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Potential factors contributing to diaphragm myopathy in congestive heart failure

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Potential factors contributing to diaphragm myopathy in congestive heart failure

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Content POTENTIAL FACTORS CONTRIBUTING TO DIAPHRAGM
MYOPATHY IN CONGESTIVE HEART FAILURE
by
Jesus F. Dominguez
A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(BIOKINESIOLOGY)
May 2002
Copyright 2002 Jesus F. Dominguez
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UNIVERSITY OF SOUTHERN CALIFORNIA
The Graduate School
University Park
LOS ANGELES. CALIFORNIA 90089-1695
This d issertatio n, w ritte n b y
J V s a s F . b onuA/tjue-z.____________
4
U nder th e d ire c tio n o f A J.5T . D issertatio n
Com m i ttee, an d approved b y a ll its m em bers,
has been p resen ted to an d accepted b y The
G raduate School , in p a rtia l fu lfillm e n t o f
requirem ents fo r th e degree o f
D O C TO R O F P H ILO S O P H Y
------------------------------- .^ . < 3 --- '-----------------------
Z ' ' De t H o f Gr a d u a t e St u di e s
D a te May 10. 2002___________
DI SSER TA n O N C O M M l
C h a i r p e r s o n
copyright owner. Further reproduction prohibited without permission.
TABLE OF CONTENTS
LIST OF TABLES................................................................................................ v
LIST OF FIGURES.............................................................................................vi
ABSTRACT........................................................................................................vii
RESEARCH QUESTION AND SPECIFIC AIMS...............................................1
CHAPTER I.
SIGNIFICANCE AND BACKGROUND...................................................3
Significance of the Research....................................................... 3
Background................................................................................... 4
Skeletal Myopathy in CHF....................................................4
Impaired Skeletal Muscle Ca2 - ” Homeostasis in CHF.............6
Apoptosis in CHF-lnduced Skeletal Myopathy.....................7
CHAPTER II.
SUMMARY OF THE EXPERIMENTAL DESIGN.................................11
CHAPTER III.
COMPARTMENTAL ANALYSIS OF STEADY-STATE
DIAPHRAGM Ca2 + KINETICS IN CHRONIC CHF.............................. 14
Introduction..................................................................................14
Methods.......................................................................................16
Development of CHF Animal Model.................................... 16
Single-Fiber Cross-Sectional Area......................................18
Analysis of Diaphragm Ca2 * Homeostasis Utilizing
4 5 Ca2 * Kinetics Technology.................................................20
Extraction and Assay for Calpain Activity............................27
Statistical Analysis............................................................. 29
Results.........................................................................................30
Characteristics of the Chronic CHF Animal Model.............. 30
Analysis of Single-Fiber Cross-Sectional Area....................32
Effects of CHF on Diaphragm 4 5 Ca2 * Efflux Data................ 32
Estimated Diaphragm Compartmental Ca2 * Contents
and Exchange Fluxes........................................................33
Relative Activity Level of Calpain within the Environment
of CHF.............................................................................. 35
Discussion...................................................................................36
Critique of Experimental Procedures...................................37
Kinetic Analysis and Fitting of Efflux Data...........................37
Assumptions...................................................................... 38
ii
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Altered Diaphragm Ca2 * Homeostasis in CHF....................40
Implications of Impaired Diaphragm Ca2 * Homeostasis
in CHF..............................................................................42
Increased Intracellular Diaphragm Ca2 *........................42
Increased Extracellular Diaphragm Ca2 *.......................45
Conclusions................................................................................ 46
CHAPTER IV.
POTENTIAL ROLE FOR APOPTOSIS IN CHF-INDUCED
DIAPHRAGM MYOPATHY...................................................................47
Introduction.................................................................................47
Methods......................................................................................48
Development of CHF Animal Model................................... 48
Quantitative Determination of PlasmaTNF-a......................50
Analysis of DNA Fragmentation in Diaphragm....................52
TUNEL...................................................................52
DNA Agarose Gel Electrophoresis...............................54
Analysis of Diaphragm Single-Fiber Architecture................57
Cross-Sectional Area.................................................57
Estimation of Myonuclear Domain Size........................ 58
Extraction and Assay for Caspase-3 Activity
in Diaphragm..................................................................... 58
Statistical Analysis.............................................................60
Results........................................................................................61
Characteristics of the Chronic CHF Animal Model.............. 61
Morphometric Analysis of Diaphragm Single Fibers............63
Plasma TNF-a Levels in CHF............................................ 64
TUNEL Analysis in Diaphragm...........................................65
DNA Laddering in Diaphragm............................................ 67
Relative Activity Level of Caspase-3 in CHF Diaphragm.... 67
Discussion.................................................................................. 68
Diaphragm Wasting in CHF ............................................ 68
Plasma TNF-a Levels Associated with CHF.......................70
Apoptotic Markers in Chronic CHF..................................... 71
Conclusions................................................................................ 74
CHAPTER V.
FINAL COMMENTS.............................................................................. 75
Limitations of the Study..............................................................75
Conclusions................................................................................ 77
REFERENCES................................................................................................. 78
CHAPTER 1 ............................................................................................78
CHAPTER II...........................................................................................86
iii
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CHAPTER III.........................................................................................86
CHAPTER IV.........................................................................................91
CHAPTER V .........................................................................................95
MASTER REFERENCE LIST.......................................................................... 97
iv
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LIST OF TABLES
Table 2.1 Development of the Chronic Animal Model and
Summary of the Experimental Design.......................................12
Table 3.1 Characteristics of the Chronic Animal Model.............................30
Table 3.2 Diaphragm Intercompartmental Ca2 * Exchange
Fluxes..........................................................................................34
Table 3.3 Diaphragm Compartmental Ca2 * Distribution............................ 35
Table 4.1 Characteristics of the Chronic Animal Model.............................62
v
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LIST OF FIGURES
Figure 3.1 Schematic Representation of 4 5 Ca2 + Efflux Experiment
Apparatus....................................................................................21
Figure 3.2 Schematic Drawing Depicting the Kinetic Model Used
to Simultaneously Fit 4 ®Ca2 + Efflux Data.................................. 22
Figure 3.3 Sensitivity of Data for Adjustable Parameters Calculated
from Partial Derivatives of Simulated Values........................... 24
Figure 3.4 Mean Diaphragm Single-Fiber CSA of SHAM and CHF
Animals........................................................................................31
Figure 3.5 Effects of CHF on Diaphragm 4 5 Ca2 + Efflux Data......................32
Figure 3.6 Effects of CHF on Diaphragm Compartmental Ca2 +
Contents......................................................................................33
Figure 3.7 Effects of CHF on Diaphragm Calpain Proteolytic
Activity......................................................................................... 36
Figure 4.1 Mean Diaphragm Single-Fiber CSA of SHAM and CHF
Animals........................................................................................62
Figure 4.2 Mean Diaphragm Single-Fiber CSA : Myonucleus Ratio
in SHAM and CHF Animals........................................................63
Figure 4.3 Effects of CHF on Mean Plasma TNF-a Levels.........................64
Figure 4.4 Interstitial Terminal Deoxynucleotidyl Transferase-
Mediated dUTP Nick End-Labeling (TUNELJ+ Nucleus
in the Diaphragm........................................................................ 65
Figure 4.5 Peripheral Myocyte Terminal Deoxynucleotidyl
Transferase-Mediated dUTP Nick End-Labeling
(TUNEL)+ Nucleus in the Diaphragm........................................66
Figure 4.6 Centralized Myocyte Terminal Deoxynucleotidyl
Transferase-Mediated dUTP Nick End-Labeling
(TUNEL)+ Nucleus in the Diaphragm........................................66
Figure 4.7 Effects of CHF on Diaphragm Caspase-3 Proteolytic
Activity.........................................................................................67
v i
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Calpain activity was increased by more than 2-fold (p<0.05) and caspase-3
activity was increased by 36% (p<0.05) in CHF diaphragm relative to SHAM.
Diaphragm from CHF animals was 27 times more likely to exhibit apoptotic-
like nuclear DNA fragmentation than diaphragm from sham-operated animals
(p<0.05, 95% Cl for OR = 2.34, 311). The present study revealed that: 1)
Diaphragm wasting occurs in the environment of CHF, 2) CHF is associated
with altered Ca2 * homeostasis that increases diaphragm compartmental Ca2 *
contents and exchange fluxes and may provide a signal for altering calpain
activity, and 3) apoptosis may play a role in CHF-associated diaphragm
wasting as suggested by increased caspase-3 activity and the presence of
apoptotic-like nuclear DNA fragmentation. The results of this study suggest
that multiple proteolytic mechanisms potentially contribute to diaphragm
wasting in CHF.
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RESEARCH QUESTION AND SPECIFIC AIMS
The hallmark of congestive heart failure (CHF) is exercise intolerance
characterized by symptoms of dyspnea and muscle fatigue. Previously,
symptomatic CHF was largely attributed to myocardial dysfunction and
circulatory insufficiency. However, it is now known that there is a poor
correlation between the degree of cardiac dysfunction and the capacity for
physical activity. Recent evidence suggests that a unique skeletal myopathy
develops within the environment of CHF that is characterized by reduced
muscle mass and impaired function. It is believed that skeletal myopathy
plays an important role in poor exercise tolerance and other disabling
symptoms that reduce the quality of life in CHF. Pathophysiologic
mechanisms underlying skeletal myopathy in CHF are not completely
understood, but it is clear that inactivity or disuse do not contribute
significantly to the skeletal muscle abnormalities. Perhaps the most
compelling example of the uncoupling between inactivity and muscle wasting
in CHF is found in the rhythmically contracting diaphragm. Yet until now,
diaphragm myopathy in CHF has not been well studied.
The purpose of this research project was to identify cellular and
molecular processes that potentially contribute to diaphragm myopathy in
CHF. Two major areas of research that have been identified as playing an
important role in cardiomyopathy and skeletal myopathy in CHF are altered
cellular Ca2 * homeostasis and apoptosis. Utilizing a coronary artery ligation
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model of CHF in the rat, diaphragm from CHF and sham-operated animals
was analyzed utilizing various morphometric, histochemical, and
immunohistochemical techniques. The specific aims of the study were to:
I: verify that diaphragm muscle mass was reduced in this
experimental animal model of CHF by measuring single-fiber
cross-sectional area and evaluating the relationship between
single-fiber cross-sectional area and myonuclei
II: characterize diaphragm Ca2 * homeostasis utilizing Ca2* tracer
kinetics technology and assess the activity status of Ca2 *-
dependent proteins (calpain)
III: explore various indicators of the apoptotic process (TUNEL,
DNA laddering) in CHF diaphragm and assess the activity
status of apoptotic proteases (caspase-3)
2
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CHAPTER I
SIGNIFICANCE AND BACKGROUND
SIGNIFICANCE OF THE RESEARCH
In the last decade, congestive heart failure (CHF) has emerged as the
most important and rapidly growing public health issue in cardiovascular
medicine. Recent reports indicate that more than 2,000,000 individuals in
the United States suffer from CHF, with approximately 400,000 new cases
diagnosed each year (67). Despite improved medical management of
coronary artery disease and hypertension, the incidence of CHF is steadily
on the rise and represents the single most common diagnosis for
hospitalization in patients over the age of 65 (25).
Dyspnea and exercise intolerance are the hallmark symptoms of CHF,
ultimately leading to marked functional impairment. A progressive decline in
exercise capacity is the cardinal manifestation of heart failure and serves as
the basis for classifying the severity of the disorder. Historically, research
focused on deterioration of hemodynamic status as the pathophysiologic
basis for the progression of CHF (13,15,20,46). Later, it was shown that the
exercise intolerance experienced by individuals with CHF correlates poorly
with indices of central hemodynamic function (19,26,49,71). It is now known
that CHF induces skeletal muscle abnormalities that may help explain the
genesis of limited exercise capacity (6,41-43,56). Maladaptive changes in
skeletal muscle ultrastructure and function include fiber atrophy, impaired
3
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contractile performance (30), myofibrillar remodeling (77), shifts in contractile
protein composition (76), alterations in metabolic capacity (11,36,48,65), and
impaired cellular Ca2 + regulation (18). Nonetheless, how these factors
interact to produce the clinical symptoms observed in CHF remains unclear.
The severity of the skeletal myopathy in CHF appears to be dictated,
in part, by the functional role of the individual muscle such that muscles
activated more frequently appear to undergo greater maladaptive changes.
Accordingly, studies have shown that CHF produces more profound
deleterious effects in the muscles of ventilation, the diaphragm in particular,
compared with limb skeletal muscles (28,30,82). Factors predisposing a
rhythmically contracting muscle to undergo structural deterioration similar to
atrophy are not clear, yet these findings form the foundation for the premise
that muscle wasting and disuse are uncoupled in CHF (70,78).
BACKGROUND
Skeletal Myopathy in CHF. Stable chronic heart failure is often
associated with the syndrome of cardiac cachexia characterized by a frail,
wasted physical appearance, marked weakness, poor endurance for activity,
and increased work of breathing (7,8,10,35,41,44,46,51). Ultrastructural
remodeling of limb skeletal muscle is now well appreciated in CHF. Most
authors agree that peripheral limb muscles from human patients and animal
models of CHF typically exhibit a decreased proportion of slow-twitch (Type
