Effects of Voluntary Physical Activity and Endurance Training on Cardiac Mitochondrial Function of Rats

Sub-Chronically Treated with Doxorubicin

Dissertation submitted to the Faculty of Sports,

University of Porto to obtain the 2nd cycle in Physical

Activity for Elderly, under the decree-law no. 74/2006

of 24 March

Supervisors. Professor Doutor António Ascensão

Professor Doutor José Magalhães

Diogo Nuno Mariani Felix

Porto, June 2013

Mariani, D. (2013). Effects of Voluntary Physical Activity and Endurance

Training on Cardiac Mitochondrial Function of Rats Sub-Chronically Treated

with Doxorubicin. Porto: D. Mariani. Master thesis presented to the Faculty of

Sport, University of Porto.

KEY-WORDS: EXERCISE; HEART; BIOENERGETICS; MITOCHONDRIAL

FUNCTION; DOXORUBICIN.

FUNDING

The present work was supported by a research grant from the FCT

(PTDC/DTP/DES/1071/2012 – FCOMP-01-0124-FEDER-028617) and from

IJUP (PP_IJUP2011 253) to António Ascensão; from Research Centre In

Physical Activity, Health And Leisure (CIAFEL) I&D UNIT (PEST-

OE/SAU/UI0617/2011).

DEDICATÓRIA

Ao papá, à mamã, ao Gonçalo e à Ely.

AGRADECIMENTOS

Agora que finalizo o mestrado, gostaria de expressar os meus profundos e

sinceros agradecimentos:

Aos Professores José Magalhães e António Ascensão, pela disponibilidade

sempre demonstrada na orientação deste estudo, pelos sábios ensinamentos

que me transmitiram, pelo incentivo, por estarem sempre presentes, pela

paciência, amabilidade e pelos momentos de carinho e amizade sempre

demonstrados.

Ao Centro de Investigação em Atividade Física, Saúde e Lazer (CIAFEL) por

todo o apoio e colaboração prestados na realização deste estudo.

A todo o grupo de trabalho em especial à Inês, à Estela e à Mané, que me

acompanharam em todos os momentos do protocolo experimental deste

estudo, pela disponibilidade, pelo incentivo e pela amizade. Inês, obrigada por

todas as explicações, pela paciência e por estares sempre disponível!

E a todas as pessoas e amigos que direta ou indiretamente contribuíram para a

concretização deste trabalho.

ix

Table of Contents  1.   Introduction .................................................................................................................... 1  

2. State of art ........................................................................................................................ 3  

2.1- Age effects on cardiac function ................................................................................. 3  

2.1.1 Aging effects on cardiac mitochondria function ....................................................... 8  

2.2. Exercise and cardioprotection ................................................................................. 16  

2.2.1 Exercise and cardiac mitochondrial adaptations ................................................... 18  

2.2.1.1. Morphological and biochemical adaptations ...................................................... 18  

2.2.1.2. Mitochondrial biogenesis ................................................................................... 20  

2.2.1.3. Oxidative stress and antioxidant capacity ......................................................... 21  

2.2.1.4. Cell death pathways .......................................................................................... 24  

2.3. Doxorubicin: therapeutic agent vs. cardiotoxicity .................................................... 25  

2.3.1 Cardiac mitochondrial toxicity induced by DOX ..................................................... 27  

2.3.1.1 Morphological evidences .................................................................................... 28  

2.3.1.2 Increased oxidative stress .................................................................................. 29  

2.3.1.3 Increased susceptibility to apoptosis .................................................................. 30  

2.4. Exercise as a therapeutic and preventive strategy against DOX-induced

cardiotoxicity. .................................................................................................................. 31  

2.4.1 Acute exercise ....................................................................................................... 31  

2.4.2 Chronic exercise .................................................................................................... 32  

3. Aim ................................................................................................................................. 37  

4. Materials and methods ................................................................................................... 39  

4.1 Reagents .................................................................................................................. 39  

4.2 Animals ..................................................................................................................... 39  

4.3 Exercise protocols .................................................................................................... 40  

4.3.1 Endurance training protocol ................................................................................... 40  

x

4.3.2 Voluntary physical activity ..................................................................................... 41  

4.4 Doxorubicin treatment .............................................................................................. 41  

4.5 Animal sacrifice, heart and soleus extraction ........................................................... 41  

4.6 Isolation of heart mitochondria ................................................................................. 42  

4.7 Mitochondrial respiratory activity .............................................................................. 43  

4.8 Mitochondrial transmembrane electric potential ....................................................... 43  

4.9 Mitochondrial osmotic swelling during MPTP induction ............................................ 44  

4.10 Mitochondrial oxidative damage ............................................................................. 44  

4.11 Soleus citrate synthase activity .............................................................................. 45  

4.12 Statistical analysis .................................................................................................. 45  

5. Results ........................................................................................................................... 47  

5.1. Characterization of animals and exercise protocols ................................................ 47  

5.2 Heart mitochondrial oxygen consumption ................................................................ 50  

5.3 Heart mitochondrial transmembrane electric potential ............................................. 51  

5.4 Mitochondrial osmotic swelling during MPTP induction ............................................ 52  

5.5 Oxidative stress markers .......................................................................................... 54  

6. Discussion ...................................................................................................................... 55  

6.1 Heart mitochondrial oxygen consumption and transmembrane electric potential .... 56  

6.2. Mitochondrial osmotic swelling during MPTP induction ........................................... 58  

6.3 Oxidative stress markers .......................................................................................... 60  

6.4 Meaning for exercise-induced cardioprotection in aging .......................................... 61  

7. Conclusion ..................................................................................................................... 63  

8. References ..................................................................................................................... 65  

xi

Figures

Figure 1. Representative scheme of reduced cardiorespiratory fitness (VO2max) in old

adults. (adapted from Oxenham and Sharpe (2003)). ......................................................... 6  

Figure 2. Mechanisms of DOX action and toxicity (adapted from Carvalho, Santos et al.

2009) .................................................................................................................................. 26  

Figure 3. Effect of exercise and DOX treatment on (A) body mass over time and (B)

distance covered per day by TM and FW groups during the 12 wks of protocol. .............. 47  

Figure 4. Effect of exercise and DOX treatment on (A) state 3 of heart mitochondrial

respiration, (B) state 4 of heart mitochondria respiration, (C) RCR and (D) ADP/O.. ........ 51  

Figure 5. Effect of exercise and DOX treatment on heart mitochondria of heart ∆ψ

fluctuations (A) maximal energization, (B) ADP-induced depolarization, (C) repolarization

and (D) ADP phosphorylation lag phase. ........................................................................... 52  

Figure 6. Effect of exercise and DOX treatment on heart mitochondria to Ca2+-induced

MPTP (A) Swelling amplitude; (B) Average swelling rate. ................................................. 53  

Figure 7. Heart mitochondrial (A) MDA and (B) reduced sulfhydryl contents.. .................. 54  

Tables

Table 1. Effects of aging on cardiovascular system ............................................................. 4  

Table 2. Summary of some described mitochondrial-related alterations associated with

DOX-induced cardiotoxicity and the modulation effect afforded by physical exercise

against DOX (adapted from Ascensao, Oliveira et al. 2012) ............................................. 34  

Table 3. TM exercise protocol ............................................................................................ 40  

Table 4. Animal data and yield of mitochondrial protein isolation ...................................... 49  

Equations

Equation 1: VO2max=Q×(A-VO2)diff .................................................................................... 5  

xiii

Resumo

O presente estudo teve como objetivo analisar o efeito de dois protocolos de

exercício crónico distintos (treino em tapete rolante - TM e atividade física

voluntária em roda livre - FW) na disfunção mitocondrial induzida pelo

tratamento sub-crónico de doxorrubicina (DOX), uma potente droga

antineoplásica bastante eficaz cuja principal limitação é a toxicidade cardíaca.

Foram utilizados 32 ratos Sprague-Dawley macho jovens divididos em seis

grupos (n = 6 por grupo): salino sedentário (SAL + SED), salino treinado (SAL +

TM, 12 semanas de treino em tapete rolante), salino roda-livre (SAL + FW 12

semanas de atividade física em roda livre), tratado com DOX sedentário (DOX

+ SED [7 semanas de tratamento sub-crónico de DOX (2mg.kg-1.wk-1)], DOX +

TM e DOX + FW. Foi analisada a funcionalidade mitocondrial cardíaca in vitro

[consumo de oxigênio, potencial transmembranar (ΔΨ) e swelling osmótico],

assim como os níveis de MDA e grupos sulfidril.

O tratamento com DOX afetou a funcionalidade mitocondrial cardíaca,

alterando o consumo de oxigénio, o potencial transmembranar assim como o

swelling osmótico durante a indução do poro de permeabilidade transitória

mitocondrial (DOX + SED vs SAL + SED). A disfunção induzida pela

administração de DOX no estado 3, no índice de controlo respiratório, ADP/O,

no ΔΨ máximo, na repolarização, na lag-phase, amplitude e taxa de swelling,

assim como no nível de MDA e no conteúdo grupos sulfidril foram revertidos

pelos dois tipos de exercício crónico.

Ambos os protocolos de exercício estudados atenuaram a disfunção

bioenergética das mitocôndrias cardíacas associada ao tratamento sub-crónico

com DOX. Os nossos resultados são mais um contributo para o estudo dos

efeitos cardioprotetores do exercício físico realizado antes, durante e após o

tratamento com DOX, não só na população adulta mas também idosa.

PALAVRAS-CHAVE: EXERCÍCIO, CORAÇÃO, BIOENERGÉTICA,

FUNCIONALIDADE MITOCONDRIAL, DOXORRUBICINA.

xv

Abstract

The effects of two distinct chronic exercise models (endurance treadmill training

– TM and voluntary free-wheel activity - FW) against mitochondrial dysfunction

induced by sub-chronic treatment of doxorubicin (DOX), a potent antineoplastic

drug known to induce a dose-related cardiac and mitochondrial toxicity, were

analyzed.

Male young Sprague-Dawley rats were divided in six groups (n=6 per group):

saline sedentary (SAL+SED), saline exercised (SAL+TM; 12-wks treadmill),

saline freewheel (SAL+FW, 12-wks voluntary free-wheel), DOX+SED [7-wks

sub-chronic DOX treatment (2mg.kg-1.wk-1)], DOX+TM and DOX+FW. In vitro

endpoints of heart mitochondrial function [oxygen consumption, membrane

potential (ΔΨ) and osmotic swelling], MDA level and sulfhydryl groups content

were evaluated.

DOX affected mitochondrial function as seen by oxygen consumption, ΔΨ

endpoints and osmotic swelling during MPTP induction (DOX+SED vs.

SAL+SED). DOX-induced impairments in state 3, respiratory control ratio,

ADP/O, maximal ΔΨ, repolarization, ADP lag-phase, swelling amplitude and

average swelling rate as well MDA level and sulfhydryl groups content were

reverted by both TM and FW.

Both studied chronic models of physical exercise reestablished heart

mitochondrial bioenergetic defects induced by sub-chronic DOX treatment. Our

results contribute to the analysis of the cardioprotective effects of exercise

performed before, during and after DOX treatment no only on adult but also in

older population.

KEY-WORDS: EXERCISE; HEART; BIOENERGETIC; MITOCHONDRIAL

FUNCTION; DOXORUBICIN.

xvii

Abbreviations and Symbols

VO2max Maximal Oxygen Uptake

A.M. Before Midday

AIF Apoptosis Inducing Factor

AMPK Adenosine Monophosphate-Activated Protein Kinase

ANT Adenosine Nucleotide Translocase

ATP Adenosine Triphosphate

Ca2+ Calcium Ion

CAT Catalase

CHF Congestive Heart Failure

CS Citrate Synthase

Cyc D Cyclophilin D

DNA Deoxyribonucleic Acid

DOX Doxorubicin

DTNB Beta Dystrobrevin

E Exercise

ER Endoplasmic Reticulum

FW Free Wheel

G Glutamate

GPX Glutathione Peroxidase

GSH Glutathione

h Hours

H2O2 Hydrogen Peroxide

HSP Heat Shock Proteins

IFM Intermyofibrillar Mitochondria

IR Ischemia Reperfusion

KCl Potassium Chloride

Kg Kilogram

KH2PO4 Monopotassium Phosphate

m Metro

M Malate

xviii

MDA Malondialdehyde

MFN Mitofusin

mg Milligram

min Minute

ml Milliliter

mM Millimolar

MnSOD Manganese Superoxide Dismutase

MPTP Mitocohndrial Permeability Transition Pore

mtDNA Mitochondrial Deoxyribonucleic Acid

MΩ Megaohm

NaCl Sodium Chloride

NADH Reduced Nicotinamide Adenine Dinucleotide

NADPH Reduced Nicotinamide Adenine Dinucleotide Phosphate

nmol Nanomol

NO Nitric Oxide

NS Non-Significant

O2 Oxygen

O2- Superoxide Radical

ºC Degree Celsius

OH. Hydroxyl Radical

PGC Proliferator-Activated Receptor Gamma

RCR Respiratory Control Ratio

RNA Ribonucleic Acid

ROS Reactive Oxygen Species

SAL Saline

SEM Standard Error Of The Mean

-SH Sulfhydryl groups

SOD Superoxide Dismutase

SSM Subsarcolemmal Mitochondria

T Treatment

t Time

TM Treadmill

TNF Tumor Necrosis Factor

xix

TPP+ Tetraphenylphosphonium

VDAC Voltage Dependent Anion Channel

(A-VO2)diff Arteriovenous oxygen difference

Δᴪ Transmembrane Electrical Potential

1

1. Introduction

Doxorubicin (DOX, or adriamycin) is a highly effective antibiotic used to treat

several types of malignancies. Unfortunately, the clinical use of DOX is limited

by the occurrence of a dose-related cardiac toxicity that results in life-

threatening cardiomyopathy. DOX-induced cardiomyocyte dysfunction is

associated with increased levels of oxidative damage involving mitochondrial

bioenergetics collapse in the process (Wallace 2007). Actually, sub-chronic

DOX treated rats reveal defects on heart mitochondrial function, which are

accompanied by compromised mitochondrial electron transport chain activity

and increased oxidative stress and damage (Berthiaume, Oliveira et al. 2005,

Santos, Moreno et al. 2002).

Among the strategies advised to counteract the cardiac side effects associated

with DOX treatment, physical exercise has been studied and recommended as

a non-pharmacological tool against myocardial injury (Ascensao, Ferreira et al.

2007, Ascensao, Lumini-Oliveira et al. 2011, Ascensao, Oliveira et al. 2012).

Previous work has suggested that the advantage of both acute (Ascensao,

Lumini-Oliveira et al. 2010, Wonders, Hydock et al. 2008) and chronic exercise

models (Ascensao, Ferreira et al. 2006, Ascensao, Magalhaes et al. 2005,

Ascensao, Magalhaes et al. 2005, Chicco, Hydock et al. 2006, Chicco,

Schneider et al. 2005, 2006) on the preconditioning of DOX-treated rats include

the protection of cardiac tissue and mitochondria against induced impairments.

Moreover, recent studies investigating the effects of exercise performed during

and following models of late-onset cardiotoxicity caused by DOX provide

evidence of exercise-induced cardioprotection in both adult and juvenile rat

models (Hayward, Lien et al. 2012, Hydock, Lien et al. 2012). However, the

cellular and molecular mechanisms underlying this protective phenotype

induced by exercise are still elusive. In particular, whether perturbations in heart

mitochondrial oxidative phosphorylation capacity and pro-oxidant redox

modifications associated with cumulative DOX administration are modulated by

long-term physical exercise performed during and after treatments is yet

unknown.

2

We therefore aimed to analyze the effects of two types of long-term exercise

with distinct characteristics, performed before and during the overall DOX

treatment, on cardiac mitochondrial bioenergetics. Heart mitochondrial

respiratory parameters associated with oxygen consumption, transmembrane

electrical potential and osmotic swelling during mitochondrial permeability

transition pore (MPTP) induction as well as markers of oxidative stress

(sulfhydryl groups (-SH) and malondialdehyde (MDA) contents) were

determined.

3

2. State of art

2.1- Age effects on cardiac function

Aging can be characterized as a time dependent decline of maximal

functionality that affects tissues and organs of the whole body (Figueiredo, Mota

et al. 2008). The average human life span has markedly increased in modern

society, a fact largely attributed to advances in medical and therapeutic

sciences that have successfully reduced the severity of several diseased

conditions (Chaudhary, El-Sikhry et al. 2011). However, elderly individuals

continue to suffer the greatest burden from cardiovascular disease, including

coronary heart disease that remains the leading cause of death in industrialized

countries (Wei 2004). That can be understood as the result of several

morphophysiological, structural and functional alterations, which we will further

address in detail (Table 1).

The aging-induced increase on vascular stiffness, septum and myocardium

thickness and fibrosis, is associated, among others, with: i) decreased myocyte

number; ii) increasing cardiomyocyte size with alterations in calcium (Ca2+)

homeostasis and iii) increased collagen fibers deposition, leading to cardiac

diastolic dysfunction, increased afterload, and loss of arterial and heart

compliance (Strait and Lakatta 2012). Importantly, these physiological-related

impairments along with unchanged cavity size and increased left ventricle

hypertrophy are the major characteristic of aging heart (Chaudhary, El-Sikhry et

al. 2011). In fact, Dai et al (2009) reported that aging left ventricle mass index

increased by around 75% compared to a young adult group, indicating the

increase prevalence of left ventricular pathological hypertrophy with age. It was

also reported reduction in diastolic function, as well as worsening of myocardial

performance index (Dai, Santana et al. 2009).

Furthermore, the conduction system also undergo some structural alterations

leading to heart dysfunction, including fat accumulation around sinoatrial node,

which creates a partial or complete separation of the node from atrial tissue

(Strait and Lakatta 2012), a marked decreased in peacemakers number cells

4

(Wei 2004) and an increased calcification in conduction system leading

ultimately to atrioventricular conduction lock. Slow and prolonged systole and

diastole are other aging-related heart features usually related to an aberrant

Ca2+ handling (Strait and Lakatta 2012). The decreased diastolic function, which

falls linearly with aging at a rate of about 6–7% per decade (Gates, Tanaka et

al. 2003), is associated to a decrease in diastolic filling reported at rest and

under stress conditions, as some daily tasks that promote an acute increase of

physical demands. In addition, the degeneration of cardiac valvular apparatus is

commonly reported, being valvular annular dilation found in the majority of older

persons and is also associated with concomitant coronary artery calcification

(Wei 2004).

Table 1. Effects of aging on cardiovascular system Morphophysiological

changes Structural Changes Functional Changes Reference

Heart and Vascular system

↓ Cardiomyocytes number

↑ Cardiomyocytes size

↑ Collagen deposition

↑ Fibers deposition

↓ Cardiomyocytes function

↑ Septum and Myocardium stiffness

↓ Heart compliance

↑ Vascular thickness

↓ Vascular compliance

↑ Systolic pressure

↓ Diastolic function

↓ max heart rate

↓ max cardiac output

↑ Afterload

Prolonged systole and diastole

Hotta and Uchida (2010), Reynolds (2004), Strait and Lakatta (2012), T.

Shioi (2012), Taylor, Cable et al.

(2004)

Conduction system

↑ Fibrosis and fat accumulation (SA node)

↓ Peacemakers cells number

↑ Conduction system calcification

↓ Ventricular compliance

↑ Heart rhythm disturbances

↑ Risk atrioventricular conduction lock

↑ Risk atrial arrhythmias,

↑ Risk atrial fibrillation

↑ Risk tachycardia

↑ Ventricular arrhythmias

Hotta and Uchida (2010), Strait and Lakatta (2012)

Valvular system

↑ Valvular annular dilatation Coronary artery calcification Reynolds (2004)

(↑) – increase; (↓) – decrease

↓ VO2 max

5

Overall, these aging related heart morphophysiological, structural and functional

alterations result in increased systolic pressure, one major risk factors for

development of atherosclerosis, hypertension and stroke, and arterial fibrillation

(North and Sinclair 2012).

As a consequence of all those age-associated alterations, cardiac function

declines maximal functionality, being that, under basal conditions this decline

may not be detected (Wessells and Bodmer 2007).

The most used standard index of cardiorespiratory fitness is the maximal

oxygen consumption (VO2max), which declines approximately 10% per decade

starting at 20 to 30 years old (Souza 2012). VO2max can be estimated by the

Fick’s equation:

Equation 1: VO2max=Q×(A-VO2)diff

Q is cardiac output, and is the product of max heart rate and stroke volume, (A-

VO2)diff is the arteriovenous difference of oxygen. As it is observed in figure 1,

based on Fick´s equation, aerobic exercise capacity depends on cardiac output

and arteriovenous difference (Seeley, Stephens et al. 2005).

Reductions in peak heart rate, peripheral oxygen utilization and stroke volume

appear to mediate the age-associated decline in VO2max. Actually, impairments

in cardiac filling and increased afterload leads to a decrease in heart rate, in a

stress situation (Taylor, Cable et al. 2004) being the major responsible for much

of the age-associated decrease in maximal cardiac output. Reductions in

muscle oxygen delivery and therefore (A-VO2)diff are mainly due to reduced

and maldistribution of cardiac output. Also, a decline in skeletal muscle

oxidative capacity with aging is reported, due in part to mitochondrial

dysfunction, which appears to play a particularly important role in old age,

where skeletal muscle VO2max is observed to decline by approximately 50%

even under conditions of similar oxygen delivery as young adult muscle (Betik

and Hepple 2008).

