle stimulation in vitro, with

little or no role played by protein carbonylation. In addition, thiol oxidation of actin,

myosin, troponin and tropomyosin have been identified as potential contributors to

muscle fatigue.

44

2.2 MATERIALS AND METHODS

2.2.1 Animals

Male Wistar rats (Rattus norvegicus) weighing an average of 140 g were kept at a constant

room temperature ranging between 22 and 24 °C, with free access to food and water. All

experiments were approved by the Animal Ethics Committee of the University of Western

Australia.

2.2.2 Intact muscle experiments

Male Wistar rats were anaesthetised with an intraperitoneal injection of pentobarbital (65

mg/kg of body weight), and both EDL muscles were carefully excised. While the first

muscle was being tested, the other was left intact inside the animals and removed once

experimentation with the first muscle was completed (≈10 min). Using a braided surgical

silk (Size 3), one end of the muscle tendon was attached to a dual mode force transducer-

servomotor (1200A Intact Muscle Test System, Aurora Scientific Inc, Canada) and the

tendon of the other end of the muscle was attached to an anchoring hook. The muscle

preparation was bathed in Kreb’s mammalian Ringer solution (KRS) (Composition: NaCl,

121 mM; KCl, 5·4 mM; MgSO4, 1.2 mM; NaHCO3, 25 mM; HEPES, 5 mM; glucose, 11.5 mM;

CaCl2, 2.5 mM at pH 7.4) gassed with 5 % CO2 and 95 % O2 at 25 °C throughout the

experiment. A temperature of 25 °C was chosen because it is within the range of

temperatures that are optimal for maintaining isometric force in vitro (25–30 °C; Segal et

al., 1985, 1986). The muscle was then stimulated with single twitches to identify the

45

optimum length producing the highest force possible before experimentation. Force

responses were elicited using a 701B muscle stimulator (Aurora Scientific Inc, Canada).

Next, maximal force responses were obtained in response to supramaximal stimulation

pulses (pulse width: 0.3 ms) delivered at 150 Hz for 500 ms with a 2 min rest between

each stimulation to stabilise the force responses. The muscles were then given a 10-min

rest before the commencement of the stimulation. The two submaximal stimulation

protocols used in the experiments on rats were based on findings from preliminary

experiments. They involved exposure to stimulation pulses (80 V and 1000 mA) delivered

at 50 Hz for 500 ms with either a 15 s (fatigue) or 60 s (sustained non-fatigue) interval

between contractions for 10 min. For all experiments, isolated muscles were maintained

for 10 min in KRS before the commencement of each experiment. In order to study the

effect of contraction on protein oxidation, isolated muscles from rats were subjected to

either a fatiguing stimulation or a sustained non-fatiguing stimulation, and the levels of

protein oxidation were compared to resting muscles not stimulated to contract. After 10

min of stimulation or rest, all muscles were frozen using aluminium clamps pre-cooled in

liquid nitrogen and stored at -80 °C. To study the effect of recovery on fatiguing

stimulation and protein oxidation, isolated muscles were subjected to two fatiguing

stimulation runs each separated by a 60 min rest. Muscles were frozen and stored at -80

°C either before the first fatiguing stimulation, after the first fatiguing stimulation, after

the 60 min recovery, or after the second fatiguing stimulation.

In order to study the effect of protein thiol reducing and protein thiol oxidising agents on

fatiguing stimulation and protein oxidation, isolated muscles from rats were maintained

46

for 10 min in KRS with 5 mM dithiothreitol (DTT) or 1 mM diamide before the

commencement of the fatiguing stimulation. Measurements made from the fatigued

muscles exposed to DTT or diamide were then compared with fatigued muscles not

exposed to any DTT or diamide during the experiment. After 10 min of fatiguing

stimulation, muscles were frozen and stored as described above.

2.2.3 Skinned fiber experiments

The skinned fiber experiments undertaken in this study were similar to those described

previously (Han et al. 2003). Briefly, EDL muscles from male Wistar rats were removed and

placed in paraffin oil. A segment of a single muscle fiber was isolated and attached to a

force transducer (SI, Heidelberg) and maintained at slack length. The fibers were then

chemically skinned by exposure to a low Ca2+ (relaxing) solution (Solution A) containing

Triton X-100 (2 % v/v) to permeabilise all cellular membrane components and eliminate

Ca2+ transport pathways. Solution A consisted of: K+ 117 mM, Na+ 36 mM, ATP 8 mM, free

Mg2+ 1 mM, creatine phosphate 10 mM, EGTA 50 mM, N-2-Hydroxyethyl-piper-azine-N'-2-

ethanesulphonic acid (HEPES) 90 mM, NaN3 1 mM at pH 7.1. Maximal force responses

were induced by exposing the muscle fibers to Solution B, which had a similar composition

to Solution A, except that the [EGTA2-] and [CaEGTA] of Solution B were 0.4 and 49.6 mM,

respectively (Lamb & Stephenson, 1990). The free [Ca2+] of the solutions was calculated

using a Kapp for EGTA of 4.78.106 (Fink et al., 1986). In these experiments, fibers were

maintained in Solution A for 2 min and then exposed to Solution B briefly to determine

maximal force. Force was then returned to baseline by transfer of the fibers to Solution A.

47

The fibers were then exposed to a similar set of solutions, containing either 5 mM diamide

or 4 mM FeSO4, 10 mM H2O2 and 25 mM ascorbate (FHA) for a similar time period, to

examine the effect of the oxidants on contractile force production. Finally, the fibers were

exposed to another similar set of Solution A and B containing 1 mM DTT, to examine

whether any effects of the oxidants on contractile force production were reversible. The

effects of the oxidising and reducing agents were measured and expressed as a

percentage of the corresponding initial control maximum contractile force. All

experiments were performed at room temperature (23 ± 2 °C).

2.2.4 Assay of proteins thiol oxidation levels

Protein thiol oxidation levels were determined using a fluorescent two-tag labelling

technique, which involved the sequential labelling of reduced and oxidised protein thiol

groups using two separate fluorescent tags on the same protein sample (Armstrong et al.,

2011). Briefly, muscles were homogenised on ice with 20 % (w/v) trichloroacetic acid

(TCA) in acetone using an Ultra-turax (IKA, Germany) and subsequently stored at -20 ˚C.

The protein pellets extracted from 20 % (w/v) TCA in acetone were re-suspended in 50 μl

of sodium dodecyl sulphate (SDS) buffer (0.5 % (w/v) SDS and 0.5 M Tris pH 7.0) in the

presence of 0.5 mM BODIPY FL-N-(2-aminoethyl) maleimide (FLm) in the dark. All samples

were sonicated and incubated for 30 min at room temperature in the dark with

continuous vortexing. After centrifugation, the labelled protein solution was precipitated

with cold ethanol to remove excess dye (FLm), and then re-dissolved in 50 µl of SDS buffer

(pH 7.0). Samples were reduced with the addition of 4 mM tris (2-carboxy-ethyl)

48

phosphine hydrochloride (TCEP) and incubated for 60 min at room temperature in the

dark. After reduction, samples were labelled with 0.25 mM Texas red-C2-maleimide

(TRm), and incubated for 30 min at room temperature in the dark. Ethanol precipitation

and re-solubilisation in SDS buffer was repeated four times to remove excess un-reacted

dye (TRm). Samples were re-solubilized in 100 μL of SDS buffer, and the intensity of the

fluorescence was measured for both FLm and TRm (FLm: excitation = 485 nm, emission =

520 nm and TRm: excitation = 595 nm, emission = 610 nm). Ovalbumin pre-labelled with a

known quantity of FLm and TRm was used as a standard to quantify labelled protein

thiols. Protein concentration (mg/ml) was determined using the Bio-Rad DC Protein Assay.

The percentage of protein thiol oxidation was expressed as the concentration of oxidised

thiols (TRm) to total protein thiols (FLm and TRm).

2.2.5 Electrophoretic separation and assay of ‘2 Tag’-labelled protein

To determine the levels of thiol oxidation of individual protein bands in the muscle

sample, polyacrylamide gel electrophoresis was performed on the fluorescently labelled

samples, prepared as described above, using the BioRad Mini Protean III system

(Armstrong et al., 2011). Gel and buffer compositions were based on those described

previously (Kohn & Myburgh, 2006), with the exceptions that a 12 % polyacrylamide gel

was used and 5 mM DTT was added to the top running buffer. Electrophoresis was carried

out at 8 mA with voltage not exceeding 150 V for 8.5 hr at 4 °C, with the buffer in the

lower chamber stirred with a magnetic stirrer throughout. Upon completion of the gel

electrophoretic separation of proteins, the gel was immediately scanned for fluorescence

49

using a Typhoon Scanner (Ammersham) at the wavelengths associated with both dyes

(FLm: excitation = 485 nm, emission = 520 nm and TRm: excitation = 595 nm, emission =

610 nm). The individual protein bands corresponding to actin, myosin, tropomyosin and

troponin were identified at molecular weight of these proteins of 42 kD, 205 kD, 33 kD

and 18 kD respectively.

2.2.6 Assay of protein carbonylation levels

Protein carbonylation levels were assayed following their reaction with 2,4-

dinitrophenylhydrazine (DNPH) to form 2,4-dinitrophenyl (DNP) hydrazone (Levine et al.,

1990; Hawkins et al., 2009). In brief, muscles were homogenised on ice with 20 % (w/v)

TCA in acetone using an Ultra-turax (IKA, Germany), then the muscle extracts were

precipitated by centrifugation at 8000 g for 10 min at 4 °C, with the resulting protein

pellets dried and re-suspended in 10 mM DNPH in 2 M HCl and incubated for 30 min at

room temperature in the dark. Proteins were later washed with ethyl acetate/ethanol

(1:1) and dissolved in 6 M guanidine hydrochloride, and absorbance was measured at 370

nm. Protein concentration (mg/ml) was determined using the Bio-Rad protein assay.

Carbonyl levels are expressed as nmol of carbonyl per mg soluble protein.

2.2.7 Statistical analysis

Results are presented as means ± standard error of the mean. All analyses were

performed using either a Student’s paired t test or a one way ANOVA followed by Fisher

LSD posteriori test, with significance at P<0.05. Sample sized is denoted by n.

50

2.3 RESULTS

2.3.1 The effect of fatiguing and sustained stimulation on protein oxidation

Isolated EDL muscles from rats were used to investigate protein oxidation levels after

muscle stimulation. Fatiguing stimulation resulted in a 51 % decline in mean contractile

force (Figure 2.1A), and a 34 % increase in mean total protein thiol oxidation level (Figure

2.1B), and no significant changes in mean protein carbonylation level (Figure 2.1C).

However, when compared with pre-fatigued muscles, there was an increase in protein

carbonylation level (Figure 2.2D). In response to sustained non-fatiguing stimulation,

mean contractile force fell by less than 5 % (Figure 2.1A), and there were no significant

changes in mean total protein thiol oxidation level (Figure 2.1B) or mean protein

carbonylation level (Figure 2.1C). There was no difference in total protein thiols (Figure

2.1D) indicating that the increase in protein thiol oxidation level measured likely

represents increased conversion of reduced protein thiols to oxidised protein thiols. These

data are consistent with protein thiol oxidation rather than oxidative protein

carbonylation contributing to the loss in muscle contractile force during fatiguing

stimulation.