I) fibers, an increased proportion of fast-twitch (Type lla, lib, and ilx) fibers,
4
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and diminished cross-sectional area (CSA) of ail fiber types proportionate to
the severity of the disease (48,70,79). Studies show that reduced activity
does not play a major role in the loss of muscle mass in CHF (70,78).
In this regard, the contracting diaphragm responds to the environment
of CHF by exhibiting global fiber atrophy and expressing a higher proportion
of Type I fibers (30,70,71,77), while demonstrating a decline in sarcomere
cross-bridge number per mm3 and single cross-bridge force (38). Blood flow
distribution to the diaphragm may actually be enhanced during low cardiac
output states, supporting the view that diminished nutritive flow is not a
contributing mechanism to diaphragm dysfunction (82). As the primary
muscle of ventilation, reduced mass and impaired function of the diaphragm
can contribute significantly to the dyspnea and exercise intolerance seen in
patients with CHF. Moreover, it has been suggested that lung stiffness
associated with CHF-induced pulmonary edema overburdens an already
wasted diaphragm, further impairing function (41).
It has been speculated that there may be common processes in CHF
that affect the functional and structural properties of both cardiac and skeletal
muscles. Chronic heart failure has been linked with decreases in mRNA and
protein expression of sarcoplasmic and endoplasmic reticulum Ca2 + -ATPase
(SERCA) in both cardiac and skeletal myocytes, which alters free
intracellular Ca2 + concentration (40,58,69). Elevated cytosolic Ca2 + is known
to be an important signaling mediator for protein and lipid degradation by
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various Ca2 + -dependent proteases, lipases, and endonucleases in
mammalian muscles (16,22,33,57,61).
Neurohormonal stimulation in CHF leads to increased sympathetic
drive and hyperactivity of the renin-angiotensin and endothelin systems, all of
which are known to play a role in the deleterious process of ventricular
remodeling (62). The renin-angiotensin-aldosterone response has also been
linked to enhanced activation of immune function in CHF, resulting in
augmentation of circulating inflammatory factors (63,83). Specifically,
excessive concentration of tumor necrosis factor-a (TNF-a) and oxygen free
radicals/reactive oxygen species have been observed during the progression
of heart failure; factors implicated as inducers of programmed cell death
(PCD) or apoptosis (23,39,50). Apoptosis is well known to occur in the
myocardium in CHF and recent evidence suggests it occurs in skeletal
muscle as well (80,81).
Impaired Skeletal Muscle Ca2 * Homeostasis in CHF. It is believed
that a major component contributing to CHF-induced skeletal myopathy is
impaired cellular Ca2 * regulation leading to contractile dysfunction. This is
due in large part to alterations in SERCA content and activity, as this protein
is responsible for the translocation of cytosolic free Ca2 + into the
sarcoplasmic reticulum (SR) following contraction. Chronic CHF is
associated with reduced SERCA mRNA transcripts and its cognate protein in
rat soleus (69). A 38% decrease in diaphragm SERCA cycling activity was
6
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reported in rats with CHF, indicating a reduced ability to sequester cytosolic
Ca2 + (58). Moreover, studies utilizing a post-infarction rabbit model revealed
a clear relationship between diaphragm contractile dysfunction and impaired
Ca2 + uptake by the SR (40).
Impaired SR function leads to elevated levels of intracellular Ca2 *
which can activate calpains (non-selective Ca2 + -activated cysteine
proteases). Calpains play an important role in mediating skeletal muscle
protein degradation, especially in the disassembly of myofibrils during the
early stages of muscle turnover (31). Moreover, calpains can play an
essential role in programmed cell death by cleaving the proapoptotic protein
Bax in the mitochondria, which in turn mediates cytochrome c release with
subsequent initiation of the apoptotic cascade (24). This observation would
seem to imply that there may be some cross-talk and integration of various
proteolytic signals during the process of muscle degradation.
Apoptosis in CHF-lnduced Skeletal Myopathy. Skeletal muscle
progenitors (i.e. myoblasts and satellite cells) have been observed to
undergo apoptosis in various experimental conditions, including transgenic
mice expressing low levels of the retinoblastoma gene (66,87), cultured
satellite cells after serum withdrawal (5,47), and cultured myoblasts after
exposure to supraphysiologic doses of the steroid stanozolol (1,2). Cellular
protein content and ATP levels initially remain unaltered, distinguishing
apoptotic cell death from the energy-independent pathway of necrosis
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(29,37,68). Normal metabolic processes are well conserved in the early
stages of PCD and are essential for its progression.
The most frequently documented indicators of apoptosis include
TUNEL + (terminal deoxynucleotidyl transferase-mediated dUTP nick end-
labeling) nuclei, DNA laddering, and altered regulation of pro- and anti-
apoptotic proteins and transcription factors, such as stress-activated protein
kinase (SAPK), c-fos, c-myc, Bax, and Bcl-2 (21,27,32,53,85,86). Because
the TUNEL method labels DNA breaks with 3’-OH terminals (a characteristic
of both necrotic and apoptotic demise, as well as DNA repair), the application
of other assays to corroborate results obtained by TUNEL analysis has
recently been suggested (17,34). Caspase (cysteinyl-aspartate-specific
proteinase)-3 is an important participant in the apoptotic cascade and its
activity is known to be increased in cardiovascular diseases (45,55,60,85). A
mammalian homologue of ced-3 (the nematode C. elegans death gene),
caspase-3 mediates apoptosis by cleaving selected intracellular proteins, as
well as proteins of the nuclear lamina, cytoskeleton, and sarcoplasmic-
endoplasmic reticulum (52).
Mature skeletal myocytes are now known to exhibit signs of apoptotic
demise without a marked inflammatory response in numerous metabolic and
degenerative muscle disorders. Nuclear condensation and TUNEL+
chromatin cleavage, expression of pro- (Bax and ICE) and anti-apoptotic
factors (Bcl-2 and Bc/-xL), and activation of caspase-3 have all been
8
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identified in the pathology of the muscular dystrophies (12,54,72,75),
amyotrophic lateral sclerosis (73), and polymyositis (9) in both human and
animal models of disease. Additionally, muscle biopsies from patients with
Duchenne’s muscular dystrophy demonstrate a mosaic pattern of Bcl-2/Bax-
positive myofibers correlating with apoptotic myonuclei, suggesting that
apoptosis does not exclusively target a specific fiber type in this disorder
(64).
Shifted Bcl-2/Bcl-xL:Bax/Fas/ICE ratios, TUNEL+ myonuclei,
decreased myonuclear content, and atrophy without evidence of whole fiber
loss have also been noted in animal models of altered activation states,
including bouts of intense exercise (59), muscle denervation/reinnervation
(74), spinal isolation (4), and hindlimb unweighting (3). Because skeletal
muscle is multi-nucleated, loss of individual myonuclei may not lead
immediately to death of the entire myofiber. By reducing myonuclear
content, the availability of templates for transcription (i.e., DNA) can be
modulated and may result in diminished muscular protein content, ultimately
impacting fiber size.
Recent evidence indicates that cytokine activity is increased in the
environment created by chronic CHF (14). Specifically, an increased
circulating concentration of TNF-a has been noted in patients with advanced
CHF and is believed to be partly responsible for the wasting syndrome of
9
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cardiac cachexia and other catabolic states (7,39,50). TNF-a has also been
shown to cause impairment of diaphragm contractility in vitro (84).
Vescovo et al (80,81) have recently demonstrated increased plasma
levels of TNF-a in human and rat models of CHF. Their work also
documented muscle atrophy, higher protein levels of caspase-3, and a
greater number of TUNEL+ nuclei in peripheral limb muscles from CHF
subjects compared to control. The increased expression of various pro-
apoptotic mediators observed within the environment of CHF offers insight
into potential cellular processes that may be involved in the development of
CHF-induced skeletal myopathy. It remains to be seen if apoptosis plays a
role in diaphragm wasting associated with this debilitating syndrome.
10
)
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CHAPTER II
SUMMARY OF THE EXPERIMENTAL DESIGN
The goals of this project were to confirm the presence of diaphragm
muscle wasting and to investigate potential mechanisms contributing to
diaphragm myopathy in a chronic animal model of CHF. To accomplish
these goals, the following areas were explored: 1) Diaphragm single-fiber
cross-sectional area, 2) Diaphragm compartmental Ca2 + kinetics, 3) Ca2 + -
dependent catabolic protease activity, and 4) Cellular and molecular
mediators and products of apoptosis. All studies were conducted using
diaphragm muscle from male Wistar rats (250-300 g, 8-10 wk old) with
chronic experimental CHF (CHF, n=12) or sham operation (SHAM, n=10) of
10 weeks duration.
The condition of CHF in experimental animals was assessed by
measuring lung wet-to-dry weight and heart weight-to-body weight ratios.
Elevated ratios have been positively correlated with reduced left ventricular
ejection fraction and increased left ventricular end-diastolic pressure (5,6).
Serum TNF-a levels were also assayed. Significantly elevated serum TNF-a
levels are consistent with the human clinical picture of CHF (1-4).
The condition of diaphragm myopathy in experimental animals was
demonstrated by comparing mean single-fiber cross-sectional area (CSA)
and mean myonuclear domain size. Reduced CSA and myonuclear domain
size in CHF animals vs. SHAM reflects an effect of CHF. Diaphragm Ca2 +
11
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Development of the Chronic Animal Model
and Summary of the Experimental Design
Week 1 ' " Week 2-9 Week 10 Week 11-*
• Pre-operative • Weekly • Sacrifice • Process
assessment physical • Harvest heart, tissue and
• Surgery assessment lungs, blood, blood:
performed: diaphragm
• Morphometric
- TNF-a
- left coronary analysis of - Calpain
artery ligation heart and - CSA
(CHF, n=12) lungs
• Fix and
- Myonuclear
domain
or
paraffin-embed
portion of
- TUNEL
- DNA
- sham operation
diaphragm
laddering
(SHAM, n=10)
• Freeze portion
of diaphragm
• Utilize portion
of diaphragm
for analysis of
Ca2 + kinetics
- Caspase-3
Table 2.1. Development of the chronic animals and summary of the experimental design.
homeostasis was evaluated by kinetic analysis of 4 5 Ca2 + tracer data collected
from a long-term 4 5 Ca2 + efflux experiment. Qualitative assessment of calpain
activity (a proteolytic enzyme activated by elevated intracellular Ca2 ") was
carried out utilizing a calpain-specific protein substrate conjugated with a
color reporter molecule. Apoptosis in diaphragm was assessed utilizing
three tools, including: a) the TUNEL method to detect apoptotic-like DNA
fragmentation via labeling of free 3’-OH ends, b) electrophoresis of isolated
DNA samples from CHF diaphragm to assess for the presence of a laddering
pattern consistent with intemucleosomal DNA fragmentation (a hallmark of
apoptosis), and c) qualitative assessment of caspase-3 activity utilizing a
12
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caspase-specific peptide substrate conjugated with a color reporter molecule.
Table 2.1 describes the development of the chronic animal model and
provides a summary of the experimental design.
1 3
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CHAPTER III
COMPARTMENTAL ANALYSIS OF STEADY-STATE
DIAPHRAGM Ca2 + KINETICS IN CHRONIC CHF
INTRODUCTION
It is well known that congestive heart failure (CHF) induces a
pathophysiologic response in skeletal muscle generally characterized by loss
of cross-sectional area (CSA) and functional impairment (33,37,38,43).
Interestingly, the degree of skeletal myopathy is different among muscles
with different functional roles. Although the reasons for this are unclear, the
literature suggests that more intense activation paradigms may predispose
muscles to greater maladaptive change (9,25,33). The most persuasive
example of this is the observation that inspiratory muscles, the diaphragm in
particular, are reportedly more severely affected by CHF than other skeletal
muscles (25,28,55). Factors predisposing a rhythmically-contracting muscle
to lose mass and strength in CHF are uncertain. In light of the data
suggesting a correlation between respiratory muscle dysfunction and
dyspnea in patients with CHF, diaphragm wasting and weakness have
important clinical implications (13,36,40).
Few investigators have examined cellular and molecular mechanisms
underlying the maladaptive response of the diaphragm to CHF. It is known
that relative shifts in fiber composition seem to occur in opposite directions in
peripheral and respiratory muscles. In heart failure, the human diaphragm
14
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exhibits a shift from fast to slow myosin heavy chain isoforms with an
increase in oxidative capacity and a decrease in glycolytic capacity (52). In a
porcine model of CHF, Howell et al (28) showed that CSA was reduced in all
fiber types, with an increased proportion of type I fibers compared with
control animals. One study utilizing a post-infarction rabbit model of CHF
revealed a relationship between diaphragm contractile dysfunction and
prolonged intracellular Ca2 + transients (35). The authors attributed the
observed Ca2+dysregulation to impaired sarcoplasmic reticulum (SR)
function. Investigations of limb skeletal muscle from chronic CHF animals
have reported altered SR Ca2 + -ATPase (SERCA) mRNA and protein
expression which affects cellular Ca2 + regulation (44,48).
Because of the ubiquitous nature of Ca2 + and its prominence as a
signaling molecule, changes in cytoplasmic Ca2 + concentration can alter or
initiate many cellular processes. Increased cytosolic Ca2 + is a well-known
trigger for protein degradation and cell death. In particular, calpain (non-
selective Ca2 + -activated cysteine protease) is known to play a significant role
in proteolytic degradation of skeletal myofibrils during normal muscle
turnover (29). However, the status of calpain in CHF-induced muscle
wasting has not been explored. Using experimental methods based on
simulation and modeling of long-term 4 5 Ca2 + efflux data, we tested the
hypothesis that discrete stores and exchange fluxes of Ca2 + in the diaphragm
are altered in response to CHF. Additionally, an assay utilizing a calpain-
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specific substrate was employed to explore the activation status of this Ca2 + -
dependent protease in CHF diaphragm.
METHODS
Development of CHF Animal Model
Left coronary artery ligation was employed to induce myocardial
infarction and CHF in experimental animals. Male Wistar rats (250-300 g, 8-
10 wk old) were anesthetized with ketamine (60 mg/kg, ip) and xylazine (9
mg/kg, ip). Animals undergoing left coronary artery ligation (CHF, n=12)
were intubated with a 20-gauge intravenous catheter utilizing
transillumination of the thorax to visualize the tracheal opening. Mechanical
ventilation (Harvard Apparatus, Holliston, MA) was initiated (3.5 cc tidal
volume, 60 Hz). A left thoracotomy was performed at the level of the 5th
intercostal space and the pericardial sac was opened. The left coronary
artery was ligated approximately 2-3 mm from its origin utilizing a 6-0 Prolene
suture bolstered by a small cardiovascular pledget to compress the vessel.
Blanching of the anterior and lateral left ventricular wall confirmed successful
ligation of the left coronary artery in all experimental animals. The incision
was closed in three layers using a 4-0 Dexon absorbable suture. Sham-
operated animals (SHAM, n=10), who served as controls, underwent a
similar procedure without ligation of the left coronary artery. The mortality
rates for the CHF and SHAM animals within the first 72 h post-surgery were
approximately 75% and 10%, respectively.
16
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After surgery, the thoracic incision was treated topically with 0.2%
