Investigation into the cardiotoxic effects of doxorubicin on contractile function and the protection afforded by cyclosporin A using the work-loop assay

Toxicology in Vitro 28 (2014) 722–731

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Investigation into the cardiotoxic effects of doxorubicin on contractile function and the protection afforded by cyclosporin A using the work-loop assay Mayel Gharanei a, Afthab Hussain a, Rob S. James a, Omar Janneh b, Helen Maddock a,⇑ a b

Department of Biomolecular and Sport Sciences, Coventry University, Cox Street, Coventry CV1 5FB, UK Division of Biomedical Sciences, St. George’s, University of London, Cranmer Terrace, Tooting, London SW17 0RE, UK

a r t i c l e

i n f o

Article history: Received 19 May 2013 Accepted 27 January 2014 Available online 7 February 2014 Keywords: Force Doxorubicin Cardiotoxicity mPTP CsA

a b s t r a c t Doxorubicin is known to cause cardiotoxicity through multiple routes including the build-up of reactive oxygen species and disruption of the calcium homeostasis in cardiac myocytes, but the effect of drug treatment on the associated biomechanics of cardiac injury remains unclear. Detecting and understanding the adverse effects of drugs on cardiac contractility is becoming a priority in non-clinical safety pharmacology assessment. The work-loop technique enables the assessment of force–length work-loop contractions, which mimic those of the pressure–volume work-loops experienced by the heart in vivo. During this study we evaluated whether the work-loop technique could potentially provide improved insight into the biomechanics associated with drug-induced cardiac dysfunction. In order to do this we investigated the cardiotoxic effects of doxorubicin and characterised the protection afforded by the coadministration of cyclosporin A (CsA). This study provides detailed biomechanical in vitro insight into the cardiac dysfunction associated with Doxorubicin treatment, including reduction in peak force, force during shortening and power output. These effects were significantly abrogated in doxorubicin-CsA co-treatment studies. Closely mimicking the in vivo pressure–volume muscle mechanics, this assay provides a quick and easy technique to gain a better understanding of the detailed biomechanics of drug-induced cardiac dysfunction. Ó 2014 Elsevier Ltd. All rights reserved.

1. Introduction It is becoming increasingly apparent that drugs have the ability to cause serious adverse cardiac dysfunction (Force and Kerkela, 2008). Cardiotoxic side effects are a major limitation in the use of anti-cancer therapies in general. Doxorubicin is one of the most effective and potent anthracycline antibiotics widely used for the treatment of a variety of cancers. However, its use has been hampered due to its cardiotoxic properties leading to congestive heart failure and irreversible dilative cardiomyopathies (Allen, 1992;

Abbreviations: ANT, adenine nucleotide translocase; CsA, cyclosporin A; Cyp-D, cyclophillin D; IR, ischaemia and reperfusion; Lmax, length at which maximum developed force was recorded; Lopt, 95% of Lmax; LWa, work required to lengthen the active muscle; LWp, work required to re-lengthen the muscle; mPTP, mitochondrial permeability transition pore; RyR, ryanodine receptor/s; SERCA, sarco/endoplasmic reticulum Ca2+-ATPase; SR, sarcoplasmic reticulum; SWa, work done by the active muscle during shortening; Wnet, net work done by the muscle. ⇑ Corresponding author. Tel.: +44 (0) 2476888170; fax: +44 02476888778. E-mail address: [email protected] (H. Maddock). http://dx.doi.org/10.1016/j.tiv.2014.01.011 0887-2333/Ó 2014 Elsevier Ltd. All rights reserved.

Lefrak et al., 1973; Minotti et al., 2004). Doxorubicin induced cardiotoxicity has been well documented and investigated in a variety of different pre-clinical and clinical models. Its cardiotoxic effects have been attributed to various factors, which stem from its ability to produce reactive oxygen species and disruption of calcium homeostasis leading to oxidative stress, mitochondrial impairment, cell necrosis and induction of apoptotic pathways (Minotti et al., 2004; Singal and Iliskovic, 1998). Doxorubicin treatment has been found to cause contractile dysfunction, affecting contractile proteins directly or by affecting cellular calcium homeostasis (Arai et al., 1998, 2000; Boucek et al., 1997; Dodd et al., 1993; Olson et al., 1988; Wang et al., 2001). Factors that contribute to the depression of contractile function in response to doxorubicin treatment may include down-regulation of sarcoplasmic reticulum (SR) calcium release channel, calcium ATPase and decrease in SR calcium loading (Arai et al., 1998; Dodd et al., 1993; Olson et al., 2005; Wang and Korth, 1995). Doxorubicin-induced oxidative stress and calcium overload leads to generation of reactive oxygen species which can lead to