6

Although controversial, some reports suggested that left ventricular systolic

function remains relatively preserved and without significant alterations in left

ventricular stroke volume and cardiac output at rest (for refs see Kappagoda

and Amsterdam 2012, Lakatta 2002, Morley and Reese 1989). However, at

peak exercise, stroke volume index is reduced in older individuals and is

thought to be the consequence of age-related reductions in β-adrenergic

stimulation, increases in vascular stiffness, aortic impedance and impaired left

ventricular diastolic function (Spina, Turner et al. 1998).

Generally, the morphophysiological and functional alterations related to

VO2max decline with aging are depicted in figure 1.

Figure 1. Representative scheme of reduced cardiorespiratory fitness (VO2max) in old adults. (adapted from Oxenham and Sharpe (2003)).

Studies have demonstrated that advancing age is also associated with

decreased heart rate variability (O'Brien, O'Hare et al. 1986, Yeragani,

7

Sobolewski et al. 1997). Heart rate variability is a reliable reflection of the many

physiological factors modulating the normal rhythm of the heart. In fact, it

provides a powerful mean of observing the interplay between the sympathetic

and parasympathetic nervous systems and it is thought to reflect the heart’s

ability to adapt to changing circumstances by detecting and quickly responding

to unpredictable stimuli (Rajendra Acharya, Paul Joseph et al. 2006). It is

particularly relevant on daily tasks, as it is important to maintain a normal

lifestyle without manifestation of heart injury. Also, it has been suggested that a

decreased responsiveness to β-adrenergic receptor in aged hearts might be

responsible for decreased heart rate variability in aged people during exercise

or stress (Chaudhary, El-Sikhry et al. 2011).

Ischemia-reperfusion (IR) injury is the primary pathological manifestation of

coronary artery disease (Lennon, Quindry et al. 2004, Powers, Demirel et al.

1998). The level of IR-induced myocardial injury can range from a small insult,

resulting in limited damage, to a large insult, culminating in major cardiac

electrical and mechanical dysfunction and ultimately in death (Powers, Quindry

et al. 2004). Nowadays, it is well established that numerous age-related cellular

and functional changes occur in the heart that could lead to ischemia-

reperfusion injury as i) alterations in cardiac gene expression; ii) increased

oxidative stress and iii) reduced ability of the heart to response to stress

(Powers, Quindry et al. 2004). In this regard, Starnes et al. (1997) reported that

immature hearts tolerate and recover from hypoxia better than adult hearts, and

that the sarcolemmal membranes of immature rat hearts seem to be less

susceptible to damage from hypoxic stress than those of older group.

It is notable that throughout life, there are many impairments originating cardiac

injuries and ultimately death. So, it is important to implement countermeasures

to attenuate and minimize those consequences, which will enhance old people

lifestyle and lifespan. Recently, several approaches have been investigated and

physical activity has been shown to be an important countermeasure to protect

against myocardial injuries (Ascensao, Ferreira et al. 2007, Kavazis 2009,

8

Kavazis, Alvarez et al. 2009, Powers, Quindry et al. 2004, Starnes and Taylor

2007).

Furthermore, among the several mechanisms or causes for cardiac dysfunction,

mitochondrial abnormalities have a central role (for refs see Braunwald and

Bristow 2000). In fact, due to the key mechanisms to which these organelles are

associated such as energy production, ion regulation, pH control, Ca2+

homeostasis, redox reactions, control of cell signaling and apoptosis

mitochondria assume a pivotal role in cellular functioning. For these reasons,

mitochondria have been suggested as reliable sensors of cellular functionality

and (dys)functional mitochondria correlates with (dys)functional heart tissue

submitted to a variety of stimuli, including age (Wallace 2010).

Because heart is primarily a postmitotic tissue that exhibits a highly aerobic

metabolism due to the abundance of large mitochondria, a dependence on

healthy mitochondria for normal organ function is implicated (Judge and

Leeuwenburgh 2007).

The following section will address the effects of aging process on the

modulation of mitochondrial function and related mechanisms.

2.1.1 Aging effects on cardiac mitochondria function

In metabolically active and thus energy-demanding tissues such as the heart,

mitochondria, as producers of most of the ATP necessary for metabolism via

oxidative phosphorylation, have a very important role in the supply of energy for

continuously contracting myocytes (Ascensão 2011). Additionally, mitochondria

play other important roles as cellular redox and ion homeostasis, oxygen

sensing, signaling and regulation of programmed cell death (for refs see

Rabinovitch 2012, Sheu 2009). Therefore, mitochondrial integrity is vital for

cellular homeostasis and cardiac performance (Chaudhary, El-Sikhry et al.

2011) being age-related heart mitochondrial dysfunction closely associated to

9

life limiting impairments, including congestive heart failure (CHF) (Chaudhary,

El-Sikhry et al. 2011, Judge and Leeuwenburgh 2007, Pashkow 2011).

In 1956, Harman et. al, proposed the free radical theory of aging, postulating

that the production of reactive oxygen species (ROS) is a major determinant of

lifespan acting as important mediators responsible for the cellular damage seen

in aged cells (Harman 1956). Later, it was defined that mitochondria are the

main source of ROS and the main target of their injury being ROS produced

within mitochondria almost 90% of the total ROS produced in the cell (Hearman

1972); so it was postulated as the “key organelles” initiating cellular processes

leading to death (Chaudhary, El-Sikhry et al. 2011). Indeed, evidences suggest

that with advanced age, mitochondrial production of ROS significantly increases

in heart tissue (Judge, Jang et al. 2005), which leads to development of

degenerative diseases (for refs see Rabinovitch 2012). Within cells, ROS are

produced in multiple compartments and by multiple enzymes including reduced

nicotinamide adenine dinucleotide phosphate (NADPH) oxidase at the plasma

membrane, respiratory chain within mitochondria, lipid oxidation within

peroxisomes and by clyclo-oxygenases and xanthine oxidase in the cytoplasm

(Dai and Rabinovitch 2009, Dai, Rabinovitch et al. 2012). Although all of these

source contribute to the overall oxidative damage, mitochondria are one of the

most contributors for ROS generation as a byproduct of electron transfer during

oxidative phosphorylation (Dai and Rabinovitch 2009). Most specifically, excess

electrons from complex I and III can be transferred directly to oxygen (O2) to

generate superoxide anion (O2-), which is then converted to hydrogen peroxide

(H2O2) and ultimately into a hydroxyl radical (OH.), the most reactive ROS

species (Dai and Rabinovitch 2009). The unbalance between ROS production

and the capacity of antioxidant machinery, favoring ROS production is usually

called oxidative stress. Mitochondria have antioxidant enzymes that play a vital

role in the protection against oxidative stress (Meng 2007), being manganese

superoxide dismutase (MnSOD), catalase (CAT) and glutathione peroxidase

(GPX) the most commonly cited (Ascensão 2003). In addition, there are non-

enzymatic antioxidant compounds, both endogenous and exogenous, such as

glutathione (GSH), vitamins C and E, and lipoic acid that play important roles in

10

ROS neutralization or in the attenuation of the effects caused by increased ROS

production (Ji, Leeuwenburgh et al. 1998). Heat shock proteins (HSPs) are

another system of cellular defense against oxidative stress (Ascensao, Ferreira

et al. 2007, Hamilton, Staib et al. 2003, Powers, Locke et al. 2001). These

“stress-induced proteins” are ubiquitous and highly conserved chaperones,

important in the folding of new synthesized, damaged or transported proteins.

Moreover, HSPs mediate mitochondrial protection against oxidative stress,

namely HSP70 have been associated with myocardial protection (Gunduz,

Senturk et al. 2004).

To understand the impact of oxidative stress, which is known to damage

proteins, lipids, and deoxyribonucleic acid (DNA) (Shioi and Inuzuka 2012),

some oxidative markers are analyzed in mitochondria as carbonil groups

(protein oxidation), malondialdehyde (MDA) (lipid peroxidation marker) or

thiobarbituric acid reactive substances (TBARS) (nonspecific marker for lipid

peroxidation), among others. The accumulation of oxidant-induced damage in

mitochondria may be a major contributing factor to the age-related alterations in

myocardial function (Chen and Knowlton 2011, Ljubicic, Menzies et al. 2010,

Pohjoismaki, Boettger et al. 2012). Nonetheless, some authors have suggested

that throughout the aging process, the antioxidant capacity can be adjusted in

response to prooxidant exposure (Ji, Leeuwenburgh et al. 1998). Localized

oxidative stress in specific organs, tissues, and organelles may stimulate

cellular uptake and synthesis of certain antioxidants under complicated genetic,

hormonal, and nutritional regulation (Harris 1992). One possibility is that aged

mitochondria produce more ROS stimulating antioxidant enzyme gene

expression. This scenario is consistent with the finding that mitochondrial

antioxidant enzyme activity showed a greater increase in the senescent

myocardium (Ji 1993, Ji, Dillon et al. 1991, Rao, Xia et al. 1990, Vertechy,

Cooper et al. 1989). Nevertheless, data seems to be controversial as no

alteration (Bejma, Ramires et al. 2000, Tian, Cai et al. 1998) or decrease in

antioxidant activity (Bagchi, Bagchi et al. 1996, Pritsos and Ma 2000) have also

been reported. So, increased antioxidant activity may be interpreted as being

beneficial because it provides better protection against oxidant-induced

11

damage, or it may be viewed as negative because it could indicate a need for

enhanced antioxidant defenses due to increased oxidant production (Beckman

and Ames 1998).

However, characterization of age-related changes in cardiac mitochondria has

been challenged due to the fact there are two distinct populations of

mitochondria in the myocardium (Fannin, Lesnefsky et al. 1999).

Subsarcolemmal mitochondria (SSM) are located beneath the plasma

membrane, whereas interfibrillar mitochondria (IFM) are found in parallel rows

between the myofibrils. There are important biochemical and functional

differences between SSM and IFM and it has been revealed that IFM are more

adversely affected with age (Fannin, Lesnefsky et al. 1999, Judge, Jang et al.

2005), although Judge et al. (2007) suggests that further studies are required to

determine the mechanisms contributing to these changes, and to further

characterize differential effects of age upon the SSM and IFM populations in the

heart.

Despite this fact, some authors report increased H2O2 production from SSM, but

not IFM, with age (Judge, Jang et al. 2005, Judge and Leeuwenburgh 2007).

This can be a consequence of the lower antioxidant capacity on this

subpopulation. Actually, whereas IFM MnSOD, GPX and CAT levels seem to

increase with age, in SSM only MnSOD and GPX, but not with CAT, seems to

be positively regulated. However, as MnSOD converts O2- radicals to H2O2,

increased MnSOD activity might also represent higher levels of H2O2. Also,

lower glutathione levels and increased oxidative damage have been suggested,

supporting that oxidant production within the matrix of “old” IFM is greater than

that in “young” IFM (Judge, Jang et al. 2005). Similarly, state 3 respiratory rates

were also lower in “old” IFM. This has a particular concern because IFM are

likely the major source of ATP production for myosin ATPase (Judge, Jang et

al. 2005). Because high levels of ATP are required for both systolic contraction

and diastolic relaxation, reduced availability of ATP as a result of IFM

dysfunction could contribute to the alterations in cardiac contractility observed

with age (Judge, Jang et al. 2005).

12

Besides nucleus, mitochondria are the only organelles in animal cells that

possess their own DNA, the mtDNA, as well as related transcriptional and

translational synthesis machinery. mtDNA is localized in matrix with physical

proximity to the mitochondrial respiratory chain (Bratic and Trifunovic 2010).

However, mitochondria cannot be synthesized de novo, instead they replicate in

the cytosolic compartment through a process of division (for refs see

Chaudhary, El-Sikhry et al. 2011). Importantly, the proteins that are encoded by

mtDNA are vital for normal mitochondrial function and mtDNA does not have

the protein protection as nuclear DNA and has less effective repair mechanisms

(Desler, Marcker et al. 2011). Therefore, mutations in mtDNA through oxidative

stress affect the expression and integrity of oxidative phosphorylation

complexes and can cause mitochondrial dysfunction and increased ROS

production (Wallace 2010). So, it is notable the interrelationship between

oxidative phosphorylation complexes, ROS levels, mtDNA mutations and

ultimately cell death. Furthermore, mutations in mtDNA and the resultant

decline in mitochondrial activity observed in aged tissues are responsible for the

increased generation of ROS, which in turn, will further negatively impact

mitochondria causing further mtDNA damage. This “vicious cycle” concept

postulated that accumulation of mtDNA mutations is exponential and associated

with massive increase in ROS production (Lenaz 1998).

Being mitochondria strategic organelles essential for cell function and

homeostasis, providing energy to the cell (Chaudhary, El-Sikhry et al. 2011),

they must undergo some dynamic mechanisms. Mitochondrial turnover and

death can occur via several processes that are suggested to be interrelated

namely apoptosis, necrosis, autophagy and related dynamics of organelles. In

this dynamic network biogenesis, fusion and fission are closely associated

mechanisms (Chen and Knowlton 2011).

In fact, apoptosis is mediated by two pathways: the extrinsic and the intrinsic

pathways, and both have been described in cardiac myocytes (Whelan,

Kaplinskiy et al. 2010). The extrinsic apoptotic pathway can be triggered by Fas

ligand or tumor necrosis factor (TNF)-α, which are expressed in cardiac

13

myocytes and have been implicated in cardiovascular pathology (Whelan,

Kaplinskiy et al. 2010).

Intrinsic pathways involve the participation of endoplasmic reticulum (ER)

and/or mitochondria (Kroemer, Galluzzi et al. 2007). Impairment in

mitochondrial integrity, dynamics or metabolic activity may result in a range of

deleterious effects to the cell, such as reduced ATP production, elevated

cytosolic Ca2+, increased ROS release, release of proapoptotic factors as

cytocrome c or caspases activation and Bax translocation, triggering cell death

(Chen and Knowlton 2011). In mammalian cells, apoptosis is regulated by a

variety of factors that are essentially either pro-life or pro-death (Goldspink,

Burniston et al. 2003). Quantitation of the expression of genes involved in the

apoptotic pathway might represent a good index of the probability for a cell to

undergo apoptosis. As the number of Bax-expressing cells dramatically

increases in left ventricular hypertrophy and left ventricular dysfunction

(Condorelli, Morisco et al. 1999, Green and Reed 1998), it is suggested that

mitochondrion is the primary organelle mediating the intrinsic apoptotic pathway

in these conditions (Chiong, Wang et al. 2011)

Also, if excessive Ca2+ enters to mitochondria and enhanced oxidative stress

conditions are present, a phenomenon known as permeability transition may

occur (Ascensao, Lumini-Oliveira et al. 2011). The mitochondrial permeability

transition is characterized by the loss of the impermeability of the mitochondrial

membranes and it is suggested that this condition is mediated by the formation

and opening of protein complex-like pores in the inner mitochondrial membrane,

the mitochondrial permeability transition pore (MPTP) (for refs see Ascensao,

Lumini-Oliveira et al. 2011). Increased pro-oxidant generation causing oxidative

stress is one condition that augmented the susceptibility for the opening of

these pores and the release of pro-apoptotic proteins within mitochondria as

cytochrome c, SMAC/DIABLO and the apoptosis inducing factor (AIF), which

will activate the caspase-related apoptotic pathways. It is suggested that the

release of these proteins is dependent on the formation and opening of MPTP

that cross the inner and the outer membranes leading to the loss of

14

mitochondrial membrane potential (Δᴪ), increased mitochondrial osmotic

swelling and rupture of the outer mitochondrial membrane, which leads to death

(Ascensao, Lumini-Oliveira et al. 2011). It is believed that the structure and

regulation of this multi-protein complex comprises the outer membrane voltage-

dependent anion channel (VDAC) as well as the inner membrane adenine

nucleotide translocator (ANT) and cyclophilin D (Cyc D). Myocyte loss has been

shown to occur in the aged rat heart and to precede the occurrence of

ventricular dysfunction (Anversa, Hiler et al. 1986), being apoptotic

cardiomyocyte death present under different conditions in humans (Haunstetter

and Izumo 1998, Narula, Haider et al. 1996, Olivetti, Abbi et al. 1997).

Among the excess of biological phenomena affected by aging, the malfunction

and decrease of biogenesis of mitochondrial biogenesis seems to exert some of

the most potent effects on the organism (Lopez-Lluch, Irusta et al. 2008). If

biogenesis is affected, it is reasonable to expect that mitochondrial turnover

must be slower and the accumulation of modified lipids, proteins and DNA must

also increase, further aggravating the conditions resulting on deficient activity of

aged mitochondria (Lopez-Lluch, Irusta et al. 2008). The precise reason for the

decrease in the rate of mitochondrial biogenesis during aging is currently

unknown. However, it seems that both, extra- and intra-cellular regulatory

factors of mitochondrial biogenesis are implicated. Specifically, peroxisome

proliferator-activated receptor gamma coactivator (PGC1-α) has been shown to

act as a common intracellular mediator during mitochondrial biogenesis induced

by hormonal factors (Weitzel, Iwen et al. 2003), and adenosine

monophosphate-activated protein kinase (AMPK) an intracellular regulator of

mitochondrial biogenesis, which activity appears to be one of the main factors

associated with deficient mitochondrial biogenesis (Reznick, Zong et al. 2007).

PGC family members have gained particular interest because of their ability to

drive virtually all mechanisms of mitochondrial biogenesis in the heart, including

mitochondrial number, mitochondrial respiration, expression of oxidative

phosphorylation and fatty acids oxidation genes, and ROS levels (Lehman,

Barger et al. 2000). Decreased PGC-1α expression has been linked to the

development of heart failure in mouse models (for refs see Moslehi, DePinho et

15

al. 2012), and decline in mitochondrial biogenesis and mitochondrial protein

quality control in cardiac muscle was found in aging (Koltai, Hart et al. 2012)

In addition, its well known that mitochondria are dynamic organelles that

constantly undergo fission and fusion and it has been found to be vibrant

organelles that continuously divide and fuse within the cell and have functions

extending beyond energy production, including cell signaling (Liesa, Palacin et

al. 2009). Disruption of fission and/or fusion can also lead to cellular dysfunction

and to apoptosis. This dynamic mechanism is regulated by proteins controlling

fission, such as hFis1 and Drp1, and fusion, such as mitofusin 1 and 2 (MFN1

and MFN2) and OPA1. The correct function of these proteins seems to be

critical for normal mitochondrial activity, and their deregulation is associated

with several pathologic conditions (Lopez-Lluch, Irusta et al. 2008). Indeed, the

impairments on fission-related protein hFis1 has been associated with the

process of senescence in mammalian cell cultures (Lee, Jeong et al. 2007).

Moreover, depletion of hFis1 by RNA interference (RNAi) induces dramatic

changes in mitochondrial structure, including the enlargement and flattening of

the organelle. Futrthermore, elimination of any of the mitochondrial fusion

proteins as MFN1, MFN2 or OPA1, induces mitochondrial fragmentation, as

expected, being that down-regulation of Opa1 expression in cells by RNAi

results in spontaneous apoptosis (for refs see Chen and Knowlton 2011).

Overall, defects in the mitochondrial fission/fusion machinery and so loss of the

symmetry between fusion and fission (Hoppins, Edlich et al. 2011) may

contribute to the decline in mitochondrial function during aging. However,

several fundamental questions remain to be answered (Bossy-Wetzel, Barsoum

et al. 2003).

In the next section, the roles of physical exercise as a strategy to improve

cardiac function in adult and old subjects as well as the mitochondrial-mediated

mechanisms associated with exercise-induced cardioprotection will be

addressed.

16

2.2. Exercise and cardioprotection

Cardiac damage is a major contributor to morbidity and mortality in

industrialized countries; so it becomes important to develop strategies that

result in cardioprotective phenotype. In this regard, several approaches have

been investigated and physical activity has been shown to be an important

countermeasure to protect against myocardial injuries (Bowles, Farrar et al.

1992, Harris and Starnes 2001, Powers, Demirel et al. 1998, Powers, Quindry

et al. 2004). In fact, cardiorespiratory fitness is inversely related to

cardiovascular and all-cause mortality and it has crucial role preventing heart

injury (Kokkinos, Myers et al. 2010). Some reports had postulated exercise-

induced benefits based on decreases of some risk factors to develop cardiac

and myocardium impairments such as bodymass index, body weight, waist

circumference, abdominal and visceral fat and consequently insulin resistance,

triglyceride levels, blood pressure and, in general, metabolic syndrome-related

parameters (for refs see Golbidi and Laher 2012). Therefore, chronic aerobic

exercise is able, not only, to improve cardiovascular function in young healthy

subjects, but also, and most importantly, in older people and those with

cardiovascular risk factors (Hambrecht, Fiehn et al. 1998).

The study of the mechanisms responsible for exercise-induced cardioprotection

has been ongoing for over decades and morphological and

biochemical/molecular alterations have been considered as putative

mechanisms of exercise-induced cardioprotection. Those include morphological

adaptations of heart and coronary arteries, induction of myocardial HSPs,

increased myocardial cyclooxygenase-2 activity, elevated ER stress proteins,

nitric oxide production, improved function of sarcolemmal and/or mitochondrial

adenosine triphosphate (ATP)-sensitive potassium channels and increased

myocardial antioxidant capacity (for refs see Kavazis 2009).