nitrofurazone solution and animals received an injection of ampicillin (0.1
g/kg body wgt, sc). Acetaminophen was placed in the drinking water (67
mg/L) for the first 48 h post surgery. The animals were housed individually
and allowed free access to water and standard rat chow ad libitum. CHF
and SHAM animals were monitored weekly for body weight fluctuation, signs
of pulmonary edema, ascites, anorexia, and lethargy. The investigation
conformed to the Guide for the Care and Use of Laboratory Animals
published by the US National Institutes of Health (NIH Publication No. 85-23,
revised 1996).
At the time of sacrifice (10 wks post-surgery) total body weight was
measured and the diaphragm, heart, and lungs were harvested. The large
airways were removed from the lungs before weighing. Heart weight-to-body
weight and lung weight-to-body weight ratios were then calculated and
multiplied by a factor of 103. In order to determine wet-to-dry ratios, the
lungs were weighed before and after drying in an oven held at 60° C for 72 h.
Consistent weights at 48 and 72 h indicated true dry weight of the lungs. The
criteria utilized for determining development of CHF in experimental animals
included significantly increased heart weight, heart weight-to-body weight
ratio, and lung wet-to-dry weight ratio which have been correlated with direct
measurements of heart failure, including depressed left ventricular systolic
pressure and elevated left ventricular end diastolic pressure (45,49).
17
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Diaphragm tissue from experimental CHF animals not meeting these criteria
was excluded from the study.
Single-Fiber Cross-Sectional Area
At the end of the 10-wk conditioning period, each rat was given a
lethal dose of Euthasol (39 mg pentobarbital sodium/5 mg phenytoin sodium)
via intraperitoneal injection. A portion of the left costal hemidiaphragm was
removed and placed in a dissecting dish containing bicarbonate-buffered
Ringer’s solution maintained at 27°C and oxygenated with a mixture of 95%
02/5% CO2. The Ringer’s solution contained 117 mM NaCI, 3.5 mM KCI, 1.2
mM KH2P04i 1.2 mM MgS04, 24 mM NaHC03 ) 1.2 mM CaCI2, and 10 mM
glucose. After trimming away the crural diaphragm and excess connective
tissue, the Ringer’s solution was replaced by 10% neutral buffered formalin.
The preparation was then allowed to fix overnight at 4°C. After the fixation
period, the tissue sample was washed in 2-3 changes (30 min/change) of
dH20 and a standard dehydration protocol was employed consisting of 2-3
changes (30 min/change) in each of the following concentrations of ethanol;
25%, 50%, 70%, 95%, and 100%. The tissue samples were then cleared in
2-3 changes (5 min/change) of xylene and then embedded in paraffin (3-4
changes, 30 min/change) in a vacuum oven set at 60°C. After cooling, the
blocks were sectioned at a thickness of 5 urn. Sampling was randomized by
taking every 10th section (representing a distance of 50 |im along the length
of the tissue block) and mounting it on a glass slide. The sections were then
18
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allowed to dry overnight on a slide warmer. Eight sections per subject were
prepared in this fashion.
The mounted specimens were deparaffinized and rehydrated by
immersing them in xylene and a series of decreasing concentrations of
ETOH. The specimens were first permeabilized by incubating them in a
solution containing 20 |.ig/ml proteinase K for 10 min at room temperature,
followed by a rinse in TBS (20 mM Tris, pH 7.6, 140 mM NaCI). Next, the
slides were covered with a blocking solution designed to quench
autofluorescence in formalin-fixed tissue (1% glycine, 2% bovine serum
albumin, 10% normal goat serum, 0.1 M PBS, 1 mM MgCh, and 0.1 mM
CaCh) for 20 min at room temperature. The slides were then incubated with
rabbit anti-laminin antibody (Sigma, St. Louis, MO) diluted 1:25 in TBS and
placed in a humidified chamber at 37°C for 1 h. Slides were rinsed in 3
changes of TBS for 5 min each and then incubated in a secondary goat-anti-
rabbit IgG conjugated with rhodamine (Sigma, St. Louis, MO) diluted 1:100 in
TBS and placed in a humidified chamber at 37°C for 1 h. The reaction was
then terminated by three, 5-min rinses in TBS at room temperature in the
dark and the slides were mounted with a coverslip using Permount.
In order to evaluate diaphragm wasting in the animal model of CHF,
single-fiber CSA was measured as an index of myofiber atrophy. Formalin
fixation and ethanol dehydration during the processing of diaphragm samples
may result in the loss of up to 30% of tissue volume (4). However, all
19
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muscles were fixed in an identical manner thus a comparison between CHF
and SHAM myofibers is nonetheless useful to assess relative differences in
single-fiber CSA (53,54). Slides were visualized by means of microscopy
fitted with a rhodamine filter and analyzed using a semi-automated,
computerized method (Bioquant Classic 98, R&M Biometrics, Nashville, TN)
to calculate single-fiber CSA (expressed as |im2). Only the area clearly
outlined by the basal lamina was identified by threshold tracing and
measured as the fiber’s CSA. Ten random magnification fields (20X) were
examined (representing approximately 300 myofibers) from each of the 8
slides prepared per subject and then averaged to yield mean single-fiber
CSA. In this way, approximately 2,400 single fibers were analyzed for each
subject in the study.
Analysis of Diaphragm Ca2 * Homeostasis Utilizing 4 5Ca2 * Kinetics
Technology
A portion of the right costal hemidiaphragm was removed and placed
in a dissecting dish containing bicarbonate-buffered Ringer’s solution
maintained at 27°C and oxygenated with a mixture of 95% 02/5% CO2. The
muscle was debrided of connective tissue and equilibrated for 1 h in an
Erlenmeyer flask containing 100 ml Ringer’s solution maintained at 27°C and
oxygenated with a mixture of 95% 02/5% CO2. The experiment was initiated
at this point by transferring the muscle to a 7-ml plastic vial containing 4 ml of
oxygenated Ringer’s solution and -5x1 O'7 disintegrations per minute
20
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(dpm)/ml 4 5 Ca2 + (half-life=164 days). 4 5 Ca2 + was loaded into the muscle for 1
h at 27°C. The exact loading solution activity was later calculated for each
experiment. Following the load, the muscle was rinsed briefly in 4 5 Ca2 + -free
Ringer’s solution and mounted at resting length in an efflux chamber
(volume=7 ml) that was perfused with a 4 5 Ca2 + -free Ringer’s solution at a
constant flow rate of 7 ml/min. The perfusate was delivered at 27°C and
continuously mixed with a fine oxygen bubble as it washed around the
muscle (Fig. 3.1).
Figure 3.1. Schematic representation of 4 S Ca2 ^ efflux experiment apparatus.
An 8-h efflux protocol was used to obtain tracer data from resting
muscles. Long-term experiments were necessary to resolve slowly
exchanging intracellular Ca2 * compartments as well as extracellular
compartments with rapid turnover kinetics (26). One ml/min of the effluent
21
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flowing from the efflux chamber was diverted to a fraction collector and 6
ml/min of the effluent was diverted to a radioactive waste receptacle. The
effluent diverted to the fraction collector was collected in 7-ml polyethylene
minivials at 2-min intervals. The exact diversion of flow and total perfusate
flow was recorded for each experiment. All effluent fractions collected were
prepared with 3 ml of liquid scintillation cocktail and analyzed in a scintillation
counter (Beckman LS 1801, Beckman Instruments, Fullerton, CA).
Ca5
(cytosol) "
Ca4 Ca7
INTRACELLULAR MEM EXTRACELLULAR EFFLUX CHAMBER
C r a t 1
Figure 3.2. Schematic drawing depicting the kinetic model used to simultaneously fit < 5 Ca2 *
efflux data, where Ca1 = efflux chamber, Ca2 = extracellular fluid (ECF), Ca3 = peripheral
sarcolemmal Ca2 * binding site, Ca4 = t-tubular membrane Ca2 * binding site, Ca5 = cytosolic
free and rapidly exchanging Ca2 *, Ca6 = terminal cistemae Ca2 * compartment, and Ca7 =
longitudinal reticulum Ca2 * compartment. Arrows represent rate constants and are
designated K{i,j) or K(into, from), indicating the fraction of Ca2 * in compartment/ that moves
into compartment / per unit of time during steady-state conditions.
22
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At the conclusion of the experiment, the tissue was removed from the
efflux chamber and weighed. The efflux data, expressed in dpm-min'1mg'1 ,
were subsequently normalized for activity of the 4 5 Ca2 + loading solution,
fraction collector flow rate, and wet weight of the muscle strip.
Efflux data were analyzed on the basis of a seven-compartment model
previously developed in conjunction with work in rodent diaphragm (26). This
model represents the minimal hypothesis consistent with the efflux data for
goodness of fit, with the least degrees of freedom, and the smallest
associated sum of squares (Fig. 3.2). To begin setting up the kinetic model,
exponential equations were fit to tracer data from single experiments to
characterize the complexity of the curve (11). A minimum of four
exponentials was necessary to achieve an adequate fit of the data,
suggesting a model of at least four compartments with independent rates of
turnover (6). Nonetheless, a four-compartment model was insufficient to
account for both the kinetic data and the in vitro physiologic system.
Therefore, the fundamental model was systematically expanded. The
stipulation of seven compartments was based on the number of distinct
exponentials in the kinetic data (Ca3, Ca4, Ca6, and Ca7), on recognized
physiology (Ca2 and Ca5), and on the methods of the experiment (Ca1).
Previous studies have shown that extracellular Ca2 * compartments
turnover quickly, whereas intracellular Ca2 * compartments exchange more
slowly (7). More recently, Howell (26) described the relative influence of
23
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each of the four exponentials representing compartments on the 8-h
4 5 Ca2 + efflux curve (Fig. 3.3). The areas of influence or sensitivity for
adjustable parameters were estimated from partial derivatives of simulated
rate constant values. Because Ca3 and Ca4 exert authority early over the
efflux curve, they were assumed to be extracellular. Accordingly, Ca6 and
Ca7 dominate the curve later and were assumed to be intracellular.
Previously, experiments using isoproterenol and caffeine have been
employed to further clarify the identities of the kinetically distinct
compartments. Both of these drugs are believed to enhance the release of
Ca2 + and/or inhibit its uptake by the SR (20,56). Addition of caffeine or
1000
Ca3
O )
100
c
E
Ca6
Ca4
Ca7
2
a.
a
600 300 400 100 200
(A
m
z
C O
<
I
tt*
3-
3
•2
MINUTES
Figure 3.3. Sensitivity of data for adjustable parameters calculated from partial derivatives of
simulated values. Sensitivity curves are superimposed on mean 4 5 Ca2 * efflux data (Ipf rom
control experiments. Influence of each kinetically distinct Ca2 + compartment on every data
point is demonstrated.
24
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isoproterenol to long-term 4 5 Ca2 * efflux experiments on rat diaphragm caused
a failure of the SR to load 4 5 Ca2 + , resulting in a dramatic reduction in the
magnitude of the latter part of the efflux curve (27). Since the latter part of
the curve is sensitive to efflux from Ca6 and Ca7, these compartments must
be SR. Moreover, Meissner (41) has resolved the SR into a heavy fraction
(representing the terminal cisternae which exhibits rapid Ca2 *-ATPase pump
activity with significant Ca2 * leak) and a light fraction (representing the
longitudinal reticulum which sequesters the maximum amount of Ca2* with
minimal Ca2 * leak). Fitting of the data in this and a previous (26) study has
indicated that Ca6 is associated with faster rate constants, while Ca7 exhibits
slower kinetic properties. It is thus likely that Ca6 represents the terminal
cisternae and Ca7 represents the longitudinal reticulum.
Caffeine has also been shown to increase the magnitude of the early
portion of the 4 5 Ca2 * efflux curve in rat diaphragm (27) and frog sartorious
(7), indicating a redistribution of Ca2* from intracellular stores to extracellular
compartments. Studies have also shown that Ca2 * is translocated from the
terminal cisternae to the t-tubule system during fatiguing stimuli (21). Ca2 *
not only resides in the t-tubule system, but is also associated with various
sarcolemmal Ca2 * binding proteins, which suggest identities for Ca3 and
Ca4. Because the t-tubule system is remote and tortuous, it would
demonstrate slower kinetic exchange of Ca2 * compared to sarcolemmal
membrane Ca2 *-binding proteins. It has been speculated that Ca3
25
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represents the sarcolemmal-bound Ca2 + since it dominates the efflux curve
early, and Ca4 would then reflect the more slowly exchanging t-tubule Ca2 *
compartment.
Data fitting was carried out using the Simulation, Analysis, and
Modeling (SAAM II) program (8). Differential equations representing models
to be tested were solved for each datum using an exponential method.
Solutions were assessed periodically for congruity with classical Rosenbrock
numerical integrators. Initial estimates of the rate constants were adjusted
by an iterative process until a least-squares fit of the data was obtained (5).
Data weighting was used in order to provide a measurement of the
confidence in a particular datum. The fractional standard deviation (FSD)
value was set at 0.10 and provided an estimate of the 95% confidence
interval for the rate constants. Once a successful hypothesis was
established, final estimates of the rate constants and their coefficients of
variation were calculated. This constituted the information extracted from the
experimental data. Statistical uncertainties associated with the final
parameter estimates were calculated by multiplying the inverse normal
equation matrix by the best estimate of the variance of the data.
All steady-state solutions for Ca2 * contents and exchange fluxes were
calculated using the matrix equation: U + RM = 0. The equation represents
unlabeled 4 0 Ca2 *. U is the vector of compartmental input rates of tracee.
There are seven Ca2 * compartments incorporated into the model so that the
26
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equation must account for seven distinct input rates. However, since Ca2 +
enters the system only from the perfusate (Ca1), six of the seven elements
are equal to 0 and the value of U is dictated solely by Ca1. R is the n x n
matrix of intercompartmental rate constants with diagonal elements, R(i,i),
defined as the negative sum of all the rate constants leaving the P
compartment. M is the vector of compartmental concentrations of tracee.
Zero (0) refers to the null matrix. The equation assumes only steady-state
and conservation of mass and enables interpretation of tracer kinetic data
[R{i,j)] into steady-state Ca2 + concentrations and fluxes (5).
Extraction and Assay for Calpain Activity
At the time of sacrifice, a portion of the left costal hemidiaphragm was
immediately placed in a dissecting dish containing bicarbonate-buffered
Ringer’s solution maintained at 27°C and oxygenated with a mixture of 95%
02/5% CO2. After removing the central tendon, the tissue was placed in a
dissecting dish containing cooled isopentane ('65-'70°C) and quick-frozen in
order to prepare it for analysis.
Enzyme extraction was achieved by utilizing a spectrophotometric
assay described by Moss et al (42). Approximately 100 mg wet wgt of the
frozen diaphragm tissue was finely minced with a clean razor blade and
placed in a mortar on dry ice. The tissue was ground into a powder and
placed in a 1.5-ml microcentrifuge tube on ice containing 0.5 ml
homogenizing buffer (20 mM Tris HCI, 1 mM EDTA, 5 mM 2-
27
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were removed from the water bath and placed on ice. 0.05 ml of 15% TCA
was added to each tube and the tubes were placed in a freezer f20°C) for 5
min and then moved to a refrigerator (4°C) for 15 min to facilitate TCA
precipitation. Afterwards, the samples were cold-centrifuged (4°C) for 10 min
at 5,000 x g. The supernatant was transferred to a new microcentrifuge tube
and 0.45 ml NaOH was added to maximize the absorbance of the azocasein
chromophore. 200 (a l from each sample was transferred to a 96-well flat
bottom microplate. The resulting reactions were read at 440 nm with a
