Contractile Function and Calpain Activity in Mouse Skeletal Muscle

during Hypoxia

by

Hui Hu

A THESIS

Submitted to

Oregon State University

in partial fulfillment of

the requirements for the

degree of

Master of Science

Presented November 7, 2007

Commencement June 2008

AN ABSTRACT OF THE THESIS OF

Hui Hu for the degree of Master of Science in Exercise and Sport Science presented

on November 7, 2007. Title: Contractile Function and Calpain Activity in Mouse

Skeletal Muscle During Hypoxia.

Abstract approved:

______________________________________

Jeffrey J. Widrick

Calcium activated proteases, or calpains, are activated in cardiac muscles under

conditions of ischemia/reperfusion and hypoxia. Their activity in skeletal muscle

under similar conditions is poorly understood. We tested the hypothesis that hypoxia

elevates calpain-mediated proteolysis in isolated glycolytic (extensor digitorum

longus, EDL) and oxidative (soleus, SOL) mouse muscles studied in vitro at 35°C.

Muscles performed a series of work loops during which time the tissue bath was

equilibrated with either 95% O2, 5% CO2, or 95% N2, 5% CO2. A sensitive substrate

of calpains, α-fodrin, and its calpain-specific 145 and 150 kDa breakdown products

were assessed by Western blot. To confirm calpain activity, similar experiments were

performed in the presence of E-64d, a calpain inhibitor, or DMSO, the E-64d solvent.

We observed that hypoxia increased the levels of α-fodrin-positive 145 and 150 kDa

peptides in both EDL and SOL. E-64d treatment during hypoxia prevented this

increase. However, inhibition of calpain activity had no effect on peak power output

and isometric force of either muscle during exposure to hypoxia. Our results provide

compelling evidence that calpain activity is elevated in actively contracting, hypoxic

EDL and SOL muscles, while unlike findings in cardiac muscles, we found no

conclusive evidence that inhibition of calpain-mediated proteolytic activity improved

skeletal muscle function during hypoxia or recovery.

@Copyright by Hui Hu

November 7, 2007

All Rights Reserved

Master of Science thesis of Hui Hu presented on November 7, 2007

APPROVED:

_____________________________________________________________________

Major Professor, representing Exercise and Sport Science

_____________________________________________________________________

Chair of the Department of Nutrition and Exercise Science

_____________________________________________________________________

Dean of the Graduate School

I understand that my thesis will become part of the permanent collection of Oregon

State University libraries. My signature below authorizes release of my thesis to any

reader upon request.

_____________________________________________________________________

Hui Hu, Author

ACKNOWLEDGEMENTS

I greatly appreciate my major professor, Dr. Jeff Widrick, for his guidance and

encouragement during my research and preparation of this thesis. My sincere

appreciation goes to all of my committee members: Dr Emily Ho, for allowing me to

use her lab equipment; Dr. Neil Forsberg, for providing knowledge on calpain and

calpain analysis; Dr. Beth Valentine, for independent study in muscle disease; Dr.

Larry Burt for serving on my committee. I would also like to thank Dr. Mike Maksud

and his generous support of my education through the George and Rachel Maksud

Fellowship. I feel honored to have received this fellowship.

Most of all I want to thank my family, who encouraged me through all these

years. Especially, I feel a deep appreciation to my husband, Zhu Zhang, for his

understanding and love.

CONTRIBUTION OF AUTHORS

Dr. Jeffrey Widrick performed the animal surgical procedures and assisted in

analysis of the data and writing of the manuscript.

TABLE OF CONTENTS

Page

Chapter 1 Introduction……………………………………………………. 1

Chapter 2 Literature Review……………………………………………… 4

Hypoxia and striated muscle function…………………………. 4

Skeletal muscle function and hypoxia…………………………. 4

Ischemia/hypoxia and myocardial function………………..…... 6

Mechanisms underlying functional changes during hypoxia….. 7

Calpain as a mediator of myocardial ischemia/reperfusion

injury…………………………………………………………… 11

Properties of calpains…………………………………………... 11

The endogenous inhibitor of calpains………………………….. 12

Activation mechanism of calpains……………………………... 13

Calpain and muscle protein degradation……………………….. 15

Hypoxia/ischemia associated calpain activity…………………. 16

Aims of this study……………………………………………… 18

References………………………………………………………. 20

TABLE OF CONTENTS (Continued)

Page

Chapter 3 Calpain Activity of Mouse Skeletal Muscles

during Hypoxia………………………………………………... 29

Abstract……………………………………………………….. 30

Introduction…………………………………………………… 31

Methods……………………………………………………….. 33

Results………………………………………………………… 40

Part I: Contractile properties…………………………….. 40

Part II: Calpain activity………………………………….. 41

Discussion…………………………………………………….. 42

References……………………………………………………… 46

Chapter 4 Conclusion…………………………………………………….. 64

LIST OF TABLES

Table Page

3.1. Characteristics of EDL and SOL muscles……………………... 53

3.2. Peak isometric force of EDL and SOL muscles………………. 54

3.3. Peak power of EDL and SOL muscles………………………… 55

LIST OF FIGURES

.

Figure Page

3.1. Representative isometric force of EDL muscle………………........ 56

3.2. Representative isometric force of SOL muscle……………………. 57

3.3. Representative power output of EDL muscle……………………... 58

3.4 Representative power output of SOL muscle……………………... 59

3.5 Isometric force of pre-treatment, post-treatment and reoxygenation

(recovery) in control (con), hypoxia (hyp), hypoxia with DMSO

(hyp-D) and hypoxia with E-64 (hyp-I) …………………………...

60

3.6 Power output of pre-treatment, post-treatment and reoxygenation

(recovery) in control (con), hypoxia (hyp), hypoxia with DMSO

(hyp-D) and hypoxia with E-64d (hyp-I) ………………………….

61

3.7 Analysis of calpain-cleaved -fodrin in EDL muscle…………….. 62

3.8 Analysis of calpain-cleaved -fodrin in SOL muscle…………….. 63

1

Contractile Function and Calpain Activity in Mouse Skeletal Muscle

During Hypoxia

Chapter 1

INTRODUCTION

Ischemia, which is characterized by reduced blood flow, deprives tissues of vital

substrates and allows metabolites to accumulate in cells. Ischemia can be difficult to

study experimentally because it is not always possible to apply drugs to isolated tissue

preparations that lack an intact blood supply. For these reasons, many studies use

hypoxia, removing all or most of the O2 from the preparation, to mimic ischemia.

Therefore, for the purposes of this project we will consider studies examining ischemia

and hypoxia.

Ischemia/hypoxia has been intensively studied in a variety of pathophysiological

situations, including heart disease (myocardial ischemia-reperfusion), respiratory disease

(such as chronic obstructive pulmonary, asthma, pulmonary fibrosis), brain infarction,

and renal disease. Most work examining the effects of ischemia or hypoxia on striated

muscle has been performed on cardiac preparations. The effects of ischemia on

myocardial function are the subject of great interest because of high prevalence of

ischemia heart disease and its high rates of morbidity and mortality. Though a number of

hypotheses have been put forward to explain the underlying mechanisms of

ischemia/reperfusion induced contractile dysfunction, most evidence suggests that the

formation of oxygen-derived free radicals and Ca2+

overload are responsible for

myocardial dysfunction during ischemia/reperfusion. However, the exact mechanisms are

not completely understood.

While most research has focused on hypoxia/ischemia and cardiac muscle function,

2

hypoxia also has important health and performance implications for skeletal muscles. For

instance, hypoxia is a common feature in several respiratory diseases, such as chronic

obstructive pulmonary disease, acute respiratory distress syndrome, and severe

pneumonia. Hypoxia-induced dysfunction of the respiratory muscles frequently occurs in

patients under these situations. Ischemia-reperfusion-induced injury in limb skeletal

muscle is also commonly encountered in a number of clinical scenarios. It may occur

following arterial clamping during arterial reconstructive surgery and following

tourniquet use during operations. Trauma may also cause acute ischemia and tissue

hypoxia. Finally, healthy muscles may experience reduced oxygen delivery during high

intensity exercise and the resulting hypoxia may be a contributing factor to the

development of fatigue.

While cardiac and skeletal muscles are similar in many ways, they have important

differences. For instance, while cardiac muscles express only slow myosin heavy chain

isoforms (α-MHC, β- or type I MHC), adult skeletal muscles express one slow isoform

(β- or type I MHC) and, depending on the species up to three fast isoforms (type IIa, IIx,

and IIb MHC). Associated with these myosin heavy chain isoforms are distinct

differences in metabolism, fatigability, and contractility. For instance, slow muscle fibers

have slow shortening velocities and develop a relatively low peak power, but because of

their dependence on aerobic metabolism, these fibers are fatigue resistant. In contrast,

fibers expressing type IIb/x MHC have a fast shortening velocity and develop high power,

but are very susceptible to fatigue because their energy conversion is based on anaerobic,

glycolytic metabolism. Fibers expressing type IIa MHC have intermediate properties in

that they have a fast shortening velocity, develop a moderate power, and are relatively

3

resistant to fatigue. These fibers work under both aerobic and anaerobic conditions.

Skeletal and cardiac muscle also have differences in their activation. In skeletal muscle, a

saturating concentration of Ca2+

is released from the sarcoplasmic reticulum (SR) upon

membrane depolarization. In cardiac muscle, Ca2+

enters the cell from extracellular pools

as well as being released from, the SR. Furthermore, the force of cardiac cells is graded

by increases in intracellular [Ca2+

] vs. the “all or none” force response of skeletal muscle.