Whether the level of protein thiol oxidation in isolated EDL muscles at 25 °C gassed with

95 % O2 and 5 % CO2 was comparable to that of the in vivo EDL muscles (freshly dissected)

was also examined. There was no significant difference between resting mean total

protein thiol oxidation level (7.5 ± 0.3 %, n=6) and in vivo mean total protein thiol

oxidation level (7.6 ± 0.2 %, n=6).

51

Figure 2.1. The effect of fatiguing and sustained stimulation on protein oxidation. Isolated EDL muscles

were stimulated at 50 Hz for 500 ms with either a 15 s (fatigue) or 60 s (sustained) interval between

contractions for a period of 10 min. Following 10 min of stimulation, mean percentage of initial force (A),

percentage of protein thiols oxidised (B), protein carbonyls (C) and total protein thiols (D) were measured.

*Significantly different measured at p<0.05 relative to resting muscle incubated for 10 min without

stimulation. n=6. Values are expressed as means ± SE.

2.3.2 The effect of recovery from fatiguing stimulation on protein oxidation

Given that protein thiol oxidation can be reversed by intracellular antioxidant pathways,

and that protein carbonylation can be short-lived, it was of interest to examine whether

muscle recovery from fatigue was associated with the normalisation of protein thiol

oxidation and/or protein carbonylation levels. Muscles were subjected to two 10 min

52

fatiguing stimulation runs each separated by a 60 min rest period. After 60 min of rest,

mean total protein thiol oxidation level returned to pre-fatigue resting level (Figure 2.2C),

but there was no recovery in mean protein carbonylation level (Figure 2.2D). Contractile

force at the beginning of the second fatiguing stimulation recovered to 92 % of the initial

contractile force attained in response to the first fatiguing stimulation (Figure 2.2A).

Following the second fatiguing stimulation, the decline in contractile force was

comparable to the first fatiguing stimulation (Figure 2.2B). Mean total protein thiol

oxidation also increased to a level comparable to that attained after the first fatiguing

stimulation (Figure 2.2C), whereas there was a further progressive increase in mean

protein carbonylation level (Figure 2.2D). Taken together, these data indicate that changes

in protein thiol oxidation, but not protein carbonylation, correlate with changes in muscle

contractile force. Overall, there was no change in total protein thiols which indicated that

reversible protein thiol oxidation could be contributing to the decrease in contractile force

(Figure 2.2E).

To investigate whether increases in protein carbonylation could lead to a decrease in

contractile function, isolated EDL were exposed to FHA to increase protein carbonylation

level. The oxidative damage reagent caused a significant increase in mean protein

carbonyls from 1.85 ± 0.2 nmol.mg-1 to 3.5 ± 0.3 nmol.mg-1 (n=6, P<0.05). This was

associated with a 24.6 ± 2.4 % (n=6, P<0.05) decline in initial contractile force. These

findings suggest that excessive protein carbonylation level can affect muscle contractile

force.

53

Figure 2.2. The effect of recovery from fatiguing stimulation on protein oxidation. Isolated EDL muscles

were fatigued by two fatiguing stimulation runs which consisted of 50 Hz for 500 ms with a 15 s interval

between contractions for a period of 10 min separated by a 60 min rest. Following a 60 min rest, the

recovered muscles were fatigued for a second

time. Mean percentage of initial force of first fatigue (A),

percentage of oxidised protein thiols (C), protein carbonyls (D), total protein thiols (E) were measured

before first fatigue (rest), after first fatigue, after recovery and after second fatigue. Percentage of mean

54

force relative to initial force of the first fatigue (△) and second fatigue (△) were measured every min for the

10 min fatiguing stimulation (B). *Significantly different measured at p<0.05. n=6. Values are expressed as

means ± SE.

2.3.3 The effect of protein thiol modification on fatiguing stimulation and protein

oxidation

To provide further evidence that reversible protein thiol oxidation contributed to the loss

of contractile force in response to 10 min of fatiguing stimulation, an investigation was

conducted to examine whether the protein thiol reducing agent, DTT could prevent the

loss in muscle contractile force. DTT did not affect the mean initial absolute force (Figure

2.3A), but following 10 min of fatiguing stimulation, the mean force of muscles exposed to

DTT was 20 % higher than muscles not exposed to DTT (Figure 2.3B). The improvement in

contractile force was associated with a 25 % reduction in mean total protein thiol

oxidation level (Figure 2.3C), but no change in mean protein carbonylation level (Figure

2.3D).

If reversible protein thiol oxidation contributed to the loss in contractile force, then the

protein thiol oxidising agent diamide should augment the loss in muscle contractile force.

Diamide did not affect the initial absolute force (Figure 2.3A), but following 10 min of

fatiguing stimulation, the mean force of muscles exposed to diamide was 19 % less than

muscles not exposed to diamide (Figure 2.3B). The greater decline in contractile force was

associated with a 43 % higher mean total protein thiol oxidation level (Figure 2.3C), but no

change in mean protein carbonylation level (Figure 2.3D). There was no change in total

55

protein thiols (Figure 2.3E). Overall, these findings are consistent with the hypothesis that

protein thiol oxidation, but not protein carbonylation, affects muscle contractile force

during fatiguing stimulation.

Figure 2.3. The effect of protein thiol modification on fatiguing stimulation and protein oxidation. Isolated

EDL muscles were fatigued by stimulating at 50 Hz for 500 ms with a 15 s interval between contractions for a

56

period of 10 min while exposed to either 5 mM DTT or 1 mM diamide. Following 10 min of fatiguing

stimulation, initial absolute force (A), mean percentage of initial force (B), percentage of oxidised protein

thiols (C), protein carbonyls (D) and total protein thiols (E) were measured. *Significantly different measured

at p<0.05 relative to fatigued muscles not exposed to either DTT or diamide. n=6. Values are expressed as

means ± SE.

2.3.4 Evidence that oxidation of contractile proteins correlates with the loss of contractile

force

In order to investigate whether oxidation of contractile proteins may be contributing to

the loss in muscle contractile force during 10 min of fatiguing stimulation, some muscles

were exposed to 10 mM caffeine at the fifth min of the 10 min fatiguing stimulation.

There was only a partial recovery in contractile force to 65.3 ± 1.6 % from 46.0 ± 2.7 %

(n=6, P<0.05) of the contractile force following the 10 min fatiguing stimulation. Since

caffeine facilitated sarcoplasmic reticulum Ca2+ channel opening (Rousseau et. al, 1988),

fatiguing effects associated with calcium availability would have been negated.

Consequently, the lack of complete recovery in contractile force was consistent with the

concept that oxidation of contractile proteins contributed to the loss of contractile force

during fatiguing stimulation. However, the duration of caffeine exposure might be

insufficient for caffeine to diffuse and reach steady state near fibers located in the muscle

core center because large muscles were used here.

Skinned muscle fibers were also used as a model to further examine whether oxidative

modification of contractile proteins affected muscle contractile force, independent of the

effects on other cellular components such as the sarcoplasmic reticulum, cytosol and

57

mitochondria. This model was adopted because skinned muscle fibers allow solutes access

to the intracellular space without the potential confounding effects caused by other non-

contractile proteins involved in muscle contraction, such as Ca2+ handling proteins

(Gutierrez-Martin et al., 2004; Aracena-Parks et al., 2006; Zissimopoulos et al., 2007).

In response to treatment with a protein thiol oxidising agent, diamide, there was a 29 %

decrease in maximal contractile force generated in skinned fibers (Figure 2.4A). When

skinned fibers were subsequently exposed to the thiol reducing agent DTT, the contractile

force generated by the fibers previously exposed to diamide recovered to 92 % of control

values (Figure 2.4A). The effect of exposing skinned muscle fibers to conditions which

increases protein carbonylation level on muscle fiber contractile force were also

examined. When skinned fibers were exposed to an oxidising reagent, FHA, which can

cause protein carbonylation, there was a 32 % decrease in contractile force (Figure 2.4B).

However, when skinned fibers were subsequently exposed to DTT, there was no recovery

in contractile force (Figure 2.4B).

Overall, these data demonstrate that oxidative stress can affect contractile force by either

reversible protein thiol oxidation or irreversible protein oxidation of contractile proteins.

Of note is that the effects of reversible protein thiol oxidation could be distinguished from

the effects of irreversible oxidative protein damage by using the thiol reducing agent, DTT.

58

Figure 2.4. The effect of protein thiol oxidation and oxidative damage on contractile force of skinned

muscle fibers. Skinned fibers were stimulated to contract after exposure to either either 5 mM diamide (A)

or 4 mM FeSO4, 10 mM H2O2 and 25 mM ascorbate (FHA, B). Following contraction, skinned fibers were

exposed to 1 mM DTT and stimulated to contract again (A, B). Mean percentage of control force were

measured after exposure to different treatments. *Significantly different measured at p<0.05 relative to

control skinned fibers not exposed to either diamide, FHA or DTT. n=8. Values are expressed as means ± SE.

2.3.5 Thiol oxidation of contractile proteins and the loss of contractile force

To investigate the effect of muscle stimulation on the thiol oxidation of contractile

proteins, thiol oxidation levels of four major contractile proteins (actin, myosin, troponin

and tropomyosin) were measured. Following 10 min of fatiguing stimulation, mean thiol

oxidation levels of actin, myosin, troponin and tropomyosin increased by 22 % (Figure

2.5A), 55 % (Figure 2.5B), 34 % (Figure 2.5C) and 29 % (Figure 2.5D), respectively. There

were no significant changes in mean actin (Figure 2.5A), troponin (Figure 2.5C) and

tropomyosin (Figure 2.5D) thiol oxidation levels after 10 min of sustained stimulation.

However, there was a 29 % (Figure 2.5B) increase in mean myosin thiol oxidation level

after 10 min of sustained stimulation which was associated with a 5 % loss in contractile

59

force (Figure 2.1A). These data are consistent with the thiol oxidation of contractile

proteins contributing to a loss of muscle contractile force in response to 10 min of

fatiguing stimulation.

Figure 2.5. The effect of fatiguing and sustained stimulation on thiol oxidation of individual proteins.

Percentage thiol oxidation of actin (A), myosin (B), troponin (C) and tropomyosin (D) were measured after

isolated EDL muscles were stimulated at 50 Hz for 500 ms with either a 15 s (fatigue) or 60 s (sustained)

interval between contractions for a period of 10 min. *Significantly different measured at p<0.05 relative to

resting muscles incubated for 10min without stimulation. n=6. Values are expressed as means ± SE.

If changes in the thiol oxidation level of a protein are responsible for changes in muscle

contractile force, any change in thiol oxidation level should be accompanied by a change

in contractile force. To investigate this further, the effect of DTT and diamide on specific

60

protein thiol oxidation and contractile force was examined. In this experiment, the

improved contractile force after incubation in DTT was associated with 20 % (Figure 2.6A),

23 % (Figure 2.6B), 22 % (Figure 2.6C) and 23 % (Figure 2.6D) decrease in mean thiol

oxidation levels of the actin, myosin, troponin and tropomyosin, respectively. The more

pronounced declined contractile force after incubation in diamide was associated with 43

% (Figure 2.7A), 25 % (Figure 2.7B), 34 % (Figure 2.7C) and 26 % (Figure 2.7D) increase in

mean thiol oxidation levels of the actin, myosin, troponin, and tropomyosin, respectively.

There were significant correlations between the degree of actin (R2=0.557, P<0.05),

myosin (R2=0.290, P<0.05), troponin (R2=0.460, P<0.05), and tropomyosin (R2=0.388,

P<0.05) thiol oxidation and muscle contractile force (Figure 2.8) suggesting that contractile

force decreases with the increase in thiol oxidation levels of contractile proteins.