correction wavelength of 540 nm and the optical densities (O.D) were
recorded. All reactions were performed in triplicate.
The results were expressed as activity of calpain in CHF diaphragm
relative to SHAM and reported in arbitrary units (AU). Additionally, if the
background control (i.e. no CaCl2 during incubation) exhibited a substantial
reading, it was subtracted from the experimental results prior to reporting
activity values.
Statistical Analysis
SHAM data were compared with data from CHF muscles utilizing a
Student’s t-test for independent samples in order to compare mean values of
organ weights, organ-to-body weight ratios, single-fiber CSA, 4 5 Ca2 + efflux
data, compartmental Ca2 + contents, Ca2 + exchange fluxes, and relative
calpain proteolytic activity. The accepted level of significance was set at
p<0.05 and all results were expressed as mean+SE.
29
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RESULTS
Characteristics of the Chronic CHF Animal Model
The initial body weight of CHF vs. SHAM animals was not different
(302.9±5.9 g vs. 296.3+4.1 g, p>0.05). Although body weight increased
significantly in both groups, CHF animals gained less weight over the 10-wk
conditioning period compared to sham-operated animals (118.1±5.9 g vs.
131.5±5.0 g, p<0.05).
Characteristics of the
Chronic Animal Model
Parameter SHAM CHF
Heart Wt (g) 1.57±0.08 2.73±0.15*
Heart Wt/Body Wt (x103) 3.63±0.17 6.63±0.42*
Lung Wet Wt/Dry Wt 4.48±0.02 5.04+0.08*
Lung Wt/Body Wt (x103) 4.18±0.10 13.20±0.97*
Table 3.1. Mean heart and lung weights and ratios in CHF and SHAM animals.
•Significantly different from SHAM (p<0.05).
As shown in Table 3.1, hearts harvested from CHF animals weighed
significantly more than hearts from sham-operated animals (2.73±0.15 g vs.
1.57+0.08 g, p<0.05) resulting in an 83% increase in heart weight/body
weight ratio (6.63+0.42 vs. 3.63±0.17, p<0.05). A large area of fibrotic scar
30
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on the anterior and lateral aspects of the left ventricle was noted in all CHF
animals, indicating myocardial infarction. Lung wet weight/dry weight ratio
was increased in CHF animals compared to SHAM (5.04+0.08 vs. 4.48+0.02,
p<0.05), and lung weight/body weight ratio was elevated more than three
fold in CHF (13.20+0.97 vs. 4.18±0.10, p<0.05). Increases in all of these
ratios have been shown to be highly correlated with elevated left ventricular
end-diastolic pressures and impaired left ventricular contractility in CHF
(45,49).
1200
1000
~ 800
E
f 600
c o
o 400
200
0
Figure 3.4. Mean diaphragm single-fiber CSA of SHAM and CHF animals. CHF was
associated with a reduction in single-fiber CSA. Error bar corresponds to 1 SE.
’ Significantly different from SHAM (p<0.05).
3 1
n SHAM
b CHF
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Analysis of Single-Fiber Cross-Sectional Area
Analysis of diaphragm myofibers from each animal in the study
revealed that mean diaphragm single-fiber CSA from rats with CHF was 12%
less than that of sham-operated rats (844+40 pm2 vs. 962±27 pm2, p<0.05,
Fig. 3.4).
□ CHF
o Sham
10'
1 0'
200 300 400 100 500 0
Time (minutes)
Figure 3.5. Effects of CHF on diaphragm 4 5 Caz + efflux data. Mean curve from CHF
diaphragm is compared with mean curve from SHAM diaphragm. Error bars are not shown
because individual efflux curves were essentially superimposable in each experimental
condition. CHF 4 5 Ca2 * efflux data (dpm min‘1g ) were significantly different from SHAM at
every time point during the 8-h collection period (p<0.05).
Effects of CHF on Diaphragm 4 5 Ca2 + Efflux Data
Analysis of the 4 5 Ca2 + efflux data revealed a significant difference in
the mean radioactivity values (dpm-min'1g"1 ) in CHF compared with SHAM at
every time point during the 8-h collection period (Fig. 3.5). During the first
32
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100 min of the efflux period, mean CHF dpmmin'1g'1 values increased to
200% of SHAM values and from 100 to 480 min CHF values were increased
by 160% (p<0.05). These data reveal that CHF was associated with a
substantial increase in diaphragm Ca2 *.
10000
>8000
o
f-6000
c
| 4000
o
% 2000
0
Ca3 Ca4 Ca6 Ca7
Diaphragm Ca2 * Compartments
Figure 3.6. Effects of CHF on diaphragm compartmental Ca2 * contents. Mean data from
CHF diaphragms are compared to SHAM. Error bars correspond to 1 SE. 'Significantly
different from SHAM (p<0.05).
Estimated Diaphragm Compartmental Ca2 * Contents and Exchange
Fluxes
The CHF kinetic data could be fit by adjusting five rate constants
compared with SHAM. K(3,2) and /c(4,2), the rate constants representing
influx to Ca3 and Ca4 respectively, were increased. This resulted in
increased Ca2 * contents (Fig. 3.6) and exchange fluxes (Table 3.2) of both
33
*
M
n SHAM
■ CHF
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extracellular compartments, accounting for elevation of the early, fast
component of the efflux curve. K(5,2) was also increased, resulting in a
Diaphragm Intercompartmental Ca2 *
Exchange Fluxes (nmol«mirr1 *g 1 )
Parameter SHAM CHF
R(3,2) 3,065±237 13,278+655*
R(4,2) 19±4 52±4*
R(5,2)
31t 61±0.1*
R(6,5) 224+4 460+6*
R(7,5) 27±1 58±1*
R(6,7) 4±0.2 7+1*
Table 3.2. Values are mean+SE. R(i,j), steady-state flux of Ca2 * into the t ih compartment
from the f h compartment. Because there is a steady-state, R(i,j) = R(j,i). Steady-state fluxes
are defined as: R(3,2), flux of Ca2 * between peripheral sarcolemma and ECF; R(4,2), flux of
Ca2 * between t-tubules and ECF; R(5,2), transmembrane flux of Ca2 * between cytosol and
ECF; R(6,5), flux of Ca2 * between terminal cisternae and cytosol; R(7,5), flux of Ca2 *
between longitudinal reticulum and cytosol; and R(6,7), flux of Ca2 * between terminal
cisternae and longitudinal reticulum. 'Significantly different from SHAM (p<0.05). rNo SE is
shown for R(5,2) because a ratio of rate constants for these two compartments was
estimated and constrained based on available information and previous studies.
Intercompartmental rate constants, K(ij), extracted from the data are not shown but may be
estimated from Ca2 * contents shown in Figure 5 and fluxes delineated in the table, using the
equation K(i,j) = R(ij)/Q(j).
predicted two-fold rise in Ca2 * flux across the sarcolemma. Indeed, if Ac(5,2)
was experimentally constrained to the value set for SHAM efflux analysis, the
CHF data curve could not be fit. Because of the increased Ca2* flux across
the plasma membrane, there was a concomitant increase in the Ca2 *
34
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contents of Ca6 and Ca7 (Fig. 3.6), as well as the exchange fluxes of both
compartments (Table 3.2). This accounted for the elevation of the late, slow
component of the efflux curve. Although the model predicted global Ca2 +
overload in CHF diaphragm, it is interesting to note that the model also
predicted an increased contribution of extracellular Ca2 + to total tissue Ca2 +
(Table 3.3).
Diaphragm Compartmental Ca2 * Distribution
(nmol*g*1 )
Parameter SHAM CHF
Total Diaphragm Ca2 * 7,785±243 20,384±1,832*
Intracellular Ca2 * 5,905±236 13,848+1,846*
Extracellular Ca2 * 1,880±107 6,535+271*
% Intracellular Ca2 * 76±1 66+3*
% Extracellular Ca2 * 24±1 34±3*
Table 3.3. Values are mean±SE. Intracellular Ca2 * Includes free Ca2 * in the cytosol and in
the components of the SR. Extracellular Ca2 * includes free Ca2 * in the ECF and bound Ca2 *
in 2 sarcolemma-associated compartments. % intracellular and % extracellular Ca2 * are
expressed as a fraction of total diaphragm Ca2 *. ’Significantly different from SHAM
(p<0.05).
Relative Activity Level of Calpain within the Environment of CHF
Qualitative analysis of calpain activity revealed a statistically
significant difference in mean O.D. readings between CHF and SHAM
diaphragm sample lysates (Fig. 3.7). This accounted for a relative calpain
35
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SHAM
Figure 3.7. Effects of CHF on diaphragm calpain proteolytic activity. Mean data from CHF
diaphragms are compared to SHAM. Error bar corresponds to 1 SE. 'Significantly different
from SHAM (p<0.05).
activity index in CHF diaphragm that was 215% of that observed in sham-
operated animals (p<0.05).
DISCUSSION
This study investigated compartmental Ca2 * kinetics in CHF-induced
diaphragm myopathy utilizing a chronic animal model. The kinetic data are
quantitatively consistent with the hypotheses that CHF leads to: 1) increased
Ca2 * contents of extracellular and intracellular diaphragm compartments; 2)
elevated total diaphragm Ca2*, 3) increased Ca2 * flux across the plasma
membrane as well as augmented Ca2 * exchange fluxes for intracellular and
extracellular diaphragm compartments. In addition, the calpain-specific
36
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assay utilized in this study revealed an increase in the proteolytic activity of
calpain in CHF diaphragm relative to SHAM.
Critique of Experimental Procedures
The rat diaphragm is appropriate for long-term in vitro studies because
it is uniformly thin and oxygen can easily diffuse to the tissue core with the
superfusion technique. Equilibration of the diaphragm in Ringer’s solution for
1 h prior to the 4 5 Ca2 + load allowed the muscle to attain a steady state.
Intuitively, a thin resting muscle in a steady state should remain viable for
many hours. Indeed, previous experiments documenting diaphragm twitch
and tetanic contractions before and after an 8-h efflux period have confirmed
that force production is essentially preserved (26). Because the diaphragm
undergoes loss of CSA in CHF (28,33), diffusion distance for nutritive factors
may actually decrease. Thus, it is unlikely that the CHF diaphragm became
hypoxic during the efflux experiments.
Kinetic Analysis and Fitting of Efflux Data
To begin analysis of the efflux data, initial estimates of all rate
constants were established. The sensitivity of parameter estimates was
optimized by limiting the amount of information to be extracted from the data.
Therefore in order to analyze the SHAM efflux data, the equations
representing the model in Figure 2 were constrained by measured,
published, or calculated values, including: 1) [Ca2 + ] in the perfusate = 1.2
mM; 2) turnover rate of the efflux chamber = 7 ml/min; 3) ECF Ca2 + content
37
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calculated as the product of the equilibrated [Ca2 + ] (1.2 mM) and the volume
estimated from preliminary sucrose washout experiments (0.32 ml/g, Howell,
unpublished observations); 4) cytosolic [Ca2 + ] = 100 nM (23); 5) cytosolic
volume = 0.26 ml/g (19); and 6) rate constants that maintained fixed
extracellular and intracellular volumes and [Ca2 + ] of the efflux chamber, ECF,
and cytosol. The rate constant representing the exchange of Ca2 + from the
ECF to the efflux chamber was necessarily set at a value that would be rapid
enough to ensure that the more remote intracellular compartments would not
appear to be mixed and unresolvable from the more rapidly exchanging
extracellular compartments. Additionally, the rate constants representing the
sequestration of Ca2 + by the terminal cisternae and longitudinal reticulum
were set at previously determined values (26,27), and their converse rate
constants (representing leak of Ca2 + back into the cytosol) were allowed to
adjust in order to fit the efflux data. The CHF efflux data was analyzed
utilizing the same stipulations, with the exception that cytosolic [Ca2 + ] was
not constrained.
Assumptions
The nature of the physiologic system under investigation in kinetic
experiments is not completely understood. Thus, it is necessary to identify
assumptions so that the effects of their violation can be assessed in terms of
interfering with the objective of the study (8). Since the tracer kinetic data
can only reflect the behavior of the tracer, certain parameters must be
38
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stipulated in order to extrapolate these findings to the behavior of the tracee.
Accordingly, the behavior of the physiologic processes under study can only
be characterized to the extent that the assumptions are validated. The
following assumptions were specified: 1) All Ca2 + enters or leaves the system
through Ca2. Not only is this physiologically appropriate and consistent with
kinetic analysis, but when other models with a second route for Ca2 +
entrance or exit were tested, the efflux data could not be fit (Howell,
unpublished observations). 2) The interaction between Ca2 + and its cytosolic
binding proteins in the resting state is negligible and does not significantly
affect long-term 4 5 Ca2 + efflux data. During steady-state resting conditions,
[Ca2 + ]j is low. In this instance, only Mg2 + competes with K+ for occupation of
bindings sites on calmodulin (the predominant intracellular binding protein for
free Ca2 '" with four metal binding sites). Since Ca2 + -bound calmodulin
comprises <1.5% of the total calmodulin species under steady-state
conditions (16), it is unlikely that free exchange of Ca2 + between
compartments would be significantly impeded. 3) The amount of the tracer is
quantitatively small and does not alter the discrete behavior of the tracee. It
has been established that the tracer exhibits kinetics that are directly
dependent on the properties of the tracee without affecting the latter’s
physiologic processes (46). 4) The system cannot distinguish between the
tracer and the tracee, but the distinct behavior of each can be resolved
experimentally. This is true given the observation that both tracer and tracee
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exhibit similar physiologic properties and behavior, while the scintillation
counter yields information only for the radioisotope (tracer) leaving the
nonradioactive material (tracee) unrecognized. 5) The properties of the
tracer (e.g. half-life = 164 days) do not impede discreet identification of
biologic processes within the system. The relatively long half-life of the
tracer ensures that the biologic processes it is intended to measure during
the 8-h efflux experiment are not artificially obscured by its rapid extinction.
Altered Diaphragm Ca2 * Homeostasis in CHF
Analysis of the kinetic data predicted that flux of Ca2 * across the
diaphragm plasma membrane was increased in CHF leading to a profound
rise in intracellular Ca2 *. The mechanism causing enhanced Ca2* flux across
the plasma membrane is unclear. Alternatively, down-regulation of SERCA
mRNA and reduced protein expression have been observed in hindlimb
muscles from rats with post-infarction CHF (48) and in the diaphragm of
spontaneously hypertensive heart failure rats (44). Depressed diaphragm
SERCA function would impair Ca2 * uptake into the SR. It is possible that
failure of the SR to resequester Ca2 * following excitation caused an initial rise
in cytosolic free Ca2 * in CHF.
It is well known that the accrual of intracellular free Ca2 * can lead to
damage of the sarcolemma. The binding of Ca2 * to calmodulin initiates a
signaling event that can subsequently activate phospholipase-A, which in
turn, cleaves phospholipids in the sarcolemma and increases membrane
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permeability (30). This may favor a rapid gradient-dependent influx of
extracellular Ca2 + into the cytosol, perpetuating an intracellular Ca2 +
overload. Thus, we speculate that diaphragm intracellular Ca2 + overload in
CHF may be the result of impaired SR Ca2 + uptake and enhanced Ca2 + influx
across a damaged sarcolemma.
Despite the probable depression of SR Ca2 + uptake in CHF, the
kinetic model actually estimated that Ca2 + exchange fluxes between the
cytosol and SR compartments (Ca6 and Ca7) were increased, leading to
elevated Ca2 + content in those compartments. The increased Ca2 + contents
of compartments Ca6 and Ca7 are not likely the result of active uptake, but
represent the corollary of Ca2 + translocation by simple mass action. It is
important to note that the Ca2 + tracer method is not sensitive enough to
resolve the relatively small amount of cytosolic Ca2+ . Intuitively then, the
predicted increases in compartmental Ca2 + contents and exchange fluxes of
Ca6 and Ca7 must reflect significant elevation of intracellular free Ca2 + to
drive the mass action.
The finding of increased extracellular Ca2 + in this study was very
interesting. A rise in extracellular Ca2 + has been reported previously in
normal in vitro rat diaphragm during exposure to a high concentration of
caffeine (27). In addition, increased extracellular Ca2 + has been observed in