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mitochondrial damage and induction of cell death pathways (Zhang et al., 2009). Furthermore, investigations using have shown that doxorubicin-induced cardiomyocytes cell death is associated with increased expression and activation of p53 tumour suppressor protein (L’Ecuyer et al., 2006; Liu et al., 2008). A large body of research is dedicated to finding strategies or pharmacological agents that potentially reduce anthracycline-induced cardiotoxicity whilst simultaneously maintaining the anti-cancer properties of anthracyclines (Minotti et al., 2004). Recently, we and others have shown that doxorubicin-induced cardiotoxicity is linked to the opening of the non-specific mitochondrial permeability transition pore (mPTP) (Gharanei et al., 2012; Marechal et al., 2011; Montaigne et al., 2010, 2011). Opening of the mPTP allows molecules (<1.5 KDa) through the mitochondrial membrane (Yellon and Hausenloy, 2007). This subsequently leads to the collapse of mitochondrial membrane potential (Crompton et al., 1999), swelling and rupture of the mitochondrial membrane, thereby releasing pro-apoptotic molecules into the cytosol (Halestrap, 2009; Halestrap and Pasdois, 2009). mPTP opening is also found to be the main cause of injury during myocardial ischaemia and reperfusion (IR). Interestingly, studies have shown that the immunosuppressant cyclosporin A (CsA) is able to reduce IR injury (Crompton et al., 1999; Griffiths and Halestrap, 1991, 1993; Hausenloy et al., 2002; Morin et al., 2009) and doxorubicin induced cardiotoxicity (Gharanei et al., 2012; Marechal et al., 2011) due to its ability to bind to and cause a conformational change to the morphology of cyclophillin D (Cyp-D) (Tanveer et al., 1996). This prevents Cyp-D from binding to adenine nucleotide translocase (ANT) and initiating mPTP opening. The beneficial effects of CsA in reducing cell death have also been documented in the liver (Kawano et al., 1989), the brain (Shiga et al., 1992) and the kidney (Yang et al., 2001). Moreover, in a small proof-of-concept clinical study, a single intravenous bolus of CsA limited myocardial infarct size development as measured by total serum creatine kinase release (Piot et al., 2008). We and others have previously reported that co-administration of doxorubicin with CsA is able to reduce doxorubicin-induced contractile dysfunction of the cardiac muscle in healthy and stressed conditions (Gharanei et al., 2012; Marechal et al., 2011; Montaigne et al., 2011). Doxorubicin-induced acute myocardial dysfunction and mitochondrial membrane dissipation were prevented by co-administration with CsA (Montaigne et al., 2010). In addition, it was also shown by the same group that CsA prevented contractile function of the human atrial trabeculae contractile function when exposed to doxorubicin (Montaigne et al., 2011). Furthermore, previous research from our laboratory demonstrated a reduction in the contractile force of rat cardiac papillary muscle when also exposed to hypoxia reoxygenation, which was prevented with co-treatment with CsA (Gharanei et al., 2013). However, the models used in these studies, e.g. isometric and force–velocity techniques, are not truly physiologically representative of the contractile biomechanics of the heart muscle. The structure and functional characteristics of the heart are responsible for the pressure–volume relationship, which regulates the rate and the potency of the complex rhythmic contractions, allowing efficient cardiac output during normal, stressed or diseased conditions (Katz and Katz, 1989). Therefore, cardiac myocytes must be flexible in adjusting their contractile properties as a consequence of the stress-strain dynamics of the heart that are affected by the ventricular pressure–volume behaviour. A variation in pre-load (diastolic filling) is one of the main dependencies of cardiac contractility and cardiac output referred to as the Frank– Starling law of the heart (Zimmer, 2002). Variation in pre-load will lead to variations in the stretch of the ventricular muscle before contraction. During contractions the cardiac muscle undergoes shortening and lengthening during cardiac filling and ejection

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(Josephson, 1985; Layland et al., 1995b). Therefore, accurate investigation of the contractile function requires the consideration of length change cycle during contraction and relaxation, which needs to be considered in studies that determine the extent of damage in the heart in clinical settings or in the basic scientific investigations. Several methods have previously been used to investigate cardiac muscle mechanics and the effects of drug treatment on cardiac contractility in the past (Gharanei et al., 2012; Layland and Kentish, 2000; Montaigne et al., 2011). In vitro isometric (constant muscle length) and force–velocity techniques (constant velocity or constant force) have been by far the most used techniques to investigate in vitro papillary or trabeculae muscle mechanics (Dekker et al., 1996; Min et al., 2000; Mudalagiri et al., 2008; Sivaraman et al., 2007). During isometric studies the muscle is held at constant length and maximally activated with performance indicated by the maximal force produced and the times taken to reach particular activation or relaxation states. Force–velocity techniques activate the muscle whilst at constant length until maximal force is reached, with muscle performance being measured during subsequent shortening at constant velocity or against constant load. Whilst such techniques provide interesting information about the intrinsic mechanical properties of muscle they do not resemble in vivo function, failing to account for the fact that muscles cannot be continuously maximally activated as time is required for relaxation, re-lengthening and activation (Caiozzo, 2002; James et al., 1996; Josephson, 1993). Therefore, force–velocity and isometric techniques greatly overestimate the contractile performance of muscle during in vivo function as these techniques do not reflect the dynamics of the heart muscle during in vivo activity as cardiac muscle undergoes lengthening and shortening during contraction and relaxation to allow ventricular filling and ejection respectively. A number of studies have shown that ventricular muscle length changes during contraction and relaxation have a near sinusoidal shape (Delhaas et al., 1993a,b; Semafuko and Bowie, 1975). Ventricular contraction and relaxation can be separated into three phases in relation to the variation in muscle strain patterns; an initial isometric phase occurs during iso-volumic contraction at the start of the ventricular systole, a shortening phase during ventricular ejection and finally re-lengthening phase that coincides with ventricular filling (Sperelakis, 2001). Thus sinusoidal length changes during muscle activation and relaxation represent a closer simulation of in vivo contractions. The work-loop technique can be used to combine sinusoidal length changes with phasic electrical stimulation, allowing power output of the muscles during contractions to be investigated under realistic physiological conditions (Caiozzo, 2002; James et al., 1996; Josephson, 1985, 1993; Syme and Josephson, 1995). By plotting the force produced by the muscle against muscle length, work and power output of the muscle can be calculated. Unlike the isometric or isotonic contraction model, the work-loop technique also takes into account the work done by the muscle during shortening as well as the work required to re-extend the muscle during lengthening (James et al., 1996; Josephson, 1985). This technique also incorporates changes in shortening velocity and the level of muscle activation during contractions as well as producing force and length changes that closely resemble papillary muscle dynamics in vivo (Hirakawa et al., 1977; Semafuko and Bowie, 1975). Thus it is clear that the work-loop technique provides a more realistic physiological representation of cardiac muscle mechanics, allowing a more accurate assessment of pharmacological compounds on cardiac muscle performance (Layland and Kentish, 1999, 2000; Layland et al., 1995b). In the current study we investigated the cardiotoxic effects of doxorubicin treatment on the contractile function of the isolated papillary muscle in vitro and the protection afforded by CsA using