Because of these differences, the results of studies performed on ischemic/hypoxic

cardiac muscle may not be entirely generalizable to skeletal muscles subjected to similar

conditions of oxygen depravation. Thus, the first goal of this study was to evaluate the

contractile function of oxidative and glycolytic skeletal muscles during hypoxic

conditions.

Calpains, calcium-activated neutral cysteine proteases, are present in many tissues

including skeletal muscles. Calpains have a Ca2+

-binding domain. Thus, calpain

proteolytic activity is activated and regulated by the intracellular Ca2+

concentration.

Recent evidence suggests that an increase in intracellular Ca2+

concentration is a common

feature of cardiac ischemia injury. As a result it might cause extensive Ca2+

-dependent

proteolytic damage to the myocardium. It has been observed that Ca2+

-dependent

calpains are activated in cardiac muscles under conditions of ischemia/reperfusion and

hypoxia. Calpain activity in skeletal muscle under similar conditions is poorly

understood. Therefore, the second goal of this study was to evaluate calpain activity in

hypoxic skeletal muscles in order to better understand the role of calpains in

hypoxia-induced impairment of skeletal muscle function.

4

Contractile Function and Calpain Activity in Skeletal Muscle of Mouse

During Hypoxia

Chapter 2

LITERATURE REVIEW

Hypoxia and Striated Muscle Function

Skeletal muscle function and hypoxia

While most research has focused on cardiac muscle function during

hypoxia/ischemia, hypoxia also has important health and performance implications for

skeletal limb and respiratory muscles. Several studies have suggested that hypoxia

impairs force generation of the diaphragm in vitro (Heunks et al., 2001; Brotto et al.,

2000; Van der Heijden et al., 1999). In the study by Van der Heijden et al. (1999) 90 min

of hypoxia (~7 kPa) resulted in a severe decrease in twitch force (Pt) and maximal tetanic

force (Po) of rat diaphragm muscle. Similar results were also reported in mouse (Seow

and Stephens., 1988) and in hamster diaphragms (Esau, 1989) and in the in situ dog

diaphragm (Bark et al., 1988). Reduction in force during hypoxia depends on the

stimulation frequency used and the degree of hypoxia, so it is often difficult to directly

compare results from different studies.

Muscular contraction results in force generation, shortening, or both. While hypoxia

is known to impair isometric force generation, its effect on shortening characteristics is

unclear. A study by Machiels et al. (2001) showed that the maximum shortening velocity

of hypoxia treated rat diaphragm muscle decreased significantly (~30% lower compared

to control) and shifted the force-velocity curve downwards. However, other studies

observed that hypoxia impaired Pt and Po but did not affect the force-velocity relationship

5

(Van der et al., 1999; Heunks et al., 2001). Peak isometric force correlates with the

number of active cross-bridges, while shortening velocity is determined by the rate of

cross-bridge cycling (Huxley, 1957) and correlates with myosin ATPase activity (Barany,

1967). Previous studies suggest that hypoxia impairs cross-bridge recruitment or reduces

the force per cross bridge but hypoxia may or may not affect the ATPase activity and the

cross-bridge cycling rate. It is important to note that these conclusions pertain only to

rodent diaphragm muscles. Whether they apply to other muscles of different fiber type

and metabolic profile is not known.

Conflicting results have been found as to whether hypoxia accelerates muscle

fatigue, with some studies indicating a lack of effect (Bark et al., 1988; Eau, 1989) and

some data indicating a detrimental effect (Salomone et al., 1991; Machiels et al. 2001;

Zhu et al., 2003). Machiels et al. (2001) observed that fatigue during isotonic

contractions was more prominent under hypoxia conditions than during hyperoxia, and

the isotonic endurance time was shorter in hypoxia. Zhu et al. (2003) reported that

hypoxia reduced muscle fatigue resistance, indicated by a decline in power output and

less fatigue endurance during repetitive contraction. The different muscles, different

species, and different models used in these studies may partly explain the inconsistency

of results. However, Bark et al. (1988) indicates in their study that the detrimental effects

of hypoxia on force during fatigue appear to be more pronounced with greater degrees of

hypoxia and with higher intensity muscle contraction. Salomone et al. (1991) also

pointed out that mild to moderate hypoxia did not further slow skeletal muscle relaxation

rate during fatigue, although the force is affected adversely by hypoxia.

Some studies have been conducted in single skinned muscle fibers obtained from

6

muscle exposed to hypoxia (Brotto et al., 2000, 2001; Ottenheijm et al., 2006). Force

generation in skinned fibers is determined by cross bridges that are formed by myosin

heavy chain heads that attach to actin (Gordon et al., 2000). Therefore, the change of Po

of single skinned fibers should be reflected by a change of the number of available cross

bridges per half sarcomere, the mean force per cross bridge in the force-generating state,

and the fraction of strongly attached cross bridges. Ottenheijm et al. (2006) reported that

30 minutes of acute hypoxia induced an ~ 32% decline of maximal Ca2+

-activated force

in skinned fibers, a reduction in the fraction of strongly attached cross bridges, and a

decline in the rate constant of force development. Brotto et al. (2000) also observed that

skinned fibers showed a 50% decrease in calcium-activated force and a decrease in Ca2+

sensitivity after hypoxia-fatigue conditions, indicating contractile protein dysfunction.

Brotto et al. (2001) further showed that detrimental effects of hypoxia-fatiguing

stimulation are correlated with the degradation of troponin T (TnT) and troponin I (TnI).

Ischemia/hypoxia and myocardial function

Myocardial ischemia-reperfusion injury, a consequence of cardiovascular disease, is

one of the most common causes of serious illness and early death in developed societies

(Allen and Orchard, 1987). Thus, the mechanisms underlying cardiac

ischemia/reperfusion, and the consequences for cardiac function, have been extensively

studied (Allen and Orchard, 1987; Lee and Allen, 1991; Carrozza et al., 1992).

The consequences of cardiac ischemia include reduced force production,

life-threatening disturbances in the cardiac rhythm, and ultimately the death of the cells

(Lee and Allen, 1991). It has been reported that when a heart is subjected to ischemia the

7

developed pressure declines rapidly (Lee and Allen, 1991). This decrease in force is

generally complete after 5-10 min with little or no re-development of force.

Mounting evidence suggests that calcium overload during ischemia and upon

reperfusion might cause extensive proteolytic damage to the myocardium, and that this is

responsible in part for the depressed cardiac function characteristic of

ischemia/reperfusion (Carrozza et al., 1992; Yoshida et al., 1995; Tsuji et al., 2001;

Singh et al., 2004). The work of Singh et al. (2004) on sarcoplasmic reticulum proteins of

the ischemia-reperfused heart suggests that the proteolytic degradation/modification of

SR proteins by calpains may be a downstream mechanism of Ca2+

overload mediated

contractile dysfunction. Selective loss or proteolysis of troponin I is also reported during

ischemia/reperfusion, and is suggested to be associated with depressed cardiac

performance (Gao et al., 1996).

Numerous investigations observe that production of reactive oxygen species (ROS)

is increased during myocardial ischemia/reperfusion and that inhibition of ROS

formation results in attenuation of subsequent contractile dysfunction (Park et al., 1991;

Bolli, 1998; Wright et al., 2005). The findings from these studies suggest that ROS

formation during hypoxia plays a certain role in myocardial dysfunction. However,

whether or not ROS formation is the actual initiator or a downstream mediator of

myocardial dysfunction and whether ROS interacts with Ca2+

during

ischemia/reperfusion remains to be clarified.

Mechanisms underlying functional changes during hypoxia

The mechanisms of hypoxia/ischemia-induced impairment of contractile

performance are not fully understood. Potential mechanisms include depletion of ATP,

8

changes in intracellular pH, proteolysis of proteins, Ca2+

overload, and free radical

damage.

a. Depletion of ATP.

Some studies suggest that depletion of adenosine triphosphate (ATP) may lead to

the decline in contractile force (Hearse, 1979). Both ischemia and hypoxia reduce the

oxygen supply to the muscle cells and reduce the rate of ATP production by oxidative

phosphorylation. It has been established that when the concentration of ATP falls below a

certain level, intracellular Ca2+

rises quite rapidly, which is assumed to be due to failure

of the various Ca2+

pumps. During early ischemia, ATP levels may be maintained by

increased glycolysis but at the expense of the limited reserves of glucose and glycogen.

Glucose utilization increases by > 10-fold in ischemic/hypoxic cardiac myocytes

(Webster et al., 1999). Shine et al. (1976) observed a substantial decline in developed

tension after one minute of ischemia of cardiac muscle, and the decline was generally

complete with 5-10 minutes. However the [ATP]i falls by only a small amount during this

period. This fact suggests that the decline in tension of cardiac muscle may not be

directly related to [ATP]i.

b. Intracellular acidosis.

Acidosis associated with hypoxia is suggested to be possible mechanism for the

decrease in muscle force production during hypoxia/ischemia (Katz and Hecht, 1969).

Accelerated glycolysis and enhanced lactic acid production are assumed to lead to a

decrease of pH. It had been observed since 1880 that acidic conditions lead to a decrease

in the amount of tension developed by the heart (Gaskell, 1880). In isolated skinned

cardiac cells, an acidosis of 0.17 pH units reduces tension production to 30-40%

9

compared to the reduction to 20% of control (Fabiato and Fabiato, 1978). Low pH also

impairs the contractile function of isolated skinned skeletal muscle fibers (Knuth et al.,

2006). A study using 31

P NMR to monitor pH in isolated hearts during hypoxia and

ischemia, showed that pH and tension both decreased during ischemia, but the decrease

in mechanical performance during ischemia was greater than that could be accounted for

by the acidosis (Jacobus et al., 1982). These studies suggest that acidosis during

hypoxia/ischemia may make some contribution to the observed decrease in tension in

cardiac muscle, but it may not be the major cause of force decline.

c. Oxidative stress.