in vitro rat diaphragm following fatiguing stimuli (26). It is well known that
both caffeine and fatigue can impair Ca2 + uptake by the SR (15,56).
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Furthermore, it has been shown that elevated intracellular Ca2 * in caffeine-
exposed or fatigued skeletal muscle is translocated into the t-tubules
(7,26,27). It has been suggested that increased intracellular Ca2* can be
translocated to the extracellular space via sarcolemmal Ca2 + ATPase (7,47).
Moreover, Cifuentes et al (12) demonstrated that the Na*-Ca2 * exchanger
participates in expelling intracellular Ca2 * from skeletal muscle following
depolarization, thus helping to restore resting cytosolic [Ca2 *]. These
transporters may play a significant role in extruding accumulated Ca2 + from
diaphragm fibers, thereby increasing extracellular Ca2 *.
Implications of Impaired Diaphragm Ca2 * Homeostasis in CHF
In the present study, CHF induced a chronic diaphragm Ca2 * overload
that pervaded every Ca2 * compartment. Prolonged elevation of Ca2 * is
known to produce deleterious effects in a cell by initiating processes that may
lead to cell damage or death by apoptosis or necrosis. It is tempting to
speculate that Ca2 + -dependent catabolic processes occur in the diaphragm
during CHF that may help explain the genesis of muscle wasting.
Increased Intracellular Diaphragm Ca2 *. Increased intracellular Ca2*
can play an important role in signaling various catabolic process, including
inflammation, proteolysis, and necrosis, by activating Ca2 *-dependent
proteases and endonucleases (14,39). Elevated intracellular Ca2 * also affect
the mitochondrial permeability transition pore, causing the pore to open. This
would result in disruption of the respiratory chain and cause release of
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multiple proapoptotic molecules (22). For example, it is well known that
cytochrome c released from the mitochondria in response to oxidative stress
interacts with ATP and pro-apoptotic factors in the cytosol to activate the
apoptotic protease caspase-3, leading to cellular demise. Increases in
intracellular Ca2 * have also been shown to activate a striated muscle
Ca2 + /Mg2 + -dependent endonuclease (DNAse X), translocating it to the
nucleus where it initiates internucleosomal DNA fragmentation (34). The fact
that apoptosis is ATP-dependent implies that some critical level of
cytochrome c remains in the mitochondria to preserve basal respiratory
function. Alternatively, if mitochondrial membrane rupture results in depletion
of cytochrome c from within the organelle, electron transport would collapse
leading to cell death by the energy-independent pathway of necrosis.
Supraphysiologic levels of [Ca2 *]; can also activate calpains (non-
selective Ca2 + -activated cysteine proteases). Increased calpain activity has
been observed in the fast-to-slow phenotype conversion of rat extensor '
digitorum longus muscle subjected to chronic low-frequency stimulation (51).
Calpains also play an important role in mediating skeletal muscle protein
degradation, especially in the disassembly of the myofibril during the early
stages of turnover (29). Downstream from proteolytic calpain activity, the
26S proteasome (proteasome) is involved in the processing of ubiquitin-
tagged myofibrillar proteins by selectively targeting these monomeric
peptides for degradation (31). Indeed, it has been suggested that the activity
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of the proteasome may be dependent upon and secondary to another
upstream protease, since the proteasome is able to degrade only monomeric
myofibrillar proteins, and not more organized myofibrillar structures (50). In
cytosolic Ca2 * overload these proteolytic processes would lead to muscle
degradation and atrophy. Moreover, calpains can play an essential role in
programmed cell death by cleaving the proapoptotic protein Bax in the
mitochondria, which in turn mediates cytochrome c release and initiates the
apoptotic cascade (18). This observation would seem to imply that there
may be some cross-talk and integration of various proteolytic signals during
the process of muscle degradation. The heightened activity of calpain in
CHF diaphragm relative to SHAM reported in the present study supports the
hypothesis that calpain-mediated proteolysis may play a part in CHF-
associated muscle wasting.
There is evidence to suggest that apoptosis occurs in rat hindlimb
muscle exposed to chronic CHF. Two studies utilizing a monocrotaline-
induced model of CHF reported an increased expression of the active form of
caspase-3 in rat tibialis anterior (54) and soleus (32) muscles, while atrophy
was observed only in tibialis anterior. The phenomenon of apoptosis
associated with skeletal muscle atrophy has received much attention recently
in conjunction with the myonuclear domain hypothesis (1-3). This hypothesis
proposes that each nucleus is responsible for the regulation of cytoplasmic
volume and proteins within its immediate vicinity (24). In light of the fact that
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skeletal muscle is multinucleated, apoptosis may serve as a means of
regulating fiber size during pathologic states or altered neuromuscular
activation by deleting individual myonuclei and their domains while sparing
the rest of the fiber. Thus, the deletion of single myonuclei by an apoptotic
process may not lead immediately to myofiber cell death.
Increased Extracellular Diaphragm Ca2 *. To our knowledge, there are
no studies examining the effects of a chronic increase in extracellular Ca2 *
on skeletal muscle, particularly in the presence of elevated intracellular Ca2 *.
A recent study reported that repeated stimulation of frog muscle led to an
acute elevation of t-tubular Ca2 * (10). It was speculated that this provided
protection against fatigue by enhancing the influx of Ca2 * to support
excitation-contraction coupling. Another study showed that recovery from
acute fatigue was accompanied by a loss of t-tubular Ca2* which probably
played a role in long-lasting fatigue (26). Ultimately, Ca2 * dysregulation in
fatigued muscles is reversible with rest. The findings in the present study
would seem to suggest that in CHF, Ca2 * dysregulation is a chronic
condition. The effects of chronic extracellular Ca2* engorgement in the
diaphragm have not been explored. In a recent investigation, osmotic shock
was induced in amphibian muscle leading to vacuole formation in the t-
tubules associated with impaired excitation-contraction coupling (17). When
this maneuver was followed by exposure to high extracellular Ca2 *,
irreversible electrophysiologic isolation of the t-tubules occurred. Whether
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similar changes play a role in CHF-induced diaphragm myopathy warrants
further study.
CONCLUSIONS
First, the 4 5 Ca2* kinetic data revealed a global diaphragm Ca2 +
overload in CHF, including; 1) increased Ca2 + contents of extracellular and
intracellular compartments; 2) increased Ca2 * exchange fluxes across the
\
plasma membrane and between compartments, 3) a global diaphragm Ca2 +
overload. Second, the assay for calpain showed a relative increase in the
proteolytic activity of this Ca2 + -dependent protease.
CHF induces a cellular Ca2 + overload in the diaphragm which alone,
can variously lead to impaired diaphragm function. Moreover, enhanced
proteolytic activity of Ca2 + -dependent calpain in CHF may play a role in
diaphragm wasting. In all probability, Ca2 + -mediated cellular processes play
a role in CHF-induced diaphragm wasting and dysfunction.
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CHAPTER IV
POTENTIAL ROLE FOR APOPTOSIS IN
CHF-INDUCED DIAPHRAGM MYOPATHY
INTRODUCTION
Symptomatic congestive heart failure (CHF) is characterized by
exercise intolerance, shortness of breath, and skeletal muscle wasting.
Although reduced exercise capacity in CHF has previously been attributed to
hemodynamic compromise, later investigations failed to correlate the degree
of exercise intolerance with the severity of left ventricular dysfunction (10,35).
This suggests there are factors other than cardiac failure impacting physical
performance. Recent reports show that CHF is associated with a specific
skeletal myopathy manifested by a wasting-like loss of muscle mass and
reduced force production (16,19-21). A distinctive feature of this process is
that the muscle wasting and dysfunction appear to be unrelated to inactivity
(32,39). Reduced muscle mass in the rhythmically-contracting diaphragm is
perhaps the most compelling example of the uncoupling of inactivity and
atrophy CHF. Yet until now, the question of how the CHF environment
effectively leads to muscle degradation in the diaphragm has not been
examined.
Several investigators have demonstrated that apoptosis contributes to
cardiomyopathy in CHF (11,28) and can be triggered by increased levels of
angiotensin II typically associated with an enhanced neurohormonal state
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(13). Recent studies have also reported the presence of apoptotic markers
such as terminal deoxynucleotidyl transferase-mediated dUTP nick end-
labeling (TUNEL) and DNA laddering within myonuclei in limb muscle of rats
and humans with CHF, as well as elevated levels of plasma TNF-a and
caspase-3 (15,38,40,41). These findings offer insight into potential cellular
processes that may be involved in the development of CHF-induced skeletal
myopathy. Using experimental methods that included gel electrophoresis,
immunohistochemistry, fluorescence microscopy, and substrate-specific
enzymatic assays, we tested the hypothesis that evidence of apoptotic
signals and products may be found in the diaphragm from a chronic animal
model of CHF.
METHODS
Development of CHF Animal Model
Left coronary artery ligation was employed to induce myocardial
infarction and CHF in experimental animals. Male Wistar rats (250-300 g, 8-
10 wk old) were anesthetized with a mixture of ketamine (60 mg/kg, ip) and
xylazine (9 mg/kg, ip). The animal was intubated with a 20-gauge
intravenous catheter by utilizing transillumination of the thorax to visualize
the vocal cords/tracheal opening and mechanical ventilation with a rodent
ventilator (Harvard Apparatus, Holliston, MA) was initiated (3.5 cc tidal
volume, 60Hz) with positive end-expiratory pressure (PEEP) of 5 cm H2 0. A
left thoracotomy was performed at the level of the 5th intercostal space and
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the pericardial sac was removed. The left coronary artery was ligated
approximately 2-3 mm from its origin utilizing a 6-0 Prolene suture and a
small cardiovascular pledget to compress the vessel. Blanching of the
antero-lateral and apical wall segments confirmed that the left coronary
artery was successfully ligated. The incision was closed in three layers using
4-0 Dexon absorbable sutures. Sham-operated animals underwent the same
procedure, but the left coronary artery was not ligated. The mortality rates
for the CHF and SHAM animals within the first 72 h post-surgery were
approximately 75% and 10%, respectively.
After surgery, the thoracic incision was treated topically with 0.2%
nitrofurazone solution and animals received an injection of ampicillin (0.1
g/kg body wgt, sc). Acetaminophen was placed in the drinking water (67
mg/L) for the first 48 h post surgery. The recovery period of the animals was
monitored for any complications or signs of distress. Body weight and
general well being of each animal was documented at regular intervals
during and at the end of the 10-wk period designated for development of
myocardial infarction-induced heart failure. The investigation conformed to
the Guide for the Care and Use of Laboratory Animals published by the US
National Institutes of Health (NIH Publication No. 85-23, revised 1996).
At the end of the 10-wk conditioning period, animals were sacrificed
by administering a lethal dose of Euthasol (39 mg pentobarbital sodium/5 mg
phenytoin sodium) via intraperitoneal injection.. This was typically done in
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the early afternoon to avoid diurnal variations in the concentrations of the
collected samples. Organ wet weights (heart and lungs with the large
airways removed) were recorded. Heart weight-to-body weight and lung
weight-to-body weight ratios were calculated. Additionally, lung wet-to-dry
weight ratios were recorded. Lung dry weight was determined by placing the
lungs in an oven held at 60°C for 72 h. Similar weights at 48 and 72 h
indicate true dry weight of the lungs. Significantly increased ratios of heart
weight-to-body weight and lung wet-to-dry weight in CHF animals compared
with SHAM indicate the presence of pulmonary congestion and cardiac
failure (29,33). Once a correlation between increased heart weight-to-body
weight and lung wet-to-dry weight ratios in CHF vs. sham-operated animals
was established, these ratios were used to verify the presence of CHF in all
experimental animals used in the study. Diaphragm tissue from experimental
CHF animals not meeting these criteria was excluded from the study.
Quantitative Determination of Plasma TNF-cc
At the time of sacrifice, approximately 3 cc of blood was collected from
the inferior vena cava of each animal by hypodermic needle and placed
directly into a vacutainer containing EDTA. The blood sample was allowed to
clot for 2 h at room temperature and then centrifuged for 20 min at 2,000 x g.
The serum was then transferred to a 1.5-ml microcentrifuge tube and stored
at T0°C until the time of analysis.
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The Quantikine M Rat TNF-a Immunoassay kit (R&D Systems,
Minneapolis, MN) was used to determine TNF-a levels in serum samples.
Per manufacturer’s instructions, 75 fJ serum was mixed with 75 ja l Diluent
RD5-17 to yield a 2-fold dilution. The TNF-a control sample was
reconstituted with 1 ml dhhO. Anti-rat TNF-a conjugate was prepared by
mixing 0.5 ml conjugate concentrate with 11 ml conjugate diluent in a sterile
container. A wash buffer was prepared by mixing 25 ml wash buffer
concentrate with 600 ml dHaO. Color Reagents A and B were mixed 1:1
(100 jal/well). Rat TNF-a standard was reconstituted with 2 ml calibrator
diluent RD5-17 (used in producing a dilution series).
For the assay, 50 fxl assay diluent RD1-41 was added to each well.
50 |il of each standard, control, and sample were also added to each well,
mixed for 1 min, and incubated for 2 h at room temperature. Each well was
aspirated, washed with 400 |a l wash buffer, reaspirated (total of 5 washes),
and blotted dry. 100 |il rat TNF-a conjugate was then added to each well
and incubated for 2 h at room temperature. Five aspiration/wash cycles were
repeated as before. 100 |il substrate solution was added to each well and
incubated for 30 min at room temperature (shielded from direct light). 100 |a i
of stop solution was then added to each well and mixed gently. All reactions
were run in triplicate. The plate was read using a microplate reader set to
450 nm with a correction wavelength set to 540 nm and the optical densities
(O.D.) were recorded. A standard curve was created by plotting the log of
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the rat TNF-a concentrations versus the log of the O.D. and the best fit line
determined by regression analysis. The concentration value determined
from the standard curve was multiplied by a factor of 2 (the initial dilution
factor) to yield the final concentration of rat TNF-a in the samples.
Analysis of DNA Fragmentation in Diaphragm
TUNEL. A portion of the right costal hemidiaphragm was removed at
the time of sacrifice and placed in a dissecting dish containing bicarbonate-
buffered Ringer’s solution maintained at 27°C and oxygenated with a mixture
of 95% 02/5% CO2. After trimming away the crural diaphragm and excess
connective tissue, the Ringer’s solution was replaced by 10% neutral
buffered formalin. The preparation was then allowed to fix overnight at 4°C.
After the fixation period, the tissue sample was washed in 2-3 changes (30
min/change) of dhhO and a standard dehydration protocol was employed
consisting of 2-3 changes (30 min/change) in each of the following
concentrations of ethanol; 25%, 50%, 70%, 95%, and 100%. The tissue
samples were then cleared in 2-3 changes (20 min/change) of xylene and
then embedded in paraffin (3-4 changes, 30 min/change) in a vacuum oven
set at 60°C. After cooling, the blocks were sectioned at a thickness of 5 p.m.