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the work-loop model, which is a better representation of the in vivo muscle mechanics. 2. Materials and methods 2.1. Chemicals Doxorubicin hydrochloride and cyclosporin A (CsA) were purchased from Tocris Cookson (Bristol, UK). Doxorubicin was dissolved in ultra-pure water and CsA was dissolved in ethanol, ensuring that the final concentration of ethanol was less than 0.01% during the experiments. The dissolved drugs were aliquotted and stored at 20 °C. All other reagents were purchased from Sigma Aldrich (UK) unless stated otherwise. 2.2. Animals Male Sprague–Dawley rats were used in all experiments (350– 400 g body mass). Experiments were conducted in accordance with the Guidelines on the operation of Animals (Scientific Procedures Act 1986). Animals were obtained from Charles River UK Limited (Margate, UK).

using a micromanipulator (Aurora Scientific, Canada) to obtain the length at which maximum developed force was recorded (Lmax). The muscles were then brought back to 95% of Lmax (Lopt), which is the optimum length at which the muscle produces maximum power during sinusoidal length change cycle (Layland et al., 1995b). Once Lopt was measured, using a microscope fitted with an eyepiece graticule, the protocol sequence was started to carry out work-loops (5 work-loops per sequence) of a cycle frequency of 6 Hz and a strain amplitude of ±6% (12% peak to peak total strain), which have previously been found to produce maximum power output in cardiac muscle (Layland et al., 1995a,b). The sequence was repeated every 5 min throughout the experimental protocol of 150 min. At the end of the experiments wet muscle mass was measured to the nearest 0.00001 g using an electronic balance (Mettler Toledo B204-S, Zurich, Switzerland). Muscle mass, fibre length and the assumed muscle density of 1060 kg m 3 (Méndez and Keys, 1960; Vinnakota and Bassingthwaighte, 2004) were used to calculate fibre cross-sectional area and the isometric muscle stress. Instantaneous power output was calculated for every data point in each work loop (1667 data points per work loop) by multiplying instantaneous force by instantaneous velocity, and then these instantaneous power output values were averaged to generate a net average power output value for each work-loop.

2.3. Work-loop protocol 2.3.1. Papillary muscle dissection Rats were sacrificed by cervical dislocation followed by rapid excision of the heart from the body. Each heart was placed in ice-cold (2–4 °C) oxygenated (95% O2 and 5% CO2) Krebs Henseleit (KH) buffer (in mM: NaCl 118.5, NaHCO3 25, KCl 4.8, MgCl 1.2, KH2PO4 1.2, CaCl2 1.7, Glucose 12) and pinned onto a Silicone Elastomer Slygard based petri dish (Farnell, UK). The petri dish was placed onto a dissecting microscope to dissect the papillary muscles under stereomicroscopic conditions. A small incision was made, starting from the apex of the heart, using micro-fine scissors to expose the papillary muscles. All papillary muscles were fully exposed and a selection was made based on the structures of the chordae tendinaea and its attachments to the myocardium. Muscles containing forking or branching were excluded as this may cause disturbance in the force production, propagation and recordings. The chosen papillary muscle was then carefully dissected and T-clips fastened onto either end (Layland et al., 1995a). The clips were fastened around the myocardium and the chordae tendinaea ensuring that the muscles were not in contact with the clips, to avoid damage and force dispersion from the muscle. The muscle preparations were then placed in a horizontal organ bath (801b, Aurora Scientific, Canada), which was connected to a 50 mN force transducer (400A, Aurora scientific, Canada) and a high speed length controller (322C-l, Aurora Scientific, Canada). The organ bath was continuously perfused with oxygenated KH buffer and maintained at 37 °C. The muscles were allowed to stabilize for at least 20 min in KH buffer before muscle length optimisation began. 2.3.2. Muscle length optimisation Preliminary studies showed that a stimulation amplitude of 60 mA produced maximum developed force (mN) when stimulation voltage was 80 mV (data not shown). The muscles were stimulated with platinum electrodes using stimulation amplitude of 60 mA and 80 mV at constant length (isometric) to produce a twitch response. The developed force was recorded using the computerised oscilloscope software (ASI6006A, Aurora Scientific, Canada) by subtracting the maximum force produced from the minimum (baseline force). The muscle was gradually stretched

2.3.3. Protocol The initial 20 min of the work-loop protocol was used to establish baseline work-loop performance. The muscles were then randomly assigned to the following groups: treatment with vehicle control (KH buffer alone) for 120 min; treatment with doxorubicin (1 lM) for 120 min; treatment with doxorubicin (1 lM) and CsA (0.2 lM) for 120 min; or treatment with CsA alone (0.2 lM) for 120 min, Fig. 1. 2.4. Statistical analysis The data were expressed as mean ± SEM. Peak forces and power output data were assessed for statistical difference using one way analysis of variance (ANOVA) with Fisher post hoc tests for each time point. p < 0.05 was considered statistically significant. 3. Results The general, pre-treatment characteristics of the papillary muscles used in this study are shown in Table 1. There was no significant difference between different experimental groups in any of the parameters shown in Table 1 at the start of the experiments.