Recent evidence shows that reactive oxygen species (ROS) may play a role in

hypoxia-induced muscle dysfunction (Heunks et al., 2001; Zhu et al., 2003). It has been

shown that hypoxia increases the generation of ROS in cardiac tissue (Park et al., 1991)

and in diaphragm muscle (Mohanraj et al., 1998). Also in the studies of Mohanraj et al.

(1998) and Wright et al. (2005) antioxidant treatments during exposure to hypoxia

resulted in improvements in force generation, suggesting hypoxia-induced ROS

formation may play a role in the impairment of muscle contractility during hypoxia.

Similar results have also been found by others (Heunks et al., 2001; Zhu et al., 2003).

The exact roles of ROS formation in hypoxia-induced impairment of muscle contractility

are unknown. A study by Eu et al. (2003) suggests that ROS may impair Ca2+

regulation

in skeletal muscle under hypoxic conditions.

d. Proteolysis as a contributing mechanism.

A body of recent studies suggests that the effects of hypoxia on proteolysis of

myofibrillar proteins may play an important role in hypoxia/ischemia-induced

10

impairment of contractile performance. Three major proteolytic pathways are considered

to be responsible for the degradation of regulatory and contractile proteins during muscle

damage: the lysosomal proteases (cathepsins), the calcium-dependent calpains, and the

ATP-dependent ubiquitin proteasome system (Lowell et al., 1986; Taillandier et al., 1996;

Huang and Forsberg, 1998; Lecker et al., 1999; Jackman et al., 2004). It has been

suggested that lysosomal proteases may not contribute significantly to the degradation of

myofibrils and other cytostolic proteins, but rather, their role is to degrade membrane

proteins, including receptors, ligands, channels, and transporters (Lowell et al., 1986).

The calcium-dependent calpain and ubiquitin-proteasome are thought to work

sequentially in muscle protein proteolysis (Hasselgren et al., 2001; Jackman et al., 2004).

It has been demonstrated that the proteasome is not able to degrade intact myofibrils. The

myofibrillar proteins must in some way be released from the sarcomere before they can

undergo ubiquitination and be degraded by the proteasome (Jagoe and Goldberg, 2001).

Calpains have been suggested to digest the Z-discs in the muscle sarcomere and release

filaments from the myofibril in order to initiate myofibrillar degradation (Solomon and

Goldberg, 1996).

Ca2+

plays a very important role in skeletal muscle. Acting as a second message,

Ca2+

controls the function of all muscles by initiating contraction. Muscle contracts when

an action potential triggers a large influx of Ca2+

into the cytoplasm, causing relatively

large but transient changes in [Ca2+

]i (Trafford et al., 2002). The Ca2+

ions then bind to

troponin C on the myofibrils, allowing the interaction of actin and myosin, the hydrolysis

of ATP, and the generation of force. The elevated [Ca2+

]i, which activates muscle

contraction, is removed mainly to the SR via sarcoplasmic reticulum Ca2+

-ATPase and

11

pumped out of the cell via the Na+/Ca

2+ exchanger (Bers, 2000).

If the permeability of the sarcolemma for Ca2+

is increased, a large influx of Ca2+

will occur, and the muscle cell may suffer Ca2+

overload. An excessive accumulation of

Ca2+

in the muscle cell, resulting from an inability of the muscle cell to control [Ca2+

]i,

will lead to muscle cell damage. Pei et al (2003) reported an impaired handling of Ca2+

by the SR from rats subjected to chronic hypoxia. Carrozza et al. (1992) also observed

functional and structural damage of the myocardium mediated by Ca2+

overload during

ischemia/reperfusion (Carrozza et al., 1992).

Calpain as a Mediator of Myocardial Ischemia/Reperfusion Injury

As reviewed in the previous section, intracellular Ca2+

overload has been proposed

as a major mechanism for contractile dysfunction during ischemia/hypoxia. However, the

exact mechanism of contractile dysfunction during ischemia/hypoxia is not clear. It has

been postulated that Ca2+

overload during hypoxia/ischemia triggers Ca2+

-dependent

calpains. These activated calpains could partially degrade contractile proteins, leading to

myofilament dysfunction. In this section, we consider the evidence supporting this

hypothesis.

Properties of calpains

Calpains (EC 3.4.22.17), which constitute a large super-family of

calcium-dependent cytoplasmic cysteine proteases, were first identified in 1968 (Huston

and Krebs, 1968). Calpains are classified as “atypical” or “typical” calpains (also called

“conventional” calpains). Typical calpains are further divided into ubiquitous and

tissue-specific calpains. Two ubiquitous calpains, µ- and m-calpain (also called calpain I

and II, respectively), are expressed in almost all animal tissues and cells. µ- and

12

m-calpain differ in their calcium sensitivity in vitro and are activated at micro- and

milli-molar Ca2+

concentrations, respectively (Goll et al., 2003). These calpains are

composed of a large catalytic subunit (80 kDa, encoded by CAPN1 and CAPN2 for

µ-and m-calpains, respectively) and a small regulatory subunit (~30 kDa, encoded by

CAPN4). The large subunit can be divided into four domains (I-IV). Domain II is the

cysteine protease domain and Domain IV is the Ca2+

-binding domain. Thus, the

Ca2+-

dependent protease activity of calpain is ascribed to the large subunit. The small

subunit also contains a Ca2+

- binding domain (domain VI) and is thought to regulate

calpain activity.

Some calpain homologues in mammals have been found to be predominantly

expressed in a limited number of organs, in contrast with the ubiquitous expression of the

“conventional” µ- and m-calpain. These tissue-specific calpains include the skeletal

muscle-specific p94 isoform (also called calpain 3), a lens-specific splicing variant of

p94, and stomach-specific calpains nCL-2 and nCL -2’. These tissue specific calpains are

suggested to be related to the specific function of the organs in which they are

predominantly expressed. The properties of calpain 3 remain an enigma. Calpain 3, by

binding to titin/connectin at the N2 line, to the M-line, and to the Z-line, might regulate

signal transduction in skeletal muscle (Spencer et al., 2002).

The endogenous inhibitor of calpains

Calpastatin is a protein that inhibits the proteolytic activity of µ- and m-calpain. This

inhibitory activity seems to be specific for calpains because no other proteases reported

so far are affected by calpastatin (Blomgren et al., 1999). µ-and m-calpain co-exist with

13

this specific endogenous inhibitor, suggesting a critical role of calpastatin in the

regulation of calpain activity. The calpastatin molecule contains four equivalent

inhibitory domains, each having three conserved regions, A, B and C, that are important

for inhibition. The region B is essential for inhibition, and region A and C are important

for potentiation of the inhibition activity. Region A and C interact with the Ca2+

-binding

domain of the calpain large subunit and small subunit, respectively, in a Ca2+

dependent

fashion.

Coexistence of the calpains with their endogeneous inhibitor may be necessary for

the fine control of calpain activity under physiological conditions. The

calpain-calpastatin interaction is reversible and calcium-dependent in that calpains bind

to the inhibitor only after binding calcium. Calpastatin can be proteolyzed by calpains as

a suicide substrate during their transient association (Blomgren et al., 1999).

Activation mechanism of calpains

Calpains exist in the cytosol as an inactive enzyme and translocate to membranes in

response to an increase in the intracellular Ca2+

level. Several other factors have been

suggested to be potential regulators of calpain activity. These include autolysis,

phospholipids, and calpastatin.

On exposure to sufficient Ca2+

, calpain molecules are assumed to undergo

significant conformational changes, and these changes involve partial unfolding of

domain II and subsequent release of constraints imposed by domain interaction to form a

functional catalytic site. In vitro activation of both µ- and m-calpain require Ca2+

concentration at levels that are hardly ever reached under physiological conditions. Thus,

activation of calpain proteases must be dependent on factors other than Ca2+

14

concentration. For instance, autolysis of calpains is suggested to lower Ca2+

requirement

for both µ-calpain and m-calpain by initiating dissociation of the large subunit from the

small subunit of calpain. The Ca2+

requirements for non-autolyzed calpains are reported

to be in the ranges of 5-50 µM Ca2+

for µ-calpain and 250-1000 µM Ca2+

for m-calpain.

These ranges are lowered by autolysis to 1-5 µM Ca2+

for µ-calpain and 100-200 µM

Ca2+

for m-calpain (Elce et al., 1997).

Although autolysis lowers the Ca2+

requirement for calpain activation, the Ca2+

concentration required for autolysis is similar to or just slightly greater than that required

for proteolytic activity (Zimmerman and Schlaepfer, 1991). A discovery that shows

certain phospholipids lower the Ca2+

concentration required for autolysis provides a

potential solution for this paradox (Coolican and Hathaway, 1984). It has been suggested

that the calpains interact with phospholipids at the membrane and that this interaction

reduces the Ca2+

requirement for autolysis (Goll et al., 2003). However, whether

phospholipids/autolysis could be the activation mechanism of calpain is still not clarified.

Several calpain activator proteins have been described to reduce the Ca2+

concentration required for calpain in vitro. Isovalerylcarnitine is reported to reduce the

Ca2+

concentration required for maximal proteolytic activity of m-calpain (Pontremoli et

al., 1990), and an activator may affect the catalytic properties of µ-calpain (Melloni et al.,

1998). The mechanism of these activators is still unclear. It has been suggested that these

calpain activators increase the catalytic activity (Vm) of the calpains without affecting the

Ca2+

concentration required for proteolytic activity.