Sampling was randomized by taking every 10th section (representing a
distance of 50 p.m along the length of the tissue block) and mounting it on a
glass slide. The sections were then allowed to dry overnight on a slide
warmer. Eight sections per subject were prepared in this fashion.
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The mounted specimens were deparaffinized and rehydrated by
immersing them in xylene and a series of decreasing concentrations of
ETOH. The Frag-EL Fluorescein TUNEL detection kit (Oncogene Research
Products/Calbiochem, San Diego, Ca.) was used for TUNEL analysis. The
specimens were first permeabilized by incubating them in a solution
containing 20 pg/ml proteinase K for 10 min at room temperature, followed
by a rinse in TBS (20 mM Tris, pH 7.6, 140 mM NaCI). The slides were
covered with a blocking solution designed to quench autofluorescence in
formalin-fixed tissue (1% glycine, 2% bovine serum albumin, 10% normal
goat serum, 0.1 M PBS, 1 mM MgC^, and 0.1 mM CaCh) for 20 min at room
temperature. The slides were then incubated with rabbit anti-laminin
antibody (Sigma, St. Louis, MO) diluted 1:25 in TBS and placed in a
humidified chamber at 37°C for 1 h. The slides were rinsed in 3 changes of
TBS for 5 min each followed by incubation in a secondary goat-anti-rabbit
IgG conjugated with rhodamine (Sigma, St. Louis, MO) diluted 1:100 in TBS
and placed in a humidified chamber at 37°C for 1 h. Terminal
deoxynucleotidyl transferase (TdT) labeling reaction mix (a combination of
TdT enzyme and fluorescein-labeled dUTP) was added in the dark and the
slides were placed in a humidified chamber at 37°C for 1.5 h. Negative
control slides were generated by substituting dHaO in place of the TdT
enzyme and were run simultaneously with all samples. The reaction was
then terminated by three, 5-min rinses in TBS at room temperature in the
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dark. This was followed by a 5-min incubation in Hoechst 33258 in the dark
and terminated by a 2-min rinse in dHaO. Finally, the slides were mounted
with a glass coverslip utilizing mounting media provided by the manufacturer
and the edges were sealed with clear nail polish. Slides were then stored in
the dark at4°C until the time of analysis (typically in 1-2 days).
In order to detect labeled nuclei, the slides were examined utilizing a
microscope fitted with a fluorescein filter (494 nm wavelength). A brightly
fluorescent green signal indicated a positively labeled nucleus. Eight
sections were analyzed per subject. An average of 150 magnification fields
(20X) were visualized to yield a sample size of approximately 15,000 nuclei
per section. Utilizing a UV filter, all nuclei were identified based upon their
incorporation of the Hoechst 33258 stain. TUNEL+ nuclei were further
distinguished as myonuclei or interstitial nuclei based upon their location
relative to the rhodamine-stained basement membrane (40X magnification).
DNA Agarose Gel Electrophoresis. A portion of the left costal
hemidiaphragm was placed in cooled isopentane ('65-'70°C), quick-frozen,
and then placed in a scintillation vial for storage at ‘70°C. At the time of
analysis, 25 mg of frozen diaphragm tissue were minced with a clean razor
blade. Collection and purification of DNA from diaphragm tissue was carried
out using a DNeasy Tissue Kit (Qiagen, Valencia, CA).
Per manufacturer instructions, the minced diaphragm tissue was
placed in a 1.5-ml microcentrifuge tube containing 180 pi Buffer ATL. 20 pi
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of Proteinase K was then added to the tube, the mixture lightly vortexed, and
the sample incubated at 55°C (occasionally vortexing during incubation to
disperse the sample) for 3 h, or until lysed. Once lysis was complete, 4 pi of
RNase A (100 mg/ml) was added to the mixture, vortexed, and incubated at
room temperature for 2 min. After a 15-sec vortex, 200 pi of Buffer AL was
added, the tube vortexed, and the sample incubated at 70°C for 10 min. 200
pi of 100% ETOH was then added, the tube vortexed, and the mixture
pipetted into a DNeasy mini column sitting in a 2-ml collection tube.
The mini column was centrifuged at > 6,000 x g (-8,000 rpm) for 1
min, removed from the 2-ml collection tube, and placed in a new 2-ml
collection tube. 500 pi Buffer AW1 was added to the mini column and
centrifuged for 1 min at > 6,000 x g. The mini column was then removed and
placed in a new 2-ml collection tube, 500 pi Buffer AW2 added, and
centrifuged for 3 min at full speed to dry the DNeasy membrane. The mini
column was then placed in a clean 1.5-ml collection tube and 100 pi Buffer
AE was pipetted directly onto the DNeasy membrane, followed by a 1-min
incubation period at room temperature. The column was centrifuged at >
6,000 x g for 1 min to elute. This elution step was repeated a second time to
maximize DNA yield.
DNA yield was determined by measuring the concentration of DNA in
the eluate by its absorbance at 260 nm (this value should fall within the range
of 0.15-1.0). The concentration of DNA in the sample was calculated by
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taking the product of the absorbance at 260 nm (A2 6 o ) and the dilution factor
(typically 1:20 or 1:50), multiplied by 50 (ig/ml (the expected concentration of
DNA with a detection path distance of 1 cm and an A2 6 o value of 1.0). The
ratio of the absorbance readings at 260 nm and 280 nm (A2 6 o/A2 8 o ) provides
an estimate of the purity of the DNA sample with respect to contaminants
that also absorb UV light, such as protein. In order to insure accuracy, the
A2 6 o /A2 s o ratio should lie between 1.8-2.0. After the total amount and
concentration of DNA was determined, the samples were stored at *20°C
until the agarose gel was prepared.
A stock 50 X Tris-acetate (TAE) buffer solution was prepared (242 g
Tris base, 57.1 ml glacial acetic acid, and 100 ml 0.5 M EDTA in 900 ml
dH2 0 ) and diluted as necessary. A 1.5% agarose gel was utilized in order to
enhance resolution and detection of apoptotic DNA fragmentation (-180-200
bp). The gel was prepared by adding 1.5 g of agarose to 100 ml of 1 X TAE
buffer in an Erlenmeyer flask and heating the mixture until dissolved. After
cooling to 60°C, the solution was poured into the gel mold, a comb positioned
at the top edge of the mold, and the gel allowed to set (-30-45 min). Enough
1 X TAE buffer was added to the well to cover the gel and the comb was then
removed.
Approximately 5 (ig of DNA from each sample was added to a gel-
loading buffer (0.25% bromophenol blue, 0.25% xylene cyanol FF, 40%
sucrose in dH2 0 ) to yield a final volume of 20 |al and loaded into individual
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lanes. A molecular weight marker (DNA Molecular Weight Marker IX, Roche
Molecular Biochemicals, Indianapolis, IN) was loaded into the first lane to
estimate band sizes. Electrical leads were attached to the gel tank and the
voltage was set at 100 V (5 V/cm for electrodes 20 cm apart). When the
bromophenol blue migrated toward the end of the gel (typically -1.5-2 h), the
electrical current was turned off and the gel bathed in ethidium bromide for
30-45 min. The gel was then examined under UV light and photographed for
analysis.
Analysis of Diaphragm Single-Fiber Architecture
Cross-Sectional Area. In order to evaluate diaphragm wasting in the
animal model of CHF, single-fiber CSA was measured as an index of
myofiber atrophy.. Formalin fixation and ethanol dehydration during the
processing of diaphragm samples may result in the loss of up to 30% of
tissue volume (6). However, all muscles were fixed in an identical manner
thus a comparison between CHF and SHAM myofibers is nonetheless useful
to assess relative differences in single-fiber CSA (40,41). Slides used for the
TUNEL analysis were visualized by means of microscopy fitted with a
rhodamine filter and analyzed using a semi-automated, computerized
method (Bioquant Classic 98, R&M Biometrics, Nashville, TN) to calculate
single-fiber CSA (expressed as pm2). Only the area clearly outlined by the
basal lamina was identified by threshold tracing and measured as the fiber’s
CSA. Ten random magnification fields (20X) were examined (representing
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approximately 300 myofibers) from each of the 8 slides prepared per subject
and then averaged to yield mean single-fiber CSA. In this way,
approximately 2,400 single fibers were analyzed for each subject in the
study.
Estimation of Myonuclear Domain Size. Recent evidence suggests
that muscle atrophy appears to be associated with the loss of myonuclei and
that the myonuclear domain size, i.e., the amount of cytoplasm per
myonucleus, is reduced following atrophy (1-3). To estimate myonuclear
domain size in the present study, the fields examined for the single-fiber CSA
measurements were also analyzed to determine the mean number of
myonuclei per single-fiber CSA as well as the mean single-fiber CSA :
myonucleus ratio. Myonuclei were identified by fluorescent microscopy on
the basis of staining by Hoechst 33258 and their location with respect to the
myofiber basal lamina. Because the method used in the present study is
based on fiber CSA and not on fiber volume, it does not represent a precise
measure of myonuclear domain size. Nevertheless, it affords a means of
evaluating the cytoplasmic area : myonucleus ratio which may provide
information on the relative cytoplasmic domain of an individual myonucleus
as an index of muscle atrophy.
Extraction and Assay for Caspase-3 Activity in Diaphragm
At the time of sacrifice, a portion of the right costal hemidiaphragm
was placed in a dissecting dish containing bicarbonate-buffered Ringer’s
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solution maintained at 27°C and oxygenated with a mixture of 95% 02/5%
CO2- After removing the central tendon, the tissue was placed in a
dissecting dish containing cooled isopentane ('65-'70°C) and quick-frozen in
order to prepare it for analysis.
Enzyme extraction was achieved by utilizing a Caspase-3 Colorimetric
Assay Kit (R&D Systems, Minneapolis, MN). Approximately 80-100 mg wet
wgt of diaphragm tissue was finely minced with a clean razor blade and
placed in a mortar on dry ice. The tissue was ground into a powder and
placed in a 1.5-ml microcentrifuge tube containing a lysis buffer as per the
manufacturer’s specifications in a ratio of 10 ml lysis buffer/1 g tissue wet wgt
(10% w/v). The tissue sample was incubated on ice for 30 min and then
cold-centrifuged (4°C) at >10,000 rpm for 10 min. The supernatant was
transferred to a new microcentrifuge tube and kept on ice. The protein
concentration of the lysates was determined using a BCA Protein Assay Kit
(Pierce Chemical Co., Rockford, IL).
Each reaction required a minimum of 50 pi of sample lysate
(approximately 100-200 pg total protein) in a 96-well flat bottom microplate.
In addition, 50 pi of 2X reaction buffer was added to each well (for each 1 ml
of 2X reaction buffer required, 10 pi of fresh DTT stock was added to the 2X
reaction buffer prior to use). If larger volumes of lysate were required to
meet the total protein requirements, the volume of 2X reaction buffer was
also proportionately increased. Finally, 5 pi of caspase-3 colorimetric
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substrate (DEVD-pNA) was added to each well. All samples were analyzed
in triplicate. The dilution buffer provided by the manufacturer was added
when necessary to keep the reaction volumes constant in all the wells. The
plate was subsequently incubated at 37°C for 1.5 h. The reaction was then
read on a microplate reader using 405 nm wavelength light with a correction
wavelength set to 540 nm and the optical densities were recorded.
Additional background controls for this assay included: a) no tissue lysate
and b) no substrate.
The results were expressed as activity of caspase-3 in CHF
diaphragm relative to SHAM and reported in arbitrary units (AU).
Additionally, if either background control (i.e. no lysate or no substrate)
exhibited a substantial reading, it was subtracted from the experimental
results prior to reporting activity values.
Statistical Analysis
SHAM data were compared with data from CHF muscles utilizing a
Student’s t-test for independent samples in order to compare mean values of
organ weights, organ-to-body weight ratios, single-fiber CSA, myonuclei per
single-fiber cross-section, C SA: myonucleus ratio, plasma TNF-a, and
relative caspase-3 activity. DNA agarose gel data was analyzed using a chi-
square test. To test if there was an association between the presence of
TUNEL+ nuclei and CHF, TUNEL data was analyzed utilizing a Fisher’s
exact probability test and the strength of any association was assessed by
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calculating the 95% confidence interval for the odds ratio. The accepted
level of significance was set at p<0.05 and results were expressed as
mean±SE when appropriate.
RESULTS
Characteristics of the Chronic CHF Animal Model
The initial body weight of CHF vs. SHAM animals was not different
(302.9±5.9 g vs. 296.3+4.1 g, p>0.05). Although body weight increased
significantly in both groups, CHF animals gained less weight over the 10-wk
conditioning period compared to sham-operated animals (118.1 ±5.9 g vs.
131.5+5.0 g, p<0.05).
As shown in Table 4.1, whole hearts harvested from CHF animals
weighed significantly more than hearts from sham-operated animals
(2.73+0.15 g vs. 1.57+0.08 g, p<0.05) resulting in an 83% increase in heart
weight/body weight ratio (6.63+0.42 vs. 3.63±0.17, p<0.05). A large area of
fibrotic scar on the anterior and lateral aspects of the left ventricle was noted
in all CHF animals, indicating myocardial infarction.
Lung wet weight/dry weight ratio was increased in CHF animals
compared to SHAM (5.04+0.08 vs. 4.48±0.02, p<0.05), and lung weight/body
weight ratio was elevated three-fold in CHF (13.20+0.97 vs. 4.18+0.10,
p<0.05). Increases in all of these ratios have been shown to be highly
correlated with elevated left ventricular end diastolic pressures and impaired
left ventricular contractility in CHF (29,33).
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Characteristics of the
Chronic Animal Model
Parameter SHAM CHF
Heart Wt (g) 1.57±0.08 2.73±0.15*
Heart Wt/Body Wt (x103) 3.63±0.17 6.63±0.42*
Lung Wet Wt/Dry Wt 4.48±0.02 5.04±0.08*
Lung Wt/Body Wt (x103) 4.18±0.10 13.20±0.97*
Table 4.1. Mean heart and lung weights and ratios in CHF and SHAM animals.
•Significantly different from SHAM (p<0.05).
1200
1000
SHAM
Figure 4.1. Mean diaphragm single-fiber CSA of SHAM and CHF animals. CHF was
associated with a smaller single-fiber CSA. Error bar corresponds to 1 SE. 'Significantly
different from SHAM (p<0.05).
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Morphometric Analysis of Diaphragm Single Fibers
Analysis of single-fiber CSA revealed a significant difference in mean
values between CHF and SHAM diaphragm (844+40 jam2 vs. 962±27 p.m2,
p<0.05). This difference resulted in a mean single-fiber CSA measurement
that was 12% less in CHF diaphragm compared with SHAM (Fig. 4.1). Mean
number of myonuclei per single-fiber CSA was not different between CHF
and SHAM diaphragm (2.73+0.12 vs. 2.68±0.15, p>0.05), but mean single-
fiber CSA : myonucleus ratio was 15% less in CHF diaphragm relative to
SHAM (312.8±21.8 vs. 369.4+13.7, p<0.05, Fig. 4.2). Although number of
500
*400
d SHAM;
■ CHF
Figure 4.2. Mean diaphragm single-fiber CSA : myonucleus ratio in SHAM and CHF
animals. CHF was associated with a smaller single-fiber CSA: myonucleus ratio. Error bar
corresponds to 1 SE. ‘ Significantly different from SHAM (p<0.05).
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nuclei per single-fiber CSA did not differ between the two groups, the smaller
predicted myonuclear domain size in CHF diaphragm relative to SHAM could
be accounted for by the observed smaller CSA noted in CHF diaphragm
muscle fibers.
Plasma TNF-a Levels in CHF
Blood samples taken from CHF rats contained markedly higher
plasma TNF-a levels compared to sham-operated rats (4.16±2.08 pg/ml vs.
0.30±0.12 pg/ml, p<0.05). This represented an almost 14-fold higher plasma
TNF -a level in CHF experimental animals relative to SHAM (Fig. 4.3).
_7
□ SHAM
g CHF
Figure 4.3. Effects of CHF on mean plasma TNF-a levels. Error bar corresponds to 1 SE.
‘Significantly different from SHAM (p<0.05).
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TUNEL Analysis in Diaphragm
The mean number of total nuclei counted per section (myonuclei +