Fig. 1. Treatment protocol; each muscle was allowed to stabilise in the Krebs solution for 20 min followed by 20 min of baseline work-loop measurements, then 120 min of drug or vehicle treatment. Dox, Doxorubicin; CsA, cyclosporin A.

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M. Gharanei et al. / Toxicology in Vitro 28 (2014) 722–731 Table 1 Characteristics of papillary muscle preparations.

Rat body mass (g) Muscle mass (mg) Lmax (mm) L95% (mm) Active force (mN) Passive force (mN) Developed force (mN) Fibre CSA at Lmax (m2) Stress at Lmax (kN m 2) Fibre CSA at L95% (m2) Stress at L95% (kN m 2)

Control

Doxorubicin

CsA

Doxorubicin + CsA

371 ± 7 0.86 ± 0.05 3.1 ± 0.3 2.95 ± 0.3 26 ± 3.4 3.7 ± 1.2 22.7 ± 2.9 2.6  10 7 ± 2.7  10 91.6 ± 13.6 2.7  10 7 ± 2.8  10 87.2 ± 12.9

384 ± 6 0.9 ± 0.05 3.5 ± 0.4 3.4 ± 0.4 26.8 ± 2.8 5 ± 1.3 21.8 ± 2.2 2.5  10 7 ± 1.8  10 89.3 ± 11.7 2.6  10 7 ± 1.9  10 85 ± 11.2

385 ± 7 0.89 ± 0.07 3.3 ± 0.2 3.1 ± 0.2 27.4 ± 2.4 5.2 ± 1 22.2 ± 2.1 2.5  10 7 ± 2.3  10 87.9 ± 8.2 2.7  10 7 ± 2.4  10 83.9 ± 7.9

377 ± 6 0.89 ± 0.08 3.2 ± 0.3 3.1 ± 0.3 28.9 ± 2.3 3.8 ± 1.5 25 ± 1.7 2.7  10 7 ± 3.9  10 103.5 ± 17.6 2.8  10 7 ± 4.1  10 98.3 ± 16.8

8

8

8

8

8

8

8

8

Lmax, length at which maximum force was produced; L95%, 95% of Lmax; Fibre CSA, fibre cross-sectional area. Values are mean ± SEM n = 4–6.

3.1. Work-loop features The relationship between force (mN), muscle length (mm) and instantaneous power (mW) are shown in Fig. 2A. The muscle needed to be activated during lengthening to maximise net work output; therefore instantaneous power became negative whilst the muscle was active and lengthening (Josephson, 1993). Force increased as muscle length increased until maximum length was reached, this coincided with the peak force. Muscle produced work during shortening; therefore the line representing instantaneous power indicates positive values during shortening, representing ventricular ejection in situ. Work-loops are produced when force is plotted against muscle length (% strain), Fig. 2B–F. The ascending limb of the work loop represents muscle activation until peak force is reached (Fig. 2B). Active force production occurred during shortening and the muscles underwent further passive re-lengthening until pre-activation length was reached (Fig. 2B). The area under the ascending limb of the work-loop represents the work that was required to lengthen the muscle whilst it was being activated (LWa; Fig. 2C), represented by the negative instantaneous power output values in Fig. 2A. Work done by the muscle occurs during shortening of the muscle therefore the entire area under the curve of muscle shortening represents the total work done by the muscle (SWa; Fig. 2D) and is represented by positive instantaneous power output values in Fig. 2A. Work required to stretch the muscle to reach its original pre-activation length is shown under the curve of passive re-lengthening (LWp; Fig. 2E), and is also represented by negative values in Fig. 2A. The area inside the loop represents the net work done by the muscle during a length change cycle contractions and it is calculated by subtracting the work required during lengthening (i.e. the work done on the muscle) from the work done by the muscle during shortening (Wnet; Fig. 2F). 3.2. The effects of doxorubicin treatment on work-loops Work-loops recorded during sinusoidal length changes are depicted in Fig. 3 by plotting force against % length change (strain). Fig. 3A shows the average work-loops of control and doxorubicintreated muscle during the baseline measurement period (10 min). There were no differences in the peak force, or work-loop shape between the two groups during the baseline measurement period (10 min, Fig. 3A), or at 50 and 80 min into the treatment protocol (Fig. 3B, C). Work-loop shapes suggest that doxorubicin treatment decreased the activation rate of the muscle as well as causing a significant reduction in peak force compared to control after 120 min and 150 min of treatment (Fig. 3D and E). Work-loop shapes indicate that doxorubicin also caused a lower force to be maintained during muscle shortening, when compared to the control. Fig. 3F shows the effects of doxorubicin treatment on average work-loop shapes at different time points over the time course of the