For instance, exercise induces vascular remodeling and so, morphological

alterations in coronary arteries through angiogenesis and arteriogenesis

(Leung, Yung et al. 2008). Here, nitric oxide (NO) assumes important roles due

its anti-inflammatory, vasodilator and platelet inhibitory effects (Landmesser and

17

Drexler 2005). Also, NO protects against inschemia-reprefusion (IR) injury in

such a way that the heart responds to ischemia using nitric oxide species in a

harmonized manner and these mechanisms could be based on inhibition of

Ca2+ influx into myocytes, antagonism of β-adrenegic stimulation, reduction in

cardiac oxygen consumption and ability to increase the expression of HSP70

(for refs see Golbidi and Laher 2011). HSPs protect cell against oxidative injury

and apoptosis (Polla, Kantengwa et al. 1996) and furthermore, enhance

recovery from acute myocardial cellular injury protecting heart for subsequent

injury (for refs see Powers, Locke et al. 2001) by promoting restoration of

dysfunctional enzymes and preventing aggregation of severely denatured

proteins. The majority of evidences indicate that members of the 70-kDa family

are the cytoprotective proteins most responsible for cell protection (for refs see

Powers, Locke et al. 2001). The expression of HSP70 in cardiomyocytes is

associated with increased cell survival and protection against ischemic damage

and its now well established that acute and chronic exercise are able to induce

increases in the expression of HSP70 (Kregel and Moseley 1996, Powers,

Demirel et al. 1998), although in a temperature-dependent “fashion”.

The improved function of sarcolemmal ATP sensitive potassium channels

(Powers, Quindry et al. 2008) and elevated ER stress proteins are other

important exercise-induced cardioprotection-related alterations and both have

special relevance during a cardiac insult (Golbidi and Laher 2012). The ER

stress proteins help cellular homeostasis by maintaining intracellular Ca2+

regulation and protein folding during IR injury (Logue, Gustafsson et al. 2005).

Considering the importance of mitochondrial machinery in the maintenance of

cardiac function, mitochondrial-mediated mechanisms have also been

associated with exercise induced cardioprotection phenomenon. This topic will

be further discussed in the next sections.

18

2.2.1 Exercise and cardiac mitochondrial adaptations

As previously mentioned, mitochondrial adaptations may play a critical role in

exercise-induced protection against cardiovascular impairments. The

mechanisms behind this phenomenon remain unclear, however it may be

related to morphological and biochemical adaptations including biogenesis,

antioxidant production or resistance to cell death pathways (Ascensao, Ferreira

et al. 2007, Kavazis, Alvarez et al. 2009, Kavazis, McClung et al. 2008). Those

will be briefly addressed in the following section.

2.2.1.1. Morphological and biochemical adaptations

Heart is a highly oxidative organ, with a low rate of cell growth and slow

turnover of proteins; so, it would be expected that heart might have a limited

ability to adapt to acute and/or chronic conditions.

Although there are scarcely addressed outcomes about exercise-induced

cardiac mitochondrial morphological adaptations in healthy hearts, the same

cannot be stated regarding hearts under deleterious conditions. In fact, under

deleterious conditions as DOX treatment, diabetes, or aging, morphological

impairments on heart mitochondria have been reported (Ascensao, Oliveira et

al. 2012, Searls, Smirnova et al. 2004). Despite this fact, in a study where

trained animals were submitted to an endurance swimming training program,

while the non-trained were not engaged in any exercise program Ascensao et

al. (2006) reported that endurance swimming training per se caused notable

changes in myocardial structure seen as an apparent increased glycogen

content, intercalated discs showing a notorious scalloped appearance and

evident signs of mitochondria biogenesis with elevated number of encroached

mitochondria per fiber area, probably resulting in an increased volume/density

of mitochondria. Also, mitochondria division, mild and focal loss of cristae

density and organization as minimal degradation by-products, were also present

in non-treated trained hearts. Another study developed by Searls et al. (2004)

19

with type I diabetes rats found in type I diabetes rats that 9 weeks of moderate

exercise is able to reverse some of the phenotype of diabetic cardiomyopathy.

Specifically, the mitochondrial quality, cytoplasmic area, and collagen cross-

sectional area returned toward non-diabetic values with exercise.

Mitochondrial biochemical adaptations to exercise were also reported and

several approaches have been trying to prove the exercise-induced

modifications of some parameters involved in mitochondrial bioenergetics.

Judge et al. (2005) reported that wheel running had no effect on mitochondrial

protein yield, rates of oxygen consumption (states 4 and 3) or RCR. This was

not entirely surprising given that other studies have shown that, unlike skeletal

muscle, oxidative capacity of cardiac muscle is not increased in response to

treadmill training, an exercise protocol that is typically much more intense than

voluntary wheel running (for refs see Judge, Jang et al. 2005). Also, Starnes et

al. (2007) reported that endurance training has no impact on mitochondrial

oxidative phosphorylation, being that all oxidative phosphorylation parameters

were similar in endurance trained and sedentary rats. Because the heart is

highly oxidative, it is not expected to be as responsive to exercise-induced

increases in oxidative capacity as skeletal muscle. Although, several reports

suggested that exercise per se is able to improve mitochondrial protein yield,

rates of oxygen consumption as well as cardiac function (for refs see Ascensao,

Ferreira et al. 2007, Ascensao, Magalhaes et al. 2006, Chicco, Schneider et al.

2006, Powers, Lennon et al. 2002). Moreover, Kavasis et al. (2009) reported

that the abundance of several proteins involved in bioenergetics was altered

following endurance exercise in both SS and IMF mitochondria. Most

specifically, the protein levels of several proteins involved in β-oxidation of fatty

acids were increased following repeated bouts of endurance exercise. It is of

especial concern as during the development of heart disease or genetic defects

in mitochondrial fatty acid β-oxidation the myocardial energy source switches

from fatty acid β-oxidation to glycolysis. Also, there are some controversial

reports about endurance exercise benefits against Ca2+-induced mitochondrial

dysfunction. In fact, while Starnes et al. (2007) found that endurance exercise

training had no influence when mitochondria was challenged with identical Ca2+

20

concentrations, being that mitochondria from trained and sedentary animals

displayed similar declines in ATP production, French et al. (2008) found that

exercise-induced protection against IR injury, in part, by attenuating IR-induced

oxidative modification of important Ca2+-handling proteins and preventing

subsequent calpain activation. In this study, MnSOD may had an important role

as, the results support that exercise-induced increases in myocardial MnSOD

activity attenuate the oxidation and degradation of Ca2+-handling proteins,

preventing calpain activation during IR. It has special concern because calpain

activation within the myocardium has been directly linked with IR-induced

necrotic and apoptotic cardiac myocyte death (French, Hamilton et al. 2008).

Furthermore, several reports suggest that exercise per se is able to enhance

cardiac mitochondria Ca2+ uptake capacity without MPTP induction (Ascensao,

Lumini-Oliveira et al. 2011, Kavazis, McClung et al. 2008, Marcil, Bourduas et

al. 2006). Therefore, exercise may induce beneficial morphological and

biochemical adaptations in mitochondria, which can be translated in a cardiac

phenotype more functional and protected against deleterious stimuli.

2.2.1.2. Mitochondrial biogenesis

Due to the high-energy demand of the heart, a decline in mitochondrial function

can result in a deterioration of cardiac performance (Li, Muhlfeld et al. 2011)

and can lead to a wide variety of pathophysiological conditions. According to

some reports, the malfunction of mitochondria and the decrease of

mitochondrial biogenesis, together with increased oxidative damage, seem to

exert some of the most deleterious effects on the organism (Guarente 2008,

Lopez-Lluch, Irusta et al. 2008). Fortunately, although controversial, heart

seems to have the ability to alter some gene expression profile and phenotype

to produce new and more functional mitochondria (Lee and Wei 2007). Regular

exercise or increased energy demand are important stimuli that lead to

increased mitochondria biogenesis (for refs see Lopez-Lluch, Irusta et al. 2008).

In fact, increases in cytosolic Ca2+ levels induced by exercise stimulates

21

calmodulin kinase which then promotes PGC1-α expression (Wu, Kanatous et

al. 2002) that has been shown to be a major regulator of mitochondrial

biogenesis (Rodgers, Lerin et al. 2005). Also, Geng et al. (2010) showed that in

skeletal muscle-specific PGC-1α knockout mice, exercise-induced mitochondrial

biogenesis and angiogenesis were significantly attenuated. However, exercise-

induced mitochondrial biogenesis on cardiac muscle is still not fully illusive. In

fact, although a significant amount of experimental data on mechanisms

involved in exercise-induced mitochondrial biogenesis has been obtained in

skeletal muscle, less information is available about the heart. In this regard, Li

et al. (2011) reported no changes in mitochondrial biogenesis on left ventricle

after 3 months of endurance exercise. Actually, in contrast to skeletal muscle in

which adaptive changes in volume fraction of mitochondria can readily occur,

this is not so frequent in cardiac muscle (Hood, Balaban et al. 1994). So, the

exercise-induced benefits on heart mitochondria biogenesis are still

controversial.

2.2.1.3. Oxidative stress and antioxidant capacity

With exercise performance, increased O2 consumption creates favorable

conditions for increased generation of ROS, an inevitable consequence that

may increase oxidative stress at the organelle, cell, and tissue (Ji,

Leeuwenburgh et al. 1998). However, if these stimuli are repeated over time, it

may have a strong modulating effect on various defense systems in cardiac

cells (Powers, Lennon et al. 2002). Also, in some deleterious conditions that

lead to ROS production, as IR-induced myocardial injury, DOX administration or

aging, physical exercise as been reported to induce benefic countermeasures.

In fact, IR-induced myocardial injury is manifested due to the complex

interaction of numerous factors but ROS generated by mitochondria during IR

injury are believed to play key role in this process (for refs see Honda, Korge et

al. 2005). During IR, mitochondrial ROS generation can lead to increased

general oxidative stress and consequently Ca2+ overload, which could be of

22

special concern due its close relation with MPTP opening (Ascensao, Lumini-

Oliveira et al. 2011), resulting in the release of proapoptotic proteins and

subsequent activation of programmed cell death (Adhihetty, Ljubicic et al.

2007). Also, an additional production of ROS has been related with aging

process and it has been also reported that ROS expression is upregulated

during cardiac hypertrophy and heart failure (Gustafsson and Gottlieb 2009),

being that, increased mitochondrial oxidant production is accepted as a cause

of myocardial cell loss via apoptosis and necrosis (Judge, Jang et al. 2005). In

this regard, exercise has been shown to provide intrinsic protection (Bowles,

Farrar, & Starnes, 1992; M. B. Harris & Starnes, 2001; Powers et al., 1998;

Powers et al., 2004) and mitochondria play an important role on this

cardioprotective phenotype (Bejma, Ramires et al. 2000, Zhu, Zuo et al. 2007).

In fact, the decreased mitochondrial oxidant damage (Judge, Jang et al. 2005)

is an important adaptive strategy within mitochondria that contribute to

cardioprotection.

Decreased oxidant damage can be understood as a result of both reduction of

oxidant production or increased of antioxidant enzymes. Several reports had

postulate that chronic endurance exercise results in a reduction of mitochondrial

oxidant production (Judge, Jang et al. 2005, Starnes, Barnes et al. 2007) and

enhanced mitochondrial antioxidant enzyme activity (Judge, Jang et al. 2005,

Kavazis, McClung et al. 2008, Starnes, Barnes et al. 2007) which could reduce

oxidative stress induced by different stress stimulators (Ascensao, Magalhaes

et al. 2006, Lennon, Quindry et al. 2004, Powers, Locke et al. 2001). In general,

most studies designed to investigate the influence of regular exercise on

cardiac antioxidant activity focus on endurance training programs, and there are

little available reports on the effect of sprint training in the modulation of

antioxidant defenses (for refs see Ascensao, Ferreira et al. 2007). However, it

seems clear that cardiac muscle tissue, when stimulated by acute exercise,

reveals increased signs of cell damage due to oxidative stress. Despite this fact,

Ascensao et al. (2011) reveal that a single bout of exercise increased SOD

activity, and so, decreased oxidative stress.

23

As previous described, GPX, SOD and CAT are included on primary enzymatic

antioxidant defenses and the relation of those with exercise has been hardly

studied. Interestingly, current data have reported increases (Lennon, Quindry et

al. 2004, Powers, Demirel et al. 1998), no changes (Ascensao, Magalhaes et al.

2005), or even decreases (Chicco, Schneider et al. 2005, Hong and Johnson

1995) in those antioxidant enzyme activities following endurance exercise

training. These differences can be a consequence of some methodological

differences, as characteristics of the animals and/or training protocols;

biochemical procedures or type of markers and enzymes considered

(Ascensao, Ferreira et al. 2007). However, the general approaches indicate that

chronic exercise is responsible for either improved or unchanged levels of

antioxidant enzymes activity.

Also, lipid peroxidation and protein oxidation following endurance training had

been hardly studied and results are still controversial. In fact, both heart lipid

peroxidation and protein oxidation have demonstrated increases (Aydin, Ince et

al. 2007), no changes (Ascensao, Magalhaes et al. 2005, Chicco, Schneider et

al. 2005) or decreases (Ascensao, Magalhaes et al. 2005) in response to

endurance training. However, major tendencies may be indicative that, in

general, endurance exercise training has some positive modulator effects on

some enzymatic and non-enzymatic antioxidant systems (for refs see

Ascensao, Ferreira et al. 2007). In addition, the rate of H2O2 production during

exercise performance has been studied, in order to analyze oxidative stress.

Judge et al. (2005) showed that exercise could decrease H2O2 production in

myocardial mitochondria of male rats. More recently, Starnes at al. (2007) also

reported a H2O2 decrease following an endurance exercise protocol.

Importantly, they found similar decreases in both IFM and SSM mitochondrial

populations. However, different results are under apparently conflict since

Marcil et al. (2006) did not find differences on H2O2 levels following an

endurance exercise protocol. It is possible that differences in sex or rat strain

are responsible for the different results (Starnes, Barnes et al. 2007). However,

it is generally accepted that endurance training induces adaptations within

myocardial mitochondria, resulting in a decreased oxidative stress.

24

Furthermore, Navarro et al. (2004) reported that moderate treadmill exercise

significantly decreased the aging-associated development of heart oxidative

stress preventing the increase in protein carbonyls and TBARS contents of

submitochondrial membranes and the decrease in antioxidant enzyme

activities.

In general, it is accepted that endurance exercise induces cardioprotection

against oxidative stress possible through an increased antioxidant enzyme

activity and/or decreased oxidant production.

2.2.1.4. Cell death pathways

As reviewed in previous sections, apoptosis is one of the pathways of cell death

and it is also the most studied phenomenon in which exercise induces

protection. Kavazis et al. (2008) reported that exercise induces a cardiac

mitochondrial phenotype that resists to apoptotic stimuli in both SSM and IMF.

In this study, cytochrome c was described with primary role in oxidative

adaptation within mitochondria and following an exercise training protocol

cytochrome c release susceptibility after ROS challenge was decreased. Also,

Cyc D and ANT levels decreased following exercise training, being that, ANT

decreased in both SSM and IMF. Furthermore, exercise induces increased

antioxidant (Ascensao, Ferreira et al. 2007, Powers, Locke et al. 2001) and

exercise is related to attenuation of the oxidation and degradation of Ca2+-

handling proteins, which can lead to caspase-3 and -9 activation, and

prevention of calpain activation (French, Hamilton et al. 2008). These reports

suggest that both subfractions of mitochondria undergo biochemical adaptations

in response to endurance exercise leading to decreased apoptotic susceptibility.

In addition to IR, the cardiomyopathy-related to and STZ-induced type I

diabetes and aging, the chronic endurance exercise also provide protection

against tissue and mitochondrial deleterious effects of DOX treatment, a topic of

special interest in the present dissertation. In the following section, the

25

mechanisms and consequences of the selective toxicity of this anti-cancer drug

for the heart will be addressed.

2.3. Doxorubicin: therapeutic agent vs. cardiotoxicity

One of the mostly used chemotherapeutic drugs is the highly effective

anthracycline DOX. However, its clinical use is limited by the dose-related and

cumulative cardiotoxicity and consequent dysfunction that can lead to apoptosis

(Ascensao, Magalhaes et al. 2005, Carvalho, Santos et al. 2009, Wallace

2003). In fact, DOX is an anthracycline prescribed alone or in combination with

other chemotherapeutics for the treatment of various neoplasms such as

leukemias, lymphomas, thyroid and lung carcinomas, several sarcomas,

stomach, breast, bone and ovarian cancers (Wallace 2003). It has been shown

that DOX antineoplastic activity is attributed to its ability to intercalate into the

DNA double helix and/or to bind covalently to proteins involved in DNA

replication and transcription, leading ultimately to cellular death through the

inhibition of DNA, ribonucleic acid (RNA) and protein synthesis (Carvalho,

Santos et al. 2009). More specifically, DOX can be classified as a

topoisomerase II poison (Fortune and Osheroff 2000). Topoisomerase II is an

essential enzyme that plays a role in virtually every cellular DNA process and as

a result of this action DOX generate high levels of enzyme-mediated breaks in

the genetic material of treated cells and ultimately trigger cell death pathways

(Fortune and Osheroff 2000). It has been shown that DOX, by simple diffusion,

enters cancer cells and binds with high affinity to a proteasome in cytoplasm

forming a DOX proteasome complex that translocates into the nucleus in an

ATP-dependent process facilitated by nuclear localization signals. Finally, DOX

dissociates from the proteasome and binds to DNA due to its higher affinity for

DNA than for the proteasome (Kiyomiya, Matsuo et al. 2001). This process is

pictured on figure 2.

26

Figure 2. Mechanisms of DOX action and toxicity (adapted from Carvalho, Santos et al. 2009).

Unfortunately, the clinical use of DOX is limited due the occurrence of a

multidirectional cytotoxic effects, being cardiotoxicity the most known

(Berthiaume and Wallace 2007, Sardao, Pereira et al. 2008). Actually, the

clinical use of DOX soon proved to be hampered by serious problems such as

the development of resistance in tumor cells or toxicity in healthy tissues and

heart seems to be a particularly susceptible organ to DOX-induced toxicity

(Carvalho, Santos et al. 2009). Several mechanisms, in which mitochondria are

involved, are proposed to be responsible for the increased heart sensitivity to

DOX-induced toxicity including: i) a special affinity of DOX by cardiolipin, which

is a major phospholipid component of the inner mitochondrial membrane in the

heart; ii) a higher mitochondrial density per unit volume in cardiomyocytes when

compared to other tissues; iii) the possible but controversial existence of a

specific and external reduced nicotinamide adenine dinucleotide (NADH)

dehydrogenase able to initiate DOX redox cycling and, consequently,

increased formation of ROS and iv) the fact that the heart has low levels of

antioxidant defenses when compared with other tissues (Lebrecht, Setzer et al.

2003, Sardao, Pereira et al. 2008, Wallace 2003).

27

However, despite the clinical manifestations of DOX-induced cardiomyopathy

and the knowledge that DOX induces several cardiac ultrastructural changes,

the mechanisms responsible for DOX-induced cardiac toxicity remain elusive.

Nevertheless, clinical manifestations of DOX-induced cardiotoxicity can be

acute and chronic but there is a wide variation in the frequency of clinical DOX-

induced cardiotoxicity (Carvalho, Santos et al. 2009). Following some reports,

acute effects in the heart, including arrhythmias, hypotension and several

electrocardiographic alterations can be clinically controlled and frequently

reversible, disappearing once the treatment ceases (van Acker, Kramer et al.

1996). On the other hand, chronic cardiotoxic effects induced by DOX are dose-

dependent and culminate in CHF (Swain, Whaley et al. 2003). In both cases,

mitochondria play an important role being classified as a primary target

organelle of DOX-induced cardiotoxicity, evidenced by morphological and

biochemical alterations. Interestingly, Hayward et al. (2012) suggested that all

patients treated with anthracyclines are at greater risk of cardiotoxicity,

particularly those individuals treated with these drugs at an early age. While it is

still unclear why children are at a greater risk, it have been proposed that

exposure to anthracyclines at a young age leads to chronically elevated

hemodynamic stress and an eventual stunting of normal cardiac growth. Also,

Huang et al. (2010) suggested that DOX exposure in juvenile rats decreased

the ability of the heart to adapt to increases in workload as adults or olds.

2.3.1 Cardiac mitochondrial toxicity induced by DOX

Mitochondria are the producers of most of the ATP necessary for metabolism

via oxidative phosphorylation. Also, they play other important roles as cellular

redox and ion homeostasis, oxygen sensing, signaling and regulation of

programmed cell death (for refs see Rabinovitch 2012, Sheu 2009). Therefore,

mitochondrial integrity is vital for cellular homeostasis and cardiac performance

(Chaudhary, El-Sikhry et al. 2011).

28

Unfortunately, mitochondria have been identified as primary DOX target

organelles, and their involvement is evidenced by the results of many studies

reporting functional and morphological alterations, as will be briefly described.

Indeed, it has been suggested that cardiomyocyte dysfunction induced by DOX

treatment is related to increased levels of ROS-induced damage and apoptotic

cellular death, involving mitochondria in the process (Jung and Reszka 2001)

culminating in the disruption of major functions (Jung and Reszka 2001).

The next sections will briefly describe some DOX-induced mitochondrial

dysfunctions such as morphological evidences, increased oxidative stress and

susceptibility to apoptosis.

2.3.1.1 Morphological evidences

As a consequence of acute or chronic DOX treatment, ultrastructural alterations

in rat cardiomyocytes have been reported (Ascensao, Ferreira et al. 2007,

Ascensao, Oliveira et al. 2012, Yilmaz, Atessahin et al. 2006, Zhou, Starkov et

al. 2001). These include nuclear swelling associated with disruption of nuclear

membrane structure, a marked interstitial and cellular edema, perinuclear

vacuolation, disorganization and degeneration of the myocardium, loss of

myofibrils, distension of the sarcoplasmic reticulum, slight enlargement of the T-

tubules and myofibrillar damage and loss (for refs see Carvalho, Santos et al.

2009, Zhou, Starkov et al. 2001). Also, in mice cardiomyocytes, DOX-induced

acute alterations in mitochondria were observed, such as vacuolization, myelin

deposition, disruption of membrane and organelle degeneration with cristae

degeneration, intramitochondrial vacuoles as well as myelin figures (Ascensao,

Ferreira et al. 2007, Ascensao, Oliveira et al. 2012, Sardao, Oliveira et al.