nuclei in the interstitium) did not differ between the CHF and SHAM groups
(14,960±1877 vs. 15,866+1213, p>0.05). Of the 12 CHF rats included in the
study, 9 (75%) demonstrated evidence of either myonuclear or interstitial
nuclear TUNEL+ staining in the diaphragm. By contrast, TUNEL+ nuclei
were noted in the diaphragm of only 1 (10%) of the 10 SHAM rats. Analysis
of the data utilizing a Fisher’s exact probability test indicated that the
proportions of diaphragms exhibiting TUNEL+ staining were not identical in
the CHF and sham-operated rats (p<0.05). Among the two groups of rats in
the study, the presence of CHF increased the incidence of TUNEL+ nuclei
(Figs. 4.4 to 4.6). Furthermore, the data suggested that the odds of
exhibiting TUNEL+ nuclei in the diaphragm were 27 times greater in rats with
Figure 4.4. Interstitial terminal deoxynucleotidyl transferase-mediated dUTP nick end-
labeling (TUNEL)+ nucleus in the diaphragm. Apoptotic nuclei within the interstitium
visualized using a fluorescein (a) and rhodamine (b) filter. Magnification = 40X. Apoptotic
nucleus is indicated by each arrow.
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CHF than in sham-operated rats (95% confidence interval for OR = 2.34,
311). In each section from the CHF animals, we observed approximately
twice as many TUNEL+ interstitial nuclei than myonuclei (3.67+0.67 vs.
1.78+0.33, p<0.05).
Figure 4.5. Peripheral myocyte terminal deoxynucleotidyl transferase-mediated dUTP nick
end-labeling (TUNEL)+ nucleus in the diaphragm. Apoptotic nucleus within the myocyte
basal lamina visualized using a fluorescein (a) and rhodamine (b) filter. Magnification = 40X.
Apoptotic nucleus is indicated by each arrow.
Figure 4.6. Centralized myocyte terminal deoxynucleotidyl transferase-mediated dUTP nick
end-labeling (TUNEL)+ nucleus in the diaphragm. Apoptotic nucleus within the myocyte
basal lamina visualized using a fluorescein (a) and rhodamine (b) filter. Magnification = 40X.
Apoptotic nucleus is indicated by each arrow.
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DNA Laddering in Diaphragm
Electrophoretic separation of DNA samples from CHF and SHAM
diaphragm did not reveal any evidence of intemucleosomal DNA cleavage in
either group (chi-square test, p>0.05, data not shown). Both groups
demonstrated nucleic acid band sizes characteristic of genomic DNA,
suggesting that if intemucleosomal DNA cleavage was indeed present, it was
likely undetectable by methods used in this study.
Relative Activity Level of Caspase-3 in CHF Diaphragm
Qualitative analysis of caspase-3 activity revealed a statistically
significant difference in mean O.D. readings between CHF and SHAM
diaphragm sample lysates (1.73+0.2 AU vs. 1.28±0.15 AU, p<0.05, Fig. 4.7).
SHAM
Figure 4.7. Effects of CHF on diaphragm caspase-3 proteolytic activity. Mean data from
CHF diaphragms are compared to SHAM. Error bar corresponds to 1 SE. 'Significantly
different from SHAM (p<0.05).
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This accounted for a relative caspase-3 activity index that was 36% higher in
CHF diaphragm compared to SHAM.
DISCUSSION
This study explored the premise that apoptosis potentially contributes
to diaphragm fiber remodeling in CHF. Ultrastructural analysis of diaphragm
from CHF and SHAM animals yielded the following observations: 1) Mean
diaphragm single-fiber CSA was smaller in CHF vs. SHAM animals. 2)
Myonuclear domain size was estimated to be smaller in CHF compared with
SHAM animals. 3) Plasma TNF-a levels were higher in CHF vs. sham-
operated animals. 4) A higher incidence of TUNEL+ nuclei was
demonstrated in experimental CHF diaphragm relative to SHAM. 5) The
proteolytic activity of caspase-3 was higher in CHF diaphragm compared with
SHAM.
Diaphragm Wasting in CHF
Loss of diaphragm CSA in the presence of CHF is now a well-
recognized phenomenon (12,21). Although limb muscle undergoes
significant wasting in models of disuse (1,27) as well as CHF (40,41), the
diaphragm exposed to CHF exhibits atrophy while maintaining its chronic,
rhythmic activation pattern. Previous studies have reported single-fiber CSA
in diaphragm and limb muscle from animals and humans with CHF to be 9-
38% less in CHF animals relative to control (12,40,41). In agreement with
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these results, the present study found mean diaphragm single-fiber CSA to
be 12% less in CHF vs. sham-operated animals (Fig. 1).
How muscle mass is reduced in the CHF environment is not well
understood. Evidence suggests that one mechanism by which muscles are
able to modulate fiber size is the regulation of myonuclear number (1,3). The
number of myonuclei per single fiber and the cytoplasmic volume :
myonucleus ratio, referred to as the myonuclear domain, offer a unique
insight into the ways in which muscle fibers can respond to factors which can
influence their structure and function. For example, Reductions of
myonuclear number and fiber CSA have been reported in rat soleus after
spaceflight (3) and in human soleus after prolonged bedrest (27), resulting in
smaller myonuclear domain size compared with control subjects. By
reducing myonuclear content, the availability of templates for transcription
(i.e., DNA) can be modulated and may result in diminished muscular protein
content, ultimately impacting fiber size.
The results of this study revealed no difference in mean number of
myonuclei per single-fiber CSA between CHF and SHAM diaphragm. This
finding may have been influenced by the methods used. Previous studies
have estimated the mean number of nuclei per fiber cytoplasmic volume
using confocal microscopy (1,27). In the present study, the mean number of
nuclei per fiber cross section was measured. This approach is obviously less
sensitive than confocal microscopy. Since the mean single-fiber CSA of CHF
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diaphragm was less than SHAM, the ratio of mean single-fiber CSA:
myonucleus (a prediction of myonuclear domain size) was also lower in CHF
diaphragm relative to SHAM (Fig. 2). Why the deletion of some myonuclei
leads to a reduction in the overall myonuclear domain size of the remaining
myonuclei is not clear. One may speculate that neighboring myonuclei share
control of overlapping areas of cytoplasm. When one myonucleus is
eliminated, the myonuclear domain of the surviving myonucleus is reduced
accordingly.
Plasma TNF-a Levels Associated with CHF
Excessive neurohormonal stimulation is now receiving attention as a
potential mediator of the complex hemodynamic, functional, and metabolic
abnormalities observed in chronic CHF (7,8). Supraphysiologic doses of
TNF-a are known to stimulate muscle proteolysis (9) and evidence suggests
that TNF-a may cause an increase in mRNA encoding for various
components of the ubiquitin-proteasome pathway that for protein degradation
(17). Proteolysis by the proteasome pathway has been characterized in
various catabolic states (23,24). Moreover, increased production of TNF-a
has been linked to the genesis of skeletal muscle cachexia (37) and is
associated with a marked activation of the renin-angiotensin system in
patients with CHF (14). These findings suggest that TNF-a may potentially
contribute to muscle wasting in CHF. Our finding of elevated plasma TNF-a
levels in experimental animals compared with sham-operated animals is in
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agreement with previous human and animal studies (4,5,14,15,41). Although
this is a non-specific finding, it is consistent with the animal model and
clinical picture of CHF.
Apoptotic Markers in Chronic CHF
Recent findings that apoptosis occurs in cardiac myocytes (26,28) and
in skeletal muscle (15,40,41) of animals and humans with CHF have
refocused research in the area of CHF-induced skeletal myopathy.
Apoptosis has now emerged as a well-recognized mechanism leading to
myocardial remodeling and is receiving attention as a potential contributing
factor to skeletal myopathy observed in CHF.
Vescovo et al (41) have identified generalized atrophy, elevated levels
of plasma TNF-a, and apoptotic-like DNA fragmentation in myofibers and
interstitial cells of rat tibialis anterior in a monocrotaline-induced model of
pulmonary hypertension and right ventricular failure. The group also
observed similar results in soleus muscle taken from the same group of
monocrotaline-treated rats (15). In both studies, there were significantly
more TUNEL+ nuclei located in the interstitium and endothelial cells than in
myocytes.
The proteolytic cascade that is the effector pathway of apoptosis can
be initiated by various signals, including; the binding of the cytokines FasL
and TNF-a to a cell surface death receptor (25), the formation of a
cytochrome c/APAF-1 complex in response to mitochondrial stress (30,43),
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and release of granzyme B from cytotoxic T-cells (34). The apoptotic
machinery is primarily composed of a group of proteolytic enzymes referred
to collectively as the caspases (cysteinyl-aspartate-specific proteinases).
Activation of caspase-3, one of the so-called executioner caspases, is a
common element to all of these signaling pathways. Once activated,
caspase-3 mediates apoptosis by cleaving selected intracellular proteins, as
well as proteins of the nuclear lamina, cytoskeleton, and sarcoplasmic-
endoplasmic reticulum (22,42). Recent findings indicate that the protein
content of the active form of caspase-3 is increased in the monocrotaline-
induced model of CHF (15,41), as well as patients with CHF (40), while the
protein content of Bcl-2 (an inhibitor of apoptosis) is decreased. In the
present study, an increase in the proteolytic activity of caspase-3 in CHF
diaphragm homogenates relative to SHAM was demonstrated utilizing an
assay containing a caspase-3-specific substrate. These data suggest
activation of a caspase-3-mediated apoptotic pathway in diaphragm from a
chronic CHF animal.
In mono-nucleated cells such as in the myocardium, apoptosis is
known to cause death of the entire cell (26,28). Since skeletal myocytes are
multi-nucleated, they represent a unique cell population in the study of
apoptotic cellular demise. Apoptosis has been implicated in the
degeneration associated with dystrophin-deficient muscular dystrophy,
although necrosis is also known to play a significant role (36). Apoptosis, in
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conjunction with necrosis, may contribute to the demise of entire skeletal
muscle fibers in muscular dystrophy. Alternatively, apoptosis of a single
myonucleus alone may not lead immediately to death of an entire skeletal
muscle fiber, but instead, result in the loss of only the associated cytoplasmic
domain which would reduce mass (1-3,41).
Results of the present study were in agreement with previous work on
skeletal myopathy associated with CHF. However, the present data
suggests a lower incidence of nuclear apoptosis in diaphragm compared with
limb skeletal muscle of rats with CHF (15,41). Several key points may help
explain the discrepancy. Vescovo et al (41) and Libera et al (15) induced
CHF in experimental rats by intraperitoneal injection of monocrotaline,
leading to pulmonary hypertension and right ventricular failure. The resulting
peripheral edema and abdominal ascites is consistent with the disorder of cor
pulmonale (31). In the present study, a myocardial infarction-induced animal
model of left ventricular failure following left coronary artery ligation was
utilized because this model approximates the most common pathogenesis of
CHF in humans. Although the toxic effects of monocrotaline are primarily
localized to the pulmonary artery, it has been suggested that it is difficult to
rule out potentially toxic effects in other organs (18).
Moreover, Vescovo et al (41) and Libera et al (15) analyzed the tibialis
anterior and soleus, which are peripheral skeletal muscles that exhibit a
spontaneous pattern of activation. In contrast, the diaphragm muscle
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possesses a chronic, rhythmic activation pattern throughout life. There are
obvious differences in the patterns of activation between the diaphragm and
the tibialis anterior/soleus. CHF-induced alterations in cellular structure and
the incidence of apoptosis in muscles possessing different activation patterns
needs to be investigated in future studies.
CONCLUSIONS
The results of this study are consistent with the hypotheses that; 1)
CHF is associated with reduced single-fiber CSA and atrophy of diaphragm
muscle, 2), mean single-fiber CSA : myonucleus ratio is decreased in CHF
diaphragm relative to SHAM, 3) plasma TNF-a levels are elevated in CHF
animals compared to sham-operated animals, 4) the frequency of apoptotic-
like nuclear fragmentation is higher in CHF diaphragm relative to SHAM, and
5) the relative activity of caspase-3 is significantly higher in CHF diaphragm
compared to SHAM. The current evidence suggests that various apoptotic
mediators are present and exhibit increased activity within the environment of
CHF and potentially contribute to diaphragm myopathy.
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CHAPTER V
FINAL COMMENTS
Limitations of the Study
The findings that lung wet weight/dry weight ratio, heart weight, and
heart weight/body weight ratio were all increased in CHF animals was used
to validate the experimental model in the present study. Increased values in
this regard are well-established post-mortem indices of chronic heart failure
that have been correlated with pulmonary congestion and increased left
ventricular end-diastolic pressure (2,3). Although we were able to confirm
the presence of CHF in our experimental animals, the severity of the cardiac
dysfunction could not be assessed. This would have been possible with the
use of neonatal/pediatric echocardiography but this technology was not
available to us during the course of the experiment. In the future,
echocardiography would allow dynamic assessment of left ventricular
function and estimation of the severity of CHF. Such capability would permit
study of diaphragm myopathy at different time points during the development
of CHF.
The experimental methodology chosen to evaluate Ca2 + homeostasis
in the diaphragm enabled us to derive estimates of compartmental Ca2 +
contents and exchange fluxes during steady-state conditions. Although the
simulation, analysis, and modeling program yields a substantial amount of
information about physiologic processes, the method is not sensitive enough
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to detect the relatively small [Ca2 + ] present in the cytosol. The inclusion of an
additional technique such as fura-2 or indo 1/AM to measure cytosolic Ca2 +
would have independently supported our contention that diaphragm
intracellular free Ca2 + is elevated in CHF.
It is well known that CHF is associated with global fiber atrophy in the
diaphragm and other skeletal muscles in animal models of CHF (1,4,5).
Thus, it was important to confirm the presence of diaphragm wasting in the
coronary artery ligation model of CHF in our hands. We randomly sampled
and measured 1,500 single diaphragm fibers from each animal in the study
to establish a mean single-fiber CSA value. Our index of diaphragm wasting
was defined as a significantly smaller mean CSA value in CHF vs. SHAM
diaphragm.
Single-fiber CSA was also used in estimating myonuclear domain size
by establishing a ratio of single-fiber CSA per myonucleus. Since the
sampling of single-fiber CSA was randomized along the entire length of the
diaphragm tissue samples, this allowed us to predict the myonuclear domain
size, which is typically estimated by calculating the fiber cytosolic volume :
myonucleus ratio. Other investigators have measured cytosolic volume by
means of confocal microscopy, allowing for the visualization of the fiber’s
longitudinal dimensions without the need for sectioning. This method yields
a more precise estimate of myonuclear domain size because the 3-
dimensional characteristics of the muscle fiber are better appreciated.
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Access to this technology in future studies will significantly improve the ability
to estimate diaphragm myonuclear domain size in our model.
Conclusions
The present study revealed the following: 1) Diaphragm wasting
occurs in the environment of CHF, 2) CHF is associated with altered Ca2 +
homeostasis that increases diaphragm compartmental Ca2 + contents and