61

Figure 2.6. The effect of protein thiol modification on thiol oxidation of individual proteins. Percentage

thiol oxidation of actin (A), myosin (B), troponin (C) and tropomyosin (D) were measured after isolated EDL

muscles were fatigued by stimulating at 50 Hz for 500 ms with a 15 s interval between contractions for a

period of 10 min while exposed to either 5 mM DTT or 1 mM diamide. *Significantly different measured at

p<0.05 relative to fatigued muscles not exposed to either DTT or diamide. n=6. Values are expressed as

means ± SE.

62

Figure 2.7. Correlation of individual protein thiol oxidation and contractile force. Correlation data of actin

(A), myosin (B), troponin (C) and tropomyosin (D) thiol oxidation plotted against mean percentage of initial

force.

To further examine the relationship between the thiol oxidation of the contractile proteins

and contractile force, the four major contractile proteins were measured after 60 min of

rest following the 10 min fatiguing stimulation. The recovery in contractile force was

associated with recovery in the mean thiol oxidation levels of actin (Figure 2.8A), myosin

(Figure 2.8B), troponin (Figure 2.8C) and tropomyosin (Figure 2.8D). However, only actin

(Figure 2.8A) and myosin (Figure 2.8B) were associated with a full recovery. When muscles

were stimulated again with the second bout of fatiguing stimulation, the decrease in mean

thiol oxidation levels of the actin, myosin, troponin and tropomyosin were comparable

63

with the effect of the first 10 min fatiguing stimulation (Figure 2.8A, B, C, D). Taken

together, these data are consistent with thiol oxidation of actin and myosin contributing

to the loss in muscle contractile force.

Figure 2.8. The effect of recovery from fatiguing stimulation on thiol oxidation of individual proteins.

Percentage thiol oxidation of actin (A), myosin (B), troponin (C) and tropomyosin (D) were measured before

first fatigue (rest), after first fatigue, after recovery and after second fatigue. *Significantly different

measured at p<0.05. n=6. Values are expressed as means ± SE.

64

2.4 DISCUSSION

Although there is considerable evidence that muscle fatigue is mediated, at least in part,

by increased oxidative stress, it is still unclear to what extent the effect of oxidative stress

on muscle contractile performance is contributed by an increase in protein thiol oxidation

or protein carbonylation levels. Using the EDL muscle preparation as the experimental

model, this study shows that although fatiguing muscle stimulation increases protein

carbonylation level and thus the degree of oxidative protein damage, it is protein thiol

oxidation rather than protein carbonylation which is most likely to contribute to the ROS-

mediated muscle fatigue associated with submaximal muscle stimulation in vitro. This

study also shows for the first time that the thiol oxidation of several muscle contractile

proteins (actin, myosin, tropoinin and tropopmyosin) increases in response to fatiguing

muscle stimulation.

The results of this study are consistent with earlier studies which have provided evidence

of increased oxidative protein damage (as measured by the carbonyl assay) following

fatiguing stimulation (Figure 2.1C). Indeed, increased oxidative protein damage has been

shown to occur in response to exercise (Reznick et al., 1992; Saxton et al., 1994; Radák et

al., 1998; Barrerio et al., 2006; Pinho et al., 2006; Matsunaga et al., 2008; Veskoukis et al.,

2008; Dubersteina et al., 2009; Agten et al., 2011). Similarly, increased oxidative damage

to muscle proteins has also been reported in response to muscle contraction in vitro

(Dalle-Donne et al., 2001; Canton et al., 2006; Coirault et al., 2007; Dalle-Donne et al.,

2007; Fedorova et al., 2010; Shao et al., 2010). Despite the association between the rise in

65

protein carbonylation level and muscle fatigue, the results of this study show that

elevated protein carbonylation and thus oxidative protein damage can occur without

causing substantial muscle fatigue. Two observations support this contention. First,

muscle contraction in the presence of the thiol reducing agent or thiol thiol oxidising

agent decreased and increased muscle fatigue, respectively, without affecting protein

carbonylation level (Figure 2.3D). Second, the recovery of muscle contractile force

following fatiguing stimulation was not associated with any fall in protein carbonylation

level (Figure 2.2D). However, these results do not necessarily imply that the fall in muscle

contractile performance during exercise cannot involve increased oxidative protein

damage. Data from EDL muscles and skinned muscle fibers experiments showed that the

use of an oxidative damage reagent increased protein carbonylation levels to

approximately 3.5 nmol.mg-1 protein, and this cause a significant decrease in contractile

force (Figure 2.4B), as opposed to the lower increase in protein carbonylation levels to

aproximately 2.3 nmol.mg-1 protein (Figure 2.1C) observed in the EDL muscles subjected

to fatiguing stimulation.

This study provides further evidence that reversible protein thiol oxidation contributes

towards the impairment in contractile performance during muscle fatigue. First, total

protein thiol oxidation level increased in response to fatiguing stimulation, but not

following sustained non-fatiguing stimulation (Figure 2.1B). Second, unlike protein

carbonylation responses, the recovery of muscle contractile performance after fatiguing

stimulation was accompanied by a return of total protein thiol oxidation to pre-

stimulation level (Figure 2.2C). Third, both muscle fatigue and total protein thiol oxidation

66

level were more pronounced when fatiguing muscle stimulation took place in the

presence of the thiol oxidising agent, diamide (Figure 2.3C). Fourth, muscle contraction

performed with the thiol reducing agent, DTT, decreased the magnitude of both muscle

fatigue and total protein thiol oxidation level (Figure 2.3C). These latter findings are

consistent with earlier reports that the non-specific reducing and oxidising agents capable

of affecting the level of protein thiol oxidation also affect muscle fatigue (Reid, 2008).

The novel finding that fatiguing muscle stimulation is associated with an increase in the

thiol oxidation levels of four contractile proteins (actin, myosin, troponin and

tropomyosin) is of interest as the functions of these proteins are known to be sensitive to

oxidative stress. Indeed, previous work has shown that the thiol oxidation of actin

(Hinshaw et al., 1991; Milzani et al., 2000; Dalle-Donne et al., 2003b; Hertelendi et al.,

2008; Prochniewicz et al., 2008b), myosin (Brooke & Kaiser, 1970; Ajtai & Burghardt, 1989;

Root & Reisler, 1992; Tiago et al., 2006; Hertelendi et al., 2008; Prochniewicz et al.,

2008a), troponin (Putkey et al., 1993; Sousa et la., 2006; Pinto et al., 2008; Pinto et al.,

2011), and tropomyosin (Williams & Swenson, 1982) affects their function. However, this

study is the first one to show that muscle contraction to fatigue increases the thiol

oxidation levels of these proteins, thus implying that muscle fatigue caused by oxidative

stress is unlikely to be a consequence of the thiol oxidation of a single protein. It should be

noted that thiol oxidation of other proteins could also be contributing to muscle fatigue in

this study. For example, the function of SR Ca2+ ATPase and the SR Ca2+ release channel

are sensitive to oxidative stress (Scherer & Deamer 1986; Abramanson & Salama, 1989;

Viner et al., 1996; Anzai et al., 2000; Pessah & Feng, 2000; Gutierrez-Martin et al., 2004;

67

Aracena-Parks et al., 2006; Zissimopoulos et al., 2007). Overall, although changes in the

thiol oxidation of several proteins might contribute to fatigue, the extent to which each of

these proteins contributes to muscle fatigue remains to be established.

Despite the evidence that the reversible rise in the thiol oxidation of muscle proteins

contributes to muscle fatigue, it is estimated that only 20 % of the overall loss of

contractile force during fatiguing stimulation was from the contribution of protein thiol

oxidation. This is suggested by the observation that although the addition of DTT returned

the total protein thiol oxidation of fatigued muscles to levels comparable to those of the

resting muscles, it improved contractile force by 20 % during fatiguing stimulation. This

calculation assumes that the extent to which all individual muscle proteins are thiol

oxidised following fatiguing stimulation in the presence of DTT was comparable to resting

muscles. This appears to be a reasonable assumption as the thiol oxidation of the

contractile proteins (actin, myosin, troponin and tropomyosin) was comparable between

DTT-treated fatigued muscles and resting muscles. These findings thus indicate that other

factors such as the depletion of muscle glycogen stores (Ortenblad et al., 2011) and

increased inorganic phosphate levels (Westerblad et al., 2002) play a more important role

in mediating muscle fatigue under the experimental conditions adopted here.

In conclusion, this study corroborates for the first time, the view that oxidative stress

contributes to muscle fatigue through protein thiol oxidation, and that oxidative protein

damage does not necessarily contribute to muscle fatigue. The relationship between

muscle fatigue and protein thiol oxidation appears to be complex as muscle fatigue was

68

associated with an increase in the thiol oxidation level of four contractile proteins (actin,

myosin, troponin and tropomyosin). This raises the interesting possibility that the thiol

oxidation of multiple proteins directly, or indirectly, associated with contractile force,

contribute to ROS-meidated muscle fatigue.

69

CHAPTER 3:


MUSCLE CONTRACTILE PERFORMANCE

AT 34 °C

70

ABSTRACT

The aim of this study was to examine whether protein thiol oxidation plays a role in ROS-

mediated muscle fatigue at 34 °C, and whether the exposure of protein thiol reducing

agent, dithiothreitol (DTT), during fatiguing stimulation at 34 °C affects protein thiol

oxidation and muscle fatigue. Isolated extensor digitorum longus (EDL) muscles from male

ARC(s) mice were incubated in Kreb’s Ringer solution at 34 °C and subjected to either a 10

min or 3.5 min fatiguing stimulation protocol consisting of 500 ms stimulation trains of 50

Hz at 15 s intervals. These muscles were compared to either muscles in a rested state or

muscles stimulated to contract while exposed to DTT. Incubation of the EDL muscles at 34

°C with no contraction was associated with 14.0 ± 0.3 % (n=6) resting mean total protein

thiol oxidation level, which was significantly higher than the 8.0 ± 0.2 % (n=6) found in

vivo. In response to 10 min of fatiguing stimulation, contractile force fell by 51.3 ± 1.2 %

(p<0.05), and this was associated with a 10.8 ± 5.1 % (p<0.05) higher mean total protein

thiol oxidation level and a 20.2 ± 3.8 % (p<0.05) higher myosin mean thiol oxidation level

compared to resting muscles. In response to 3.5 min of fatiguing stimulation, the force

generated by muscles exposed to DTT increased progressively with time and was 87.4 ±

7.8 % (p<0.05) higher compared to stimulated muscles not exposed to DTT. The

improvement in contractility was associated with a 27.3 ± 2.8 % (p<0.05) lower mean total

protein oxidation and 22 ± 0.8 % (p<0.05), 25% ± 3.2 % (p<0.05), 12% ± 2.2 % (p<0.05) and

16% ± 3.2 % (p<0.05) lower actin, myosin, troponin and tropomyosin mean thiol oxidation

levels, respectively, in DTT-treated muscles than DTT-untreated muscles. In conclusion,

the effects of DTT on muscle contractile performance are consistent with muscle

71

contractile force being highly responsive to changes in the thiol oxidation of muscle

proteins at physiological temperature, with proteins such as myosin and maybe actin,

troponin and tropomyosin likely to be involved. Since the thiol oxidation of muscle

proteins in muscle preparations at rest is well above normal, this suggests that DTT is

beneficial to muscle contractile performance in part because it normalises muscle function

rather than reduces muscle fatigue per se.