Calpastatin, as an endogenous calpain inhibitor, has an important role in regulating

activity of µ-calpain and m-calpain. How calpastatin regulates activity of calpain in

15

living cells is still a mystery. Further study on the regulation of calpain/calpastatin

interaction will be needed to better understand the mechanisms regulating calpain

activity.

Calpain and muscle protein degradation

Muscle tissues contains three classes of proteins: the myofibrillar or contractile

muscle proteins (~55-60% of total muscle proteins), the sarcoplasmic or cytoplasmic

proteins (~30-35% of total muscle protein), and membrane proteins (~10-15% of total

muscle protein) (Goll et al., 2003). It seems clear that calpains mainly target the

myofibrillar or the cytoskeletal proteins in muscle tissue. As stated previously, the

myofibrillar proteins must be released before they are degraded by the proteasome

(Williams et al., 1999; Jagoe and Goldberg, 2001). There is evidence that release of

myofilaments from the myofibrils is mediated by the calcium-dependent calpain activity

(Williams et al., 1999; Jackman et al., 2004). In muscle protein degradation, it has been

suggested that the Z-disc is the preferential site of calpain proteolysis and many calpains

substrates are components of the Z-disc. Calpain cleaves titin and nebulin filaments near

the Z-disc, severing the attachments of Z-disc proteins, such as �-actinin, to the myofibril,

and releasing thick and thin filaments from the myofibril (Taylor et al., 1995; Williams et

al., 1999; Goll et al., 2003). Calpain also cleaves the intermediate filament protein

desmin, the thin filament regulatory proteins troponins T and I, and the thick filament

C-protein, resulting in dissociation of the thick and thin filaments into myosin and actin

molecules (Taylor et al., 1995; Williams et al., 1999). Myosin and actin molecules, as

well as and other degradation fragments produced by calpain activity, can be further

degraded by the proteosome or by the lysosomal system (Williams et al., 1999; Jackman

16

et al., 2004).

Hypoxia/ischemia associated calpain activity

The physiological function of the calpain system is still not completely understood.

It has been suggested that the calpains play a number of different roles in living cells

(Taylor et al., 1995; Williams et al., 1999; Goll et al., 2003), such as cleavage of

cytoskeletal/membrane attachments, proteolytic modification of molecules involved in

signal transduction pathways (including many kinases, phosphatases, cytoskeletal

proteins), degradation of enzymes controlling the cell cycle, regulation of gene

expression by cleaving several transcription factors (including p53, c-Jun, c-Fos), and

substrate degradation in apoptosis.

The calpains have also been demonstrated to be involved in many pathological

situations in cell or tissues. The purported role of the calpains in various pathologic

conditions is not well understood, but it is widely assumed that loss of Ca2+

homeostasis

in some way activates the calpain system, leading to the increased protein degradation.

Results from many studies have shown that increases in intracellular Ca2+

concentration

are a common feature of hypoxia/ischemia injury. As a result, calcium-dependent

intracellular pathways and proteases are activated, culminating in cell injury

and

eventually cell death (Fredsted et al., 2005; Jones et al., 1984). The calcium-activated

calpains have been shown to be mediators of hypoxia/ischemia injury in several tissues,

including the brain (Lee et al., 1991), myocardium (Iizuka et al., 1991), and renal tubular

cells (Gengaro et al., 1995). Calpain activation also was reported in liver tissue after

reperfusion of transplanted human livers (Aguilar et al., 1997).

Lee et al. (1991) observed that calcium-activated proteolysis was initiated during

17

ischemia of hippocampal neurons and did not depend wholly on tissue reoxygenation

because hypoxia alone was a sufficient stimulus to elicit calcium-activated proteolysis.

They also demonstrated that inhibition of calpain proteolysis protects hippocampal

neurons from ischemia. This indicates that calpain activation is an important event of the

post-ischemic neurodegenerative response and that suppression of calpain proteolysis

may be a feasible strategy for preserving both the structure and function of vulnerable

neurons.

Studies with hypoxic-induced myocardial cell injury found that calpain was

activated during hypoxic myocardial cell injury, and that calcium antagonists and calpain

inhibitor-1 prevented protein degradation during hypoxic cell injury (Iizuka et al., 1991).

Studies with hypoxia-induced renal tubular injury demonstrated that intracellular free

Ca2+

was significantly increased after 2 min of hypoxia in proximal tubules, implying

that an increase in Ca2+

may play an initiating role in hypoxia injury. A study by

Edelstein et al. (1995) also found that calpain activity increased after hypoxia and that

cysteine protease inhibition attenuated and prevented renal tubular injury during hypoxia.

This suggests that calpains play a role as a mediator of hypoxia-induced proximal tubular

injury.

In skeletal and cardiac muscle, several calpain substrates have been shown to be

degraded in the response to hypoxia/ischemia. A study by Gao et al. (1997) observed that

calpain-I induced selective troponin-I (TnI) degradation in the post-ischemic stunned

myocardium, leading to a decrease in Ca2+

sensitivity of the myofibrillar system. A

similar study by Brotto et al. (2001) found that hypoxia-fatigue induced degradation of

troponins TnI and TnC in skeletal muscle and this disruption of the troponin complex

18

appears to be similar to that experienced by myocardial muscle under ischemic

conditions. Degradation and loss of various cytoskeletal elements, such as β-calspectin

(spectrin), �-actinin and desmin were also reported in myocardial cell studies (Van Eyk,

1998; Yoshida et al., 1995). Papp et al. (2000) demonstrated that during

ischemia/reperfusion calpain 1 induced both a major reduction in maximal force and a

degradation of desmin. Desmin is the constitutive protein of the intermediate filaments,

closely associated to �-actinin and actin filaments at the level of Z bands (Tokuyasu et al.,

1983). Intermediate filaments of desmin link Z bands with the plasmalemma and the

nucleus which contribute to the organization of myofibrils and maintenance of cell shape

(Tokuyasu et al., 1983). In the studies of the mechanical role of intermediate filaments,

desmin has been showed to be indispensable for normal development and viability, force

generation and maintenance of passive tension (Lazarides, 1980). In addition, Singh et al.

(2004) observed an ischemia-reperfusion-induced depression in the major SR

Ca2+

-handling proteins, such as RyR and sarcoplasmic reticulum Ca2+

-ATPase (SERCA).

They suggest that proteolytic degradation/modification of SR proteins by calpains may

play an important role in contractile dysfunction in IR hearts.

Aims of this study

While most research has focused on hypoxia/ischemia and cardiac muscle function,

hypoxia also has important health and performance implications for skeletal muscles.

Since cardiac and skeletal muscles are different in metabolism, fatigability and

contractility, as well as in their activation, the results of studies performed on ischemic

cardiac muscles may not be entirely generalizable to skeletal muscles. Therefore, it is

important to understand contractile function in skeletal muscle during hypoxia.

19

Skeletal muscle fibers are classified as fast-twitch glycolytic (FG), fast-twitch

oxidative glycolytic (FOG) and slow-twitch oxidative (SO). These various fiber types

differ in their contractile and metabolic properties. However, limited data are available

regarding the contractile response of different muscle fiber types to decreased delivery of

oxygen to muscle (ischemia/hypoxia). Often, the results regarding this issue are

contradictory. Furthermore, previous studies examining skeletal muscle function during

hypoxia have focused mainly on changes in isometric force. However, isometrically

contracting muscles perform no external work and it is the capacity to perform work per

unit time, or power, which is arguably the most important determinant of skeletal muscle

performance.

To the best of our knowledge, no previous study has evaluated the effects of hypoxia

on the capacity of skeletal muscle to produce power during cyclical lengthening and

shortening. Therefore, we prepared extensor digitorum longus (EDL) and soleus (SOL)

muscles from mice in order to investigate the contractile function of these muscles under

hypoxic conditions. EDL and SOL have often been used experimentally as representative

glycolytic and oxidative muscles, respectively.

In order to test the hypothesis that calpain activity is increased in skeletal muscle

during hypoxia, calpain activity was evaluated by the appearance of �-fodrin breakdown

products (145/150 kDa). To further understand the role of calpain in hypoxic skeletal

muscle function, calpain activity was also examined in the presence of E-64d, a calpain

inhibitor.

20

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29

Chapter 3

Calpain Activity of Mouse Skeletal Muscles during Hypoxia

Hui Hu and Jeffrey J. Widrick

Department of Nutrition and Exercise Science

30

ABSTRACT

Calcium activated proteases, or calpains, are activated in cardiac muscles under

conditions of ischemia/reperfusion and hypoxia. Their activity in skeletal muscle under

similar conditions is poorly understood. We tested the hypothesis that hypoxia elevates

calpain-mediated proteolysis in isolated glycolytic (extensor digitorum longus, EDL) and

oxidative (soleus, SOL) mouse muscles studied in vitro at 35°C. Muscles performed a

series of work loops during which time the tissue bath was equilibrated with either 95%

O2, 5% CO2 or 95% N2, 5% CO2. A sensitive substrate of calpains, α-fodrin, and its

calpain-specific 145 and 150 kDa breakdown products were assessed by Western blot. To

confirm calpain activity, similar experiments were performed in the presence of E-64d, a

calpain inhibitor, or DMSO, the E-64d solvent. We observed that hypoxia increased the

levels of α-fodrin-positive 145 and 150 kDa peptides in both EDL and SOL. E-64d

treatment during hypoxia prevented this increase. However, inhibition of calpain activity

had no effect on peak power output and isometric force of either muscle during exposure

to hypoxia. These findings suggest that calpains are activated in skeletal muscles exposed

to hypoxia, but unlike cardiac muscle, calpain-mediated proteolysis does not appear to be

a major cause of skeletal muscle dysfunction under these conditions.