2009).

These ultrastructural alterations are dose-dependent being that higher DOX

concentrations promote more profound cellular alterations (Carvalho, Santos et

29

al. 2009) and those ultrastructural injuries are not repaired after cessation of

DOX treatment becoming even more extensive.

2.3.1.2 Increased oxidative stress

One of the most known DOX-induced side effects is the increased oxidative

stress and it seems to be an event that occurs both acutely and chronically

(Ascensao, Oliveira et al. 2012, Wallace 2003). It has been suggested that DOX

accumulation in mitochondria leads to a redox cycling by complex I of the

mitochondrial respiratory chain, where single electrons are transferred to DOX

(Ascensao, Oliveira et al. 2012). As previously mentioned, the possible but

controversial existence of a specific NADH dehydrogenase able to start this

redox cycling may have an important role. DOX enters mitochondria and reacts

with mitochondrial complex I to form semiquinone radical intermediates, which

is a short-lived metabolite and can react with O2 producing ROS (for refs see

Carvalho, Santos et al. 2009). Then, ROS can react with mitochondrial

biomolecules in the vicinity, which include lipids, proteins and nuclei acids.

Furthermore, increased cardiac oxidative stress associated with DOX toxicity

leads to the depletion of reducing equivalents, impairment in oxidative

phosphorylation with consequent decline in ATP, and interference with cellular

Ca2+ homeostasis (Wallace 2003). Also, DOX is known to react with

mitochondrial mtDNA forming adducts that interfere with proteins expression,

lipid oxidation and normal mitochondrial function, which in turn further increases

ROS production (Sardao, Pereira et al. 2008). This “vicious cycle” postulated

that accumulation of mtDNA mutations is exponential and associated with

massive increase in ROS production (Lenaz 1998). As previously described,

heart mitochondria are important target of DOX, accumulating the drug at

relatively high concentrations. So, it is not surprising that these organelles are

especially susceptible to DOX-induced oxidative damage, and at the same time,

they are also important sources of DOX-induced ROS (Ascensao, Oliveira et al.

2012). On other hand, studies show an upregulation of antioxidant defenses,

30

suggesting an adaptive response of cells to oxidative unbalance promoted by

DOX (Yilmaz, Atessahin et al. 2006). Interestingly, the antioxidant capacity

seems to increase significantly following DOX treatment in young but not in old

Fischer suggesting an increase in DOX-induced oxidative damage with age

(Pritsos and Ma 2000). Despite this fact, is notable that, although the heart has

relatively low antioxidant capacity, it shows an upregulation as an adaptive

response to this oxidative unbalance. Moreover, as a response to this cellular

stress, HSP 60 and HSP 70 increased in hearts and mitochondria from DOX

treated animals and cells (Ascensao, Magalhaes et al. 2005, Kavazis, Smuder

et al. 2010).

2.3.1.3 Increased susceptibility to apoptosis

Another important DOX-induced alteration on cell is the increased susceptibility

to trigger apoptosis. DOX toxicity leads to impairments on cellular Ca2+

homeostasis, which leads to increased susceptibility to the MPTP opening

(Oliveira and Wallace 2006) that is characterized by the loss of the

impermeability that characterizes the inner mitochondrial membrane, as it was

previously described. This complex process is mediated by the formation and

opening of protein complex-like pores, the MPTP (Ascensao, Lumini-Oliveira et

al. 2011, Lumini-Oliveira, Magalhaes et al. 2011). As already mentioned, one of

the consequences of the MPTP induction, besides the disturbance of cell and

mitochondrial Ca2+ homeostasis, is the release of cytochrome c and some

others pro-apoptotic proteins, with consequent initiation of apoptotic cascades.

Also, the increased oxidative stress induced by DOX, is able to interfere with

mitochondrial functionality, which leads to apoptosis. Indeed, DOX-induced

cardiomyocytes apoptosis has been suggested to occur both acutely and

chronically (Carvalho, Santos et al. 2009).

As previously mentioned, its important to develop a strategy that results in a

cardioprotective phenotype against DOX-induced impairments. In this regard,

physical exercise in its various forms has been shown to be an effective

31

intervention that can counteract the acute and chronic deleterious insults for the

myocardium, which includes DOX treatment. The following sections will briefly

discuss this issue.

2.4. Exercise as a therapeutic and preventive strategy against DOX-induced cardiotoxicity.

Exercise has been considered the most effective strategy to promote a healthy

lifestyle and its benefits against DOX-related impairments are evident

(Ascensao, Lumini-Oliveira et al. 2011, Ascensao, Magalhaes et al. 2005,

Ascensao, Oliveira et al. 2012, Chicco, Schneider et al. 2005, Emter and

Bowles 2008, Hayward, Lien et al. 2012, Kavazis, Smuder et al. 2010, Powers,

Lennon et al. 2002, Wonders, Hydock et al. 2008). Although the most studied

form of exercise in DOX treated animals is the chronic exercise, acute exercise

also seems to provide beneficial effects. Mitochondrial adaptations may play

critical role in exercise-induced protection against DOX-induced cardiac

impairments (Ascensao, Ferreira et al. 2007, Ascensao, Oliveira et al. 2012).

The effects of exercise on DOX-treated mitochondria are highlighted in table 2.

2.4.1 Acute exercise

Acute exercise broadly refers to a single bout of exercise performed only once

and it was already proved to be effective against DOX-induced impairments. In

fact, Wonders et al. (2008) reported that an acute bout of treadmill running

performed 24 hours prior to DOX injection attenuated the hemodynamic

impairment observed after acute DOX administration and reduced left

ventricular lipid peroxidation. In addition, Ascensao et al. (2011) showed that an

acute bout of treadmill exercise protects against cardiac mitochondrial

dysfunction, preserving mitochondrial phosphorylation capacity and attenuating

DOX-induced decreased tolerance to MPTP induction. In the same study, it was

32

observed that acute exercise prevented the decreased cardiac mitochondrial

function detected as impaired state 3, phosphorylative lag-phase and maximal

transmembrane potential. Also, acute exercise prevented the inhibitory effects

of DOX treatment on the activity of cardiac mitochondrial respiratory chain

complexes I and V, and on increased caspase-3 and -9 activities.

Furthermore, it has also been described that acute exercise may contribute to

diminished free radical production (for refs see Ascensao, Oliveira et al. 2012).

However, further research is needed to clarify the exact mechanisms by which

an acute exercise induces a protective phenotype in DOX-treated cardiac

mitochondria.

2.4.2 Chronic exercise

Unlike acute exercise, chronic exercise has been hardly studied and the results

consistently demonstrate that is able to antagonize DOX-induced cardiac

impairments (Ascensao, Magalhaes et al. 2005, Chicco, Schneider et al. 2005,

Hayward, Lien et al. 2012, Hydock, Lien et al. 2008, Kavazis, Smuder et al.

2010). Those alterations induced by chronic exercise can be seen at

morphological, functional and biochemical levels and the role of mitochondria is

pivotal in this process (Ascensao, Oliveira et al. 2012, Kavazis, Smuder et al.

2010). Ascensao et al. (2005) reported the involvement of mitochondria in

cardioprotection afforded by endurance training against DOX treatment,

demonstrating prevention of acute DOX-induced mitochondrial alterations

regarding oxidative stress, respiration, and Ca2+ loading capacity.

At ultrastructural level, it has been reported that cardiac alterations induced by

DOX treatment are also attenuated by previous chronic exercise (Ascensao,

Magalhaes et al. 2005). In fact, when compared with saline (SAL) group,

sedentary (SED) DOX-treated animals showed myocardial damage (Ascensão,

Magalhães et al. 2006). The observed morphological alterations consisted of

mitochondrial damage with extensive degeneration and loss of cristae, swelling

33

and abnormal size and shape, intramitochondrial vacuoles and notorious myelin

figures that probably resulted in the formation of secondary lysosomes. All of

these alterations were attenuated in trained animals treated with DOX

(Ascensão, Magalhães et al. 2006).

At functional level, different authors reported the protective effects of chronic

exercise. In fact, Hydock et al. (2008) suggested that exercise training in rats

before DOX treatment attenuated DOX-induced cardiac dysfunction, through

the maintenance of fractional shortening, developed pressure and contractility.

Also, Chicco et al. (2005) reported that both low intensity exercise training and

an endurance training protocol with gradually increased intensity attenuated the

adverse effects of DOX by preventing DOX-induced decline in cardiac function

through maintenance of left ventricular diastolic pressure, rate of left ventricular

pressure development and rate of left ventricular relaxation.

At biochemical level, as depicted on table 2, several authors had proved the

beneficial effects of chronic exercise. Importantly, some of the most studied

biochemical parameters associated with exercise and DOX are the antioxidant

capacity, oxidative stress markers and apoptotic susceptibility (for refs see

Ascensao, Oliveira et al. 2012). As previous referred, HSP have an important

role as antioxidants molecules contributing to normal cellular integrity and are

overexpressed after endurance training. However, Chicco et al (2006) and

Kavasis et al. (2010) also showed that exercise had no influence on HSP or that

it is not determinant on cardioprotection being that exercise in cold vs. normal

temperatures may also display other types of differences regarding alteration of

mitochondrial physiology, besides alteration in the expression of HSPs.

Furthermore, the upregulation of mitochondrial manganese superoxide

dismutase (MnSOD) seems contribute for cardioprotection. In fact, as

mitochondrial DOX toxicity has been largely attributed to increased oxidative

stress, increased antioxidant activity may be important to explain how

endurance training counteracts some of DOX-induced myocardial damage.

Chicco et al. (2006) associated low-intensity treadmill exercise training-induced

cardioprotection to the inhibition of apoptotic signaling and the increased activity

34

of GPX. Also, the effect of training on preventing activation of cardiac apoptotic

pathways has been described being that, training decreased the susceptibility of

appearance of apoptotic markers in the hearts of DOX-treated animals, as

increased mitochondrial Bax, Bax-to-Bcl2 ratio and tissue caspase 3 activity

(Ascensao, Magalhaes et al. 2005). In fact, it has been suggested that chronic

exercise stimulation may also afford protection against the increased

susceptibility to the MPTP as the deleterious effects of Ca2+ on heart

mitochondrial respiration of DOX-treated animals were attenuated in trained

group treated with DOX (Ascensao, Magalhaes et al. 2005). As previous

described, MPTP is related to oxidative damage and therefore, it is possible that

increased resistance of cardiac mitochondria from trained animals to the MPTP

can be related to increased antioxidant defenses. Accordingly, the higher levels

of reduced sulfhydryl groups in trained mitochondria than in sedentary groups

may be indicative of enhanced antioxidant capacity and/or of more elevated

sulfhydryl-donors, such as GSH, in mitochondria from trained animals (for refs

see Ascensao, Oliveira et al. 2012). However, further studies are necessary in

order to better understand this issue.

Table 2. Summary of some described mitochondrial-related alterations associated with DOX-induced cardiotoxicity and the modulation effect afforded by physical exercise against DOX (adapted from Ascensao, Oliveira et al. 2012).

DOX effect Exercise effect against DOX

ROS production ↑ ↓

Oxidative damage markers

Lipid peroxidation ↑ ↓

Protein oxidation ↑ ↓

DNA oxidation ↑

Aconitase activity ↓ or = ↑

Apoptotic signaling

35

(↑) – increase; (↓) – decrease; (=) – no alterations

Despite the extensive number of studies on this topic, the effects of both

endurance treadmill training and voluntary free-wheel running activity performed

Bax-Bcl-2 ratio ↑ ↓

Cytochrome c release ↑

Caspase 9 activation ↑ ↓

Respiratory endpoints

State 3 ↓ ↑ or =

State 4 ↑ or = or ↓ = or ↓

RCR ↓ = or ↑

ADP/O ratio = or ↓

Uncoupled respiration ↓ ↑

Creatine-stimulated respiration ↓

Maximal ΔΨ ↓ or = ↑

Ca2+-induced MTPT ↑ ↓

ANT content and functioning ↓

Mitochondrial chaperones ↑ ↑

Mitochondrial antioxidants

Thiols ↓ ↑

Vitamin E ↓ or =

Enzymes = or ↑ ↑

Coenzyme Q isoenzymes =

ETC complex activity

Complex I ↓ ↑

Complex II ↓

Complex III =

Complex IV = or ↓

Complex V ↓ ↑

36

before and during sub-chronic DOX treatment schedule on cardiac

mitochondrial bioenergetics are yet to be elucidated.

This is of particular importance in the context of exercise-induced protection

against DOX-related cardiac mitochondriopathy, as cancer patients undergoing

DOX treatment may be advised to exercise for many reasons including to

counteract physical fatigue and to improve performance, and also to mitigate

cardiac damage as a result of chemotherapy.

This master sports science course in which this work is inserted is the context of

physical activity and elderly. Despite developed with adult rats, the present work

can contribute to extend the knowledge in this particular area representing

preliminary findings in adult population and considering that DOX-based

chemotherapeutic treatments against several types of malignances are more

prevalent with increasing age.

37

3. Aim

The aim of the present study was to analyze the effect of two types of physical

exercise (treadmill endurance training (TM) and free-wheel voluntary physical

activity (FW)) against heart mitochondrial dysfunction induced by sub-chronic

treatment of DOX.

We can define as specific purposes of this work the analysis of the adaptations

induced by both types of exercise on heart mitochondria from DOX treated

animals on:

• Mitochondrial respiratory function;

• Mitochondrial electrical transmembrane potential;

• MPTP susceptibility;

• Mitochondrial oxidative damage.

39

4. Materials and methods

4.1 Reagents

Deionized water (18.7 MΩ) from an arium®611VF system (Sartorius, Göttingen,

Deutschland) was used. Doxorubicin hydrochloride, commercial/clinic use, was

obtained from Ferrer Farma (Barcelona, Spain), prepared in a sterile saline

solution, sodium chloride (NaCl) 0.9% (pH 3.0) and stored at 4ºC for no longer

than five days upon rehydration. All other chemicals were purchased from

Sigma Aldrich (Sintra, Portugal).

4.2 Animals

All experiments involving animals were conducted in accordance with the

European Convention for the Protection of Vertebrate Animal Used for

Experimental and Other Scientific Purposes (CETS no. 123 of 18 march 1986

and 2005 revision) and the Commission Recommendation of 18 June 2007 on

guidelines for the accommodation and care of animals used for experimental

and other scientific purposes (C (2007) 2525). Supervisors of this work are

accredited by the Federation of Laboratory Animal Science Associations

(FELASA) for animal experimentation. The Ethics Committee of the Faculty of

Sport approved this experimental protocol.

Thirty-six male Sprague-Dawley rats (aged 2 weeks old) were obtained from

Charles River Laboratories (L’Arbresle, France) and were randomly divided into

six groups (n=6 per group): Saline sedentary (SAL+SED), saline treadmill

endurance training (SAL+TM), saline free wheel voluntary physical activity

(SAL+FW), doxorubicin sedentary (DOX+SED) doxorubicin treadmill endurance

training (DOX+TM) and doxorubicin free wheel voluntary physical activity

(DOX+FW). Only male rats were used to avoid hormone-dependent influence in

drug mitochondrial function/toxicity. During the experimental protocol, animals

were housed in collective cages (two rats per cage) and were maintained in a

40

room at normal atmosphere (21–22 ºC; 50–60% humidity) receiving food

(Scientific Animal Food and Engineering, A04) and water ad libitum in 12-h

light/dark cycles.

4.3 Exercise protocols

4.3.1 Endurance training protocol

The animals from TM groups were exercised 5 days/week (Monday–Friday) in

the morning (between 10:00 and 12:00 A.M.), for 12 weeks on a LE8700 motor

driven treadmill (Panlab, Harvard, U.S.A). The treadmill speed was gradually

increased over the course of the 12-week training period. The protocol included

5 days of habituation to the treadmill with 10 min of running at 15 m/min, with

daily increases of 5-10 minutes (min) until 30 min was achieved (week 0).

Habituation was followed by one consecutive week of continuous running (30

min/day) at 15 m/min and was gradually increased until 60 min/day on the week

1. The velocity increased gradually from 18 m/min to 27 m/min (week 7).

SAL+TM continue to increase velocity to 30 m/min while DOX+TM animals

gradually decreased running velocity to 20 m/min (Table 3).

Table 3. TM exercise protocol

Week 0 1 2 3 4 5 6 7 8 9 10 11 12

SAL+ TM

TM Velocity (m/min) 15 18 20 22 24 25 25 27 27 28 28 30 30

DOX + TM

TM Velocity (m/min) 15 18 20 22 24 25 25 27 27 25 25 22 20

Exercise Duration (min/day) 30 60 60 60 60 60 60 60 60 60 60 60 60

41

4.3.2 Voluntary physical activity

The animals from FW groups were housed in a polyethylene cage equipped

with a running wheel [perimeter=1,05m, Type 304 Stainless steel (2154F0106-

1284L0106) Technicplast, Casale Litta, Italy)]. The rats were allowed to

exercise ad libitum with an unlimited access to the running wheel 24h/day.

Running distance was recorded using ECO 701 from Hengstler (Lancashire,

U.K.).

4.4 Doxorubicin treatment

After the 5th week of endurance training or free wheel exercise, the animals

were sub-chronically treated with DOX (2 mg/Kg of body weight) or sterile saline

solution NaCl 0.9% (SAL, 2 mg/kg of body weight) intraperitoneal injection/week

during 7 weeks. The animals assigned to the TM groups received DOX or SAL

injections during the weekend in a day-off training.

4.5 Animal sacrifice, heart and soleus extraction

Forty-eight hours after the last TM exercise bout, non-fasted rats were

euthanized by cervical dislocation between 9:00 and 10:00 AM to eliminate

possible effects due to diurnal variation. After quickly opening the chest cavity,

rat hearts were rapidly excised, rinsed, carefully dried, and weighed. Right

soleus muscle was also rapidly extracted and weighed. Portions of

approximately 50 milligrams (mg) of one soleus muscle were homogenized in

homogenization buffer (200 minimolar (mM) Tris, 137 mM NaCl, 0.2 mM EDTA,

0.5 mM EGTA, 1% triton X-100, tissue: buffer ratio of 100 mg/mL, pH 7.4) using

a Teflon pestle on a motor driven Potter-Elvehjem glass homogenizer at 0–4°C

three to five times for 5 s at speed low setting, with a final burst at a higher

speed setting. Homogenates were centrifuged (2 min at 3000 xg, 4°C, in order

to eliminate cellular debris) and the resulting supernatants were stored at −80°C

42

for later determinations, as detailed bellow. Protein content from soleus

homogenates were spectrophotometrically determined using the biuret method

and bovine serum albumin as standard (Gornall, Bardawill et al. 1949).

4.6 Isolation of heart mitochondria

Heart mitochondria were daily prepared using conventional methods of

differential centrifugation (Bhattacharya, Thakar et al. 1991) as follows. Briefly,

the animals were sacrificed as stated above and the heart was immediately

excised and finely minced in an ice-cold isolation medium containing 250 mM

sucrose, 0.5 mM EGTA, 10 mM HEPES (pH 7.4), and 0.1% defatted bovine

serum albumin (BSA, Sigma, cat. no. A-7030). The minced blood-free tissue

was then resuspended in 40 mL of isolation medium containing 0,75 mg/mL

protease subtilopeptidase A Type III (Sigma P-5380) and homogenized with a

tightly fitted homogenizer (Teflon: glass pestle). The suspension was incubated

for 1 min (4°C) and then re-homogenized. The homogenate was then

centrifuged at 13,000 xg for 10 min. The supernatant was decanted and the

pellet, essentially devoid of protease, was gently re-suspended with a loose-

fitting homogenizer. The suspension was centrifuged at 750 xg for 10 min and

the resulting supernatant was centrifuged at 12,000 xg for 10 min. The pellet

was re-suspended using a paintbrush and re-pellet at 12,000 xg for 10 min.

EGTA and defatted BSA were omitted from the final washing medium.

Mitochondrial protein content was determined by the Biuret method calibrated

with BSA (Gornall, Bardawill et al. 1949). The isolation procedures were

performed within approximately 1 h at 0–4°C. Aliquots of isolated mitochondria

were separated and frozen at −80°C for later determination of oxidative

damage. The remaining mitochondrial were used within 2–3 h after the excision

of the heart and was maintained on ice (0–4 ºC) throughout this period.

43

4.7 Mitochondrial respiratory activity

Mitochondrial respiratory function was measured polarographically at 30ºC

using a Biological Oxygen Monitor System (Hansatech Instruments) and a

Clarktype oxygen electrode (Hansatech DW1, Norfolk, UK). The reactions were

conducted in a 0.75 mL closed, thermostatted and magnetically stirred glass

chamber containing 0.5 mg/mL of mitochondrial protein in a respiration buffer

containing 50 mM KCl, 130 mM sucrose, 2.5 mM KH2PO4, and 0.5 mM Hepes,

pH 7.4. After 1-min equilibration period, mitochondrial respiration was initiated

by adding glutamate/malate (G/M) to a final concentration of 5 and 2.5 mM,

respectively. State 3 respiration was determined after adding ADP (150 nmol);

state 4 was measured as the rate of oxygen consumption after ADP

phosphorylation. The RCR (state 3/state 4) and the ADP/O (the number of nmol

ADP phosphorylated by atom of oxygen consumed) ratios were calculated

according to Estabrook (1967).