Keywords: Reactive oxygen species, protein thiols, dithiothreitol, muscle fatigue, 34 °C

72

3.1 INTRODUCTION

Reactive oxygen species (ROS) have been shown to be involved in the development of

muscle fatigue (Ferreira et al., 2009; Kelly et al., 2009; Reardon & Allen, 2009a; Reardon &

Allen, 2009b; Agten et al., 2011). One mechanism whereby ROS might contibute to muscle

fatigue involves the reversible thiol oxidation of muscle proteins. The thiol moiety (-SH) of

the cysteine residues of proteins can undergo reversible covalent reactions with ROS to

form disulfide bonds (Eaton, 2006), which in turn can alter protein structure and function.

Since muscle fatigue is also a reversible process, the reversible thiol oxidation of muscle

proteins is generally considered to explain how ROS contribute to fatigue (Ferreira & Reid,

2008). In this regard, numerous muscle proteins can undergo reversible thiol oxidation

when incubated in vitro with ROS. For instance, the thiol oxidation of actin can disrupt the

organisation of the actin filaments, leading to reductions in the force generating ability of

the sarcomere (Milzani et al., 2000). Peroxynitrite-induced oxidation of myosin has been

shown to inhibit myosin ATPase activity, thus potentially impairing the force generating

transition during the actomyosin ATPase cycle (Tiago et al., 2006). The function of

troponin is also altered when oxidised (Putkey et al., 1993). Finally, the thiol oxidation of

tropomyosin affects the ATPase activity of the actomyosin complex (Williams & Swenson,

1982).

Although the thiol oxidation of muscle proteins can affect their structure and function, it is

important to stress that the aforementioned effects of thiol oxidation on individual muscle

proteins have been based mainly on studies performed on purified proteins, and that

73

none of the studies published so far has examined whether the thiol oxidation of muscle

proteins actually does increase with muscle fatigue. The first direct evidence that ROS-

mediated muscle fatigue might be contributed to by the reversible thiol oxidation of

several muscle proteins including myosin, actin, troponin and tropomyosin was provided

by the results from Chapter 2. However, this work suggested that the reversible thiol

oxidation of muscle proteins plays only a moderate role in muscle fatigue since the

normalisation of the thiol oxidation of muscle proteins with the protein thiol reducing

agent, dithiothreitol (DTT) during fatiguing stimulation improved contractile force

generation by only 20 % (Chapter 2). Nonetheless, one limitation with these findings is

that they were based on experiments performed at 25 °C on isolated extensor digitorum

longus (EDL) from rats. Since this temperature is markedly different from that of working

muscles; which may start as low as 30 °C and increase to 38 °C and even up to 40.8 °C

when heat exhaustion occurs, (Allen et al., 2008), and considering that the importance of

other mechanisms of fatigue (e.g. acidosis) is temperature sensitive (Adams et al., 1991;

Pate et al., 1995; Westerblad et al., 1997), this raises the question of the importance of

the thiol oxidation of muscle proteins contributing to muscle fatigue at physiological

muscle temperature. For this reason, the primary objective of this study was to examine

the extent to which protein thiol oxidation contributes to muscle fatigue at 34 °C, and the

exposure to DTT during fatiguing stimulation at 34 °C decreases the magnitude of both

protein thiol oxidation and muscle fatigue.

74


3.2.1 Animals

Isolated EDL muscle preparations from eight weeks old male ARC(s) mice (n=24) were

adopted as the experimental model. EDL muscles from mice were used to avoid potential

issues with oxygen delivery to the muscle core (Lui et al., 2010), an important issue as

oxygen consumption increases with temperature. All mice used in this study were

obtained from the Animal Resources Centre (Murdoch University, Western Australia) and

their mean weight was 38.1 ± 0.59 g. All animals were kept at a constant room

temperature ranging between 22 and 24 °C, with free access to food and water. All

procedures described here have been approved by the Animal Ethics Committee of the

University of Western Australia.

3.2.2 Intact muscle preparation

All mice were anaesthetised with an intraperitoneal injection of pentobarbital (65 mg/kg

of body weight), and both EDL muscles were carefully excised. While the first muscle was

being tested, the other was left intact inside the animals and removed once

experimentation with the first muscle was completed (≈10 min). Using a braided surgical

silk (Size 5), one end of the muscle tendon was attached to a dual mode force transducer-

servomotor (1200A Intact Muscle Test System, Aurora Scientific Inc, Canada) and the

tendon at the other end of the muscle was attachedto an anchoring hook. The muscle

preparation was then bathed in Kreb’s mammalian Ringer solution (KRS) (Composition:

75

NaCl, 121 mM; KCl, 5·4 mM; MgSO4, 1.2 mM; NaHCO3, 25 mM; HEPES, 5 mM; glucose,

11.5 mM; CaCl2, 2.5 mM at pH 7.4) gassed with 5 % CO2 and 95 % O2 and maintained at 34

°C throughout the experiment. The muscle was then stimulated with single twitches to

attain the optimum length producing the highest force possible before experimentation.

Force responses were elicited using a 701B muscle stimulator (Aurora Scientific Inc,

Canada). Next, maximal force responses were obtained in response to supramaximal

stimulation pulses (pulse width: 0.3 ms) delivered at 150 Hz for 500 ms with a 2-min rest

between each stimulation to stabilise the force responses. The muscles were then given a

10 min rest before the commencement of the stimulation. The two submaximal fatiguing

stimulation protocols used in the experiments on mice were pre-determined in the

preliminary experiments. They involved exposure to stimulation pulses delivered (80 V

and 1000 mA) at 50 Hz for 500 ms with a 15 s interval between contractions for either 3.5

min or 10 min.

3.2.3 Fatiguing stimulation protocol

In order to study the effect of fatiguing stimulation on protein thiol oxidation at 34 °C,

isolated muscles were subjected to a 10 min fatiguing stimulation. Measurement made in

these muscles were compared with resting muscles at 34 °C, but not stimulated to

contract. After 10 min of fatiguing stimulation or rest, all muscles were frozen using

aluminium clamps pre-cooled in liquid nitrogen and stored at -80 °C. In order to study the

effect of the protein thiol reducing agent, DTT on fatiguing stimulation and protein thiol

oxidation at 34 °C, isolated muscles were maintained in KRS with 25 mM DTT (n=6; Sigma

76

Chemical, Australia) during the initial 10 min resting period and an additional 25 mM of

DTT was added into the bath immediately before the commencement of the 3.5 min

fatiguing stimulation to replace the oxidised DTT dose with a freshedly reduced DTT. A 3.5

min fatiguing stimulation time was adopted because preliminary experiments showed that

two thirds of the fall in contractile force took place within 3.5 min, with a slow rate of fall

in contractile force afterwards. Measurements made in these muscles were compared

with fatigued muscles not exposed to DTT. After 3.5 min of fatiguing stimulation, all

muscles were immediately frozen using aluminum clamps pre-cooled in liquid nitrogen

and stored at -80 ˚C until analysed. In addition, muscles maintained in KRS with and

without DTT were also frozen before the commencement of fatiguing stimulation.












77














3.2.5 Electrophoretic separation and assay of ‘2 Tag’-labeled protein








78










performed using either a Student’s paired t test or a one way ANOVA followed by Fisher

LSD posteriori test , with significance at P<0.05. Sample sized is denoted by n.

79

3.3 RESULTS

3.3.1 The effect of fatiguing stimulation on protein thiol oxidation at 34 °C

To establish whether protein thiol oxidation increased in response to fatiguing muscle

stimulation at a physiological temperature, isolated EDL muscles were subjected to a

fatiguing stimulation for 10 min at 34 °C. Mean contractile force fell by approximately 51

% (Figure 3.1A) following the fatiguing stimulation, and this was associated with an 11 %

increased in mean total protein thiol oxidation level (Figure 3.1B). There was no difference

in total protein thiols (Figure 3.1C), indicating that the increase in protein thiol oxidation

level likely represents increased oxidation of reduced protein thiols to oxidised protein

disulfides. Incubation of the EDL muscles at 34 °C with no stimulation was associated with

resting mean total protein thiol oxidation level of 14.0 ± 0.3 % (n=6) which was

significantly higher than the in vivo mean total protein thiol oxidation level of 8.0 ± 0.2 %

(n=6).

80

Figure 3.1. The effect of fatiguing stimulation on protein thiol oxidation at 34 °C. Isolated EDL muscles

were fatigued by stimulating at 50 Hz for 500 ms with a 15 s interval between contractions for a period of 10

min. Following 10 min of stimulation, mean percentage of initial force (A), percentage of oxidised protein

thiols (B) and total protein thiols (C) were measured. *Significantly different measured at p<0.05 relative to

resting muscle incubated for 10 min without stimulation. n=6. Values are expressed as means ± SE.

3.3.2 The effect of the protein thiol reducing agent, DTT on fatiguing stimulation and

protein thiol oxidation at 34 °C

In order to provide further evidence that protein thiol oxidation contributed to the loss in

contractile force at 34 °C, fatiguing stimulation was also performed in the presence or

absence of the protein thiol reducing agent, DTT. Following 3.5 min of fatiguing

stimulation in the absence of DTT, the final contractile force of fatigued muscles

81

decreased to 66 % of the initial mean contractile force (Figure 3.2B). The mean total

protein thiol oxidation level of fatigued muscles not exposed to DTT was 15 % higher than

pre-fatigue level (Figure 3.2C). In contrast, fatiguing stimulation for 3.5 min in the

presence of DTT increased the final contractile force of fatigued muscles to 124 % of the

initial contractile force (Figure 3.2B). The mean total protein thiol oxidation level of

fatigued muscles exposed to DTT was 12 % lower than the pre-fatigue level (Figure 3.2C).

The initial 10 min incubation with DTT did not affect pre-fatigue mean total protein thiol

oxidation level (Figure 3.2C). However, the effect of adding the second dose of DTT before

the start of fatiguing stimulation on contractile force was evident after 0.5 min (Figure

3.2A).

The 87 % improvement in contractile force in fatigued muscles exposed to DTT was

associated with a 27 % lower mean total protein thiol oxidation level than the fatigued

muscles not exposed to DTT (Figure 3.2C). Given there was no difference in total protein

thiols (Figure 3.2D), the decrease in protein thiol oxidation likely represents decreased

oxidation of reduced protein thiols to oxidised protein thiols. These data therefore, show

that although incubation with DTT did not affect pre-fatigue protein thiol oxidation level,

exposure of DTT during fatiguing stimulation prevented the loss in contractile force, a

finding consistent with protein thiol oxidation affecting muscle contractile force.

82

Figure 3.2. The effect of protein thiol reducing agent, DTT on fatiguing stimulation and protein thiol

oxidation at 34 °C. Isolated EDL muscles were fatigued by stimulating at 50 Hz for 500 ms with a 15 s interval

between contractions while exposed to 2 doses of 25 mM DTT for a period of 10 min before and at the start

of stimulation or not exposed to DTT. Mean percentage of initial force (B), percentage of oxidised protein

thiols (C) and total protein thiols (D) were measured at the start of fatigue in the absence or presence of DTT

and after fatigue in the absence and presence of DTT. Percentage of mean force relative to initial force of

fatigue in the absence of DTT (□) and fatigue in the presence of DTT (△) were also measured every 0.5 min

during the 3.5 min fatiguing stimulation (A). *Significantly different measured at p<0.05 relative to fatigued

muscles not exposed to DTT. n=6. Values are expressed as means ± SE.