Keywords: hypoxia, contractile function, calpain activity, calpain inhibitor

31

INTRODUCTION

Calcium-activated neutral cysteine proteases, or calpains, play important roles in

cells of many tissues, including cleavage of cytostolic and membrane proteins,

proteolytic modification of molecules in signal transduction pathways, and

degradation of enzymes controlling the cell cycle (Goll et al., 2003). Calpains are also

involved in many pathological situations in cells or tissues (Goll et al., 2003). The

purported role of the calpains in various pathologic conditions is not well understood.

It is suggested that loss of Ca2+

homeostasis results in an elevation of intracellular

Ca2+

concentrations. This increased Ca2+

concentrations may then activate the calpain

system, leading to the increased protein degradation in these situations.

An increase in intracellular Ca2+

concentration is a common feature of cardiac

ischemia injury (Allen and Orchard., 1987), and may lead to extensive

Ca2+

-dependent proteolytic damage to the myocardium (Iizuka et al., 1991; Carrozza

et al., 1992; Gao et al., 1996; Matsumura et al., 1996). Ca2+

-activated calpain has been

shown to play a deleterious role in the heart during ischemia-reperfusion (Carrozza et

al., 1992; Singh et al., 2004) and calpain inhibition significantly attenuates myocardial

injury (Iizuka et al., 1991; Yoshikawa et al., 2005). Calpains tend to be concentrated

in the Z-disk, the site where disassembly of the sarcomere is initiated (Kumamoto et

al., 1992; Huang and Forsberg, 1998). Compelling evidence show that calpains

initiate degradation of Z line associated cytoskeletal proteins, including α-actinin, as

well as the regulatory proteins troponin C and troponin I, in the ischemic/reperfused

heart (Matsumura et al., 1996; McDonough et al., 1999; Papp et al., 2000).

32

While calpain activity has been extensively studied in cardiac muscle during

ischemia/hypoxia, its activity in skeletal muscle under similar conditions is poorly

understood. Ischemia and hypoxia have important health and performance

implications for skeletal muscles. For instance, hypoxia frequently occurs in several

respiratory diseases, including chronic obstructive pulmonary disease, acute

respiratory distress syndrome, and severe pneumonia (Zhu et al., 2003; Franssen et al.,

2002; Jardim et al., 1981), and may impair respiratory muscle function. It may also

occur following arterial clamping during reconstructive surgery or during tourniquet

use in surgery (Barry et al., 1997; Wakai et al., 2001). Finally, healthy muscles may

experience reduced oxygen delivery during high intensity exercise and the resulting

hypoxia may be a contributing factor to the development of fatigue (Bylund-Fellenius

et al., 1981; Howlett and Hogan, 2007). Since cardiac and skeletal muscles have

significant differences in their mechanisms of activation, metabolism, fatigability and

contractility, the results of studies performed on ischemic cardiac muscles may not be

generalizable to skeletal muscles.

In the present study, we tested the hypothesis that calpains are activated in

oxidative and glycolytic skeletal muscles subjected to hypoxic conditions. We further

hypothesized that if calpains were activated, this activation would contribute to

impaired muscle function. Calpain activity is difficult to measure directly in vivo.

Therefore, we used degradation of a highly sensitive and specific calpain substrate,

�-fodrin, as an index of in vivo calpain activity. To assess muscle function we

quantified power output as the muscle contracted and relaxed during cyclical changes

33

in its length. This procedure is designed to mimic muscle activity during locomotion,

ventilation, and other repetitive activities (Josephson, 1999; James et al., 1995, 1996).

The “work loops” produced by this procedure are dependent on the muscle

length-tension relationship, its force-velocity relationship, and its ability to activate

and deactivate the contractile proteins (Askew and Marsh, 1998).

METHODS

Animals and muscle preparation

This study was approved by the Oregon State University Institutional Animal

Care and Use Committee. Adult male ICR mice (average body mass 26.0 ± 1.3 g)

were obtained from Harlan (Harlan Sprague Dawley, Indianapolis, IN). All animals

were housed in university facilities (22°C with a 12:12 hour light-dark cycle).

Animals were maintained on a standard diet of rodent chow and tap water.

Animals were anesthetized with an i.p. injection of sodium pentobarbital (40

mg/g body mass). Soleus (SOL) and extensor digitorum longus (EDL) muscles were

removed and the animals euthanatized. Muscles were placed in a bicarbonate

Krebs-Ringer solution (composition in mM: 137 NaCl, 5 KCl, 1.25 CaCl2, 1.0 MgSO4,

1.0 NaH2PO4, 24 NaHCO3, 11 glucose and 0.025 tubocurarine chloride) maintained at

room temperature and aerated with 95% O2, 5% CO2. Silk suture was attached to both

tendons and the muscle was transferred to a water-jacketed chamber containing the

Krebs-Ringer solution (35oC, aerated with 95% O2, 5% CO2). Each muscle was

mounted vertically between two parallel platinum electrodes by attaching the

34

proximal tendon to a stationary hook, and the distal tendon to the lever arm of a

dual-mode muscle lever system (model 300B-LR or model 305C, Aurora Scientific,

Aurora, Ontario, Canada). In general, two EDL and two SOL muscles were studied

from each animal. EDL muscles were always studied before the SOL muscles because

like others (James.et al., 2004), we found that several hours of incubation at room

temperature had minimal effects on SOL contractile function.

Experimental Design

Pairs of muscles were studied simultaneously (i.e. two SOL muscles, or two

EDL muscles), with each muscle subjected to a different treatment. All muscles

followed a similar experimental protocol: 1) thermoequilibration for 20 min under

standard conditions, i.e. aerated with 95% O2, 5% CO2 , 2) determination of optimal

muscle length (Lo), 3) assessment of pre-treatment isometric force and power output,

4) administration of the experimental treatment for 60 min, 5) evaluation of

post-treatment isometric force and power output under the same condition as the

experimental treatment, 6) recovery for 30 min under standard conditions, and 7)

evaluation of post-recovery isometric force and power output at the standard

conditions.

The experimental treatments were as follows. The control treatment (con)

consisted of the standard condition, ie., bicarbonate Ringer aerated with 95% O2-5%

CO2. For the hypoxia treatment (hyp), the Ringer was aerated with 95% N2-5% CO2.

To verify calpain activity during hypoxia, a stock solution of E-64d (Peptide Institute

Inc., Osaka, Japan) was made by dissolving this selective calpain inhibitor in dimethyl

35

sulfoxide (DMSO, D2650, Sigma-Aldrich, St. Loius, MO). This stock solution was

added to the Ringer solution immediately before the experimental treatment to give

final E-64d and DMSO concentrations of 120 µM and 0.6%, respectively (hyp-E-64d).

The fourth experimental treatment consisted of Ringer solution containing the same

concentration of DMSO (0.6%) added to the Ringer solution immediately before

changing the gas to 95% N2-5% CO2 (hyp-DMSO).

Assessment of Muscle Function

Muscles were stimulated supramaximally by a constant current muscle

stimulator (model 701, Aurora Scientific, Inc.). Stimulus duration was 200 µs. For

EDL muscles, an isometric tetanic contraction was elicited by subjecting the muscle

to a 150 ms train of stimuli at 300 Hz. Corresponding parameters for SOL muscles

were 500 ms and 200 Hz. Muscle length was adjusted to obtain maximal isometric

tetanic force (Po), and this muscle length was defined as optimal muscle length (Lo).

Optimal muscle length was measured with a fine caliper. Stimulation current was

adjusted to ~ 25% above that which elicited peak isometric force. Current level and Lo

were maintained for all subsequent measurements. A rest period of 3 min was given

between each tetanic contraction.

Cyclical power output was measured using the work loop technique. The work

loop technique provides a realistic approach for studying the capacity of skeletal

muscle to produce mechanical work and power during cyclical changes in length that

mimic those that occur during locomotion (Josephson, 1999; James et al., 1995, 1996).

Both EDL and SOL were subjected to sinusoidal changes in length at an amplitude of

36

± 5% of fiber length (FL). Fiber length was calculated from Lo assuming a FL/Lo of

0.44 for mouse EDL (Warren et al., 1994 ) and 0.71 for mouse SOL (Warren et al.,

1994 ; Widrick and Barker, 2006). Sinusoidal changes in length were administered to

SOL muscles at a frequency of 5 Hz and to EDL muscles at a frequency of 8 Hz. The

strain amplitude and cycle frequency used here optimizes cyclical power output of

these muscles (James et al., 1995, 2004; Askew et al., 1997).

The treatment protocol for EDL muscles consisted of 20 sets of work loops, with

each set consisting of one passive work loop and 5 active work loops. Sets were

repeated every three minutes. SOL muscles were also subjected to 20 sets of work

loops with each set repeated once every 3 min. However, each set was comprised of

one passive loop followed by 15 active work loops. Like the EDL, each set was

repeated once every three minutes. SOL muscles were subjected to more work loops

in each set to more closely equate the energetic demands placed upon the EDL and

SOL muscles (Barclay et al., 1993). The pre-treatment work loop protocol was

comprised of a set of 1 or 3 active cycles for EDL and SOL, respectively. In

preliminary experiments we, like others, observed that peak power of the SOL was

not attained until the second work loop (Askew and Marsh, 1998), while the peak

power output of EDL was attained on the first cycle (James, et al., 1995). Thus, the

second work loop of each set was used to represent peak power output for the SOL,

while the first work loop was used for the EDL.