experiment (10 min, 50 min, 80 min, 120 min and 150 min). The shape of these work-loops suggest that a decrease in rate of muscle force generation and force maintained during shortening accompanied the significant drop in peak muscle force during the time course of doxorubicin treatment (Fig. 3F). 3.3. Instantaneous power The graphs depicting the changes in instantaneous power output over time in control and doxorubicin treated muscles are shown in Fig. 4. No differences between instantaneous power output readings were observed from the work-loop shape during the baseline measurement period (10 min, Fig. 4A) between the doxorubicin-treated and the non-treated muscles. Doxorubicin treatment caused a reduction in the instantaneous power output required to stretch the muscle, while it was active, as compared to control from 50 min into the treatment protocol as observed from the work-loop shape (Fig. 4B). This effect is more pronounced at later time points (80–150 min, Fig. 4C–E, respectively). Timedependent effects of doxorubicin on instantaneous power output are shown in Fig. 4F. Doxorubicin caused a time-dependent reduction in the both the instantaneous power output required to stretch the active muscle as well as the instantaneous power output produced by the muscle during shortening (Fig. 4E). 3.4. Peak force Peak force over time for all the treatment groups is shown in Fig. 5. Doxorubicin treatment significantly reduced the peak force produced during sinusoidal length changes from 100 min onwards as compared to time matched controls (at 120 min; 61.6 ± 4.4% vs. 74.6 ± 6.1% of baseline measurement, p < 0.05, Fig. 5), as can also been seen from the work-loop shapes in Fig. 3. Co-treatment with CsA abrogated the effects of doxorubicin alone on peak force (at 120 min; 75.5 ± 3% vs. 61.6 ± 4.4%, p < 0.05, Fig. 5). CsA treatment alone did not cause a significant change in the peak force when compared to control. 3.5. Power output Net power output is shown in Fig. 6 as a percentage of power during stabilisation period. Doxorubicin treatment caused a significant reduction in power output from 110 min into the protocol as compared to time-matched controls (at 120 min; 49.5 ± 5.0% vs. 64.7 ± 6.7% of baseline measurement, p < 0.05, Fig. 6). Co-administration of doxorubicin with CsA significantly reduced the detrimental effects on power output seen with doxorubicin treatment alone (at 120 min; 68.5 ± 5.4% vs. 49.5 ± 5.0% of doxorubicin alone, p < 0.05, Fig. 6). Treatment with CsA did not show a significant difference in the power output when compared to control.

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Fig. 2. Work-loop features and characteristics. (A) Typical raw data for power, length and force during a work-loop sequence. (B) demonstrates the work-loop by plotting force against muscle length (% strain) during muscle shortening and passive re-lengthening; the ascending limb corresponds to muscle activation; peak force is also shown. (C) The area, LWa, under the curve of the ascending limb of the work-loop corresponds to the work required to lengthen the muscle when active. (D) The area, SWa, under the shortening curve corresponds to the work done by the muscle during shortening. (E) The work, LWp, required to re-lengthen the muscle back to its starting length. (F) The net work done, Wnet, during the work-loop cycle; the net work done by the muscle is calculated by subtracting the work required for lengthening from the work done during muscle shortening.

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Fig. 3. Doxorubicin-induced effects on average work-loop shape of control and doxorubicin treatment during baseline (A), 50 min (B), 80 min (C), 120 min (D) and 150 min (E) into the treatment protocol. (F) Comparison of the effect of doxorubicin on work loop shape over time. p < 0.05 vs. peak force control. Values presented are mean of n = 4–6.

4. Discussion In this study we use the work-loop technique to investigate the effects of doxorubicin treatment on the biomechanical contractile function of cardiac papillary muscle. Using sinusoidal muscle

length changes during contraction and subsequent re-lengthening, this model assesses mechanical muscle performance of a cardiac muscle (Josephson, 1985). It is of particular importance that this method considers the positive work done by a muscle during shortening and the negative work required to re-extend the muscle

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Fig. 4. The effects of doxorubicin on instantaneous power output during work-loop cycles. Work is done on the muscle during lengthening hence negative instantaneous power output values. Muscle does work during shortening, hence positive instantaneous power output values. Effects of drug treatment during baseline (A), 50 min (B), 80 min (C), 120 min (D) and 150 min (E) into the treatment protocol. The effect of doxorubicin treatment over time (F). Values presented are mean of n = 4–6.

(Fig. 4). Clearly this is an important technique that allows the measurement of cardiac muscle performance in healthy, stressed and pathological conditions while the muscle is undergoing near realistic physiological length changes (Layland and Kentish, 2000; Layland et al., 1995b). Traditional cardiac muscle performance experiments have used isometric contraction or after-load isotonic contraction studies. However, neither of these mimic in vivo muscle function. The cardiac muscle is dynamic and undergoes lengthening and shorting during ventricular filling and ejection, respectively. Therefore, isometric or isotonic contraction studies will not give a true representation of power produced by

the muscle during contractions as during the contractile process, muscles are not able to continue to shorten indefinitely and produce positive work, instead during each cycle, the muscle needs to be re-extended, which involves negative work, i.e. work being done on the muscle to stretch it (Josephson, 1993). In theory to maximise work production the work loop should be as large and as square as possible, however, the shape of the work loop is constrained by activation rate, force–velocity properties, relaxation rate and passive resistance to stretch (James et al., 1996; Josephson, 1993; Caiozzo, 2002). Therefore, investigation of work loop shape allows us to identify the differing effects of

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Fig. 5. The effects of control, doxorubicin, cyclosporin A (CsA) and doxorubicin + CsA on peak force plotted as a % of mean peak force during baseline measurements. p < 0.05 doxorubicin vs. time-matched control, #p < 0.05 vs. timematched doxorubicin. Values presented are mean ± SEM of n = 4–6.