83

3.3.3 Thiol oxidation of contractile proteins and the loss of contractile force at 34 °C

In order to determine whether the thiol oxidation of individual contractile proteins might

contribute to the loss of contractile force during the 10 min fatiguing stimulation at 34 °C,

thiol oxidation levels were examined in four important contractile proteins (actin, myosin,

troponin and tropomyosin). Following 10 min of fatiguing stimulation, the mean thiol

oxidation level of myosin was 20 % higher (Figure 3.3B), with no change in actin, troponin

and tropomyosin mean thiol oxidation levels (Figure 3.3A, C, D). These data are consistent

with the thiol oxidation of myosin contributing to the loss of contractile force during the

10 min of fatiguing stimulation at 34 °C.

84

Figure 3.3. The effect of fatiguing stimulation on thiol oxidation of individual proteins at 34 °C. Percentage


muscles were stimulated to fatigue at 50 Hz for 500 ms with a 15 s interval between contractions for a

period of 10 min. *Significantly different measured at p<0.05 relative to resting muscles incubated for 10

min without stimulation. n=6. Values are expressed as means ± SE.

An investigation of whether thiol oxidation of individual contractile proteins could be

affected by exposure to DTT during the 3.5 min fatiguing stimulation at 34 °C was carried

out. The mean thiol oxidation levels of actin, myosin, troponin and tropomyosin were 58

% (Figure 3.4A), 37 % (Figure 3.4B), 36 % (Figure 3.4C) and 33 % (Figure 3.4D) higher,

respectively, in muscles incubated at 34 °C relative to in vivo muscle.

85

Following 3.5 min of fatiguing stimulation in the absence of DTT, mean thiol oxidation

levels of actin and myosin were 7 % (Figure 4A) and 13 % (Figure 3.4B) higher, respectively

compared to pre-fatigue levels. In contrast, fatiguing stimulation for 3.5 min in the

presence of DTT was associated with a 15 % (Figure 3.4A), 11 % (Figure 3.4B), 8 % (Figure

3.4C) and 10 % (Figure 3.4D) lower mean thiol oxidation levels of actin, myosin, troponin

and tropomyosin, respectively, compared to pre-fatigue levels.

The 87 % improvement in contractile force of fatigued muscles exposed to DTT was

associated with a 22 % (Figure 3.4A), 25 % (Figure 3.4B), 12 % (Figure 3.4C) and 16 %

(Figure 3.4D) decrease in mean actin, myosin, troponin and tropomyosin thiol oxidation

levels, respectively compared to fatigued muscles not exposed to DTT. Taken together,

these data suggest that increased protein thiol oxidation and thiol oxidation of myosin

contributed to the loss in contractile force during fatigue at 34 °C.

86

Figure 3.4. The effect of protein thiol reducing agent, DTT on thiol oxidation of individual proteins at 34°C.

Percentage thiol oxidation of actin (A), myosin (B), troponin (C) and tropomyosin (D) were measured in vivo,

before the start of fatigue, after fatigue in the absence of DTT and after fatigue in the presence of DTT.

*Significantly different measured at p<0.05. n=6. Values are expressed as means ± SE.

87

3.4 DISCUSSION

The first direct evidence that ROS-mediated muscle fatigue associated with submaximal

contraction in vitro is mediated, in part, by the reversible thiol oxidation of muscle

contractile proteins was provided in Chapter 2. However, muscle contraction performed in

the presence of thiol reducing agent, DTT at levels that normalised the thiol oxidation of

muscle proteins improved muscle contractile performance only to a moderate extent

(Chapter 2), suggesting that the reversible thiol oxidation of contractile proteins plays only

a moderate role in muscle fatigue. Since these findings were based on experiments

performed at 25 °C rather than at a more physiological temperature, and that other

mechanisms of muscle fatigue, such as those associated with acidosis, have been reported

to play little role at physiological temperature (Adams et al., 1991; Pate et al., 1995;

Westerblad et al., 1997). The issue of whether reversible protein thiol oxidation is an

important contributor of muscle fatigue at physiological temperature was raised.

Using EDL muscle preparations from mice as the experimental model, this study shows

that high levels of DTT not only prevents a fall in the contractile force of the muscles

tested at physiological temperature, but actually results in a progressive increase in

muscle contratile force, reaching 87 % more than that attained in fatigued muscles not

exposed to DTT (Figure 3.2B). Since the intensity of the contraction protocol was

submaximal, it is unclear whether the large DTT-mediated improvement in contractile

force resulted from an increase in maximal force or a shift in the force-frequency toward

higher frequencies due to decreased fusion of twitch (or individual) contractions. The

88

marked benefit of such DTT treatment on muscle contractile performance was associated

with the absence of a rise in the total thiol oxidation of muscle proteins (Figure 3.2C),

including myosin, actin, tropomyosin and troponin (Figure 3.4) compared to no DTT

treatment. However, since the thiol oxidation of muscle proteins were above normal in

resting muscles at 34 °C compared to in vivo muscles (freshed dissected), these findings do

not allow one to evaluate the importance of protein thiol oxidation in ROS-mediated

muscle fatigue since part of the effect of DTT treatment is to normalise muscle function

rather than preventing muscle fatigue. Nevertheless, irrespective of the extent to which

DTT affects muscle fatigue, this is the first study to show that changes in the thiol

oxidation of muscle proteins can be associated with major effects on muscle contractile

performance at 34 °C.

The findings of this study suggest that muscle contractile force at physiological

temperature is highly sensitive to the thiol oxidation of muscle proteins. Firstly, this is

supported by the observation that total protein thiol oxidation level increased in response

to fatiguing stimulation (Figure 3.1B). Secondly, acute exposure to the thiol reducing

agent, DTT improved muscle contractile force with time (Figure 3.1A) while decreasing the

thiol oxidation levels of muscle proteins including that of myosin, actin, troponin and

tropomyosin (Figure 3.4). These results not only corroborate earlier findings from Chapter

2, that reversible protein thiol oxidation affects muscle contractile force (Chapter 2), but

also are consistent with the many earlier reports that nonspecific reducing agents can

affect muscle contractile performance (Reid, 2008). It is important to stress, however, that

although many of these studies have implicated oxidative stress as a mediator of muscle

89

contractile performance, none of these studies have examined whether reversible thiol

oxidation of muscle proteins has the capacity to affect muscle contractile force.

The marked increase in the thiol oxidation level of muscle proteins in response to fatiguing

stimulation raises the obvious issue of the identity of the proteins involved. Close

inspection of the responses of individual protein bands separated by gel electrophoresis

suggests that the thiol oxidation of myosin might contribute to muscle fatigue in the in

vitro model investigated here at a more physiological temperature, whereas the absence

of changes in actin, troponin and tropomyosin thiol oxidation suggests that these proteins

are not involved. However, since DTT treatment and associated improvement in muscle

contractile performance were accompanied by a fall in the thiol oxidation levels of

myosin, actin, troponin, and tropomyosin, it is possible that the thiol oxidation of actin,

troponin and tropomyosin also affects muscle contractile force. This interpretation that

myosin and other contractile proteins affect muscle contractile force is consistent with

earlier reports that the thiol oxidation of myosin causes a marked decrease in myosin

ATPase activity (Root & Reisler, 1992) due to a decrease in the fraction of myosin heads

undergoing the force-generating transition step during the actomyosin ATPase cycle

(Tiago et al., 2006). Others have also shown that changes in the thiol oxidation of actin,

troponin and tropomyosin affect not only their structures, but also their functions (Putkey

et al., 1993; Williams & Swenson, 1982; Milzani et al., 2000).

The benefits of DTT treatment on muscle contractile performance suggests that the thiol

oxidation of muscle proteins might play some role in ROS-mediated muscle fatigue at

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physiological temperature. However, a higher than normal thiol oxidation level of muscle

proteins prior to contraction together with the progressive temporal increase in muscle

contractile force exposed to DTT during stimulation suggests that DTT is beneficial, in part,

because it normalises muscle function rather than only opposing muscle fatigue. This is

likely the case here as many studies have reported an increased production of ROS with

increased muscle temperature (Arbogast & Reid, 2004; Edwards et al., 2007), and a

marked decrease in muscle performance above 35 °C (Segal et al., 1985, 1986; Pedersen

et al., 2003). In addition, there is strong evidence that the generation of ROS explains the

reduced longevity of in vitro skeletal muscle preparations at temperatures ranging

between 37-43 °C (Zuo et al., 2000; Edwards et al., 2007; van der Poel et al., 2002;

Arbogast & Reid, 2004; Moopanar & Allen, 2005; van der Poel et al., 2007).

These findings thus highlight some of the limitations with using isolated muscle

preparations as experimental models to investigate ROS-mediated muscle fatigue at

physiological temperature due to the associated abnormal increase in protein thiol

oxidation level compared to muscle in vivo. Investigating muscle fatigue at a lower and

unphysiological temperature is clearly not a suitable alternative despite being associated

with normal protein thiol oxidation (Chapter 2). This is because of the possibility that the

importance of some of the mechanisms of muscle fatigue is temperature sensitive. This is

best illustrated by the finding that acidosis is an important mediator of fatigue in muscles

incubated at room temperature, but only plays a marginal role at physiological

temperature (Adams et al., 1991; Pate et al., 1995; Westerblad et al., 1997). Thus, future

studies investigating the physiological importance of the thiol oxidation of muscle proteins

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in muscle fatigue in vitro requires the introduction of better models that do not affect

protein thiol oxidation at physiological temperature.

In conclusion, the effects of DTT on muscle contractile performance are consistent with

muscle contractile force being highly responsive to changes in the thiol oxidation of

muscle proteins at physiological temperature, with proteins such as myosin and maybe

actin, troponin and tropomyosin likely to be involved. Since the thiol oxidation of muscle

proteins in muscle preparations at rest is well above normal, these results at physiological

temperature are consistent with the possibility that DTT is beneficial in part, to muscle

contractile performance. This is because it normalises muscle function rather than reduces

muscle fatigue per se, thus leaving without a definitive answer the importance of the role

played by protein thiol oxidation in ROS-mediated muscle fatigue at physiological

temperature. This chapter, however, again clearly shows for the first time that the thiol

oxidation of muscle proteins is associated with large changes in muscle contractile

performance. This raises the possibility that protein thiol oxidation has the potential to be

an important mediator of ROS-mediated muscle fatigue, and is worth further

investigation.

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CHAPTER 4:


MUSCLE CONTRACTILE FUNCTION IN

INS2AKITA DIABETIC MICE

93

ABSTRACT

The aim of this study was to examine whether the thiol oxidation level of muscle proteins

is increased before and after muscle contraction in diabetic Ins2Akita mice, and whether

this is associated with lower muscle contractile performance. Isolated extensor digitorum

longus muscles from male Ins2Akita mice were subjected to a fatiguing stimulation that

consisted of 10 min of submaximal tetanic contraction of 500 ms duration at 15 s intervals

with a stimulation frequency of 50 Hz. These muscles were compared with muscles from

non-diabetic mice subjected to the same fatiguing stimulation. Initial specific force did not

differ between the different muscles; however the initial mean thiol oxidation levels of

total proteins, actin and tropomyosin in muscles from diabetic mice compared to non-

diabetic mice were 16.5 ± 0.2 % (p<0.05), 16.3 ± 0.4 % (p<0.05) and 7.6 ± 0.4 % (p<0.05)

higher, respectively. Following 10 min of fatiguing stimulation, contractile force of muscles

from diabetic mice fell by 27.4 ± 1.2 % (p<0.05) more than non-diabetic mice and this was

associated with a 21.8 ± 0.3 % (p<0.05), 15.5 ± 0.5 % (p<0.05) and 18.9 ± 0.4 % (p<0.05)

higher mean thiol oxidation of total proteins, actin, and tropomyosin in muscles from

diabetic mice compared to non-diabetic mice, respectively. It is concluded that initial

specific force of the muscles from diabetic mice compared to non-diabetic mice is not

reduced despite higher total protein thiol oxidation in muscles from diabetic mice.