Preliminary experiments were conducted to determine the appropriate time for

initiation and termination of stimulus trains in order to maximize power output.

37

Cyclical power of SOL muscles was maximized with a 65 ms stimulus train initiated

35 ms before the muscle attained maximal length. Cyclical power of EDL muscles

was maximized with a 60 ms stimulus train initiated 15 ms before the muscle attained

maximal length. Thus, SOL muscles received 15 stimuli every complete loop while

EDL muscles received 5. These stimulation timing patterns are in agreement with

literature values (James, et al., 1995, 2004; Askew et al., 1997).

Experiments were controlled by a PC running a customized program written in

our laboratory (using LabView 67.1, National Instruments). Force and length data

were recorded for further analysis.

After completion of the contractile measurements, the muscles were removed

from the instrumentation, the sutures and tendons trimmed and the muscle mass

determined. Muscles were then quickly frozen in liquid nitrogen and stored at -80° for

Western blot analysis.

Gel Electrophoresis

The frozen muscles were homogenized using a high shear tissue homogenizer

(TH, Onmi international, Marictta, GA) in a Tris-HCL buffer (pH 7.4) containing 20

mM Tris-HCl, 5 mM EDTA, 5 mM EGTA, 1 mM DTT, and a protease inhibitor

cocktail (EGTA-Free Protease Inhibitor, Boehringer Mannheim, Indianapolis, IN).

The homogenate was centrifuged at 10,000 g for 10 min. The protein concentration of

the supernatant was determined using the Bio-Rad RC DC Protein Assay Kit II

(500-0122, Bio-Rad, Hercules, CA) with bovine serum albumin as a standard.

Muscle proteins were diluted to a concentration of 0.8 ug/ul using 4X LDS

38

(lithium dodecyl-sulfate) sample buffer (Invitrogen, Carlsbad, CA) and denatured by

heating at 70° for 10 min. Proteins (12 µg/lane) were separated on 4-12% NuPAGE

Novex Bis-Tris gels (Invitrogen, Carlsbad, CA). Electrophoresis was carried out at a

constant voltage (120 V) for 5 hr at room temperature.

Western Blots

To assess calpain activity, calpain-specific cleavage products of α-fodrin were

analyzed using Western blots. Calpains are the only known proteases that degrade

intact 240 kDa fodrin into 145 and 150 kDa positive peptides, and the appearance of

145/150 kDa α-fodrin positive signal by Western blot is considered one of the most

conclusive indicators of in vivo calpain activity (Goll et al., 2003). These 145/150 kDa

α-fodrin breakdown products retain their immunoreactivity to the primary antibody

used in this study.

Proteins were transferred from gels to nitrocellulose membranes (0.2 µm,

Invitrogen, Carlsbad, CA) in buffer containing 25 mM Bis-Tris, 25 mM Bicine, and

1mM EDTA at a constant voltage (45 V) for 2 hr at room temperature. Following

transfer, the membrane was blocked for 1 hr with 5% nonfat dry milk in a Tween-Tris

buffered saline solution (TTBS: 50 mM tris-HCl, 150 mM NaCl, 0.05% Tween-20,

pH 7.4), followed by incubation with mouse anti-spectrin antibody (1:1000,

MAB1622, Chemicon, Temecula, CA) either overnight at 4oC or 2.5 hr at room

temperature. Blots were then incubated for 1 hr with an anti-mouse IgG-horseradish

peroxidase (HRP)-conjugated secondary antibody (1:2,000, Bio-Rad, Hercules, CA).

Immunoreactive bands were visualized using the Pierce SuperSignal West Femto

39

Maximum Sensitivity Substrate Kit (34094, Pierce Biotechnology, Inc., Rockford, IL)

and imaged on an Alpha Innotech imaging system (San Leandro, CA).

Data Reduction

A plot of the force attained during each complete cycle of shortening and

lengthening against length produced a “work loop.” The area bounded by the work

loop represents the net work produced by the muscle. The net work equals the work

done by the muscle during shortening (positive work) minus the work done on the

muscle during lengthening (negative work). The net power output of muscle was

calculated as the net work performed per second. The peak stress during shortening

was determined from the force records.

Muscle cross-sectional area was calculated by dividing muscle mass by the

product of fiber length and muscle density (assumed to be 1.06 mg/mm3, Mendez and

Keys, 1960). Peak force and cyclical power output were normalized to cross-sectional

area and muscle mass, respectively.

The optical density of the intact α-fodrin band and the 145/150 kDa breakdown

fragments were quantified by Image J software. To reduce variability between blots,

the optical density of each lane was normalized to the optical density of the control

treatments on the same blot. Values were then expressed as a percentage of the

control treatment. Thus, control samples would have a normalized 145/150 kDa

optical density of 100% and samples expressing greater 145/150 kDa optical density

would have a normalized density value greater than this. Because two control

treatments were present on each blot, the final optical density value of each sample

40

was the mean of two normalized values.

Statistical Methods

All data are presented as means ± SE. For EDL muscles, the optical density of

the combined 145/150 α-fodrin positive bands was analyzed using a one-way

ANOVA. For SOL muscles, a one-way Kruskal-Wallis ANOVA was employed

because the optimal density measurements were not normally distributed. Peak force

and power were analyzed at the pre-, post-treatment, and post recover time points

using a two-way ANOVA with a main effect of experimental treatment and a repeated

effect of time. In all cases, significant main effects and interactions were further

analyzed using the Student-Newman-Keuls post hoc test. Statistical analysis was

performed using SigmaStat 3.0 with a type I error rate of P < 0.05.

RESULTS

Part I. Contractile properties

The physical properties of the mouse EDL and SOL muscles are presented in

Table 3.1. Prior to treatment, no inter-treatment differences were found in muscle

mass, Lo, or physiological cross-section area (CSA) across the four experimental

treatments for either EDL or SOL muscles.

The mean pre-treatment maximum isometric force was 224.6 ± 3.8 kN/m2

for

EDL and 235.6 ± 4.8 kN/m2 for SOL muscles (Table 3.2). These are similar to the

values of 233 and 224 kN/m2 reported by James et al. (1995) for mouse EDL and SOL

muscles, respectively. The mean pre-treatment maximum net power output was 37.8 ±

41

1.2 W/kg (at 5 Hz) and 108.5 ± 3.3 W/kg (at 8 Hz) for SOL and EDL muscles,

respectively (Table 3.3). Our value for the SOL is slightly higher than the maximum

power output of 33.2 W/kg reported by Askew et al. (1997) at 37oC and 34.02 W/kg

obtained by James et al. (1995) at 35oC. Our value for the EDL is slightly higher than

the value of ~98 W/kg reported by James et al. (1995) at 8 Hz and similar to the value

of 107.2 W/kg that these authors obtained at 10-12 Hz. We opted to study the EDL at

8 Hz because 8 Hz represents the upper limit of the limb cycle frequency of mice

during treadmill running (James et al., 1995).

Small, but significant losses in Po (~12%) and peak power (~16%) occurred

across the control treatment for EDL muscles (Table 3.2, 3.3; Fig 3.1, 3.3, 3.5, 3.6).

Most of this appeared to be due to run-down of the preparations, as there was no

recovery following the treatment. In contrast to the EDL, power and Po of the control

SOL remained relatively stable during the treatment period.

Hypoxia reduced EDL (p<0.001) and SOL (p<0.001) peak isometric force and

this response was not significantly affected by E-64d or DMSO treatments (Table 3.2,

Fig 3.1, 3.2, 3.5). Peak power output was also significantly reduced after 60 min of

hypoxia treatment in EDL (p<0.001) and SOL muscles (p<0.001) and again, this was

not significantly altered by the presence of E-64d or DMSO (Table 3.3, Fig 3.3, 3.4,

3.6). Both Po and peak power output showed only partial recovery following 30 min

of reoxygenation.

Part II. Calpain Activity

As illustrated in Figs 3.7 and 3.8, hypoxia induced a significant increase in the

42

normalized optical density of 145/150-kDa α-fodrin fragments in both EDL (P<0.05)

and SOL (P<0.05). Importantly, the hypoxia-induced increase in 145/150-kDa

α-fodrin fragments was prevented by the E-64 treatment for both muscles. The

addition of 0.6% DMSO to the tissue bath had no effect on the normalized optical

density of 145/150 bands under hypoxic conditions, indicating that it was the E-64

and not the DMSO in the E-64 trial that prevented α-fodrin degradation. No

significant differences in normalized optical density of the intact (240 kDa) α-fodrin

bands were found among four experimental groups.

DISCUSSION

The main findings of this study are, 1) the increase in α-fodrin-positive 145 and

150 peptides after contractile activity under hypoxia conditions, and 2) the

disappearance of these peptides after treatment with the cysteine protease inhibitor

E-64d. Alpha-fodrin (also termed non-erythroid α-spectrin or α-spectrin-II), is a well

characterized calpain proteolytic substrate (Huang and Forsberg, 1998; Goll et al.,

2003). Alpha-fodrin 145/150 kDa breakdown fragments are widely accepted as a

sensitive marker of in vivo proteolytic activity of calpains (Goll et al., 2003; Huang

and Forsberg, 1998; Tsuji et al, 2001). Our finding that 145/150 kDa α-fodrin positive

fragments are elevated after the hyp treatment is evidence that calpains are activated

in skeletal muscles exposed to hypoxia.