Fig. 6. The effects of control, doxorubicin, cyclosporin A (CsA) and doxorubicin + CsA on power output as a % of power output during baseline measurements. Showing the effects of control, doxorubicin, cyclosporin A (CsA) and doxorubicin + CsA. p < 0.05 doxorubicin vs. time-matched control, #p < 0.05 doxorubicin + CsA vs. time-matched doxorubicin. Values presented are mean ± SEM of n = 4–6.

pharmacological treatment on these various mechanical properties of cardiac muscle. Our data demonstrate detrimental effects of doxorubicin treatment on cardiac muscle mechanics. In particular, a significant decrease in the work and peak force produced along with changes in the work-loop shapes that indicate a reduced rate of muscle activation and reduced force production during shortening (Figs. 3 and 5). Cardiac action potential results in the entry of calcium into the cytosol of the cell through the sarcolemmal membrane, which in turn causes further calcium-induced calcium release from the SR, increasing cytosolic calcium concentrations (Carafoli, 1985; Stehle and Iorga, 2010). Calcium in the cytosol is free to bind to cardiac troponin C, which initiates the contractile process of the cardiac cells (Stehle and Iorga, 2010). Previous studies have shown that treatment with doxorubicin can lead to changes affecting the contractile response and properties of cardiac

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myocytes (Boucek et al., 1997; de Beer et al., 2000). Although the main cause of doxorubicin-induced cardiotoxicity is thought to be due to build of reactive oxygen species, there is ample evidence to suggest a role for doxorubicin-induced imbalance of calcium homeostasis (Arai et al., 1998, 2000; Boucek et al., 1997; Dodd et al., 1993; Olson et al., 1988; Wang et al., 2001). Chronic doxorubicin treatment has been found to cause a down-regulation of SR calcium release channel density (ryanodine receptors, RyR) and down-regulation of gene expression of RyR and calcium ATPase (SERCA) accompanied by a decrease in cardiac output and fractional shortening (Arai et al., 1998; Dodd et al., 1993; Olson et al., 2005). Furthermore, acute effects of doxorubicin treatment have also shown transcriptional repression of SERCA and activation of mitogen-activated protein kinases (Arai et al., 2000). A decrease in SR calcium loading and a consequent decrease in calcium-induced calcium release has been observed with doxorubicin treatment in isolated guinea pig ventricular myocytes (Wang and Korth, 1995). A decrease in calcium release should result in a reduced rate of activation during doxorubicin treatment, as highlighted in our work loop shapes, and a consequent decreased peak force, as only a finite time was available for force increase before the muscle was shortened. During sinusoidal work loop experiments, regardless of pharmacological treatment, muscle shortening usually causes force to decrease in the first half of the work loop, in line with the force–velocity relationship, whilst the velocity of shortening is increasing. Therefore, as peak force achieved during doxorubicin treatment is lower the force achieved during shortening is also likely to be lower, as demonstrated in the work loop shapes. Studies investigating the effects of doxorubicin on cross bridge kinetics have shown that both acute and chronic doxorubicin administration result in impairment of cross bridge kinetics in skinned cardiac trabeculae (de Beer et al., 2000). Delayed uncoupling of the attached cross bridges were observed with acute treatment, resulting in higher isometric tension, while chronic treatment resulted in impairment of both attachment and detachment of cross bridges (de Beer et al., 2000). By using near physiological length changes implemented in the work-loop technique, we demonstrate the detrimental effects of doxorubicin on muscle activation, peak force, force during shortening and power output. Furthermore, we have demonstrated that the detrimental effects of doxorubicin on papillary muscle biomechanics can be prevented by the protective effects of CsA in doxorubicin-induced effects on cardiac papillary muscle function. Co-administration with CsA improved the peak force and cardiac output of doxorubicin-treated muscles. The cardio-protective effects of CsA are generally attributed to its inhibitory effects of the mPTP (Halestrap et al., 1997; Hausenloy et al., 2010). In addition, investigations into isometric contractile function of rat papillary muscle and human trabeculae muscle have also shown that the detrimental effects of doxorubicin treatment were prevented when co-treated with CsA (Gharanei et al., 2012; Montaigne et al., 2011). In contrast, CsA has also been found to cause concentration-dependent cardiodepressive effects when used in muscle preparations from human failing hearts (Janssen et al., 2000). To our knowledge this is the first study to investigate the cardiotoxic effects of doxorubicin in papillary muscle contraction using the work-loop technique. We definitively show reductions in the rate of muscle activation, peak force, and maintenance of force during shortening and hence muscle power output with doxorubicin treatment. We provide evidence that inhibition of the mitochondrial permeability transition pore with CsA protects against the detrimental effects of doxorubicin on contractile function. Collectively, this work-loop assay could serve as a better representation of in vivo muscle mechanics when compared to the traditional isometric or isotonic contraction studies. Therefore

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using the work-loop technique can provide a better understanding of cardiac muscle mechanics in healthy, stressed or pathological conditions. It serves as a relevant pre-clinical testing method allowing a more accurate characterisation of the pathophysiological effects of compounds on cardiac contractile function. Conflict of Interest The authors declare that there are no conflicts of interest. Transparency Document The Transparency document associated with this article can be found in the online version.