4.8 Mitochondrial transmembrane electric potential

Mitochondrial transmembrane electric potential (∆ψ) was monitored indirectly

based on the activity of the lipophilic cation tetraphenylphosphonium (TPP+)

using a TPP+ selective electrode prepared in our laboratory as previously

described (Ascensao, Lumini-Oliveira et al. 2011). Reactions were carried out in

1 mL of reaction buffer containing 50 mM KCl, 130 mM sucrose, 2.5 mM

KH2PO4, and 0.5 mM Hepes, pH 7.4 supplemented with 3 µM TPP+ and 0.5

mg/mL of mitochondrial protein. For the measurements of ∆ψ with complex I-

linked substrates, energization was carried out with G/M (5 mM and 2.5 mM,

respectively) and ADP phosphorylation was achieved by adding 150 nmol ADP.

The lag phase, which reflects the time needed to phosphorylate the added ADP,

was also measured during experiments.

44

4.9 Mitochondrial osmotic swelling during MPTP induction

Mitochondrial osmotic volume changes were followed by monitoring the classic

decrease of absorbance at 540 nm with a Jasco V-630 spectrophotometer.

Swelling amplitude and rate of decreased absorbance upon Ca2+ addition were

considered as MPTP susceptibility indexes. The reaction was continuously

stirred and the temperature was maintained at 25 ºC. The assays were

performed in 1 ml of reaction medium containing 200 mM sucrose, 10 mM

HEPES, 5 mM KH2PO4, 10 µM EGTA , pH 7.4, supplemented with 1.5 µM

rotenone, 8 mM succinate and a single pulse of 80 nmol of Ca2+ with 0.5 mg/ml

protein. Control trials were performed by using 1 µM of cyclosporin-A, the

selective MPTP inhibitor (Broekemeier, Dempsey et al. 1989).

4.10 Mitochondrial oxidative damage

Before analysis, mitochondrial membranes were disrupted by several freeze–

thawing cycles to allow free access to substrates. The extent of lipid

peroxidation in heart mitochondria was determined by measuring MDA contents

by colorimetric assay, according to a modified procedure described previously

(Buege and Aust 1978). Suspended mitochondria were centrifuged at 12,000 xg

for 10 min and re-suspended in 150 µL of a medium containing 175 mM KCl

and 10 mM Tris-HCl, pH 7.4. Subsequently, mitochondria from the six groups

were mixed with 2 volumes of trichloroacetic acid (10%) and 2 volumes of

thiobarbituric acid (1%). The mixtures were heated at 80–90 ºC for 10 min and

re-cooled in ice for 10 min before centrifugation (4,000 xg for 10 min). The

supernatants were collected and the absorbance measured at 535 nm. The

amount of MDA content formed was expressed as nanomoles of MDA per

milligram of protein (ε535=1.56 x 10−5 M−1 cm−1)

The basal mitochondrial content of oxidative modified -SH groups, including

GSH and other -SH containing proteins, was quantified by spectrophotometric

measurement according to Hu (1990). Briefly, a mitochondrial suspension

45

containing 5 mg/mL protein was mixed with 0.25 M Tris buffer pH 8.2 and 10

mM DTNB and the volume was adjusted to 1 mL with absolute methanol.

Subsequently, the samples were incubated for 30 min in the dark at room

temperature and centrifuged at 3000 xg for 10 min. The colorimetric assay of

supernatant was performed at 414 nm against a blank test. Total -SH content

was expressed in nanomoles per milligrams of mitochondrial protein (ε414=13.6

mM−1 cm−1).

4.11 Soleus citrate synthase activity

Soleus CS activity was measured using the method proposed by Coore et al.

(1971). The principle of assay was to initiate the reaction of acetyl-CoA with

oxaloacetate and link the release of CoA-SH to 5,5-dithiobis (2-nitrobenzoate)

at 412 nm.

4.12 Statistical analysis

All data are expressed as the mean±SEM (Standard Error of the Mean).

Statistical analyses were performed using GraphPad Prism (version 6.0) or

Statistical Package for the Social Sciences (SPSS version 21.0). Three-way

repeated-measures ANOVA for body weight and distance cover by exercised

groups to verify the effect of exercise and treatment over time. Two-way

analysis of variance ANOVA were used to examine possible effect of treatment

and/or exercise. To determine specific group differences, the two-way ANOVA

were followed by Bonferroni post-hoc tests. In all cases, the significance level

was set at p≤0.05.

47

5. Results

5.1. Characterization of animals and exercise protocols

Body weight alterations and distances covered by the animals during the entire

protocol are shown in figure 3. No significant differences in the mean body

weight of the animals from the beginning of the protocol until the 5th week, when

sub-chronical DOX treatment was initiated, were found. Body weights of DOX

treated animals were lower than SAL counterparts at the end of the protocol

(DOX+SED vs. SAL+SED; DOX+TM vs. SAL+TM; DOX+FW vs. SAL+FW;

p≤0.05). No differences in body weight between exercised groups, were found.

TM and FW decreased body weight at 12th and 9th week, respectively (SAL+TM

and SAL+FW vs. SAL+SED; p≤0.05). DOX treatment combined with TM

decreased body weight from the 5th week (DOX+TM vs. DOX+SED; p≤0.05),

whereas no significant differences were found between FW and SED treated

groups (Figure 3A). After the 5th week DOX treated groups consumed less food

than their SAL counterparts (data non shown, p≤0.05). Water consumption

increased in FW groups (SAL and DOX) compared with SAL+SED and

DOX+SED groups (data non shown, p≤0.05).

Figure 3. Effect of exercise and DOX treatment on (A) body mass over time and (B) distance covered per day by TM and FW groups during the 12 wks of protocol. Significant differences (p≤0.05) are mentioned in the text. Significant (p≤0.05) effects of Exercise (E), Treatment (T), time (t) or their interaction (E x T x t) are shown; Non Significant (NS, p>0.05).

48

As can be seen in Figure 3B, voluntary running distance decreased significantly

in DOX+FW after the 5th week and remained lower until the end of the protocol

(p≤0.05). Animals from TM group ran at the same velocity throughout the 8

weeks of the protocol. Running velocity and distance covered diminished in

DOX+TM group at 11th and 12th week compared to SED+TM (p≤0.05).

Body, heart absolute weights, heart weight and femur length to body weight

ratios, mitochondrial protein yielding as well as the activity of soleus citrate

synthase in the six groups are shown in Table 4. Final body, heart weight and

ratio of heart weight to body weight significantly decreased with DOX treatment

(SAL+SED vs DOX+SED). Both chronic exercise types decreased final body

weight, increased heart weight and the heart to body ratio (SAL+TM and

SAL+FW vs SAL+SED). DOX treatment combined with TM and FW exercise

induced a significant increase in heart weight and heart weight to body weight

ratio compared with their DOX+SED counterparts. No significant differences

were observed between groups regarding the Initial body weight, femur length

to body weight ratio and yield of mitochondria isolation. TM induced a significant

increase in the activity of soleus citrate synthase in both SAL and DOX treated

animals (SAL+SED vs SAL+TM and DOX+SED vs DOX+TM).

49

Table 4. Animal data and yield of mitochondrial protein isolation

SAL+SED SAL+TM SAL+FW DOX+SED DOX+TM DOX+FW P*

Initial body weight (g) 207±3.91a 214±3.76a 212±1.55a 209±4.68a 207±4.63a 209±2.60a NS

Final body weight (g) 598±10.58a 522±9.87b 498±5.98b 438±6.22c 426±10.79c 429±16.95c ExT

Heart weight (g) 1.44±0.03a 1.92±0.08b 1.85±0.10b 1.10±0.03c 1.45±0.05a 1.55±0.11a E, T

Heart weight/body weight (mg.g−1) 2.32±0.08a 3.53±0.10bc 3.68±0.12c 2.62±0.11d 3.34±0.08b 3.34±0.12b ExT

Femur length/body weight (mm.g-1) 0.08±0.00a 0.08±0.00a 0.09±0.00a 0.08±0.01a 0.09±0.00a 0.09±0.00a E, T

Mitochondrial protein yielding (mg protein/g tissue)

18.58±0.62a 15.15±1.22a 15.01±1.50a 16.96±0.83a 18.60±4.68a 17.84±1.57a NS

soleus citrate synthase activity (nmol. min-1.mg-1) 10.94±2.07a 23.34±1.79b 11.22±2.65a 8.69±1.15a 22.22±1.97b 10.27±1.32a E, T

Values (mean ± SEM). Different letters are significantly different (p≤0.05). * Significant (p≤0.05) effects of Exercise (E), Treatment (T), or their interaction (E x T) are shown; Non Significant (NS, p> 0.05).

50

5.2 Heart mitochondrial oxygen consumption

Mitochondrial respiratory activity in both SAL and DOX treated groups was

measured to identify exercise-dependent effects (Figure 4). DOX treatment

decreased heart mitochondrial respiration during state 3 and increased state 4

in SED animals (DOX+SED vs SAL+SED). Importantly, TM and FW exercise

per se increased State 3 respiration in both SAL and DOX (SAL+TM and

SAL+FW vs. SAL+DOX; DOX+TM and DOX+FW vs. DOX+SED). The coupling

between oxygen consumption and ADP phosphorylation (RCR) was

significantly affected by DOX treatment (DOX+SED vs. SAL+SED). TM

significantly increased RCR in both SAL and DOX groups (SAL+TM vs.

SAL+SED; DOX+TM vs. DOX+SED). FW increased RCR in DOX treated

animals (DOX+FW vs. DOX+SED). Also, both TM and FW increased ADP/O in

DOX group (DOX+SED vs. DOX+TM and DOX+FW).

51

Figure 4. Effect of exercise and DOX treatment on (A) state 3 of heart mitochondrial respiration, (B) state 4 of heart mitochondria respiration, (C) RCR and (D) ADP/O. Data are means±SEM for heart mitochondria (0.5 mg/mL protein) obtained from different mitochondrial preparations for each experimental group. Oxidative phosphorylation was measured polarographically at 30ºC in a total volume of 0.75 mL. Respiration medium and other experimental details are provided in methods. RCR, respiratory control ratio (state 3/state 4); ADP/O, number of nmol ADP phosphorylated by atom of oxygen consumed. Different letters are significantly different (P≤0.05). Significant (p≤0.05) effects of Exercise (E), Treatment (T), or their interaction (E x T) are shown; Non Significant (NS, p>0.05).

5.3 Heart mitochondrial transmembrane electric potential

Heart mitochondrial variations in maximal ∆ψ and during ADP phosphorylation

were determined using G/M as substrates. DOX treatment significantly affected

the maximal ∆ψ, repolarization and ADP lag-phase (Figure 5). FW but not TM,

increased maximal ∆ψ and repolarization, whereas both types of exercise

decreased the ADP lag phase (SAL+TM and SAL+FW vs. SAL+SED). Both

exercise protocols were able to counteract the DOX harmful effect normalizing

maximal ∆ψ, repolarization and ADP lag phase.

52

Figure 5. Effect of exercise and DOX treatment on heart mitochondria ∆ψ fluctuations (A) maximal energization, (B) ADP-induced depolarization, (C) repolarization and (D) ADP phosphorylation lag phase. Data are mean±SEM for heart mitochondria (0.5 mg/mL protein) obtained from different mitochondrial preparations for each experimental group. Figure shows the average response of maximal mitochondrial membrane potential developed with glutamate (5 mM) plus malate (2.5 mM), the decrease in membrane potential after ADP addition (depolarization), the repolarization value after ADP phosphorylation, and the lag phase. Mitochondrial transmembrane potential was measured using a TPP+-selective electrode at 30ºC in a total volume of 1 mL. Reaction medium and other experimental details are provided in methods. Different letters are significantly different (p≤0.05). * Significant (p≤0.05) effects of Exercise (E), Treatment (T), or their interaction (E x T) are shown; Non Significant (NS, p>0.05)

5.4 Mitochondrial osmotic swelling during MPTP induction

The effects of both types of exercise training and DOX treatment on in vitro

susceptibility to Ca2+-induced MPTP opening were investigated. The addition of

Ca2+ on mitochondria suspension resulted in a decrease in absorbance with

three distinct phases. Initially, an increase in absorbance was observed, which

most likely results from the formation of opaque Ca2+ crystals inside

mitochondria (Andreyev, Fahy et al. 1998). Upon MPTP opening, a decrease of

53

absorbance with a slow followed by a fast kinetic rate is usually observed in

cardiac mitochondria. Incubation of mitochondrial suspension with cyclosporine

A, a specific MPTP inhibitor (Broekemeier, Dempsey et al. 1989), limits the

absorbance decrease after Ca2+ addition, which demonstrate the association

with MPTP opening.

Figure 6 shows different end-points measured from the recordings obtained,

namely (A) swelling amplitude (the difference between the initial and the final

absorbance value) and (B) the average swelling rate. The results demonstrate

that DOX treatment significantly increased susceptibility to Ca2+-induced MPTP

opening (DOX + SED vs. SAL + SED). Heart mitochondria isolated from

SAL+TM group, but not SAL+FW were less susceptible to Ca2+-induced MPTP

opening (SAL + TM vs. SAL + SED). Both types of exercise were able to

mitigate DOX-induced increased susceptibility to MPTP opening (DOX + TM

and DOX + FW vs. DOX + SED).

Figure 6. Effect of exercise and DOX treatment on heart mitochondria to Ca2+-induced MPTP (A) Swelling amplitude; (B) Average swelling rate. Data are mean ± SEM. The absorbance of mitochondrial suspension was followed at 540 nm. Mitochondria were incubated as described in methods. A 80 nmol of Ca2+ pulse (160 nmol/mg protein) was added to 0.5 mg of mitochondrial protein in order to attain the cyclosporin A-sensitive swelling, indicating that the decreased optical density corresponding to the increased swelling was due to MPTP opening. Different letters are significantly different (p≤0.05). * Significant (p≤0.05) effects of Exercise (E), Treatment (T), or their interaction (E x T) are shown; Non Significant (NS, p>0.05)

54

5.5 Oxidative stress markers

The next step was to ascertain whether exercise and DOX treatment-modulated

mitochondrial oxidative stress markers. According to the protective phenotype

seen in Fig.7 DOX treatment increased MDA levels and decreased –SH content

(SAL + SED vs. DOX + SED). Exercise, particularly TM decreased

mitochondrial MDA levels (SAL + TM vs. SAL + SED). Both types of exercise

were able to revert DOX-induced alterations in MDA level and –SH content

(DOX + SAL vs. DOX + TM and DOX + FW).

Figure 7. Heart mitochondrial (A) MDA and (B) reduced sulfhydryl contents. Data are means±SEM for heart mitochondria obtained from different mitochondrial preparations for each experimental group. Different letters are significantly different (p≤0.05). Significant (p<0.05) effects of Exercise (E), Treatment (T), or their interaction (E x T) are shown; Non Significant (NS, p> 0.05).

55

6. Discussion

The current study provided additional support to understand the effects of both

endurance treadmill training and voluntary wheel running activity performed

before, during and after sub-chronic DOX treatment schedule on cardiac

mitochondrial bioenergetics. Only male rats were used to avoid hormone-

dependent influence in drug-induced mitochondrial toxicity (Lagranha,

Deschamps et al. 2010). Rats were sub-chronically treated with DOX in an

attempt to mimic human’s treatment, a protocol previously used by others

(Pereira, Pereira et al. 2012, Santos, Moreno et al. 2002). Moreover, in the

present study 1st DOX injection was administrated 5 weeks after the beginning

of the exercise protocol with subsequent weekly injection until the end of

protocol. This set up can be understood with both a preconditioning (preventive)

and a therapeutic strategy against DOX treatment schedules. Two different

types of exercise were analyzed: voluntary free wheel run and treadmill run.

Considerable attention has been focused on the efficacy of exercise protocols

since it has been speculated that further stress induced by exercise could be

potentially detrimental, exacerbating the impairments induced by DOX (Emter

and Bowles 2008). In fact, patients undergoing chemotherapy experience

severe fatigue or exercise intolerance. Consequently, the intensity and duration

of exercise they are able to tolerate is likely to be severely limited (Emter and

Bowles 2008). For these reason, and because we wanted to analyze the

response on heart mitochondria of DOX treated rats at distinct intensity and

duration, both free wheel and run treadmill were performed.

The results on mitochondrial function obtained in the present study confirm at

least in part, the cardiac protection afforded by both endurance treadmill training

and voluntary free wheel running (for refs see Ascensao, Oliveira et al. 2012)

against DOX toxicity. Cardiac dysfunction and defective mitochondrial function

in DOX-treated animals has been studied previously (Ascensao, Lumini-Oliveira

et al. 2011, Ascensão, Magalhães et al. 2006, Chicco, Schneider et al. 2005,

Kavazis, Smuder et al. 2010, Sardao, Pereira et al. 2008, Wallace 2003). The

present study confirmed that sub-chronic DOX administration in heart

56

mitochondria results in: (i) worsening heart mitochondrial respiration; (ii)

decreased maximally developed ΔΨ, repolarization and increased

phosphorylative lag phase; (iii) decreased ability of heart mitochondria to

accumulate Ca2+ before MPTP induction and (iv) increased MDA levels and

decreased -SH content. Both types of exercise performed before and during

DOX treatment resulted in attenuation or complete prevention of the heart

mitochondrial impairments induced by DOX.

6.1 Heart mitochondrial oxygen consumption and transmembrane electric potential

Previous studies have shown that exercise attenuates DOX-induced cardiac

damage, diminishing the increased biochemical and morphological signs of

toxicity induced (Ascensao, Magalhaes et al. 2005, Ascensao, Oliveira et al.

2012, Chicco, Schneider et al. 2006, Hydock, Lien et al. 2008, Kanter, Hamlin et

al. 1985, Kavazis, Smuder et al. 2010); however, no data are available

concerning the cross-tolerance effect of both voluntary free wheel running and

endurance training on sub-chronic DOX treatment mitochondrial malfunction.

Present results demonstrate that sub-chronic treatment of DOX induces

impairments on mitochondrial respiration, and that 12 weeks of endurance

running training and voluntary free wheel running prevented the inhibition of

mitochondrial respiration. Alterations in mitochondrial oxidative phosphorylation

induced by DOX relies in several factors including the decreased aconitase

activity; increased ROS production and activity; and the decreased activity,

content or organization of the electron transport chain complexes or proteins of

the phosphorylation system (Ascensao, Lumini-Oliveira et al. 2011). Also, the

depressed activity of mitochondrial complexes I and II caused by DOX (Santos,

Moreno et al. 2002) could partially justify the diminished electron transport

through electron transport chain (ETC) in the SAL+DOX group. Thus, the

unaltered state 3 respiration observed in exercised groups suggest that, among

other possible effects, training probably prevented the inactivation of complex I

57

and II in DOX-treated heart mitochondria. The enhanced ETC functionality

could also be due to an up regulation of oxido-redutase activity or increased

availability of reduced equivalents formation, consequently increasing the

supply of electrons to the ETC (Nulton-Persson and Szweda 2001) or even to

enhanced capability of phosphorylative system due an upregulation of Krebs

cycle enzymes (Holloszy, Oscai et al. 1970). Furthermore, our experiments

reveal that mitochondria isolated from hearts of animals treated with DOX

exhibited impaired coupling (i.e., lower RCR). RCR is known as a respiratory

parameter associated with mitochondrial functionality and structural integrity

(Brand and Nicholls 2011). As exercise training prevent DOX-induced

uncoupled cardiac mitochondrial respiration, this might suggest that exercise

training enhance mitochondrial respiratory activity due increased

phosphorilative system functionality. Interestingly, regarding RCR, FW running

protocol was more effective at counteracting DOX-induced impairments, which

may be consequence of a slight decreased in sate 4 observed in DOX+FW

group. Concerning ADP/O, both TM and FW groups reverted the values of

DOX+SED group, which suggest that training prevented the heart mitochondrial

impairments in oxidative phosphorylation capacity in DOX rats.

Because mitochondrial complexes rely on enzymatic machinery (Bernstein,

Bucher et al. 1978), they can become prone to impairments induced by ROS,

resulting in accumulation of products of protein oxidation. DOX-induced

impairments in oxidative damage markers, such as MDA level and -SH content

were consistent with alterations in mitochondrial respiratory function, which can

suggest that exercise may counteract DOX-induced impairments in redox

homeostasis. One possible justification for these alterations is that exercise

induced up-regulation of mitochondrial defenses including HSPs or SOD

contributing to the up-regulation of mitochondrial tolerance against DOX effects

(Ascensao, Ferreira et al. 2007). The up-regulation of other antioxidants such

as GSH and CAT has also been described with exercise and DOX (Ascensao,

Magalhaes et al. 2005, Kavazis, Smuder et al. 2010).

58

The complementary study of Δψ is indispensable for a complete analysis of

mitochondrial function being that it reflects the basic energetic relation to

cellular homeostasis maintenance. In fact, the electrochemical gradient due the

pumping of protons through the inner membrane (Murphy and Brand 1988) is

indispensable to ADP phosphorylation (Stock, Leslie et al. 1999). Moreover,

when cytosolic concentration of Ca2+ increases, mitochondria act as Ca2+

buffers due its ability to uptake and accumulate Ca2+ (Gunter, Yule et al. 2004).

It has been suggested that intramitochondrial Ca2+ concentration, whose flow is

directed in accordance with the protomotriz gradient, has a controlling function

in metabolic rate of oxidative energy production through the activation of Ca2+-

sensitive dehydrogenases, F0F1ATPase and ANT (Glancy, Willis et al. 2013).

Our results showed that DOX decreased maximal Δψ, repolarization and

increased the lag phase. The lag phase represents the time elapsed to

phosphorylate ADP. In the present study, exercise led to increased maximal

Δψ, repolarization and decreased time to restore membrane potential after

addition and consequent ADP phosphorylation in the DOX+FW and DOX+TM

groups. One possible explanation for the observed protective effect of exercise

may be associated with the preservation of mitochondrial complex activity,

namely complex I and V in exercised groups (Ascensao, Lumini-Oliveira et al.