We inhibited calpain activity with the low-molecular weight selective inhibitor

E64-d. E-64, or trans–L-epoxysuccinyl-leucylamido (4guanidino) butane, and its

derivates E-64c and E-64d, irreversibly inactivate cysteine proteases by forming a

43

covalent bond with the active site thiolate (Tamai et al., 1981). E-64 is therefore a

potent inhibitor of the calpains as well as cathepsins B, H, and L (Barrett et al., 1982).

It is important to note that while cathepsins can degrade α-fodrin, the 145 kDa

α-fodrin positive fragment is produced only by the proteolytic activity of calpain

(Nath et al., 1996). Thus, our finding that E-64d inhibited the appearance of the

145/150 kDa α-fodrin positive fragment strengthens our conclusion that calpains are

activated in skeletal muscles under the hypoxic conditions studied here.

Taken together, these findings support our hypothesis that calpains are activated

in active glycolytic and oxidative skeletal muscles upon exposure to hypoxia.

Calpains have been found to be activated in several other tissues subjected to

physiological stress, including the brain (Lee et al., 1991) and renal tubular cells

(Weinberg, 1985) as well as the myocardium (Iizuka et al., 1991; Matsumura et al.,

1995; Yoshikawa et al., 2005; Singh et al., 2004), The present results are one of the

first instances in which in vivo calpain proteolytic activity has been demonstrated in

skeletal muscles under physiological stress.

The second finding of this study is that inhibition of calpain activity had no

effect on peak power output and isometric force at the end of the 60 min of hypoxic

treatment, or after 30 min of recovery under standard conditions, for either the EDL or

the SOL. Thus, we reject our hypothesis that calpain inhibition would prevent

contractile dysfunction during hypoxia.

Our conclusion differ’s from results obtained on ischemic or hypoxic cardiac

muscle. For instance, Matsumura et al. (1995) found that the protease inhibitor

44

leupeptin protected the myocardium against ischemia. Others have reported that

hearts treated with calpain inhibitors showed greater recovery of contractile function

following ischemia/reperfusion or Ca2+

overload (Tsuji et al., 2001; Singh et al., 2004;

Yoshikawa et al., 2005).

There are several possibilities why the present results differ from those

conducted on cardiac muscle. First, much of the cardiac work has been conducted

using models of ischemia/reperfusion. Thus, inherent differences between models of

ischemic/reperfusion and hypoxia could contribute to why the present findings differ

from studies on cardiac muscles.

Second, calpain inhibition may have had a positive effect on function, but our

protocol was unable to detect this difference. For instance, E-64d was dissolved in

dimethyl sulfoxide (DMSO). Because DMSO is a scavenger of hydroxyl radicals

(Byler et al., 1994), and may by itself protect against hypoxia/ischemia-induced injury

(Mohanraj et al., 1998), we included a DMSO-only trial in the present study. We

found that the DMSO only treatment appeared to have a slight detrimental effect on

power output during hypoxia, though it did not reach statistical significance. Reid and

Moody (1994) concluded that DMSO may cause inhibition of the

excitation-contraction coupling process and DMSO has been shown to reduce the

utilization of oxygen by inhibiting mitochondrial function (Ghosh et al., 1976).

These small effects of DMSO on function may be important because power

appeared to be somewhat elevated in the hypI treatment vs. its negative control, the

hypD treatment. In fact, a one-way ANOVA performed on the post-treatment hypD

45

and hypI peak power values indicates a significant difference at the p < 0.10 level.

Thus, it is possible that the solvent DMSO confounded our interpretation of results. It

is important to note that DMSO had no effect on the appearance of α-fodrin -positive

145/150 kDa peptides during hypoxia.

A third reason for the differences between our results and findings in cardiac

preparations is that calpain could have differential effects on the processes of

activation in these muscles. In cardiac cells, ischemia-reperfusion causes a decrease in

sarcoplasmic reticulum Ca2+

-induced Ca2+

release (Temsah et al., 1999). This

ligand-gated mechanism of calcium release is thought to be of primary importance in

the activation of cardiac muscle but only of secondary importance in skeletal muscle.

Hypoxia has also been shown to result in degradation and loss of thin filament

regulatory proteins in skeletal muscle (Brotto et al., 2001) and in post-ischemic

stunned myocardium (Gao et al., 1997). The functional impact of a loss of thin

filament regulatory proteins would be expected to be greater in cardiac muscle, where

activation during contraction is sub-maximal, compared to skeletal muscle cells,

where intracellular Ca2+

levels saturate the thin filament regulatory proteins. Thus,

differences between cardiac and skeletal muscle intracellular Ca2+

levels and

mechanisms of Ca2+

release could be responsible for the differential effects of calpain

on striated muscle function during hypoxia.

In summary, the present work provides strong evidence for calpain activation in

glycolytic and oxidative muscles subjected to hypoxia. In this regard, our results are

similar to previous work conducted on cardiac muscle preparations. However, unlike

46

these previous findings, we found no conclusive evidence that inhibition of

calpain-mediated proteolytic activity improved skeletal muscle function during

hypoxia or recovery.

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Table 3.1: Characteristics of EDL and SOL muscles

Values are mean ± SE, (N= number of muscle). Abbreviations: Lo, optimal length; CSA, physiological cross-sectional area; con, control

group; hyp, hypoxia treatment group; hyp-D, hypoxia treatment plus dimethyl sulfoxide (DMSO); hyp-I, hypoxia treatment plus calpain

inhibitor ( E-64d) plus DMSO..

Treatment

Muscle Variable

con hyp hyp-D hyp-I

mean

Muscle mass, mg 9.8 ± 0.6 (N=6) 9.5 ± 0.3 (N=7) 9.5 ± 0.1 (N=6) 9.8 ± 0.4 (N=8) 9.6 ± 0.2

Lo, mm 12.4 ± 0.4 11.9 ± 0.4 11.7 ± 0.2 12.5 ± 0.4 12.1 ± 0.2 EDL

CSA , mm2 1.69 ± 0.06 1.72 ± 0.04 1.75 ± 0.03 1.67 ± 0.03 1.70 ± 0.02

Muscle mass, mg 6.3 ± 0.4 (N=6) 6.8 ± 0.6 (N=7) 6.8 ± 0.3 (N=7) 7.0 ± 0.4 (N=8) 6.7 ± 0.2

Lo, mm 11.2 ± 0.3 11.3 ± 0.5 11.1 ± 0.6 11.4 ± 0.6 11.3 ± 0.1 SOL

CSA , mm2 0.74 ± 0.04 0.80 ± 0.06 0.81 ± 0.02 0.81 ± 0.03 0 .79 ± 0.01

54

Table 3.2: Peak isometric force of EDL and SOL muscles

Normalized peak isometric force (force per physiological cross sectional area, kN/m2) in control (con), hypoxia (hyp), hypoxia with

DMSO (hyp-D) and hypoxia with E-64d (hyp-I) are shown. Values are mean ± SE. (N= number of muscle).

* p<0.001 compared with initial value; # p<0.001 compared with control group

Muscle Time con hyp hyp-D hyp-I

pre-treatment 237.3 ± 6.5 (N=6) 221.1 ± 7.4 (N=7) 214.2 ± 3.5 (N=6) 225.9 ± 9.0 (N=8)

post-treatment 208.0 ± 7.0 * 45.3 ± 4.9 *# 34.3 ± 2.4 *# 52.3 ± 6.8 *# EDL

Post-recovery 209.1 ± 6.9 * 85.2 ± 6.0 *# 69.6 ± 4.3 *# 98.7 ± 12.5*#

pre-treatment 240.2 ± 13.1 (N=6) 245.2 ± 8.6 (N=7) 231.8 ± 8.1 (N=7) 227.3 ± 9.5 (N=8)

post-treatment 238.8 ± 13.4 138.8 ± 4.4 *# 119.3 ± 11.1 *# 130.5 ± 6.5 *# SOL

Post-recovery 233.8 ± 13.5 167.5 ± 6.7 *# 141.0 ± 13.7*# 153.8 ± 8.0 *#

55

Table 3.3: Peak power of EDL and SOL muscles

Peak power output per kg muscle (W/kg) in control (con), hypoxia (hyp), hypoxia with DMSO (hyp-D) and hypoxia with E-64d (hyp-I)

are shown here. Values are mean ± SE.(N= number of muscle).

* p<0.001 compared with initial value; # p<0.001 compared with control group.