However, the higher thiol oxidation level of myosin in muscles from diabetic mice exposed

to fatiguing stimulation might explain the lesser resistance of these muscles to fatigue.

Keywords: Reactive oxygen species, protein thiols, Ins2Akita mice, muscle fatigue

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4.1 INTRODUCTION

Regular exercise provides a number of well documented health benefits for individuals

with Type 1 diabetes mellitus (T1DM) including improvements in cardiovascular function

(Choi & Chisholm, 1996; Fuchsjager-Mayrl et al., 2002), blood lipid profile (Lehmann et al.,

1997; Laaksonen et al., 2000), weight control (Lehmann et al., 1997) and quality of life

(Zoppini et al., 2003). Unfortunately, people with T1DM have reduced cardiopulmonary

fitness (Raev, 1994; Niranjan et al., 1997; Komatsu et al., 2005, Nadeau et al., 2010)

compared to their healthy non-diabetic peers. This is unfortunate considering that

premature cardiovascular disease significantly shortens the average lifespan of individuals

with T1DM (Libby et al., 2005).

One factor that has the potential to limit the ability of T1DM individuals to engage in an

active life style compared to their peers is their reduced motor function. Diabetic

peripheral neuropathies are one cause of this reduced motor function as isokinetic muscle

strength in long-term T1DM patients is reduced to an extent that increases with the

severity of neuropathy (Andersen et al., 1996; Andersen et al., 1997; Andreassen et al.,

2006). Motor function is also reduced in neuropathy-free T1DM individuals (Andersen,

1998; Andersen et al., 2005; Almeida et al., 2008). These individuals have lower motor

conduction velocities, and the onset of fatigue is much earlier than non-diabetic

individuals (Almeida et al., 2008). Finally, young T1DM individuals have reduced resistance

to skeletal muscle fatigue compared to non-diabetic individuals (Ramiere et al., 1993).

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There is indirect evidence that the poor muscle contractile performance in diabetes might

be related to the increase in oxidative stress associated with this disease (Nishikawa et al.,

2001; Evans et al., 2005; Kaneki et al., 2007), a condition characterised by an imbalance

between the production of reactive oxygen species (ROS; e.g. superoxide, hydroxyl

radicals) and removal by antioxidant defences. For instance, marked increase in ROS

production has also been reported during exercise and pharmacologic antioxidants has

been shown to improve muscle fatigue (Reid, 2008). These evidences together with the

results of Chapters 2 and 3 that changes in the thiol oxidation level of muscle proteins can

acutely affect muscle contractile performance raise the question of whether impaired

contractile performance associated with diabeties might be contributed, at least in part,

by an increase in the thiol oxidation level of skeletal muscle proteins. Using isolated

extensor digitorum longus (EDL) muscle preparations from diabetic Ins2Akita mice and

wild type non-diabetic mice as controls, the primary objective of this study was to

examine whether the thiol oxidation levels of muscle proteins was increased before and

after muscle contraction in diabetic mice, and whether this was associated with lower

muscle contractile performance.

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4.2.1 Animals

Male Ins2Akita mice (n=5) bred on a C57BL/6J background and non-diabetic C57BL/6Js

mice (n=8) at 40 weeks of age and non-diabetic C57BL/6Js mice (n=10) at 13 weeks of age

were housed under controlled temperature of 22 °C with a 12:12hr light/cycle (lights on at

0600hr), with food and water available ad libitum. Ins2Akita mice were used as they

undergo mutation resulting in a single amino acid substitution in the insulin 2 gene, which

causes misfolding of the insulin protein leading to significant hyperglycemia, as early as

four weeks of age (Barber et al., 2005). All experiments were approved by the Animal

Ethics Committee of the University of Western Australia.

4.2.2 Intact muscle preparation

Following a 12 hr overnight fast, mice were anaesthetised with an intraperitoneal

injection of pentobarbital (65 mg/kg of body weight). Blood glucose and glycated

haemoglobin levels (HbA1c) were then immediately measured using Accu-Check Performa

Blood Glucose Meter (Roche, Sydney, Australia) and DCA 2000®+ Analyser (Siemens, USA),

respectively. Next, both EDL muscles were carefully removed one after the other for

experiments. While the first muscle was being tested, the other was left intact inside the

animals and removed once experimentation with the first muscle was completed (≈10

min). After removing the muscles, one end of the muscle tendon was attached to a dual

mode force transducer-servomotor (1200A Intact Muscle Test System, Aurora Scientific

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Inc, Canada) and the other tendon to an anchoring hook using a braided surgical silk (Size

5). The muscle preparation was bathed in Kreb’s mammalian Ringer solution (KRS)

(Composition: NaCl, 121 mM; KCl, 5·4 mM; MgSO4, 1.2 mM; NaHCO3, 25 mM; HEPES, 5

mM; glucose, 11.5 mM; CaCl2, 2.5 mM at pH 7.4) gassed with 5 % CO2 and 95 % O2 at 25 °C

throughout the experiment. The muscle was then stimulated with single twitches to attain

the optimum length producing the highest force possible before experimentation. Force

responses were then elicited using a 701B muscle stimulator (Aurora Scientific Inc,

Canada). Next, maximal force responses were obtained in response to supramaximal

stimulation pulses (pulse width: 0.3 ms) delivered at 150 Hz for 500 ms with a 2 min rest

between each stimulation to stabilise the force responses. The muscles were then given a

10-min rest before the commencement of the stimulation. The submaximal stimulation

protocol used in the experiments on mice was pre-determined in the preliminary

experiments. It involved exposure to stimulation pulses (80 V and 1000 mA) delivered at

50 Hz for 500 ms with a 15 s interval between contractions for 10 min.

4.2 3 Fatiguing stimulation protocol

For all experiments, isolated muscles were maintained for 10 min in KRS before

stimulation. In order to study the effect of contraction on the protein thiol oxidation of

skeletal muscles, isolated muscles from diabetic and non-diabetic mice were subjected to

a 10 min fatiguing stimulation. Immediately after stimulation, all muscles were frozen

using an aluminium clamp pre-cooled in liquid nitrogen and stored at -80 °C. In addition,

resting EDL muscles obtained from the contralateral leg of each of the tested animals

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were also frozen before the commencement of the fatiguing stimulation. In order to study

the effect of protein thiol modification on muscle contraction and protein thiol oxidation,

isolated muscles from non-diabetic mice were maintained for 10 min in KRS with 15 mM

dithiothreitol (DTT) before being subjected to a single contraction that consisted of

submaximal tetanic contraction of 500 ms duration at a stimulation frequency of 50 Hz.

These muscles were compared with contracting muscles not exposed to DTT. After

contraction, all muscles were frozen using aluminium clamps pre-cooled in liquid nitrogen

and stored at -80 °C.














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4.2.5 Electrophoretic separation and assay of ‘2 Tag’-labeled protein










100








performed using a Student’s unpaired t test or a two way ANOVA followed by Fisher LSD

posteriori test, with significance at P<0.05. Sample sized is denoted by n.

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4.3 RESULTS

4.3.1 Body mass, fasting plasma glucose level and fasting glycated hemoglobin of non-

diabetic mice and diabetic mice

The mean body mass of non-diabetic mice was 55 % (Figure 4.1A) higher than that of

diabetic mice. Mean fasting blood glucose levels and mean HbA1c levels were 137 % (Figure

4.1B) and 84 % (Figure 4.1C) higher, respectively, in the diabetic mice compared to the

non-diabetic mice. These results indicate that the Ins2akita mice used in this study

showed clear signs of diabetes.

Figure 4.1. Body mass, fasting plasma glucose level and fasting glycated hemoglobin of non-diabetic mice

(CON) and diabetic mice (DM). Body mass (A), fasting plasma glucose level (B) and fasting glycated

haemoglobin (C) in non-diabetic mice (n=8) and diabetic mice (n=5). *Significantly different measured at

p<0.05. Values are expressed as means ± SE.

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4.3.2 The relationship between protein thiol oxidation of non-diabetic mice and diabetic

mice on associated muscle functions

Initial specific force at the start of fatiguing stimulation did not differ between the muscles

from non-diabetic mice and diabetic mice (Figure 4.2A). Initial pre-fatigue mean total

protein thiol oxidation level was 16 % higher in the muscles of diabetic mice than non-

diabetic mice (Figure 4.2C). Following 10 min of fatiguing stimulation, contractile force of

the muscles from diabetic mice fell by 27 % more than muscles from non-diabetic mice

(Figure 4.2B), and mean total protein thiol oxidation levels were 10 % and 15 % higher in

muscles of non-diabetic mice and diabetic mice, respectively (Figure 4.2C). The 27 % lower

contractile force in muscles from diabetic mice was associated with a 22 % (Figure 4.2C)

higher mean total protein thiol oxidation level. There was no difference in total protein

thiols (Figure 4.2D) so the increase in protein thiol oxidation level likely represents

increased conversion of reduced protein thiols to oxidised protein thiols.

The resting mean total protein thiol oxidation levels of the muscles from non-diabetic

mice and diabetic mice incubated at 25 °C with no contraction were 15.7 ± 0.3 % (n=8) and

18.3 ± 0.3 % (n=5), which were significantly higher than 7.5 ± 0.2 % (n=5) and 10.7 ± 0.3 %

(n=5) measured in in vivo muscles from non-diabetic mice and diabetic mice, respectively,

frozen immediately after dissection, indicating that incubation resulted in an increase in

protein thiol oxidation level.

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Figure 4.2. The relationship between protein thiol oxidation of non-diabetic mice (CON) and diabetic mice

(DM) on associated muscle functions. Isolated EDL muscles (CON, n=8; DM, n=5) were fatigued by

stimulating at 50 Hz for 500 ms with a 15 s interval between contractions for a period of 10 min. At the start

of the stimulation, initial specific force (A) was measured. Following 10 min of stimulation, mean percentage

of initial force (B), percentage of protein thiols oxidised (C) and total protein thiols (D) were measured.

*Significantly different measured at p<0.05 relative to pre-fatigue muscles (CON, n=8; DM, n=5). Values are

expressed as means ± SE.

