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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 Materials and methods ……………………………………………………………………. 75
3.2.1 Animals ………………………………………………………………………………. 75
3.2.2 Intact muscle preparation …………………………….……………………. 75
3.2.3 Fatiguing stimulation protocol ……………………………………………. 76
3.2.4 Assay of proteins thiol oxidation levels ………………………………. 77
3.2.5 Electrophoretic separation and assay of ‘2 Tag’-labelled
protein ………………………………………………………………………………..
78
3.2.6 Statistical analysis ………………………………………………………………. 79
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
3.3.3 Thiol oxidation of contractile proteins and the loss of
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 Materials and methods ……………………………………………………………………. 97
4.2.1 Animals ………………………………………………………………………………. 97
viii
4.2.2 Intact muscle preparation …………………………….……………………. 97
4.2.3 Fatiguing stimulation protocol ……………………………………………. 98
4.2.4 Assay of proteins thiol oxidation levels ………………………………. 99
4.2.5 Electrophoretic separation and assay of ‘2 Tag’-labelled
protein ………………………………………………………………………………..
100
4.2.6 Statistical analysis ………………………………………………………………. 101
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
4.3.3 Thiol oxidation of contractile proteins and the loss of
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)
Located in both mitochondria and cytosol; removes
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)
Located in both mitochondria and cytosol; removes
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
Fatigue for 5 min at 30 Hz using
electrically stimulated
diaphragm muscle (In vitro)
20
Diaz et al (1994) 4 mg/ml N-
acetylcysteine
Fatigue for 4 min at 40 Hz using
electrically stimulated
diaphragm muscle (In vitro)
35
Khawli & Reid
(1994)
10 mM N-
acetylcysetine
Fatigue for 10 min at 30-40 Hz
using electrically stimulated
diaphragm muscle (In vitro)
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)
Fatigue for 5 min at 50 Hz using
electrically stimulated
diaphragm muscle (In vitro)
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)
150 mg/kg/hr NAC for
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
Time to exhaustion from
repetitive knee extensions
5
Medved et al
(2004)
150 mg/kg/hr NAC for
15 min then 25
mg/kg/hr NAC pre-
exercise and during
exercise
Time to exhaustion from cycling 26
Matuszczak et al
(2005)
150 mg/kg/hr NAC for
45 min pre-exercise
Time to exhaustion from
repetitive isometric handgrip
15
Mckenna et al
(2006)
125 mg/kg/hr NAC for
15 min then 25
mg/kg/hr NAC pre-
exercise and during
exercise
Time to exhaustion from cycling 24
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
Time to exhaustion from cycling 13
Nieman et al
(2010)
1000 mg/day
quercetin × 14 days
Treadmill running for 12 min
3
Bailey et al
(2011)
125 mg/kg/hr NAC for
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.,
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.
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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.
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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.
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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.
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CHAPTER 3:
PROTEIN THIOL OXIDATION AND
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 MATERIALS AND METHODS
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.
3.2.4 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
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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)
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).
3.2.5 Electrophoretic separation and assay of ‘2 Tag’-labeled 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
78
lower chamber stirred with a magnetic stirrer throughout. Upon completion of the gel
electrophoretic separation of proteins, the gel was immediately scanned for fluorescence
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.
3.2.6 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.
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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
thiol oxidation of actin (A), myosin (B), troponin (C) and tropomyosin (D) were measured after isolated EDL
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.
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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.
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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
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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
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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:
PROTEIN THIOL OXIDATION AND
MUSCLE CONTRACTILE FUNCTION IN
INS2AKITA DIABETIC MICE
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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 MATERIALS AND METHODS
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
98
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.
4.2.4 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)
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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).
4.2.5 Electrophoretic separation and assay of ‘2 Tag’-labeled 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
100
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.
4.2.6 Statistical analysis
Results are presented as means ± standard error of the mean. All analyses were
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
fatiguing stimulation.
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
fatiguing stimulation.
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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
thiol oxidation of actin (A), myosin (B), troponin (C) and tropomyosin (D) were measured after isolated EDL
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
113
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
117
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
118
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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