Muscle Time con hyp hyp-D hyp-I

pre-treatment 108.5 ± 3.3 (N=6) 102.0 ± 2.6 (N=7) 98.3 ± 2.0 (N=6) 101.1 ± 4.2 (N=8)

post-treatment 92.3 ± 4.1 * 10.7 ± 1.9 *# 5.7 ± 1.3 *# 11.9 ± 2.8 *# EDL

Post-recovery 96.7 ± 3.6 * 33.8 ± 1.8 *# 26.6 ± 1.5 *# 36.0 ± 4.6 *#

pre-treatment 39.2 ± 2.9 (N=6) 40.8 ± 2.0 (N=7) 37.0 ± 2.1 (N=7) 34.6 ± 2.3 (N=8)

post-treatment 38.0 ± 2.9 13.4 ± 3.3 *# 10.7 ± 1.2 *# 10.8 ± 1.3 *# SOL

Post-recovery 38.7 ± 2.8 26.0 ± 1.5 *# 20.8 ± 2.1 *# 21.2 ± 1.3 *#

56

time (sec)

0.0 0.1 0.2 0.3 0.4 0.5

iso

me

tric

fo

rce (

mN

))

0

100

200

300

400

pre

post

rec

time (sec)

0.0 0.1 0.2 0.3 0.4 0.5

iso

metr

ic f

orc

e (

mN

)

0

100

200

300

400

pre

post

rec

time (sec)

0.0 0.1 0.2 0.3 0.4 0.5

iso

metr

ic f

orc

e (

mN

)

0

100

200

300

400

pre

post

rec

time (sec)

0.0 0.1 0.2 0.3 0.4 0.5

iso

metr

ic f

orc

e (

mN

)

0

100

200

300

400

pre

post

rec

Fig 3.1 Representative isometric

force of EDL muscle during pre-,

post-treatment and recovery

periods in control (con), hypoxia

(hyp), hypoxia with DMSO

(hyp-D) and hypoxia with

inhibitor (hyp-I) groups.

con hyp

hyp-D hyp-I

57

time (sec)

0.0 0.2 0.4 0.6 0.8 1.0

iso

me

tric

fo

rce (

mN

)

0

50

100

150

200

250

pre

post

rec

time (sec)

0.0 0.2 0.4 0.6 0.8 1.0

iso

metr

ic f

orc

e (

mN

)

0

50

100

150

200

250

pre

post

rec

time (sec)

0.0 0.2 0.4 0.6 0.8 1.0

iso

metr

ic f

orc

e (

mN

)

0

50

100

150

200

250

pre

post

rec

time (sec)

0.0 0.2 0.4 0.6 0.8 1.0

iso

metr

ic f

orc

e (

mN

)

0

50

100

150

200

250

pre

post

rec

Fig 3.2 Representative isometric

force of SOL muscle during pre-,

post-treatment and recovery

periods in control (con), hypoxia

(hyp), hypoxia with DMSO

(hyp-D) and hypoxia with

inhibitor (hyp-I) groups.

hyp-D hyp-I

con hyp

58

length (mm)

-0.3 -0.2 -0.1 0.0 0.1 0.2 0.3

forc

e (

mN

)

0

50

100

150

200

250

300

350

pre

post

rec

length (mm)

-0.3 -0.2 -0.1 0.0 0.1 0.2 0.3

forc

e (

mN

)

0

50

100

150

200

250

300

pre

post

rec

length (mm)

-0.3 -0.2 -0.1 0.0 0.1 0.2 0.3

forc

e (

mN

)

0

50

100

150

200

250

300

pre

post

rec

length (mm)

-0.3 -0.2 -0.1 0.0 0.1 0.2 0.3

forc

e (

mN

)

0

50

100

150

200

250

300

pre

post

rec

Fig 3.3 Representative peak

power output of EDL muscle

during pre-, post-treatment and

recovery periods in control

(con), hypoxia (hyp), hypoxia

with DMSO (hyp-D) and

hypoxia with inhibitor (hyp-I)

groups.

hyp-D

hyp con

hyp-I

59

length (mm)

-0.6 -0.4 -0.2 0.0 0.2 0.4 0.6

forc

e (

mN

)

0

20

40

60

80

100 prepost

rec

length (mm)

-0.6 -0.4 -0.2 0.0 0.2 0.4 0.6

forc

e (

mN

)

0

20

40

60

80

100

120pre

post

rec

length (mm)

-0.6 -0.4 -0.2 0.0 0.2 0.4 0.6

forc

e (

mN

)

0

20

40

60

80

100

120

140pre

post

rec

length (mm)

-0.6 -0.4 -0.2 0.0 0.2 0.4 0.6

forc

e (

mN

)

0

20

40

60

80

100

120pre

post

rec

Fig 3.4 Representative peak

power output of SOL muscle

during pre-, post-treatment

and recovery periods in

control (con), hypoxia (hyp),

hypoxia with DMSO (hyp-D)

and hypoxia with inhibitor

(hyp-I) groups.

60

Time

pre post recovery

Po

(%

of

init

ial)

0

20

40

60

80

100

120

con (n=6) hyp (n=7)hyp-D (n=6)hyp-I (n=8)

Time

pre post Recovery

Po

(%

of

init

ial)

0

20

40

60

80

100

120

con (n=6)hyp (n=7)hyp-D (n=7) hyp-I (n=8)

Fig 3.5 Isometric force of pre-treatment, post-treatment and reoxygenation (recovery)

in control (con), hypoxia (hyp), hypoxia with DMSO (hyp-D) and hypoxia with

E-64d (hyp-I). A: EDL muscles. B: SOL muscles. Data are means ± SE and expressed

as a percentage of the initial value. * p<0.001 compared with pre-treatment value

B

A

* * *

* * *

* * *

* * *

61

Time

pre post Recovery

Pe

ak

po

we

r (%

of

init

ial)

0

20

40

60

80

100

120

con (n=6) hyp (n=7)hyp-D (n=6) hyp-I (n=8)

Time

pre post Recovery

Pe

ak

po

we

r (%

of

init

ial)

0

20

40

60

80

100

120

con (n=6)

hyp (n=7)

hyp-D (n=7)

hyp-I (n=8)

Fig 3.6 Power output of pre-treatment, post-treatment and reoxygenation (recovery)

in control (con), hypoxia (hyp), hypoxia with DMSO (hyp-D) and hypoxia with

E-64d (hyp-I). A: EDL muscles. B: SOL muscles. Data are means ± SE and expressed

as a percentage of the initial value. * p<0.001 compared with pre-treatment value

B

A

* * *

* * *

* * *

* * *

62

Treatment

con hyp hyp-D hyp-I

14

5/1

50

kD

a %

of

Co

ntr

ol

0

50

100

150

200

250

300

Fig 3.7 Analysis of calpain-cleaved α-fodrin in EDL muscle. A: representative

western blot illustrating intact α-fodrin (240 kDa) and its 145/150 kDa breakdown

products in control (con), hypoxia (hyp), hypoxia with DMSO (hyp-D) and hypoxia

with inhibitor (hyp-I) groups. B: Semi-quantitative analysis of α-fodrin breakdown

products, expressed as percentage of combined 145/150 kDa fragment to the control

value. * Significantly different from the control (p<0.05)

A

B

* * *

150 kDa

145 kDa

240 kDa

63

Treatment

con hyp hyp-D hyp-I

14

5/1

50 k

Da

% o

f C

on

tro

l

0

100

200

300

400

Fig 3.8 Analysis of calpain-cleaved α-fodrin in SOL muscle. A: representative

western blot illustrating intact α-fodrin (240 kDa) and its 145/150 kDa breakdown

products in control (con), hypoxia (hyp), hypoxia with DMSO (hyp-D) and hypoxia

with inhibitor (hyp-I) groups. B: Semi-quantitative analysis of α-fodrin breakdown

products, expressed as percentage of combined 145/150 kDa fragment to the control

value. * Significantly different from the control (p<0.05)

*

240 kDa

150 kDa

145 kDa

B

A

*

64

Contractile Function and Calpain Activity in Mouse Skeletal Muscle

During Hypoxia

Chapter 4

CONCLUSION

This project examined the contractile function of skeletal muscles subjected to

hypoxia and whether changes in their function could be attributed to the activation of

the calcium-dependent protease calpain. Function was evaluated using the work loop

technique and calpain mediated proteolysis was evaluated by the appearance of two

�-fodrin breakdown peptides (145/150 kDa) on Western blots prepared following

each experimental condition. To further understand the role of calpain after

contractile activity under hypoxic conditions, �-fodrin breakdown was also evaluated

in the presence of E-64d, a calpain inhibitor. Isolated EDL and SOL muscles from

mice were studied because these represent glycolytic and oxidative muscles,

respectively. Hypoxia was implemented by equilibrating the isolated muscles with

95% N2, 5% CO2 compared with the standard condition of 95% O2, 5% CO2.

We tested a possible mechanism responsible for the impaired muscle function

observed during hypoxia. In cardiac muscle, it has been proposed that the

Ca2+

-dependent protease calpain contributes to contractile dysfunction under hypoxic

conditions. Therefore, in order to understand the role of calpains in skeletal muscle

during hypoxia, we evaluated calpain activity after hypoxia treatment.

65

Degradation of the calpain substrate α-fodrin was used as a marker of in vivo

calpain activity. In order to test that calpain activity had an effect on contractile

function, we conducted additional trials in which muscles were exposed to E-64d, a

selective calpain inhibitor. DMSO was used to dissolve E-64d, and because DMSO

may have independent effects on muscle function, a DMSO only trial was included in

the experimental design.

Our results demonstrated that hypoxia caused severe reductions in isometric

force and sustained cyclical power output of both EDL and SOL muscles. Hypoxia

also induced proteolysis of the sensitive and specific calpain substrate α-fodrin in

both muscles. In both muscles, proteolysis of α-fodrin was inhibited by treatment

with the cysteine protease inhibitor E-64d. However, treatment with E-64d had little

effect on the changes in force and power that occurred during hypoxia. Thus, we

concluded that calpain activity increases under hypoxic conditions but this activity

does not contribute to skeletal muscle contractile dysfunction.

A limitation to our approach is that DMSO appeared to have a slight depressive

effect on muscle function. This may have confounded our interpretation of results.

Future studies may want to use other calpain inhibitors that eliminate this possibility.

In summary, this project demonstrated that degradation of α-fodrin is increased in

oxidative and glycolytic skeletal muscles exposed to hypoxia. The observed α-fodrin

breakdown fragments were consistent with calpain-mediated degradation of this

protein, and because these fragments were reduced in muscles exposed to the calpain

66

inhibitor E-64d, our findings suggest that calpains are activated in skeletal muscles

exposed to hypoxia. However, unlike cardiac muscle, calpain-mediated proteolysis

did not appear to be a major cause of skeletal muscle dysfunction under these

conditions.