4.3.3 Thiol oxidation of contractile proteins and the loss of contractile force

In order to identify the proteins likely to be affected by muscle contraction to fatigue,

muscle extracts labelled for protein thiol oxidation were subjected to gel electrophoresis

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to examine the behavior of actin, myosin, troponin and tropomyosin. Immediately before

the start of fatiguing stimulation, the mean thiol oxidation levels of actin and tropmyosin

were 16 % (Figure 4.3A) and 8 % (Figure 4.3D) higher, respectively, in muscles from

diabetic mice than non-diabetic mice. There were no significant differences in the mean

thiol oxidation levels of myosin (Figure 4.3A) and troponin (Figure 4.3C) before the start of


Following 10 min of fatiguing stimulation, the mean thiol oxidation levels of actin in

muscles from non-diabetic mice and diabetic mice were 11 % and 10 % (Figure 4.3A)

higher compared with pre-fatigue levels, respectively. The mean thiol oxidation levels

myosin in muscles from non-diabetic mice and diabetic mice were 13 % and 28 % (Figure

4.3B) higher compared with pre-fatigue levels, respectively. There were no significant

differences in the mean thiol oxidation levels of troponin (Figure 4.3C) and tropomyosin

(Figure 4.3D) following 10 min of fatiguing stimulation compared with pre-fatigue levels.

The 27 % lower contractile force in muscles from diabetic mice (Figure 4.2B) was

associated with 15 % (Figure 4.3A) and 19 % (Figure 4.3B) higher actin and myosin mean

thiol oxidation levels respectively compared to muscles from non-diabetic mice after the


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Figure 4.3. The effect of fatiguing stimulation on thiol oxidation of individual proteins in non-diabetic mice

(CON) and diabetic mice (DM). Percentage thiol oxidation of actin (A), myosin (B), troponin (C) and

tropomyosin (D) were measured before and after EDL muscles (CON, n=8; DM, n=5) were fatigue by

stimulating at 50 Hz for 500 ms with a 15 s interval between contractions for a period of 10 min.

*Significantly different measured at p<0.05. Values are expressed as means ± SE.

4.3.4 The effect of protein thiol modification on muscle contraction and protein thiol

oxidation

Given that muscle incubation in KRS for 10 min was associated with an increase in protein

thiol oxidation level, the effect of the protein thiol reducing agent, DTT, on contractile

106

force was examined. To this end, EDL muscles from non-diabetic mice were titrated with

different levels of DTT, and 15 mM of DTT was found to significantly increase the specific

contractile force. The mean contractile force of muscles exposed to DTT was 48 % higher

than those not exposed to antioxidant (Figure 4.4A). The improvement in contractile force

was associated with a 34 % lower mean total protein thiol oxidation level (Figure 4.4B).

There was no difference in total protein thiols (Figure 4.4C).

The stimulatory effect of DTT on muscle contraction was accompanied by a significant

change in the thiol oxidation levels of the four major contractile proteins examined here.

The improvement in contractile force of muscles incubated with DTT was associated with

33 % (Figure 4.5A), 33 % (Figure 4.5B), 31 % (Figure 4.5C) and 30 % (Figure 4.5D) decrease

in the mean thiol oxidation levels of the actin, myosin, troponin and tropomyosin,

respectively compared to muscles not exposed to with antioxidant.

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Figure 4.4. The effect of protein thiol modification on single muscle contraction and protein thiol

oxidation. Isolated EDL muscles from non-diabetic mice were stimulated once at 50 Hz for 500 ms while

exposed to 15 mM DTT. Following the single contraction, specific force (A), percentage of oxidised protein

thiols (B) and total protein thiols (C) were measured. *Significantly different measured at p<0.05 relative to

contracting muscles not exposed to DTT. n=5. Values are expressed as means ± SE.

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Figure 4.5. The effect of protein thiol modification on thiol oxidation of individual proteins. Percentage


muscles from non-diabetic mice were stimulated once at 50 Hz for 500 ms while exposed to 15 mM DTT.

*Significantly different measured at p<0.05 relative to contracting muscles not exposed to DTT. N=5. Values

are expressed as means ± SE.

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4.4 DISCUSSION

This study examined whether the lower muscle strength and lower resistance to muscle

fatigue associated with diabetes might be contributed, at least in part, by a higher than

normal thiol oxidation level of muscle proteins. In order to determine whether this is the

case, the thiol oxidation levels of muscle proteins and muscle contractile function were

compared between diabetic Ins2Akita mice and wild type non-diabetic mice. Contrary to

expectations, there was no difference in maximal specific force despite the higher total

mean protein thiol oxidation in muscles from diabetic mice. However, muscles from

diabetic mice were less resistant to fatigue, and had a higher mean total protein thiol

oxidation level compared to muscles from non-diabetic mice.

The absence of a difference in initial specific force between the non-diabetic mice and

diabetic mice (Figure 4.2A) does not support past findings that diabetes impairs muscle

strength. It has been reported that isokinetic muscle strength is reduced in T1DM patients

(Andersen et al., 1996; Andersen et al., 1997; Andreassen et al., 2006) as well as in other

animal models of diabetes. In particular, insulin-untreated STZ-diabetes in rats results in a

fall in the specific tension of the EDL (Cotter et al., 1989; Cotter et al., 1993; McGuire &

MacDermott, 1999). Similarly, alloxan-induced diabetes reduces the tetanic force of the

EDL (Paulus & Grossie, 1983). Finally, skinned-fibers from the EDL of STZ-diabetic rats have

lower maximal specific tension compared to those of non-diabetic animals (Stephenson et

al., 1994).

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However, the absence of difference in initial specific force might be explained, at least in

part, by the possible differences between muscle preparations from these mice. These

preparations were exposed to similar physiological glucose concentrations, unlike the

different glucose concentrations to which the muscles are exposed in the intact animals.

Exposing the muscles from diabetic mice to the high glucose concentrations normally

found in these mice would have possibly affected their contractile function compared to

muscles from non-diabetic animals. This is suggested by the observation that acute

hyperglycaemia in healthy individuals with T1DM is associated with lower maximal

isometric voluntary contraction compared to euglycaemia (Andersen et al., 2005). It is

important to stress, however, that some marked differences in muscle specific force were

expected between diabetic and non-diabetic animals even when exposed to matching

glucose concentrations because studies performed by others on isolated muscle

preparation from rats reported impaired muscle tension in STZ-diabetic compared to non-

diabetic animals despite being tested a identical glucose levels (Paulus & Grossie, 1983;

Stephenson et al., 1994).

The finding that total protein thiol oxidation from diabetic mice was increased compared

to non-diabetic mice (Figure 4.2C) is consistent with the observations of others that

diabetes is associated with an increase in oxidative stress (Nishikawa et al., 2001; Evans et

al., 2005; Kaneki et al., 2007). However, since the increase in total protein thiol oxidation

was not associated with lower muscle maximal specific tension in muscles from diabetic

mice compared to non-diabetic mice, this would appear not only to refute the hypothesis

of a link between muscle protein thiol oxidation in diabetes and impaired muscle tension,

111

but would also suggest that muscle contractile performance is unresponsive to muscle

protein thiol oxidation. This latter interpretation, however, is inconsistent with the

findings in Chapters 2 and 3 which showed a link between muscle contractile force and

the thiol oxidation of muscle proteins, and the findings here that muscle tension increases

and the thiol oxidation levels of muscle proteins falls in the presence of DTT.

The observation that elevated total protein thiol oxidation level of muscles from diabetic

mice was not accompanied by a fall in maximal specific force could be explained on the

grounds that although total protein thiol oxidation level in the muscles of diabetic mice

was higher than muscle of non-diabetic mice, this might not be the case at the level of

specific proteins known to affect muscle contractile function. That this might be the case

here, is suggested by the findings in pre-fatigue muscles that although the thiol oxidation

levels of actin and tropomyosin were higher in fatigued muscles of diabetic mice (Figure

4.3A,D), no differences were detected in the thiol oxidation levels of myosin and troponin

(Figure 4.3B,C) in pre-fatigue and fatigued muscles of diabetic mice. Given the role played

by myosin in muscle contraction and previous findings that the oxidation of myosin

inhibits myosin ATPase activity thus potentially impairing the force generating transition

during the actomyosin ATPase cycle (Tiago et al., 2006), the absence of a difference in the

thiol oxidation levels of myosin and other contractile proteins between muscles from non-

diabetic mice and diabetic mice might explain their similar maximal specific force.

Arguably, this interpretation holds as long as myosin plays a more important role in

determining contractile force compared with other contractile proteins such as actin,

troponin and tropomyosin. Although further work is required to test this interpretation,

112

the findings with myosin highlight the importance of examining the response to individual

proteins compared to total proteins.

The finding that resistance to muscle fatigue is lower in muscles in diabetic mice (Figure

4.2B) concurs with experiments performed in T1DM individuals. Non-diabetic individuals

are more resistant to fatigue than T1DM individuals, particularly those in poor glycaemic

control (Ramiere et al., 1993; Almeida et al., 2008). This is also the case in T1DM

individuals free of diabetic complications (Ramiere et al., 1993). However, this is not the

case in animal models of diabetes, resistance to fatigue is not affected by STZ-induced

diabetes in the EDL muscle of insulin-untreated STZ-diabetic rats although muscle specific

tension is impaired as mentioned above (Cotter et al., 1989; Cotter et al., 1993; McGuire &

MacDermott, 1999). The greater fall in force in muscles from diabetic mice could be

attributed to their higher total protein thiol oxidation level compared to muscles from

non-diabetic mice. However, one difficulty with this interpretation is that the total protein

thiol oxidation level of muscle proteins attained after fatiguing stimulation in muscles

from non-diabetic mice were comparable to those of diabetic mice prior to contraction

(Figure 4.2C). This suggests that changes in protein thiol oxidation might not play an

important role in fatigue.

A different interpretation arises when the thiol oxidation levels of individual proteins are

compared between muscles from non-diabetic mice and diabetic mice. Although the thiol

oxidation levels of actin, troponin and tropomyosin did not differ between fatigued

muscles from non-diabetic mice and pre-fatigue muscles from diabetic mice (Figure

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4.3A,C,D), the thiol oxidation level of myosin was lower in pre-fatigue muscles from

diabetic mice compared to fatigued muscles from non-diabetic mice (Figure 4.3B; p<0.05).

If the thiol oxidation of myosin was to play a major role in muscle force generation and

fatigue, this observation alone would explain why resistance to muscle fatigue is lower in

muscles from diabetic mice compared to muscles from non-diabetic mice despite

comparable protein thiol oxidation levels. However, a cautionary approach should be

adopted with this interpretation given that myosin is only one of many proteins whose

function is affected by thiol oxidation. For instance, the thiol oxidation of actin can disrupt

the organisation of the actin filaments, leading to reductions in the force generating ability

of the sarcomere (Milzani et al., 2000). The function of troponin is also altered when

oxidised (Putkey et al., 1993). Finally, the thiol oxidation of tropomyosin affects the

ATPase activity of actomyosin (Williams & Swenson, 1982). Although further work is

required to corroborate this interpretation, these findings highlight further the

importance of investigating the response of the thiol oxidation of individual proteins in

addition to that of total proteins.

It is important to note that one potentially confounding factor with the muscle model

adopted here is that the mean total protein thiol oxidation of resting muscles prior to

contraction was higher than those measured in muscles which were immediately frozen

after being removed from mice. This could represent a limitation with performing studies

using in vitro muscle preparations given the evidence that the thiol oxidation of muscle

proteins affects muscle contractile function (Chapters 2 and 3). Since the muscle

preparations used here were already under increased oxidative stress compared to those

114

in intact animals, and considering the improvement in contractile force associated with

DTT treatment, it is highly likely that muscle contractile performances measured in

muscles from non-diabetic mice and diabetic mice were suboptimal.