2011). It is however important to note that the Δψ values above −200 mV in all

experimental groups do not seem to compromise ATP synthase flow or the

transport of ions and metabolites. In fact, the range of the Δψ is −120 to −220

mV. For instance, regarding the driving force for ATP generation, it has been

shown that the kinetics of the ATP synthase follow a sigmoid pattern in

response to Δψ, reaching saturation at approximately − 100 mV (Kaim and

Dimroth 1999).

6.2. Mitochondrial osmotic swelling during MPTP induction

In addition to their role in energy supply, mitochondria are also considered

determinant players in the establishment of cytosolic Ca2+ homeostasis,

59

uptaking and accumulation of Ca2+ in the matrix, in a process that is favored by

the electrochemical gradient formed across the inner mitochondrial membrane

(Gustafsson and Gottlieb 2008). However, mitochondria have a finite capability

to accumulate Ca2+ before undergoing Ca2+-dependent MPTP opening, and

thereafter to the release of pro-apoptotic proteins, which in turn results in

apoptosis (Ascensao, Lumini-Oliveira et al. 2011). In this regard, the study of

exercise in the context of MPTP modulation may assume an important clinical

relevance. Also, it has been described that, among others, a characteristic of

MPTP after Δψ loss, is the increased osmotic swelling amplitude induced by

Ca2+ in vitro (Gunter, Yule et al. 2004). Furthermore, endogenous Ca2+ levels in

matrix are greatly higher in oxidative tissue, limiting heart ability to uptake Ca2+

before MPTP induction (Picard, Csukly et al. 2008).

In the present study, only TM exercise per se was able to increase the

mitochondrial capability to accumulate Ca2+ after MPTP induction. However,

both protocols afforded protection against DOX impairments. Briefly, our results

showed that DOX per se decreased the mitochondrial Ca2+ tolerance

(SED+SAL vs. SED+DOX) and both exercise protocols counteract DOX-

induced impairments, possibly activating some defense mechanisms that might

contribute to prevent the increased DOX-induced MPTP opening susceptibility

(Marcil, Bourduas et al. 2006).

As MPTP is known to be formed/regulated by several proteins including ANT,

hexokinase VDAC, phosphate carrier or Cyp D (Ascensao, Lumini-Oliveira et al.

2011, Crompton 1999, Halestrap and Brenner 2003), it is possible that exercise

may positively modulate the expression and activity of those proteins. Also,

given the refereed close relationship between increased mitochondrial oxidative

stress and the susceptibility to MPTP induction (Kowaltowski, Castilho et al.

2001), it is possible that the up-regulation and modulation of some mechanisms

involving stress chaperones, as HSPs antioxidants or other defense systems

(Ascensao, Ferreira et al. 2007), as well as the decreased heart mitochondrial

free radical production found in rats undergoing regular exercise (Judge, Jang

et al. 2005) may contribute to these protective effects. Moreover, the possible

60

increased functionality of the phosphorylative system in general, and the ETC in

particular, induced by both exercise protocols may have some implications in

Ca2+ uptake capacity. However, to better understand this phenomenon, further

studies need to be addressed.

6.3 Oxidative stress markers

Prevailing hypotheses suggest that myocardial oxidative stress is a primary

event in DOX-induced cardiotoxicity and it is believed to initiate several of the

deleterious cellular events reported following DOX treatment (Zucchi and

Danesi 2003). In fact, at present the principal mechanism of DOX-induced

cardiotoxicity is believed to be increased mitochondrial oxidant production

leading to protease activation and induction of apoptosis (Ascensão, Magalhães

et al. 2006, Ascensao, Magalhaes et al. 2005, Chicco, Hydock et al. 2006,

Chicco, Schneider et al. 2005). Accordingly, oxidative injury of fatty acids at

subcellular level measured by increased levels of lipid peroxidation products

has been frequently reported following DOX exposure (for refs see Chicco,

Schneider et al. 2005).

The present results show that TM, but not FW per se was able to decrease

MDA level and increase -SH groups. In accordance to previous reports, DOX

induced a significant decrease in -SH, indicating increased disulfide linkages

from both proteins and GSH. As polyunsaturated fatty acids are considered

highly susceptible to ROS attack, the increased oxidative stress caused by DOX

led to peroxidative modification of lipid membranes affecting membrane integrity

and permeability, which leads to decoupled mitochondria, altering normal

mitochondrial respiratory function.

Myocardial antioxidant enzymes defend the heart against the damaging effects

of ROS and have been hypothesized to play an important role in exercise-

induced resistance to oxidative stress (Powers, Lennon et al. 2002) and in the

attenuation of DOX cardiotoxicity (Singal, Iliskovic et al. 1997). In particular,

61

some studies suggested that the presence of myocardial SOD might be

important for the prevention of DOX cardiotoxicity (Ascensao, Lumini-Oliveira et

al. 2011, Sarvazyan, Askari et al. 1995, Yen, Oberley et al. 1996). This is

reasonable, as SOD dismutates superoxide into H2O2, thereby providing the

first line of defense against DOX-induced oxidative stress. Furthermore,

increasing evidence suggest that myocardial HSP72 induction plays a pivotal

role in exercise-induced cardioprotection against oxidative stress (Powers,

Lennon et al. 2002, Powers, Locke et al. 2001, Taylor and Starnes 2003).

6.4 Meaning for exercise-induced cardioprotection in aging

Considering the present results in the context of exercise-induced

cardioprotection in advanced age, they can be interpreted as preliminary.

Indeed, it can be carefully speculated that the observed protective phenotype

caused by both chronic models of exercise against DOX can also be observed

in aged rats. In fact, Quindry et al. (2005) reported that aged rats submitted to

exercise training ameliorate cardiac hemodynamic response with significant

improvements in the apoptotic levels and signaling caused by IR injury.

Furthermore, the authors observed that trained old rats increased MnSOD

activity, which can be interpreted as a sign of cardiac mitochondrial adaptations

induced by chronic exercise in old rats compared to their young counterparts.

Similar results were found by Starnes et al. (2003), which suggest that, although

observing cardioprotective protein phenotype alterations with age, exercise can

enhance cardioprotection regardless of elderly. Furthermore, physical exercise

has the ability to positively modulate some gene expression associated with

improved heart function in aged rats. In fact, heart is known for its ability to

produce energy from fatty acids because of its important β-oxidation equipment,

which capacity is reduced with age (Starnes, Beyer et al. 1983). Confirming the

potential beneficial effects of physical exercise on cardiac metabolism in elderly,

Iemitsu et al. (2002) reported that exercise training improved the aging-induced

decreased expression of peroxisome proliferator-activated receptor, which

regulates genes related to fatty acid metabolism in the heart. Giving those

62

alterations reported in aged hearts, it is possible to speculate that the results of

the present work could also be observed in aged rats; affording protection and

mitigating the deleterious consequences associated with sub-chronic DOX

treatment schedules.

63

7. Conclusion

In summary, the data from the present work provide additional support about

the effect of two types of physical exercise (treadmill endurance training and

free-wheel voluntary physical activity) against heart mitochondrial dysfunction

induced by sub-chronic treatment of Doxorubicin (DOX). Our results showed us

that:

• Regarding mitochondrial respiratory function both types of exercise

reverted the effects induced by DOX on state 3, RCR and ADP/O.

Interestingly, free wheel voluntary physical activity was more efficient at

counteracting DOX-induced defects on RCR;

• Both types of exercise were able to counteract DOX-induced

impairments in mitochondrial transmembrane endpoints. Importantly, free

wheel voluntary physical activity was also more efficient at normalizing

DOX-induced increases in lag phase;

• Regarding mitochondrial osmotic swelling during MPTP induction, both

exercise protocols reverted DOX-induced impairments. In fact, both

types of exercise mitigated DOX-induced increases in swelling amplitude

and average swelling rate. However, once again free wheel voluntary

physical activity was more efficient at counteracting Ca2+-induced MPTP

induction

• Exercise protocols were able to revert DOX-induced increases in MDA

content and decrease in sulfhydryl groups.

The mechanisms by which treadmill endurance training and free-wheel

voluntary physical activity seems to confer additional protection against DOX

remain elusive and further studies need to be addressed in order to

comprehend the role of the different systems, such as those related to

mitochondria, in this process.

65

8. References

Adhihetty,   P.   J.,   V.   Ljubicic   and   D.   A.   Hood   (2007).   Effect   of   chronic   contractile   activity   on   SS   and   IMF  mitochondrial  apoptotic  susceptibility  in  skeletal  muscle.  Am  J  Physiol  Endocrinol  Metab,292(3):  E748-­‐755.  

Andreyev,   A.   Y.,   B.   Fahy   and   G.   Fiskum   (1998).   Cytochrome   c   release   from   brain   mitochondria   is  independent  of  the  mitochondrial  permeability  transition.  FEBS  Lett,439(3):  373-­‐376.  

Anversa,  P.,  B.  Hiler,  R.  Ricci,  G.  Guideri  and  G.  Olivetti  (1986).  Myocyte  cell  loss  and  myocyte  hypertrophy  in  the  aging  rat  heart.  J  Am  Coll  Cardiol,8(6):  1441-­‐1448.  

Ascensão,  A.  (2003).  Exercício  e  Stress  Oxidativo  Cardíaco.  Rev  Port  Cardiol,22(5).  

Ascensão,   A.   (2011).   Mitochondria   as   a   target   for   exercise-­‐induces   cardioprotection.   Current   Drug  Targets,12(6):  860-­‐871.  

Ascensao,   A.,   R.   Ferreira   and   J.   Magalhaes   (2007).   Exercise-­‐induced   cardioprotection-­‐-­‐biochemical,  morphological  and  functional  evidence  in  whole  tissue  and  isolated  mitochondria.   International  Journal  of  Cardiology,117(1):  16-­‐30.  

Ascensao,   A.,   R.   Ferreira   and   J.   Magalhaes   (2007).   Exercise-­‐induced   cardioprotection-­‐-­‐biochemical,  morphological  and  functional  evidence  in  whole  tissue  and  isolated  mitochondria.  Int  J  Cardiol,117(1):  16-­‐30.  

Ascensao,   A.,   R.   Ferreira,   P.   J.  Oliveira   and   J.  Magalhaes   (2006).   Effects   of   endurance   training   and   acute  Doxorubicin   treatment   on   rat   heart   mitochondrial   alterations   induced   by   in   vitro   anoxia-­‐reoxygenation.  Cardiovasc  Toxicol,6(3-­‐4):  159-­‐172.  

Ascensao,  A.,  J.  Lumini-­‐Oliveira,  N.  G.  Machado,  R.  M.  Ferreira,  I.  O.  Goncalves,  A.  C.  Moreira,  F.  Marques,  V.  A.  Sardao,  P.   J.  Oliveira  and   J.  Magalhaes   (2010).  Acute  exercise  protects  against  calcium-­‐induced  cardiac  mitochondrial  permeability   transition  pore  opening   in  doxorubicin-­‐treated  rats.  Clin  Sci   (Lond),120(1):  37-­‐49.  

Ascensao,  A.,  J.  Lumini-­‐Oliveira,  N.  G.  Machado,  R.  M.  Ferreira,  I.  O.  Goncalves,  A.  C.  Moreira,  F.  Marques,  V.  A.  Sardao,  P.   J.  Oliveira  and   J.  Magalhaes   (2011).  Acute  exercise  protects  against  calcium-­‐induced  cardiac  mitochondrial  permeability   transition  pore  opening   in  doxorubicin-­‐treated  rats.  Clin  Sci   (Lond),120(1):  37-­‐49.  

Ascensao,  A.,  J.  Lumini-­‐Oliveira,  P.  J.  Oliveira  and  J.  Magalhaes  (2011).  Mitochondria  as  a  target  for  exercise-­‐induced  cardioprotection.  Curr  Drug  Targets,12(6):  860-­‐871.  

Ascensao,   A.,   J.   Lumini-­‐Oliveira,   P.   J.   Oliveira   and   J.   Magalhaes   (2011).   Mitochondria   as   a   Target   for  Exercise-­‐Induced  Cardioprotection.  Current  Drug  Targets,12(6):  860-­‐871.  

Ascensão,  A.,  J.  Magalhães,  J.  Soares,  R.  Ferreira,  M.  Neuparth,  F.  Marques  and  J.  Duarte  (2006).  Endurance  exercise  training  attenuates  morphological  signs  of  cardiac  muscle  damage  induced  by  doxorubicin  in  male  mice.  

Ascensao,  A.,  J.  Magalhaes,  J.  Soares,  R.  Ferreira,  M.  Neuparth,  F.  Marques,  J.  Oliveira  and  J.  Duarte  (2005).  Endurance  training  attenuates  doxorubicin-­‐induced  cardiac  oxidative  damage  in  mice.   Int  J  Cardiol,100(3):  451-­‐460.  

Ascensao,   A.,   J.  Magalhaes,   J.  M.   Soares,   R.   Ferreira,  M.   J.   Neuparth,   F.  Marques,   P.   J.  Oliveira   and   J.   A.  Duarte  (2005).  Moderate  endurance  training  prevents  doxorubicin-­‐induced  in  vivo  mitochondriopathy  and  reduces  the  development  of  cardiac  apoptosis.  Am  J  Physiol  Heart  Circ  Physiol,289(2):  H722-­‐731.  

Ascensao,   A.,   J.  Magalhaes,   J.  M.   Soares,   R.   Ferreira,  M.   J.   Neuparth,   F.  Marques,   P.   J.  Oliveira   and   J.   A.  Duarte  (2006).  Endurance  training  limits  the  functional  alterations  of  rat  heart  mitochondria  submitted  to  in  vitro  anoxia-­‐reoxygenation.  Int  J  Cardiol,109(2):  169-­‐178.  

66

Ascensao,   A.,   P.   J.   Oliveira   and   J.   Magalhaes   (2012).   Exercise   as   a   beneficial   adjunct   therapy   during  Doxorubicin  treatment-­‐-­‐role  of  mitochondria  in  cardioprotection.  Int  J  Cardiol,156(1):  4-­‐10.  

Aydin,  C.,   E.   Ince,   S.  Koparan,   I.   T.  Cangul,  M.  Naziroglu  and  F.  Ak   (2007).  Protective  effects  of   long   term  dietary  restriction  on  swimming  exercise-­‐induced  oxidative  stress  in  the  liver,  heart  and  kidney  of  rat.  Cell  Biochem  Funct,25(2):  129-­‐137.  

Bagchi,   M.,   D.   Bagchi,   E.   B.   Patterson,   L.   Tang   and   S.   J.   Stohs   (1996).   Age-­‐related   changes   in   lipid  peroxidation  and  antioxidant  defense  in  Fischer  344  rats.  Ann  N  Y  Acad  Sci,793:  449-­‐452.  

Beckman,  K.  B.  and  B.  N.  Ames  (1998).  The  free  radical  theory  of  aging  matures.  Physiol  Rev,78(2):  547-­‐581.  

Bejma,   J.,   P.   Ramires   and   L.   L.   Ji   (2000).   Free   radical   generation   and   oxidative   stress   with   ageing   and  exercise:  differential  effects  in  the  myocardium  and  liver.  Acta  Physiol  Scand,169(4):  343-­‐351.  

Bernstein,   J.  D.,   J.  R.  Bucher  and  R.  Penniall   (1978).  Origin  of  mitochondrial  enzymes.  V.  The  polypeptide  character  and  the  biosynthesis  of  rat  liver  cytochrome  c  oxidase  polypeptides  by  mitochondria.  J  Bioenerg  Biomembr,10(1-­‐2):  59-­‐74.  

Berthiaume,   J.   M.,   P.   J.   Oliveira,   M.   W.   Fariss   and   K.   B.   Wallace   (2005).   Dietary   vitamin   E   decreases  doxorubicin-­‐induced   oxidative   stress   without   preventing   mitochondrial   dysfunction.   Cardiovasc  Toxicol,5(3):  257-­‐267.  

Berthiaume,  J.  M.  and  K.  B.  Wallace  (2007).  Adriamycin-­‐induced  oxidative  mitochondrial  cardiotoxicity.  Cell  Biol  Toxicol,23(1):  15-­‐25.  

Betik,   A.   C.   and   R.   T.   Hepple   (2008).   Determinants   of   VO2   max   decline   with   aging:   an   integrated  perspective.  Appl  Physiol  Nutr  Metab,33(1):  130-­‐140.  

Bhattacharya,   S.   K.,   J.   H.   Thakar,   P.   L.   Johnson   and   D.   R.   Shanklin   (1991).   Isolation   of   skeletal   muscle  mitochondria   from   hamsters   using   an   ionic   medium   containing   ethylenediaminetetraacetic   acid   and  nagarse.  Anal  Biochem,192(2):  344-­‐349.  

Bossy-­‐Wetzel,   E.,   M.   J.   Barsoum,   A.   Godzik,   R.   Schwarzenbacher   and   S.   A.   Lipton   (2003).   Mitochondrial  fission  in  apoptosis,  neurodegeneration  and  aging.  Curr  Opin  Cell  Biol,15(6):  706-­‐716.  

Bowles,   D.   K.,   R.   P.   Farrar   and   J.   W.   Starnes   (1992).   Exercise   training   improves   cardiac   function   after  ischemia  in  the  isolated,  working  rat  heart.  Am  J  Physiol,263(3  Pt  2):  H804-­‐809.  

Brand,  M.  D.  and  D.  G.  Nicholls  (2011).  Assessing  mitochondrial  dysfunction  in  cells.  Biochem  J,435(2):  297-­‐312.  

Bratic,   I.   and   A.   Trifunovic   (2010).   Mitochondrial   energy   metabolism   and   ageing.   Biochim   Biophys  Acta,1797(6-­‐7):  961-­‐967.  

Braunwald,  E.  and  M.  R.  Bristow  (2000).  Congestive  heart  failure:  fifty  years  of  progress.  Circulation,102(20  Suppl  4):  IV14-­‐23.  

Broekemeier,  K.  M.,  M.  E.  Dempsey  and  D.  R.  Pfeiffer  (1989).  Cyclosporin  A  is  a  potent  inhibitor  of  the  inner  membrane  permeability  transition  in  liver  mitochondria.  J  Biol  Chem,264(14):  7826-­‐7830.  

Buege,  J.  A.  and  S.  D.  Aust  (1978).  Microsomal  lipid  peroxidation.  Methods  Enzymol,52:  302-­‐310.  

Carvalho,   C.,   R.   X.   Santos,   S.   Cardoso,   S.   Correia,   P.   J.   Oliveira,   M.   S.   Santos   and   P.   I.   Moreira   (2009).  Doxorubicin:  the  good,  the  bad  and  the  ugly  effect.  Curr  Med  Chem,16(25):  3267-­‐3285.  

Chaudhary,   K.   R.,   H.   El-­‐Sikhry   and   J.   M.   Seubert   (2011).   Mitochondria   and   the   aging   heart.   J   Geriatr  Cardiol,8(3):  159-­‐167.  

Chen,  L.  and  A.  A.  Knowlton  (2011).  Mitochondrial  dynamics  in  heart  failure.  Congest  Heart  Fail,17(6):  257-­‐261.  

67

Chicco,  A.   J.,  D.  S.  Hydock,  C.  M.  Schneider  and  R.  Hayward  (2006).  Low-­‐intensity  exercise  training  during  doxorubicin  treatment  protects  against  cardiotoxicity.  J  Appl  Physiol,100(2):  519-­‐527.  

Chicco,  A.  J.,  C.  M.  Schneider  and  R.  Hayward  (2005).  Voluntary  exercise  protects  against  acute  doxorubicin  cardiotoxicity  in  the  isolated  perfused  rat  heart.  Am  J  Physiol  Regul  Integr  Comp  Physiol,289(2):  R424-­‐R431.  

Chicco,   A.   J.,   C.   M.   Schneider   and   R.   Hayward   (2006).   Exercise   training   attenuates   acute   doxorubicin-­‐induced  cardiac  dysfunction.  J  Cardiovasc  Pharmacol,47(2):  182-­‐189.  

Chiong,  M.,  Z.  V.  Wang,  Z.  Pedrozo,  D.  J.  Cao,  R.  Troncoso,  M.  Ibacache,  A.  Criollo,  A.  Nemchenko,  J.  A.  Hill  and   S.   Lavandero   (2011).   Cardiomyocyte   death:   mechanisms   and   translational   implications.   Cell   Death  Dis,2:  e244.  

Condorelli,   G.,   C.   Morisco,   G.   Stassi,   A.   Notte,   F.   Farina,   G.   Sgaramella,   A.   de   Rienzo,   R.   Roncarati,   B.  Trimarco   and   G.   Lembo   (1999).   Increased   cardiomyocyte   apoptosis   and   changes   in   proapoptotic   and  antiapoptotic  genes  bax  and  bcl-­‐2  during   left   ventricular  adaptations   to   chronic  pressure  overload   in   the  rat.  Circulation,99(23):  3071-­‐3078.  

Coore,   H.   G.,   R.  M.   Denton,   B.   R.  Martin   and   P.   J.   Randle   (1971).   Regulation   of   adipose   tissue   pyruvate  dehydrogenase  by  insulin  and  other  hormones.  Biochem  J,125(1):  115-­‐127.  

Crompton,  M.   (1999).   The  mitochondrial   permeability   transition   pore   and   its   role   in   cell   death.  Biochem  J,341  (  Pt  2):  233-­‐249.  

Dai,   D.   F.   and   P.   S.   Rabinovitch   (2009).   Cardiac   aging   in   mice   and   humans:   the   role   of   mitochondrial  oxidative  stress.  Trends  Cardiovasc  Med,19(7):  213-­‐220.  