Acknowledgements We would like to thank Coventry University for funding this research and thank Mark Bodycote, Bethan Grist and Samuel Pollard for technical support. References Allen, A., 1992. The cardiotoxicity of chemotherapeutic drugs. Semin. Oncol. 19, 529–542. Arai, M., Tomaru, K., Takizawa, T., Sekiguchi, K., Yokoyama, T., Suzuki, T., Nagai, R., 1998. Sarcoplasmic reticulum genes are selectively down-regulated in cardiomyopathy produced by doxorubicin in rabbits. J. Mol. Cell. Cardiol. 30, 243–254. Arai, M., Yoguchi, A., Takizawa, T., Yokoyama, T., Kanda, T., Kurabayashi, M., Nagai, R., 2000. Mechanism of doxorubicin-induced inhibition of sarcoplasmic reticulum Ca(2+)-ATPase gene transcription. Circ. Res. 86, 8–14. Boucek Jr., R.J., Dodd, D.A., Atkinson, J.B., Oquist, N., Olson, R.D., 1997. Contractile failure in chronic doxorubicin-induced cardiomyopathy. J. Mol. Cell. Cardiol. 29, 2631–2640. Caiozzo, V.J., 2002. Plasticity of skeletal muscle phenotype: mechanical consequences. Muscle Nerve 26, 740–768. Carafoli, E., 1985. The homeostasis of calcium in heart cells. J. Mol. Cell. Cardiol. 17, 203–212. Crompton, M., Virji, S., Doyle, V., Johnson, N., Ward, J.M., 1999. The mitochondrial permeability transition pore. Biochem. Soc. Symp. 66, 167–179. de Beer, E.L., Bottone, A.E., van Der Velden, J., Voest, E.E., 2000. Doxorubicin impairs crossbridge turnover kinetics in skinned cardiac trabeculae after acute and chronic treatment. Mol. Pharmacol. 57, 1152–1157. Dekker, L.R., Fiolet, J.W., VanBavel, E., Coronel, R., Opthof, T., Spaan, J.A., Janse, M.J., 1996. Intracellular Ca2+, intercellular electrical coupling, and mechanical activity in ischemic rabbit papillary muscle. Effects of preconditioning and metabolic blockade. Circ. Res. 79, 237–246. Delhaas, T., Arts, T., Bovendeerd, P.H., Prinzen, F.W., Reneman, R.S., 1993a. Subepicardial fiber strain and stress as related to left ventricular pressure and volume. Am. J. Physiol. 264, H1548–H1559. Delhaas, T., Arts, T., Prinzen, F.W., Reneman, R.S., 1993b. Relation between regional electrical activation time and subepicardial fiber strain in the canine left ventricle. Pflugers Arch. 423, 78–87. Dodd, D.A., Atkinson, J.B., Olson, R.D., Buck, S., Cusack, B.J., Fleischer, S., Boucek Jr., R.J., 1993. Doxorubicin cardiomyopathy is associated with a decrease in calcium release channel of the sarcoplasmic reticulum in a chronic rabbit model. J. Clin. Invest. 91, 1697–1705. Force, T., Kerkela, R., 2008. Cardiotoxicity of the new cancer therapeutics– mechanisms of, and approaches to, the problem. Drug Discov. Today 13, 778– 784. Gharanei, M., Hussain, A., Janneh, O., Maddock, H.L., 2012. Doxorubicin induced myocardial injury is exacerbated following ischaemic stress via opening of the mitochondrial permeability transition pore. Toxicol. Appl. Pharmacol. Gharanei, M., Hussain, A., Janneh, O., Maddock, H.L., 2013. Doxorubicin induced myocardial injury is exacerbated following ischaemic stress via opening of the mitochondrial permeability transition pore. Toxicol. Appl. Pharmacol. 268, 149– 156. Griffiths, E.J., Halestrap, A.P., 1991. Further evidence that cyclosporin A protects mitochondria from calcium overload by inhibiting a matrix peptidyl–prolyl cis– trans isomerase. Implications for the immunosuppressive and toxic effects of cyclosporin. Biochem. J. 274 (Pt 2), 611–614. Griffiths, E.J., Halestrap, A.P., 1993. Protection by Cyclosporin A of ischemia/ reperfusion-induced damage in isolated rat hearts. J. Mol. Cell. Cardiol. 25, 1461–1469.