exchange fluxes and may provide a signal for altering calpain activity, and 3)
apoptosis may play a role in CHF-associated diaphragm wasting and
dysfunction as suggested by increased caspase-3 activity and the presence
of TUNEL+ nuclei. The results of this study suggest that multiple proteolytic
mechanisms potentially contribute to diaphragm wasting within the
environment of CHF.
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42. Mancini DM, Henson D, LaManca J, et al. Evidence of reduced
respiratory muscle endurance in patients with heart failure. JACC 24:
972-981, 1994.
43. Mancini DM, LaManca J, Donchez L, et al. The sensation of dyspnea
during exercise is not determined by the work of breathing in patients
with heart failure. JACC 28: 391-395, 1996.
8 1
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44. Mancini DM, Walter G, Reichek N, et al. Contribution of skeletal
muscle atrophy to exercise intolerance and altered muscle metabolism
in heart failure. Circulation 85(4): 1364-1373, 1992.
45. Mancini M, Nicholson DW, Roy S, et al. The caspase-3 precursor has
a cytosolic and mitochondrial distribution: implications for apoptotic
signaling. J Cell Bio 140(6): 1485-1495, 1998.
46. Mankak JS and McConnell TR. Pulmonary manifestations of chronic
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47. Manpuru LJ, Chen S-J, Kalenik JL, et al. Analysis of events
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48. Massie BM, Simonini A, Sahgal P, et al. Relation of systemic and local
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67. Schocken DD, Arrieta Ml, Leverton PE, et al. Prevalence and mortality
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78. Vescovo G, Serafini F, Facchin L, et al. Specific changes in skeletal
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85. Yue T-L, Wang C, Romanic AM, et al. Staurosporine-induced
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Cardiol 30:495-507,1998.
86. Yue T-L, Xin-Liang M, Wang X, et al. Possible involvement of stress-
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CHAPTER II
1. Ferrari R. The role of TNF in cardiovascular disease. Pharmacol Res
40(2): 97-105, 1999.
2. Ferrari R, Bachetti T, Confortini R, et al. Tumor necrosis factor soluble
receptors in patients with various degrees of congestive heart failure.
Circulation 92(6): 1479-1486, 1995.
3. Levine B, Kalman J, Mayer L, et al. Elevated circulating levels of
tumor necrosis factor in severe chronic heart failure. N Engl J Med
323: 236-241, 1990.
4. McMurray J, Abdullah I, Dargie HJ, et al. Increased concentrations of
tumor necrosis factor in “cachectic" patients with severe chronic heart
failure. Br. Heart J 66: 356-358, 1991.
5. Randhawa AK and Singal PK. Pressure overload-induced cardiac
hypertrophy with and without dilation. J Am Coll Cardiol 20:1569-
1575, 1992.
6. Sjaastad I, Sejersted OM, llebekk A, et al. Echocardiographic criteria
for detection of postinfarction congestive heart failure in rats. J Appl
Physiol 89: 1445-1454, 2000.
CHAPTER III
1. Allen DL, Linderman JK, Roy RR, et al. Apoptosis: a mechanism
contributing to remodeling of skeletal muscle in response to hindlimb
unweighting. Am J Physiol 273 (Cell Physiol. 42): C579-C587, 1997.
2. Allen DL, Monke SR, Talmadge RJ, et al. Plasticity of myonuclear
number in hypertrophied and atrophied mammalian skeletal muscle
fibers. J Appl Physiol 78 (5): 1969-1976, 1995.
3. Allen DL, Yasui W, Tanaka T, et al. Myonuclear number and myosin
heavy chain expression in rat soleus single muscle fibers after
spaceflight. J Appl Physiol 81 (1): 145-151, 1996.
4. Baker JR. Principles of Biological Microtechnique. London: Methuen,
1960.
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24. Hall ZW and Ralston E. Nuclear domains in muscle cells. Cell 59: 771-
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25. Hammond MD, Bauer KA, Sharp JT, et al. Respiratory muscle
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27. Howell S, Fitzgerald RS, and Phair RD. Long term calcium-45 efflux
kinetics from rat diaphragm. In: Sieck GC, ed. Respiratory Muscles
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28. Howell S, Maarek J-M, Fournier M, et al. Congestive heart failure:
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29. Huang J and Forsberg NE. The role of calpain in skeletal-muscle
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31. Lecker SH, Solomon V, Mitch WE, et al. Muscle protein breakdown
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disease states. J Nutr 129: 227S-237S, 1999.
32. Libera LD, Zennaro R, Sandri M, et al. Apoptosis and atrophy in rat
slow skeletal muscles in chronic heart failure. Am J Physiol 277 (Cell
Physiol 46): C982-C986, 1999.
33. Lindsay DC, Lovegrove CA, Dunn MJ, et al. Histological abnormalities
of muscle from limb, thorax, and diaphragm in chronic heart failure.
Eur Heart J 17:1239-1250, 1996.
34. Los M, Neubuser D, Coy JF, et al. Functional characterization of
DNase X, a novel endonuclease expressed in muscle cells.
Biochemistry 39(25): 7365-7373, 2000.
35. MacFarlane NG, Darnley GM, and Smith GL. Cellular basis for
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heart failure. Am J Physiol Cell Physiol 278: C739-C746, 2000.
36. Mancini DM, Henson D, LaManca J, et al. Respiratory muscle function
and dyspnea in patients with chronic congestive heart failure.
Circulation 86: 909-918, 1992.
37. Mancini DM, Henson D, LaManca J, et al. Evidence of reduced
respiratory muscle endurance in patients with heart failure. J Am Coll
Cardiol 24: 972-981, 1994.
89
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48. Simonini A, Chang K, Yue P, et al. Expression of skeletal muscle
sarcoplasmic reticulum calcium-ATPase is reduced in rats with
postinfarction heart failure. Heart 81 (3): 303-307, 1999.
49. Sjaastad I, Sejersted OM, llebekk A, et al. Echocardiographic criteria
for detection of postinfarction congestive heart failure in rats. J Appl
Physiol 89:1445-1454, 2000.
50. Solomon V and Goldberg AL. Importance of the ATP-ubiquitin-
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51. Sultan KR, Dittrich BT, and Pette D. Calpain activity in fast, slow,
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Cell Physiol 279: C639-C647, 2000.
52. Tikunov B, Levine S, and Mancini D. Chronic congestive heart failure
elicits adaptations of endurance exercise in diaphragmatic muscle.
Circulation 95: 910-916,1997.
53. Vescovo G, Volterrani M, Zennaro R, et al. Apoptosis in the skeletal
muscle of patients with heart failure: investigation of clinical and
biochemical changes. Heart 84(4): 431-437, 2000.
54. Vescovo G, Zennaro R, Sandri M, et al. Apoptosis of skeletal muscle
myofibers and interstitial cells in experimental heart failure. J Mol Cell
Cardiol 30: 2449-2459, 1998.
55. Walsh JT, Andrews R, Johnson P, et al. Inspiratory muscle endurance
in patients with chronic heart failure. Heart 76(4): 332-336, 1996.
56. Weber A. The mechanism of the action of caffeine on sarcoplasmic
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CHAPTER IV
1. Allen DL, Linderman JK, Roy RR, et al. Apoptosis: a mechanism
contributing to remodeling of skeletal muscle in response to hindlimb
unweighting. Am J Physiol 273 (Cell Physiol. 42): C579-C587,1997.
2. Allen DL, Monke SR, Talmadge RJ, et al. Plasticity of myonuclear
number in hypertrophied and atrophied mammalian skeletal muscle
fibers. J Appl Physiol 78 (5): 1969-1976, 1995.
91
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
3. Allen DL, Yasui W, Tanaka T, et al. Myonuclear number and myosin
heavy chain expression in rat soleus single muscle fibers after
spaceflight. J Appl Physiol 81 (1): 145-151, 1996.
4. Anker SD, Chua TP, Ponikowski P, et al. Hormonal changes and
catabolic/anabolic imbalance in chronic heart failure and their
importance for cardiac cachexia. Circulation 96(2): 526-534, 1997.
5. Anker SD, Volterrani M, Egerer KR, et al. Tumour necrosis factor
alpha as a predictor of impaired peak leg blood flow in patients with
chronic heart failure. QJM 91(3): 199-203, 1998.
6. Baker JR. Principles of Biological Microtechnique. London: Methuen,
1960.
7. Ferrari R, Bachetti T, Confortini R, et al. Tumor necrosis factor soluble
receptors in patients with various degrees of congestive heart failure.
Circulation 92(6): 1479-1486, 1995.
8. Ferrari R, Ceconi C, Curello S et al. Activation of the neuroendocrine
response in heart failure: adaptive or maladaptive process?
Cardiovasc Drug Ther 10: 623-629, 1996.
9. Goodman MN. Tumor necrosis factor induces skeletal muscle protein
breakdown in rats. Am J Physiol 260: E727-E730, 1991.
10. Gordon A, Voipio-Pulkki, and Liisa-Maria. Crosstalk of the heart and
periphery: skeletal and cardiac muscle as therapeutic targets in heart
failure. Ann Med 29(4): 327-331,1997.
11. Hamet P, Lucie R, Dam T-V, et al. Apoptosis in target organs of
hypertension. Hypertension 26(4): 642-648, 1995
12. Howell S, Maarek J-M, Fournier M, et al. Congestive heart failure:
differential adaptation of the diaphragm and latissimus dorsi. J Appl
Physiol 79 (2): 389-397, 1995.
13. Kajstura J, Cigola E, Malhotra A, et al. Angiotensin II induces
apoptosis of adult ventricular myocytes in vitro. J Mol Cell Cardiol 29:
859-870, 1997.
14. Levine B, Kalman J, Mayer L, et al. Elevated circulating levels of
tumor necrosis factor in severe chronic heart failure. N Engl J Med
323:236-241, 1990.
92
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15. Libera LD, Zennaro R, Sandri M, et al. Apoptosis and atrophy in rat
slow skeletal muscles in chronic heart failure. Am J Physiol 277 (Cell
Physiol 46): C982-C986, 1999.
16. Lindsay DC, Lovegrove CA, Dunn MJ, et al. Histological abnormalities
of muscle from limb, thorax, and diaphragm in chronic heart failure.
Eur Heart J 17:1239-1250, 1996.
17. Llovera M, Garcia-Martinez C, Agell N, et al. TNF can directly induce
the expression of ubiquitin-dependent proteolytic system in rat soleus
muscles. Biochem Biophys Res Commun 230: 238-241, 1997.
18. Lunde PK, Sjaastad I, Schiotz Thorud H-M, et al. Skeletal muscle
disorders in heart failure. Acta Physiol Scand 171(3): 277-294, 2001.
19. Mancini DM, Henson D, LaManca J, et al. Evidence of reduced
respiratory muscle endurance in patients with heart failure. J Am Coll
Cardiol 24: 972-981, 1994.
20. Mancini DM, LaManca J, Donchez L, et al. The sensation of dyspnea
during exercise is not determined by the work of breathing in patients
with heart failure. JACC 28: 391-395, 1996.
21. Mancini DM, Walter G, Reichek N, et al. Contribution of skeletal
muscle atrophy to exercise intolerance and altered muscle metabolism
in heart failure. Circulation 85:1364-1373, 1992.
22. Mignotte B and Vayssiere J-L. Mitochondria and apoptosis. Eur J
Biochem 252:1-15, 1998.
23. Mitch WE, Bailey JL, Wang X, et al. Evaluation of signals activating
ubiquitin-proteasome proteolysis in a model of muscle wasting. Am J
Physiol 276 (Cell Physiol 45): C1132-C1138, 1999.
24. Mitch WE and Price SR. Protein degradation by proteasomes:
molecular mechanisms of muscle catabolism. Nephrol Dial Transplant
12: 13-15, 1997.
25. Muzio M, Chinaiyan AM, Kischkel FC, et al. Flice, a novel fadd-
homologous ice/ced-3-like protease, is recruited to the cd95 (fas/apo-
1) death-inducing signaling complex. Cell 85(6): 803-815, 1996.
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26. Narula J, Pandey P, Arbustini E, et al. Apoptosis in heart failure:
release of cytochrome c from mitochondria and activation of caspase-
3 in human cardiomyopathy. Proc Natl Acad Sci USA 96(14): 8144-
SI 49, 1999.
27. Ohira Y, Yoshinaga T, Ohara M, et al. Myonuclear domain and
myosin phenotype in human soleus after bed rest with or without
loading. J Appl Physiol 87(5): 1776-1785, 1999.
28. Olivetti G, Abbi R, Quaini F, et al. Apoptosis in the failing human heart.
N Engl J Med 336:1131-1141, 1997.
29. Randhawa AK and Singal PK. Pressure overload-induced cardiac
hypertrophy with and without dilation. J Am Coll Cardiol 20:1569-
1575, 1992.
30. Reed JC and Patemostro G. Postmitochondrial regulation of apoptosis
during heart failure. Proc Natl Acad Sci USA 96(14): 7614-7616, 1999.
31. Schultze AE and Roth RA. Chronic pulmonary hypertension- the
monocrotaline model and involvement of the hemostatic system. J Tox
Environ Health Part B, Crit Rev 1: 271-346, 1998.
32. Simonini A, Long CS, Dudley GA, et al. Heart failure in rats causes
changes in skeletal muscle morphology and gene expression that are
not explained by reduced activity. Circ Res 79:128-136, 1996.
33. Sjaastad I, Sejersted OM, llebekk A, et al. Echocardiographic criteria
for detection of postinfarction congestive heart failure in rats. J Appl
Physiol 89: 1445-1454, 2000.
34. Stennicke HR, Jugensmeier JM, Shin H et al. Pro-caspase-3 is a
major physiologic target of caspase-8. J Biol Chem 273: 27084-27090,
1998.
35. Strassijns G, Lysens R, and Decramer M. Peripheral and respiratory
muscles in chronic heart failure. Eur Respir J 9: 2161-2167,1996.
36. Tidball JG, Albrecht DE, Lokensgard BE, et al. Apoptosis precedes
necrosis of dystrophin-deficient muscle. J Cell Science 108: 2197-
2204, 1995.
94
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37. Tracey KJ, Morgello S, Koplin B, et al. Metabolic effects of
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38. Vescovo G, Ambrosio GB, and Dalla Libera L. Apoptosis and changes
in contractile protein pattern in the skeletal muscle in heart failure.
Acta Physiol Scand 171(3): 305-310, 2001.
39. Vescovo G, Serafini F, Facchin L, et al. Specific changes in skeletal
muscle myosin heavy chain composition in cardiac failure: differences
compared with disuse atrophy as assessed on microbiopsies by high
resolution electrophoresis. Heart 76: 337-343, 1996.
40. Vescovo G, Volterrani M, Zennaro R, et al. Apoptosis in the skeletal
muscle of patients with heart failure: investigation of clinical and
biochemical changes. Heart 84(4): 431-437, 2000.
41. Vescovo G, Zennaro R, Sandri M, et al. Apoptosis of skeletal muscle
myofibers and interstitial cells in experimental heart failure. J Mol Cell
Cardiol 30: 2449-2459, 1998.
42. Wolf BB, Schuler M, Echeverri F, et al. Caspase-3 is the primary
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45/inhibitor of caspase-activated dnase inactivation. J Biol Chem 274:
30651-30656, 1999.
43. Yue T-L, Wang C, Romanic AM, et al. Staurosporine-induced
apoptosis in cardiomyocytes: a potential role of caspase-3. J Mol Cell
Cardiol 30:495-507, 1998.
CHAPTER V
1. Howell S, Maarek J-M, Fournier M, et al. Congestive heart failure:
differential adaptation of the diaphragm and latissimus dorsi. J Appl
Physiol 79 (2): 389-397, 1995.
2. Randhawa AK and Singal PK. Pressure overload-induced cardiac
hypertrophy with and without dilation. J Am Coll Cardiol 20:1569-
1575, 1992.
95
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
3. Sjaastad I, Sejersted OM, llebekk A, et al. Echocardiographic criteria
for detection of postinfarction congestive heart failure in rats. J Appl
Physiol 89: 1445-1454, 2000.
4. Vescovo G, Volterrani M, Zennaro R, et al. Apoptosis in the skeletal
muscle of patients with heart failure: investigation of clinical and
biochemical changes. Heart 84(4): 431-437, 2000.
5. Vescovo G, Zennaro R, Sandri M, et al. Apoptosis of skeletal muscle
myofibers and interstitial cells in experimental heart failure. J Mol Cell
Cardiol 30: 2449-2459, 1998.
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
96
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catabolic/anabolic imbalance in chronic heart failure and their importance for
cardiac cachexia. Circulation 96(2): 526-534,1997.
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Asset Metadata

Creator Dominguez, Jesus Felipe (author)

Core Title Potential factors contributing to diaphragm myopathy in congestive heart failure

Contributor Digitized by ProQuest (provenance)

Degree Doctor of Philosophy

Degree Program Biokinesiology

Degree Conferral Date 2002-05

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag biology, animal physiology,biology, cell,biology, molecular,health sciences, medicine and surgery,health sciences, pathology,OAI-PMH Harvest

Language English

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c16-217048

Unique identifier UC11339250

Identifier 3073770.pdf (filename),usctheses-c16-217048 (legacy record id)

Legacy Identifier 3073770.pdf

Dmrecord 217048

Document Type Dissertation

Rights Dominguez, Jesus Felipe

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the au...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus, Los Angeles, California 90089, USA