In conclusion, this study shows that the maximal specific force of the EDL preparation is

not reduced in Ins2Akita diabetic mice compared to non-diabetic mice despite higher thiol

oxidation level of muscle proteins in diabetic mice. These findings suggest that the higher

thiol oxidation level of proteins such as myosin in diabetic mice exposed to fatiguing

stimulation might explain their lesser resistance to fatigue. However, given the limitations

with using in vitro muscle preparations in this study, any attempt to extrapolate these

findings to T1DM in humans or Ins2Akita diabetic mice should be performed with caution.

115

CHAPTER 5:

GENERAL DISCUSSION

116

5.1 GENERAL DISCUSSION

In recent years, there has been mounting evidence suggesting that reactive oxygen

species (ROS) play some role in the development of skeletal muscle fatigue. However, the

intracellular mechanism(s) by which ROS contribute to muscle fatigue is still unclear. One

mechanism whereby ROS might play a role in muscle fatigue involves the reversible thiol

oxidation of muscle proteins. Another mechanism implicates ROS-induced protein

carbonylation which has recently been shown to be reversible in some situations

(Matsunaga et al., 2008; Wong et al., 2008, 2010). Although the functions of several

muscle proteins have been reported to be affected by these processes, it still remains to

be determined whether the thiol oxidation levels of these and other proteins increase

with fatiguing muscle stimulation. For this reason, the primary objective of Chapter 2 was

to use the extensor digitorum longus (EDL) muscle preparation from rats to establish

whether protein thiol oxidation level increases with muscle fatigue and to determine the

extent to which this contributes to muscle fatigue. Given the importance of controlling for

muscle damage and the possibility that short-lived transient rise in protein carbonylation

might also contribute to fatigue, the second objective of Chapter 2 was to examine the

response of protein carbonylation to fatiguing muscle stimulation and to evaluate the

contribution of this process to fatigue.

The findings in Chapter 2 provide compelling evidence that reversible protein thiol

oxidation rather than protein carbonylation is one of the mechanisms responsible for

muscle fatigue under the experimental conditions adopted in this thesis. Firstly, total

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protein thiol oxidation level increases in response to fatiguing stimulation, but not

following sustained non-fatiguing stimulation. Secondly, unlike protein carbonylation

responses, the recovery of muscle contractile performance after fatiguing stimulation is

accompanied by a return of total protein thiol oxidation to pre-stimulation levels. Thirdly,

both muscle fatigue and total protein thiol oxidation are more pronounced when fatiguing

stimulation takes place in the presence of the thiol oxidising agent, diamide. Fourthly,

muscle contraction performed with the thiol reducing agent, DTT decreases the

magnitude of both muscle fatigue and total protein thiol oxidation. Among the many

muscle proteins likely to be targeted by reversible protein thiol oxidation, the responses

of individual proteins separated by gel electrophoresis suggest that the thiol oxidation of

the muscle contractile proteins, myosin and actin might contribute to muscle fatigue.

Despite the rise in protein carbonylation levels with muscle fatigue, the results of Chapter

2 show that protein carbonylation and thus oxidative protein damage is unlikely to

contribute to muscle fatigue under the experimental conditions adopted in this study.

Firstly, muscle contraction in the presence of the thiol reducing agent or thiol thiol

oxidising agent decreased and increased muscle fatigue without affecting protein

carbonylation level. Secondly, the recovery of contractile following fatiguing stimulation is

not associated with any fall in protein carbonylation level. It is important to stress,

however, that these results do not necessarily imply that the fall in muscle contractile

performance during exercise cannot involve an increase in oxidative protein damage as

the extent to which proteins were damaged in this study might have been too low to

affect muscle contractile performance. This is supported by the data on skinned muscle

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fibers showing that a marked rise in protein carbonylation level can impair muscle

contractile performance.

Although the reversible rise in the thiol oxidation level of muscle proteins contributes to

muscle fatigue, it is estimated that only 20 % of the overall loss of contractile force during

fatiguing stimulation is from the contribution of protein thiol oxidation. This is suggested

by the observation that the normalisation of the thiol oxidation level of muscle proteins

with the antioxidant, DTT during fatiguing stimulation improves muscle contractile force

by only 20 %. This finding thus indicates that other factors play a more important role in

muscle fatigue. One limitation, however, with these results is that they are based on

experiments performed at 25 °C on isolated EDL muscles from rats. Since this temperature

is markedly different from that of working muscles, and considering that the importance

of other mechanisms of muscle fatigue (e.g. acidosis) have been shown to be temperature

sensitive (Adams et al., 1991; Pate et al., 1995; Westerblad et al., 1997), this raises the

question of the importance of the thiol oxidation of muscle proteins as a contributor of

fatigue at physiological muscle temperature. For this reason, the objective of Chapter 3

was to determine whether protein thiol oxidation plays an important role in muscle

fatigue at 34 °C due to greater ROS production and whether the exposure of protein thiol

reducing agent, DTT during fatiguing stimulation at 34 °C affects protein thiol oxidation

and muscle fatigue.

The results of Chapter 3 show that the thiol oxidation level of muscle proteins has a large

effect on muscle contractility, thus suggesting that this mechanism has the potential to

119

play an important role in muscle fatigue at physiological temperature. Firstly, this is

supported by the observation that total protein thiol oxidation level increases in response

to fatiguing stimulation. Secondly, acute exposure to the thiol reducing agent, DTT

improves muscle contractile performance with time while decreasing the thiol oxidation

level of total muscle proteins including that of myosin, actin, troponin and tropomyosin.

These results differ, however, from those in Chapter 2 in that muscle contractility can

increase by as much as 87 % in the presence of DTT, thus suggesting that reversible

protein thiol oxidation has the potential to be a major contributor to muscle fatigue at

physiological temperature. Unfortunately, the importance of this mechanism in

contrinuting to fatigue could not be ascertained in this study. This is because the

observation that total protein thiol oxidation levels prior to contraction in resting muscles

were far above those found in vivo and together with the progressive temporal increase in

muscle contractile force exposed to DTT during stimulation suggests that DTT is beneficial,

in part, because it normalises muscle function rather than only opposing muscle fatigue.

The increase in the thiol oxidation of muscle proteins in response to fatiguing stimulation

raises the obvious issue of the identity of the proteins involved. Close inspection of the

responses of individual protein bands separated by gel electrophoresis suggests that the

thiol oxidation of myosin might affect muscle contractile performance at physiological

temperature, whereas the absence of changes in actin, troponin and tropomyosin

suggests that these proteins are not involved. However, since DTT treatment and

associated improvement in muscle contractile performance is accompanied by a fall in the

thiol oxidation levels of myosin, actin, troponin, and tropomyosin, it is possible that the

120

thiol oxidation of actin, troponin and tropomyosin also play a role in muscle fatigue. These

findings in mice at physiological temperature are thus consistent with those obtained in

rats in Chapter 2.

The results of Chapters 2 and 3 that changes in the thiol oxidation level of muscle proteins

can affect acutely muscle contractile performance raise the question of whether some of

the medical conditions associated with impaired muscle function might also be linked with

an increased in the thiol oxidation level of muscle proteins. Since there is compelling

evidence for oxidative stress to be increased (Nishikawa et al., 2001; Evans et al., 2005;

Kaneki et al., 2007) and muscle contractile performance to be impaired (Andersen et al.,

1996; Andersen et al., 1997; Andreassen et al., 2006) in individuals with Type 1 diabetes

mellitus (T1DM) as well as in animal models of diabetes (Chapter 4), the objectives of

Chapter 4 were to determine whether the thiol oxidation level of muscle proteins is

increased in the diabetic Ins2Akita mice model and to examine whether there is an

association between muscle contractile performance and protein thiol oxidation level in

this animal model.

Against expectations, there was no difference in maximal specific force between muscles

from non-diabetic mice and diabetic mice despite the higher mean total protein thiol

oxidation in muscles from diabetic mice. This might be taken as evidence that difference in

the thiol oxidation level of muscle proteins are not necessarily associated with

corresponding differences in muscle contractile performance. In particular, the

observation that the thiol oxidation levels of actin and tropomyosin were higher in

121

muscles of diabetic mice suggests that the degree to which they were thiol oxidised was

not high enough to affect muscle function. However, one important finding is that the

thiol oxidation of myosin and troponin did not differ between muscles from non-diabetic

mice and diabetic mice. Given the role played by myosin in muscle contraction, and

previous findings that the oxidation of myosin inhibits myosin ATPase activity (Tiago et al.,

2006), the absence of difference in the thiol oxidation of myosin between muscles from

non-diabetic and diabetic mice might possibly explain their similar maximal specific force.

Arguably, this interpretation holds as long as myosin responsiveness to thiol oxidation

plays a more important role in determining contractile force. Although further work is

required to determine whether this is the case, the findings with myosin and troponin

highlight the importance of examining the response of individual proteins in addition to

that of total proteins.

Despite the absence of difference in maximal specific force between the muscles of non-

diabetic and diabetic mice, the latter was less resistant to fatigue. Interestingly, the

finding that the total thiol oxidation level of muscle proteins attained after fatiguing

stimulation in muscles from non-diabetic mice are comparable to that of muscles from

diabetic mice prior to contraction could be taken as evidence that changes in protein thiol

oxidation level may not play an important role in explaining the increased fatigue in

diabetic mice. However, a different interpretation emerges when the thiol oxidation levels

of individual proteins are compared between muscles from non-diabetic and diabetic

mice. Although the thiol oxidation levels of actin, troponin and tropomyosin did not differ

between fatigued muscles from non-diabetic mice and pre-fatigue muscles from diabetic

122

mice, the thiol oxidation of myosin is lower in pre-fatigue muscles from diabetic mice

compared to fatigued muscles from non-diabetic mice. If the thiol oxidation of myosin

were to play a major role in muscle force generation and fatigue, this observation alone

would explain why resistance to muscle fatigue is lower in muscles from diabetic mice

compared to muscles from non-diabetic mice despite comparable total protein thiol

oxidation level. However, more work is required to corroborate this interpretation.

In conclusion, this thesis shows for the first time that the thiol oxidation level of muscle

proteins increases in response to fatiguing muscle stimulation, and identifies experimental

conditions where oxidative stress contributes to muscle fatigue through protein thiol

oxidation rather than oxidative protein damage. This thesis also suggests that reversible

protein thiol oxidation has the potential to be a major mechanism of fatigue at

physiological temperature and that muscle fatigue is likely a consequence of the thiol

oxidation of multiple proteins such as the contractile proteins. This might be part of a

redox braking strategy, where protein thiol oxidation decreases contractile force in order

to protect the cell against dangerous increases in ROS production. However, the findings

of this thesis also highlight the limitations with using isolated muscle preparations as

experimental models. In particular, there is a need for better models that normalise the

protein thiol oxidation level of isolated muscle preparations at physiological temperature.

Also, and more importantly, the findings described in this thesis raise the issue of

investigating the physiological importance of the thiol oxidation of muscle proteins in

muscle fatigue not only in healthy humans, but also in individuals afflicted by medical

conditions known to impair muscle contractile function such as type 1 diabetes,

123

fibromyalgia, and chronic obstructive diseases to name a few. Finally, this thesis raises the

future challenge of evaluating the processes that lead to the changes in the thiol status of

muscle proteins and also the critical sulfhydryls in these proteins that affect their

activities. In addition, the relative contributions of the different thiol oxidised proteins to

muscle fatigue as well as the relative importance of thiol-mediated fatigue compared to

the many other mechanisms of muscle fatigue remain to be investigated.

124

CHAPTER 6:

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