Dai,  D.  F.,  P.  S.  Rabinovitch  and  Z.  Ungvari  (2012).  Mitochondria  and  cardiovascular  aging.  Circ  Res,110(8):  1109-­‐1124.  

Dai,   D.   F.,   L.   F.   Santana,  M.   Vermulst,   D.  M.   Tomazela,  M.   J.   Emond,  M.   J.  MacCoss,   K.   Gollahon,   G.  M.  Martin,   L.   A.   Loeb,   W.   C.   Ladiges   and   P.   S.   Rabinovitch   (2009).   Overexpression   of   catalase   targeted   to  mitochondria  attenuates  murine  cardiac  aging.  Circulation,119(21):  2789-­‐2797.  

Desler,  C.,  M.  L.  Marcker,  K.  K.  Singh  and  L.  J.  Rasmussen  (2011).  The  importance  of  mitochondrial  DNA  in  aging  and  cancer.  J  Aging  Res,2011:  407-­‐536.  

Emter,   C.   A.   and   D.   K.   Bowles   (2008).   Curing   the   cure:   utilizing   exercise   to   limit   cardiotoxicity.  Med   Sci  Sports  Exerc,40(5):  806-­‐807.  

Estabrook,  R.  W.  (1967).  Mitochondrial  respiratory  control  and  the  polarographic  measurement  of  ADP/O  ratios.  Meth.  Enzymol.,10:  41-­‐57.  

Fannin,  S.  W.,  E.  J.  Lesnefsky,  T.  J.  Slabe,  M.  O.  Hassan  and  C.  L.  Hoppel  (1999).  Aging  selectively  decreases  oxidative  capacity  in  rat  heart  interfibrillar  mitochondria.  Arch  Biochem  Biophys,372(2):  399-­‐407.  

Figueiredo,   P.   A.,  M.   P.  Mota,   H.   J.   Appell   and   J.   A.   Duarte   (2008).   The   role   of  mitochondria   in   aging   of  skeletal  muscle.  Biogerontology,9(2):  67-­‐84.  

Fortune,   J.  M.   and  N.  Osheroff   (2000).   Topoisomerase   II   as   a   target   for   anticancer  drugs:  when  enzymes  stop  being  nice.  Prog  Nucleic  Acid  Res  Mol  Biol,64:  221-­‐253.  

French,  J.  P.,  K.  L.  Hamilton,  J.  C.  Quindry,  Y.  Lee,  P.  A.  Upchurch  and  S.  K.  Powers  (2008).  Exercise-­‐induced  protection   against   myocardial   apoptosis   and   necrosis:   MnSOD,   calcium-­‐handling   proteins,   and   calpain.  FASEB  J,22(8):  2862-­‐2871.  

Gates,  P.  E.,  H.  Tanaka,  J.  Graves  and  D.  R.  Seals  (2003).  Left  ventricular  structure  and  diastolic  function  with  human  ageing.  Relation  to  habitual  exercise  and  arterial  stiffness.  Eur  Heart  J,24(24):  2213-­‐2220.  

Geng,  T.,  P.  Li,  M.  Okutsu,  X.  Yin,  J.  Kwek,  M.  Zhang  and  Z.  Yan  (2010).  PGC-­‐1alpha  plays  a  functional  role  in  exercise-­‐induced  mitochondrial   biogenesis   and   angiogenesis   but   not   fiber-­‐type   transformation   in  mouse  skeletal  muscle.  Am  J  Physiol  Cell  Physiol,298(3):  C572-­‐579.  

68

Glancy,   B.,   W.   T.   Willis,   D.   J.   Chess   and   R.   S.   Balaban   (2013).   Effect   of   calcium   on   the   oxidative  phosphorylation  cascade  in  skeletal  muscle  mitochondria.  Biochemistry,52(16):  2793-­‐2809.  

Golbidi,   S.   and   I.   Laher   (2011).  Molecular   mechanisms   in   exercise-­‐induced   cardioprotection.   Cardiol   Res  Pract,2011:  972807.  

Golbidi,  S.  and  I.  Laher  (2012).  Exercise  and  the  cardiovascular  system.  Cardiol  Res  Pract,2012:  210852.  

Goldspink,  D.  F.,  J.  G.  Burniston  and  L.  B.  Tan  (2003).  Cardiomyocyte  death  and  the  ageing  and  failing  heart.  Exp  Physiol,88(3):  447-­‐458.  

Gornall,  A.  G.,  C.   J.  Bardawill   and  M.  M.  David   (1949).  Determination  of   serum  proteins  by  means  of   the  biuret  reaction.  J  Biol  Chem,177(2):  751-­‐766.  

Green,  D.  R.  and  J.  C.  Reed  (1998).  Mitochondria  and  apoptosis.  Science,281(5381):  1309-­‐1312.  

Guarente,  L.  (2008).  Mitochondria-­‐-­‐a  nexus  for  aging,  calorie  restriction,  and  sirtuins?  Cell,132(2):  171-­‐176.  

Gunduz,  F.,  U.  K.  Senturk,  O.  Kuru,  B.  Aktekin  and  M.  R.  Aktekin  (2004).  The  effect  of  one  year's  swimming  exercise  on  oxidant  stress  and  antioxidant  capacity  in  aged  rats.  Physiol  Res,53(2):  171-­‐176.  

Gunter,  T.  E.,  D.  I.  Yule,  K.  K.  Gunter,  R.  A.  Eliseev  and  J.  D.  Salter  (2004).  Calcium  and  mitochondria.  FEBS  Lett,567(1):  96-­‐102.  

Gustafsson,   A.   B.   and   R.   A.   Gottlieb   (2008).   Heart   mitochondria:   gates   of   life   and   death.   Cardiovasc  Res,77(2):  334-­‐343.  

Gustafsson,  A.  B.  and  R.  A.  Gottlieb  (2009).  Autophagy  in  ischemic  heart  disease.  Circ  Res,104(2):  150-­‐158.  

Halestrap,   A.   P.   and   C.   Brenner   (2003).   The   adenine   nucleotide   translocase:   a   central   component   of   the  mitochondrial  permeability  transition  pore  and  key  player  in  cell  death.  Curr  Med  Chem,10(16):  1507-­‐1525.  

Hambrecht,   R.,   E.   Fiehn,   C.  Weigl,   S.   Gielen,   C.   Hamann,   R.   Kaiser,   J.   Yu,   V.   Adams,   J.   Niebauer   and   G.  Schuler  (1998).  Regular  physical  exercise  corrects  endothelial  dysfunction  and  improves  exercise  capacity  in  patients  with  chronic  heart  failure.  Circulation,98(24):  2709-­‐2715.  

Hamilton,  K.  L.,  J.  L.  Staib,  T.  Phillips,  A.  Hess,  S.  L.  Lennon  and  S.  K.  Powers  (2003).  Exercise,  antioxidants,  and  HSP72:  protection  against  myocardial  ischemia/reperfusion.  Free  Radic  Biol  Med,34(7):  800-­‐809.  

Harman,  D.  (1956).  Aging:  a  theory  based  on  free  radical  and  radiation  chemistry.  J  Gerontol(11): 298-­‐300  Harris,  E.  D.  (1992).  Regulation  of  antioxidant  enzymes.  FASEB  J,6(9):  2675-­‐2683.  

Harris,  M.  B.  and  J.  W.  Starnes  (2001).  Effects  of  body  temperature  during  exercise  training  on  myocardial  adaptations.  Am  J  Physiol  Heart  Circ  Physiol,280(5):  H2271-­‐2280.  

Haunstetter,   A.   and   S.   Izumo   (1998).   Apoptosis:   basic   mechanisms   and   implications   for   cardiovascular  disease.  Circ  Res,82(11):  1111-­‐1129.  

Hayward,  R.,  C.  Y.  Lien,  B.  T.   Jensen,  D.  S.  Hydock  and  C.  M.  Schneider   (2012).  Exercise   training  mitigates  anthracycline-­‐induced   chronic   cardiotoxicity   in   a   juvenile   rat  model.  Pediatric   blood  &   cancer,59(1):   149-­‐154.  

Hayward,  R.,  C.  Y.  Lien,  B.  T.   Jensen,  D.  S.  Hydock  and  C.  M.  Schneider   (2012).  Exercise   training  mitigates  anthracycline-­‐induced  chronic  cardiotoxicity  in  a  juvenile  rat  model.  Pediatr  Blood  Cancer,59(1):  149-­‐154.  

Harman,  D.  (1972).  The  biologic  clock:  the  mitochondria?  J  Am  Geriatr  Soc(20): 145-­‐7  Holloszy,   J.   O.,   L.   B.   Oscai,   I.   J.   Don   and   P.   A.   Mole   (1970).   Mitochondrial   citric   acid   cycle   and   related  enzymes:  adaptive  response  to  exercise.  Biochem  Biophys  Res  Commun,40(6):  1368-­‐1373.  

Honda,  H.  M.,  P.  Korge  and  J.  N.  Weiss  (2005).  Mitochondria  and  ischemia/reperfusion  injury.  Ann  N  Y  Acad  Sci,1047:  248-­‐258.  

69

Hong,  H.  and  P.  Johnson  (1995).  Antioxidant  enzyme  activities  and  lipid  peroxidation  levels  in  exercised  and  hypertensive  rat  tissues.  Int  J  Biochem  Cell  Biol,27(9):  923-­‐931.  

Hood,   D.   A.,   A.   Balaban,   M.   K.   Connor,   E.   E.   Craig,   M.   L.   Nishio,   M.   Rezvani   and   M.   Takahashi   (1994).  Mitochondrial  biogenesis  in  striated  muscle.  Can  J  Appl  Physiol,19(1):  12-­‐48.  

Hoppins,  S.,   F.  Edlich,  M.  M.  Cleland,  S.  Banerjee,   J.  M.  McCaffery,  R.   J.  Youle  and   J.  Nunnari   (2011).  The  soluble   form  of   Bax   regulates  mitochondrial   fusion   via  MFN2  homotypic   complexes.  Mol   Cell,41(2):   150-­‐160.  

Hotta,   H.   and   S.   Uchida   (2010).   Aging   of   the   autonomic   nervous   system   and   possible   improvements   in  autonomic   activity   using   somatic   afferent   stimulation.   Geriatrics   &   Gerontology   International,10:   S127-­‐S136.  

Hu,  M.  L.  (1990).  Measurement  of  protein  thiol  groups  and  GSH  in  plasma.  Methods  in  Enzymology.  Parker,  L.  San  Diego,  CA:  Academic:  380-­‐385.  

Huang,  C.,  X.  Zhang,  J.  M.  Ramil,  S.  Rikka,  L.  Kim,  Y.  Lee,  N.  A.  Gude,  P.  A.  Thistlethwaite,  M.  A.  Sussman,  R.  A.  Gottlieb  and  A.  B.  Gustafsson  (2010).  Juvenile  exposure  to  anthracyclines  impairs  cardiac  progenitor  cell  function  and  vascularization  resulting   in  greater  susceptibility   to  stress-­‐induced  myocardial   injury   in  adult  mice.  Circulation,121(5):  675-­‐683.  

Hydock,  D.   S.,   C.   Y.   Lien,   B.   T.   Jensen,   T.   L.   Parry,   C.  M.   Schneider   and  R.  Hayward   (2012).   Rehabilitative  exercise   in   a   rat   model   of   doxorubicin   cardiotoxicity.   Experimental   biology   and  medicine,237(12):   1483-­‐1492.  

Hydock,  D.  S.,  C.  Y.  Lien,  C.  M.  Schneider  and  R.  Hayward  (2008).  Exercise  preconditioning  protects  against  doxorubicin-­‐induced  cardiac  dysfunction.  Med  Sci  Sports  Exerc,40(5):  808-­‐817.  

Iemitsu,  M.,  T.  Miyauchi,  S.  Maeda,  T.  Tanabe,  M.  Takanashi,  Y.  Irukayama-­‐Tomobe,  S.  Sakai,  H.  Ohmori,  M.  Matsuda  and  I.  Yamaguchi  (2002).  Aging-­‐induced  decrease  in  the  PPAR-­‐alpha  level  in  hearts  is  improved  by  exercise  training.  American  Journal  of  Physiology-­‐Heart  and  Circulatory  Physiology,283(5):  H1750-­‐H1760.  

Ji,  L.  L.  (1993).  Antioxidant  enzyme  response  to  exercise  and  aging.  Med  Sci  Sports  Exerc,25(2):  225-­‐231.  

Ji,  L.  L.,  D.  Dillon  and  E.  Wu  (1991).  Myocardial  aging:  antioxidant  enzyme  systems  and  related  biochemical  properties.  Am  J  Physiol,261(2  Pt  2):  R386-­‐392.  

Ji,   L.   L.,   C.   Leeuwenburgh,   S.   Leichtweis,  M.   Gore,   R.   Fiebig,   J.   Hollander   and   J.   Bejma   (1998).   Oxidative  stress  and  aging.  Role  of  exercise  and  its  influences  on  antioxidant  systems.  Ann  N  Y  Acad  Sci,854:  102-­‐117.  

Judge,  S.,  Y.  M.  Jang,  A.  Smith,  T.  Hagen  and  C.  Leeuwenburgh  (2005).  Age-­‐associated  increases  in  oxidative  stress   and   antioxidant   enzyme   activities   in   cardiac   interfibrillar   mitochondria:   implications   for   the  mitochondrial  theory  of  aging.  FASEB  J,19(3):  419-­‐421.  

Judge,  S.,  Y.  M.  Jang,  A.  Smith,  C.  Selman,  T.  Phillips,  J.  R.  Speakman,  T.  Hagen  and  C.  Leeuwenburgh  (2005).  Exercise   by   lifelong   voluntary   wheel   running   reduces   subsarcolemmal   and   interfibrillar   mitochondrial  hydrogen  peroxide  production  in  the  heart.  Am  J  Physiol  Regul  Integr  Comp  Physiol,289(6):  R1564-­‐1572.  

Judge,  S.  and  C.  Leeuwenburgh  (2007).  Cardiac  mitochondrial  bioenergetics,  oxidative  stress,  and  aging.  Am  J  Physiol  Cell  Physiol,292(6):  C1983-­‐1992.  

Jung,  K.  and  R.  Reszka   (2001).  Mitochondria  as   subcellular   targets   for  clinically  useful  anthracyclines.  Adv  Drug  Deliv  Rev,49(1-­‐2):  87-­‐105.  

Kaim,   G.   and   P.   Dimroth   (1999).   ATP   synthesis   by   F-­‐type   ATP   synthase   is   obligatorily   dependent   on   the  transmembrane  voltage.  EMBO  J,18(15):  4118-­‐4127.  

Kanter,  M.  M.,  R.  L.  Hamlin,  D.  V.  Unverferth,  H.  W.  Davis  and  A.  J.  Merola  (1985).  Effect  of  exercise  training  on  antioxidant  enzymes  and  cardiotoxicity  of  doxorubicin.  J  Appl  Physiol,59(4):  1298-­‐1303.  

70

Kappagoda,  T.  and  E.  A.  Amsterdam  (2012).  Exercise  and  heart  failure  in  the  elderly.  Heart  Fail  Rev,17(4-­‐5):  635-­‐662.  

Kavazis,  A.  N.  (2009).  Exercise  preconditioning  of  the  myocardium.  Sports  Med,39(11):  923-­‐935.  

Kavazis,   A.   N.,   S.   Alvarez,   E.   Talbert,   Y.   Lee   and   S.   K.   Powers   (2009).   Exercise   training   induces   a  cardioprotective  phenotype  and  alterations  in  cardiac  subsarcolemmal  and  intermyofibrillar  mitochondrial  proteins.  Am  J  Physiol  Heart  Circ  Physiol,297(1):  H144-­‐152.  

Kavazis,  A.  N.,  J.  M.  McClung,  D.  A.  Hood  and  S.  K.  Powers  (2008).  Exercise  induces  a  cardiac  mitochondrial  phenotype  that  resists  apoptotic  stimuli.  Am  J  Physiol  Heart  Circ  Physiol,294(2):  H928-­‐935.  

Kavazis,  A.  N.,  A.  J.  Smuder,  K.  Min,  N.  Tumer  and  S.  K.  Powers  (2010).  Short-­‐term  exercise  training  protects  against  doxorubicin-­‐induced  cardiac  mitochondrial  damage  independent  of  HSP72.  Am  J  Physiol  Heart  Circ  Physiol,299(5):  H1515-­‐1524.  

Kiyomiya,  K.,  S.  Matsuo  and  M.  Kurebe  (2001).  Mechanism  of  specific  nuclear  transport  of  adriamycin:  the  mode  of  nuclear  translocation  of  adriamycin-­‐proteasome  complex.  Cancer  Res,61(6):  2467-­‐2471.  

Kokkinos,  P.,  J.  Myers,  C.  Faselis,  D.  B.  Panagiotakos,  M.  Doumas,  A.  Pittaras,  A.  Manolis,  J.  P.  Kokkinos,  P.  Karasik,  M.  Greenberg,  V.  Papademetriou  and  R.  Fletcher   (2010).  Exercise  capacity  and  mortality   in  older  men:  a  20-­‐year  follow-­‐up  study.  Circulation,122(8):  790-­‐797.  

Koltai,  E.,  N.  Hart,  A.  W.  Taylor,  S.  Goto,  J.  K.  Ngo,  K.  J.  Davies  and  Z.  Radak  (2012).  Age-­‐associated  declines  in  mitochondrial   biogenesis   and   protein   quality   control   factors   are  minimized   by   exercise   training.  Am   J  Physiol  Regul  Integr  Comp  Physiol,303(2):  R127-­‐134.  

Kowaltowski,   A.   J.,   R.   F.   Castilho   and   A.   E.   Vercesi   (2001).   Mitochondrial   permeability   transition   and  oxidative  stress.  FEBS  Lett,495(1-­‐2):  12-­‐15.  

Kregel,   K.   C.   and   P.   L.   Moseley   (1996).   Differential   effects   of   exercise   and   heat   stress   on   liver   HSP70  accumulation  with  aging.  J  Appl  Physiol,80(2):  547-­‐551.  

Kroemer,   G.,   L.   Galluzzi   and   C.   Brenner   (2007).  Mitochondrial  membrane   permeabilization   in   cell   death.  Physiol  Rev,87(1):  99-­‐163.  

Lagranha,   C.   J.,   A.   Deschamps,   A.   Aponte,   C.   Steenbergen   and   E.  Murphy   (2010).   Sex   differences   in   the  phosphorylation   of   mitochondrial   proteins   result   in   reduced   production   of   reactive   oxygen   species   and  cardioprotection  in  females.  Circ  Res,106(11):  1681-­‐1691.  

Lakatta,  E.  G.  (2002).  Age-­‐associated  cardiovascular  changes  in  health:  impact  on  cardiovascular  disease  in  older  persons.  Heart  Fail  Rev,7(1):  29-­‐49.  

Landmesser,   U.   and   H.   Drexler   (2005).   The   clinical   significance   of   endothelial   dysfunction.   Curr   Opin  Cardiol,20(6):  547-­‐551.  

Lebrecht,  D.,  B.  Setzer,  U.  P.  Ketelsen,  J.  Haberstroh  and  U.  A.  Walker  (2003).  Time-­‐dependent  and  tissue-­‐specific   accumulation   of   mtDNA   and   respiratory   chain   defects   in   chronic   doxorubicin   cardiomyopathy.  Circulation,108(19):  2423-­‐2429.  

Lee,  H.  C.  and  Y.  H.  Wei  (2007).  Oxidative  stress,  mitochondrial  DNA  mutation,  and  apoptosis  in  aging.  Exp  Biol  Med  (Maywood),232(5):  592-­‐606.  

Lee,  S.,  S.  Y.  Jeong,  W.  C.  Lim,  S.  Kim,  Y.  Y.  Park,  X.  Sun,  R.  J.  Youle  and  H.  Cho  (2007).  Mitochondrial  fission  and  fusion  mediators,  hFis1  and  OPA1,  modulate  cellular  senescence.  J  Biol  Chem,282(31):  22977-­‐22983.  

Lehman,   J.   J.,   P.   M.   Barger,   A.   Kovacs,   J.   E.   Saffitz,   D.   M.   Medeiros   and   D.   P.   Kelly   (2000).   Peroxisome  proliferator-­‐activated   receptor   gamma   coactivator-­‐1   promotes   cardiac   mitochondrial   biogenesis.   J   Clin  Invest,106(7):  847-­‐856.  

Lenaz,  G.  (1998).  Role  of  mitochondria  in  oxidative  stress  and  ageing.  Biochim  Biophys  Acta,1366(1-­‐2):  53-­‐67.  

71

Lennon,  S.  L.,  J.  C.  Quindry,  J.  P.  French,  S.  Kim,  J.  L.  Mehta  and  S.  K.  Powers  (2004).  Exercise  and  myocardial  tolerance  to  ischaemia-­‐reperfusion.  Acta  Physiol  Scand,182(2):  161-­‐169.  

Leung,   F.   P.,   L.   M.   Yung,   I.   Laher,   X.   Yao,   Z.   Y.   Chen   and   Y.   Huang   (2008).   Exercise,   vascular   wall   and  cardiovascular  diseases:  an  update  (Part  1).  Sports  Med,38(12):  1009-­‐1024.  

Li,   L.,   C.   Muhlfeld,   B.   Niemann,   R.   Pan,   R.   Li,   D.   Hilfiker-­‐Kleiner,   Y.   Chen   and   S.   Rohrbach   (2011).  Mitochondrial  biogenesis  and  PGC-­‐1alpha  deacetylation  by  chronic  treadmill  exercise:  differential  response  in  cardiac  and  skeletal  muscle.  Basic  Res  Cardiol,106(6):  1221-­‐1234.  

Liesa,  M.,  M.   Palacin   and   A.   Zorzano   (2009).  Mitochondrial   dynamics   in  mammalian   health   and   disease.  Physiol  Rev,89(3):  799-­‐845.  

Ljubicic,   V.,   K.   J.   Menzies   and   D.   A.   Hood   (2010).   Mitochondrial   dysfunction   is   associated   with   a   pro-­‐apoptotic  cellular  environment  in  senescent  cardiac  muscle.  Mech  Ageing  Dev,131(2):  79-­‐88.  

Logue,   S.   E.,   A.   B.   Gustafsson,   A.   Samali   and   R.   A.   Gottlieb   (2005).   Ischemia/reperfusion   injury   at   the  intersection  with  cell  death.  J  Mol  Cell  Cardiol,38(1):  21-­‐33.  

Lopez-­‐Lluch,  G.,  P.  M.  Irusta,  P.  Navas  and  R.  de  Cabo  (2008).  Mitochondrial  biogenesis  and  healthy  aging.  Exp  Gerontol,43(9):  813-­‐819.  

Lumini-­‐Oliveira,   J.,   J.   Magalhaes,   C.   V.   Pereira,   A.   C.   Moreira,   P.   J.   Oliveira   and   A.   Ascensao   (2011).  Endurance  training  reverts  heart  mitochondrial  dysfunction,  permeability  transition  and  apoptotic  signaling  in  long-­‐term  severe  hyperglycemia.  Mitochondrion,11(1):  54-­‐63.  

Marcil,  M.,   K.   Bourduas,   A.   Ascah   and   Y.   Burelle   (2006).   Exercise   training   induces   respiratory   substrate-­‐specific  decrease  in  Ca2+-­‐induced  permeability  transition  pore  opening  in  heart  mitochondria.  Am  J  Physiol  Heart  Circ  Physiol,290(4):  H1549-­‐1557.  

Meng,  R.  R.  Y.  T.  W.  J.  C.  Q.  (2007).  Age-­‐related  changes  in  mitochondrial  function  and  antioxidative  enzyme  activity  in  fischer  344  rats.  Mechanisms  of  Ageing  and  Development(128): 286-­‐92  Morley,  J.  E.  and  S.  S.  Reese  (1989).  Clinical  implications  of  the  aging  heart.  Am  J  Med,86(1):  77-­‐86.  

Moslehi,   J.,   R.   A.   DePinho   and   E.   Sahin   (2012).   Telomeres   and   mitochondria   in   the   aging   heart.   Circ  Res,110(9):  1226-­‐1237.  

Murphy,  M.  P.   and  M.  D.  Brand   (1988).  Membrane-­‐potential-­‐dependent   changes   in   the   stoichiometry  of  charge  translocation  by  the  mitochondrial  electron  transport  chain.  Eur  J  Biochem,173(3):  637-­‐644.  

Narula,  J.,  N.  Haider,  R.  Virmani,  T.  G.  DiSalvo,  F.  D.  Kolodgie,  R.  J.  Hajjar,  U.  Schmidt,  M.  J.  Semigran,  G.  W.  Dec  and  B.  A.  Khaw  (1996).  Apoptosis  in  myocytes  in  end-­‐stage  heart  failure.  N  Engl  J  Med,335(16):  1182-­‐1189.  

Navarro,  A.,  C.  Gomez,  J.  M.  Lopez-­‐Cepero  and  A.  Boveris  (2004).  Beneficial  effects  of  moderate  exercise  on  mice   aging:   survival,   behavior,   oxidative   stress,   and  mitochondrial   electron   transfer.   Am   J   Physiol   Regul  Integr  Comp  Physiol,286(3):  R505-­‐511.  

North,   B.   J.   and   D.   A.   Sinclair   (2012).   The   intersection   between   aging   and   cardiovascular   disease.   Circ  Res,110(8):  1097-­‐1108.  

Nulton-­‐Persson,  A.  C.  and  L.  I.  Szweda  (2001).  Modulation  of  mitochondrial  function  by  hydrogen  peroxide.  J  Biol  Chem,276(26):  23357-­‐23361.  

O'Brien,  I.  A.,  P.  O'Hare  and  R.  J.  Corrall  (1986).  Heart  rate  variability  in  healthy  subjects:  effect  of  age  and  the  derivation  of  normal  ranges  for  tests  of  autonomic  function.  Br  Heart  J,55(4):  348-­‐354.  

Oliveira,   P.   J.   and   K.   B.   Wallace   (2006).   Depletion   of   adenine   nucleotide   translocator   protein   in   heart  mitochondria  from  doxorubicin-­‐treated  rats-­‐-­‐relevance  for  mitochondrial  dysfunction.  Toxicology,220(2-­‐3):  160-­‐168.  

72

Olivetti,  G.,  R.  Abbi,  F.  Quaini,  J.  Kajstura,  W.  Cheng,  J.  A.  Nitahara,  E.  Quaini,  C.  Di  Loreto,  C.  A.  Beltrami,  S.  Krajewski,   J.  C.  Reed  and  P.  Anversa   (1997).  Apoptosis   in   the   failing  human  heart.  N  Engl   J  Med,336(16):  1131-­‐1141.  

Oxenham,  H.  and  N.  Sharpe  (2003).  Cardiovascular  aging  and  heart  failure.  Eur  J  Heart  Fail,5(4):  427-­‐434.  

Pashkow,  F.  J.  (2011).  Oxidative  Stress  and  Inflammation  in  Heart  Disease:  Do  Antioxidants  Have  a  Role  in  Treatment  and/or  Prevention?  Int  J  Inflam,2011:  514623.  

Pereira,  G.  C.,  S.  P.  Pereira,  C.  V.  Pereira,  J.  A.  Lumini,  J.  Magalhaes,  A.  Ascensao,  M.  S.  Santos,  A.  J.  Moreno  and  P.   J.  Oliveira   (2012).  Mitochondrionopathy  phenotype   in  doxorubicin-­‐treated  Wistar   rats  depends  on  treatment  protocol  and  is  cardiac-­‐specific.  PLoS  One,7(6):  e38867.  

Picard,  M.,  K.  Csukly,  M.  E.  Robillard,  R.  Godin,  A.  Ascah,  C.  Bourcier-­‐Lucas  and  Y.  Burelle  (2008).  Resistance  to   Ca2+-­‐induced   opening   of   the   permeability   transition   pore   differs   in  mitochondria   from   glycolytic   and  oxidative  muscles.  Am  J  Physiol  Regul  Integr  Comp  Physiol,295(2):  R659-­‐668.  

Pohjoismaki,   J.   L.,   T.   Boettger,   Z.   Liu,   S.   Goffart,  M.   Szibor   and   T.   Braun   (2012).   Oxidative   stress   during  mitochondrial  biogenesis  compromises  mtDNA  integrity  in  growing  hearts  and  induces  a  global  DNA  repair  response.  Nucleic  Acids  Res,40(14):  6595-­‐6607.  

Polla,   B.   S.,   S.   Kantengwa,   D.   Francois,   S.   Salvioli,   C.   Franceschi,   C.   Marsac   and   A.   Cossarizza   (1996).  Mitochondria   are   selective   targets   for   the   protective   effects   of   heat   shock   against   oxidative   injury.  Proc  Natl  Acad  Sci  U  S  A,93(13):  6458-­‐6463.  

Powers,  S.  K.,  H.  A.  Demirel,  H.  K.  Vincent,  J.  S.  Coombes,  H.  Naito,  K.  L.  Hamilton,  R.  A.  Shanely  and  J.  Jessup  (1998).   Exercise   training   improves  myocardial   tolerance   to   in   vivo   ischemia-­‐reperfusion   in   the   rat.  Am   J  Physiol,275(5  Pt  2):  R1468-­‐1477.  

Powers,   S.   K.,   S.   L.   Lennon,   J.   Quindry   and   J.   L.  Mehta   (2002).   Exercise   and   cardioprotection.  Curr   Opin  Cardiol,17(5):  495-­‐502.  

Powers,   S.   K.,   Locke   and   H.   A.   Demirel   (2001).   Exercise,   heat   shock   proteins,   and  myocardial   protection  from  I-­‐R  injury.  Med  Sci  Sports  Exerc,33(3):  386-­‐392.  

Powers,   S.   K.,   J.   Quindry   and   K.   Hamilton   (2004).   Aging,   exercise,   and   cardioprotection.   Ann   N   Y   Acad  Sci,1019:  462-­‐470.  

Powers,  S.  K.,  J.  C.  Quindry  and  A.  N.  Kavazis  (2008).  Exercise-­‐induced  cardioprotection  against  myocardial  ischemia-­‐reperfusion  injury.  Free  Radic  Biol  Med,44(2):  193-­‐201.  

Pritsos,   C.  A.   and   J.  Ma   (2000).   Basal   and  drug-­‐induced  antioxidant   enzyme  activities   correlate  with   age-­‐dependent  doxorubicin  oxidative  toxicity.  Chem  Biol  Interact,127(1):  1-­‐11.  

Quindry,   J.,   J.   French,   K.   Hamilton,   Y.   Lee,   J.   L.   Mehta   and   S.   Powers   (2005).   Exercise   training   provides  cardioprotection   against   ischemia-­‐reperfusion   induced   apoptosis   in   young   and   old   animals.   Exp  Gerontol,40(5):  416-­‐425.  

Rabinovitch,  P.  (2012).  Mitochondria  and  cardiovascular  aging.  American  Heart  Association,  Inc.(110): 1109-­‐24.  

Rajendra  Acharya,  U.,  K.  Paul  Joseph,  N.  Kannathal,  C.  M.  Lim  and  J.  S.  Suri  (2006).  Heart  rate  variability:  a  review.  Med  Biol  Eng  Comput,44(12):  1031-­‐1051.  

Rao,  G.,  E.  Xia  and  A.  Richardson   (1990).  Effect  of  age  on   the  expression  of  antioxidant  enzymes   in  male  Fischer  F344  rats.  Mech  Ageing  Dev,53(1):  49-­‐60.  

Reynolds,   D.   W.   (2004).   Understanding   the   aging   cardiovascular   system.   Geriatrics   and   Gerontology  International(4): 298–303.  

 

73

Reznick,  R.  M.,  H.  Zong,  J.  Li,  K.  Morino,  I.  K.  Moore,  H.  J.  Yu,  Z.  X.  Liu,  J.  Dong,  K.  J.  Mustard,  S.  A.  Hawley,  D.  Befroy,  M.  Pypaert,  D.  G.  Hardie,  L.  H.  Young  and  G.  I.  Shulman  (2007).  Aging-­‐associated  reductions  in  AMP-­‐activated  protein  kinase  activity  and  mitochondrial  biogenesis.  Cell  Metab,5(2):  151-­‐156.  

Rodgers,  J.  T.,  C.  Lerin,  W.  Haas,  S.  P.  Gygi,  B.  M.  Spiegelman  and  P.  Puigserver  (2005).  Nutrient  control  of  glucose  homeostasis  through  a  complex  of  PGC-­‐1alpha  and  SIRT1.  Nature,434(7029):  113-­‐118.  

Santos,  D.  L.,  A.  J.  Moreno,  R.  L.  Leino,  M.  K.  Froberg  and  K.  B.  Wallace  (2002).  Carvedilol  protects  against  doxorubicin-­‐induced  mitochondrial  cardiomyopathy.  Toxicol  Appl  Pharmacol,185(3):  218-­‐227.  

Sardao,   V.   A.,   P.   J.   Oliveira,   J.   Holy,   C.   R.   Oliveira   and   K.   B.   Wallace   (2009).   Morphological   alterations  induced   by   doxorubicin   on   H9c2   myoblasts:   nuclear,   mitochondrial,   and   cytoskeletal   targets.   Cell   Biol  Toxicol,25(3):  227-­‐243.  

Sardao,  V.  A.,  S.  L.  Pereira  and  P.  J.  Oliveira  (2008).  Drug-­‐induced  mitochondrial  dysfunction  in  cardiac  and  skeletal  muscle  injury.  Expert  Opin  Drug  Saf,7(2):  129-­‐146.  

Sarvazyan,  N.  A.,  A.  Askari  and  W.  H.  Huang  (1995).  Effects  of  doxorubicin  on  cardiomyocytes  with  reduced  level  of  superoxide  dismutase.  Life  Sci,57(10):  1003-­‐1010.  

Searls,  Y.  M.,  I.  V.  Smirnova,  B.  R.  Fegley  and  L.  Stehno-­‐Bittel  (2004).  Exercise  attenuates  diabetes-­‐induced  ultrastructural  changes  in  rat  cardiac  tissue.  Med  Sci  Sports  Exerc,36(11):  1863-­‐1870.  

Seeley,  R.,  T.  Stephens  and  P.  Tate  (2005).  Anatomy  &  Physiology,  Martin  J.  Lange.  

Sheu,   S.   (2009).  Morphological  Dynamics  of  Mitochondria-­‐  A   Special   Emphasis  on  Cardiac  Muscle  Cells.   J  Mol  Cell  Cardiol,46(6).  

Shioi,  T.  and  Y.  Inuzuka  (2012).  Aging  as  a  substrate  of  heart  failure.  J  Cardiol: 423-­‐8.  Singal,   P.   K.,   N.   Iliskovic,   T.   Li   and   D.   Kumar   (1997).   Adriamycin   cardiomyopathy:   pathophysiology   and  prevention.  FASEB  J,11(12):  931-­‐936.  

Souza,   T.   M.   S.   B.   F.   V.   A.   (2012).   Reprodutibilidade   do   VO2Máx   estimado   na   corrida   pela   frequência  cardíaca  e  consumo  de  oxigênio  de  reserva.  Rev.  bras.  educ.  fís.  esporte  26(1).  

Spina,  R.  J.,  M.  J.  Turner  and  A.  A.  Ehsani  (1998).  Beta-­‐adrenergic-­‐mediated  improvement  in  left  ventricular  function  by  exercise  training  in  older  men.  Am  J  Physiol,274(2  Pt  2):  H397-­‐404.  

Starnes,  J.  W.,  B.  D.  Barnes  and  M.  E.  Olsen  (2007).  Exercise  training  decreases  rat  heart  mitochondria  free  radical  generation  but  does  not  prevent  Ca2+-­‐induced  dysfunction.  J  Appl  Physiol,102(5):  1793-­‐1798.  

Starnes,  J.  W.,  R.  E.  Beyer  and  D.  W.  Edington  (1983).  Myocardial  adaptations  to  endurance  exercise  in  aged  rats.  Am  J  Physiol,245(4):  H560-­‐566.  

Starnes,  J.  W.,  D.  K.  Bowles  and  K.  S.  Seiler  (1997).  Myocardial  injury  after  hypoxia  in  immature,  adult  and  aged  rats.  Aging  (milano),9(4):  268-­‐276.  

Starnes,  J.  W.  and  R.  P.  Taylor  (2007).  Exercise-­‐induced  cardioprotection:  endogenous  mechanisms.  Med  Sci  Sports  Exerc,39(9):  1537-­‐1543.  

Starnes,  J.  W.,  R.  P.  Taylor  and  Y.  Park  (2003).  Exercise  improves  postischemic  function  in  aging  hearts.  Am  J  Physiol  Heart  Circ  Physiol,285(1):  H347-­‐351.  

Stock,  D.,  A.  G.  Leslie  and  J.  E.  Walker  (1999).  Molecular  architecture  of  the  rotary  motor  in  ATP  synthase.  Science,286(5445):  1700-­‐1705.  

Strait,   J.   B.   and   E.   G.   Lakatta   (2012).   Aging-­‐Associated   Cardiovascular   Changes   and   Their   Relationship   to  Heart  Failure.  Heart  Failure  Clinics,8(1):  143-­‐+.  

Strait,  J.  B.  and  E.  G.  Lakatta  (2012).  Aging-­‐associated  cardiovascular  changes  and  their  relationship  to  heart  failure.  Heart  Fail  Clin,8(1):  143-­‐164.  

74

Swain,   S.   M.,   F.   S.   Whaley   and   M.   S.   Ewer   (2003).   Congestive   heart   failure   in   patients   treated   with  doxorubicin:  a  retrospective  analysis  of  three  trials.  Cancer,97(11):  2869-­‐2879.  

T.  Shioi,  Y.  I.  (2012).  Aging  as  a  substrate  of  heart  failure.  Journal  of  Cardiology: 423-­‐8.  Taylor,  A.  H.,  N.  T.  Cable,  G.  Faulkner,  M.  Hillsdon,  M.  Narici  and  A.  K.  van  Der  Bij  (2004).  Physical  activity  and   older   adults:   a   review   of   health   benefits   and   the   effectiveness   of   interventions.   Journal   of   Sports  Sciences,22(8):  703-­‐725.  

Taylor,   R.   P.   and   J.   W.   Starnes   (2003).   Age,   cell   signalling   and   cardioprotection.   Acta   Physiologica  Scandinavica,178(2):  107-­‐116.  

Tian,   L.,   Q.   Cai   and   H.   Wei   (1998).   Alterations   of   antioxidant   enzymes   and   oxidative   damage   to  macromolecules  in  different  organs  of  rats  during  aging.  Free  Radic  Biol  Med,24(9):  1477-­‐1484.  

van  Acker,  S.  A.,  K.  Kramer,  E.  E.  Voest,  J.  A.  Grimbergen,  J.  Zhang,  W.  J.  van  der  Vijgh  and  A.  Bast  (1996).  Doxorubicin-­‐induced  cardiotoxicity  monitored  by  ECG  in  freely  moving  mice.  A  new  model  to  test  potential  protectors.  Cancer  Chemother  Pharmacol,38(1):  95-­‐101.  

Vertechy,  M.,  M.  B.  Cooper,  O.  Ghirardi  and  M.  T.  Ramacci   (1989).  Antioxidant  enzyme  activities   in  heart  and  skeletal  muscle  of  rats  of  different  ages.  Exp  Gerontol,24(3):  211-­‐218.  

Wallace,  D.  C.  (2010).  Mitochondrial  DNA  mutations  in  disease  and  aging.  Environ  Mol  Mutagen,51(5):  440-­‐450.  

Wallace,   K.   B.   (2003).   Doxorubicin-­‐induced   cardiac   mitochondrionopathy.   Pharmacol   Toxicol,93(3):   105-­‐115.  

Wallace,   K.   B.   (2007).   Adriamycin-­‐induced   interference   with   cardiac  mitochondrial   calcium   homeostasis.  Cardiovasc  Toxicol,7(2):  101-­‐107.  

Wei,  J.  Y.  (2004).  Understanding  the  aging  cardiovascular  system.  Geriatrics  &  Gerontology  International,4:  S298-­‐S303.  

Weitzel,  J.  M.,  K.  A.  Iwen  and  H.  J.  Seitz  (2003).  Regulation  of  mitochondrial  biogenesis  by  thyroid  hormone.  Exp  Physiol,88(1):  121-­‐128.  

Wessells,  R.  J.  and  R.  Bodmer  (2007).  Cardiac  aging.  Semin  Cell  Dev  Biol,18(1):  111-­‐116.  

Whelan,   R.   S.,   V.   Kaplinskiy   and   R.   N.   Kitsis   (2010).   Cell   death   in   the   pathogenesis   of   heart   disease:  mechanisms  and  significance.  Annu  Rev  Physiol,72:  19-­‐44.  

Wonders,   K.   Y.,   D.   S.   Hydock,   C.   M.   Schneider   and   R.   Hayward   (2008).   Acute   exercise   protects   against  doxorubicin  cardiotoxicity.  Integr  Cancer  Ther,7(3):  147-­‐154.  

Wu,  H.,   S.  B.  Kanatous,   F.  A.   Thurmond,  T.  Gallardo,   E.   Isotani,  R.  Bassel-­‐Duby  and  R.   S.  Williams   (2002).  Regulation  of  mitochondrial  biogenesis  in  skeletal  muscle  by  CaMK.  Science,296(5566):  349-­‐352.  

Yen,   H.   C.,   T.   D.   Oberley,   S.   Vichitbandha,   Y.   S.   Ho   and   D.   K.   St   Clair   (1996).   The   protective   role   of  manganese   superoxide   dismutase   against   adriamycin-­‐induced   acute   cardiac   toxicity   in   transgenic  mice.   J  Clin  Invest,98(5):  1253-­‐1260.  

Yeragani,  V.  K.,  E.  Sobolewski,  J.  Kay,  V.  C.  Jampala  and  G.  Igel  (1997).  Effect  of  age  on  long-­‐term  heart  rate  variability.  Cardiovasc  Res,35(1):  35-­‐42.  

Yilmaz,   S.,   A.   Atessahin,   E.   Sahna,   I.   Karahan   and   S.   Ozer   (2006).   Protective   effect   of   lycopene   on  adriamycin-­‐induced  cardiotoxicity  and  nephrotoxicity.  Toxicology,218(2-­‐3):  164-­‐171.  

Zhou,  S.,  A.  Starkov,  M.  K.  Froberg,  R.  L.  Leino  and  K.  B.  Wallace  (2001).  Cumulative  and  irreversible  cardiac  mitochondrial  dysfunction  induced  by  doxorubicin.  Cancer  Res,61(2):  771-­‐777.  

75

Zhu,  X.,  L.  Zuo,  A.  J.  Cardounel,  J.  L.  Zweier  and  G.  He  (2007).  Characterization  of  in  vivo  tissue  redox  status,  oxygenation,   and   formation   of   reactive   oxygen   species   in   postischemic   myocardium.   Antioxid   Redox  Signal,9(4):  447-­‐455.  

Zucchi,  R.  and  R.  Danesi  (2003).  Cardiac  toxicity  of  antineoplastic  anthracyclines.  Curr  Med  Chem  Anticancer  Agents,3(2):  151-­‐171.