Halestrap, A.P., 2009. What is the mitochondrial permeability transition pore? J. Mol. Cell. Cardiol. 46, 821–831. Halestrap, A.P., Pasdois, P., 2009. The role of the mitochondrial permeability transition pore in heart disease. Biochim. Biophys. Acta 1787, 1402–1415. Halestrap, A.P., Connern, C.P., Griffiths, E.J., Kerr, P.M., 1997. Cyclosporin A binding to mitochondrial cyclophilin inhibits the permeability transition pore and protects hearts from ischaemia/reperfusion injury. Mol. Cell Biochem. 174, 167– 172. Hausenloy, D.J., Maddock, H.L., Baxter, G.F., Yellon, D.M., 2002. Inhibiting mitochondrial permeability transition pore opening: a new paradigm for myocardial preconditioning? Cardiovasc. Res. 55, 534–543. Hausenloy, D.J., Lim, S.Y., Ong, S.G., Davidson, S.M., Yellon, D.M., 2010. Mitochondrial cyclophilin-D as a critical mediator of ischaemic preconditioning. Cardiovasc. Res. 88, 67–74. Hirakawa, S., Sasayama, S., Tomoike, H., Crozatier, B., Franklin, D., McKown, D., Ross Jr., J., 1977. In situ measurement of papillary muscle dynamics in the dog left ventricle. Am. J. Physiol. 233, H384–H391. James, R.S., Young, I.S., Cox, V.M., Goldspink, D.F., Altringham, J.D., 1996. Isometric and isotonic muscle properties as determinants of work loop power output. Pflugers Arch. 432, 767–774. Janssen, P.M., Zeitz, O., Keweloh, B., Siegel, U., Maier, L.S., Barckhausen, P., Pieske, B., Prestle, J., Lehnart, S.E., Hasenfuss, G., 2000. Influence of cyclosporine A on contractile function, calcium handling, and energetics in isolated human and rabbit myocardium. Cardiovasc. Res. 47, 99–107. Josephson, R.K., 1985. Mechanical power output from striated muscle during cyclic contraction. J. Exp. Biol., 493–512. Josephson, R.K., 1993. Contraction dynamics and power output of skeletal muscle. Annu. Rev. Physiol. 55, 527–546. Katz, A.M., Katz, P.B., 1989. Homogeneity out of heterogeneity. Circulation 79, 712– 717. Kawano, K., Kim, Y.I., Kaketani, K., Kobayashi, M., 1989. The beneficial effect of cyclosporine on liver ischemia in rats. Transplantation 48, 759–764. Layland, J., Kentish, J.C., 1999. Positive force- and [Ca2+]i-frequency relationships in rat ventricular trabeculae at physiological frequencies. Am. J. Physiol. 276, H9– H18. Layland, J., Kentish, J.C., 2000. Effects of 1- or -adrenoceptor stimulation on workloop and isometric contractions of isolated rat cardiac trabeculae. J. Physiol. 524 (Pt 1), 205–219. Layland, J., Young, I.S., Altringham, J.D., 1995a. The effect of cycle frequency on the power output of rat papillary muscles in vitro. J. Exp. Biol. 198, 1035–1043. Layland, J., Young, I.S., Altringham, J.D., 1995b. The length dependence of work production in rat papillary muscles in vitro. J. Exp. Biol. 198, 2491–2499. L’Ecuyer, T., Sanjeev, S., Thomas, R., Novak, R., Das, L., Campbell, W., Heide, R.V., 2006. DNA damage is an early event in doxorubicin-induced cardiac myocyte death. Am. J. Physiol. Heart Circ. Physiol. 291, H1273–H1280. Lefrak, E.A., Pitha, J., Rosenheim, S., Gottlieb, J.A., 1973. A clinicopathologic analysis of adriamycin cardiotoxicity. Cancer 32, 302–314. Liu, J., Mao, W., Ding, B., Liang, C.S., 2008. ERKs/p53 signal transduction pathway is involved in doxorubicin-induced apoptosis in H9c2 cells and cardiomyocytes. Am. J. Physiol. Heart Circ. Physiol. 295, H1956–H1965. Marechal, X., Montaigne, D., Marciniak, C., Marchetti, P., Hassoun, S.M., Beauvillain, J.C., Lancel, S., Neviere, R., 2011. Doxorubicin-induced cardiac dysfunction is attenuated by ciclosporin treatment in mice through improvements in mitochondrial bioenergetics. Clin. Sci. (Lond.) 121, 405–413. Méndez, J., Keys, A., 1960. Density and composition of mammalian muscle. Metabolism 9, 184–188. Min, J.Y., Hampton, T.G., Wang, J.F., DeAngelis, J., Morgan, J.P., 2000. Depressed tolerance to fluorocarbon-simulated ischemia in failing myocardium due to impaired [Ca(2+)](i) modulation. Am. J. Physiol. Heart Circ. Physiol. 278, H1446–H1456. Minotti, G., Menna, P., Salvatorelli, E., Cairo, G., Gianni, L., 2004. Anthracyclines: molecular advances and pharmacologic developments in antitumor activity and cardiotoxicity. Pharmacol. Rev. 56, 185–229. Montaigne, D., Marechal, X., Baccouch, R., Modine, T., Preau, S., Zannis, K., Marchetti, P., Lancel, S., Neviere, R., 2010. Stabilization of mitochondrial membrane potential prevents doxorubicin-induced cardiotoxicity in isolated rat heart. Toxicol. Appl. Pharmacol. 244, 300–307. Montaigne, D., Marechal, X., Preau, S., Baccouch, R., Modine, T., Fayad, G., Lancel, S., Neviere, R., 2011. Doxorubicin induces mitochondrial permeability transition and contractile dysfunction in the human myocardium. Mitochondrion 11, 22– 26. Morin, D., Assaly, R., Paradis, S., Berdeaux, A., 2009. Inhibition of mitochondrial membrane permeability as a putative pharmacological target for cardioprotection. Curr. Med. Chem. 16, 4382–4398. Mudalagiri, N.R., Mocanu, M.M., Di Salvo, C., Kolvekar, S., Hayward, M., Yap, J., Keogh, B., Yellon, D.M., 2008. Erythropoietin protects the human myocardium against hypoxia/reoxygenation injury via phosphatidylinositol-3 kinase and ERK1/2 activation. Br. J. Pharmacol. 153, 50–56. Olson, R.D., Mushlin, P.S., Brenner, D.E., Fleischer, S., Cusack, B.J., Chang, B.K., Boucek Jr., R.J., 1988. Doxorubicin cardiotoxicity may be caused by its metabolite, doxorubicinol. Proc. Natl. Acad. Sci. USA 85, 3585–3589. Olson, R.D., Gambliel, H.A., Vestal, R.E., Shadle, S.E., Charlier Jr., H.A., Cusack, B.J., 2005. Doxorubicin cardiac dysfunction: effects on calcium regulatory proteins, sarcoplasmic reticulum, and triiodothyronine. Cardiovasc. Toxicol. 5, 269–283. Piot, C., Croisille, P., Staat, P., Thibault, H., Rioufol, G., Mewton, N., Elbelghiti, R., Cung, T.T., Bonnefoy, E., Angoulvant, D., Macia, C., Raczka, F., Sportouch, C.,

M. Gharanei et al. / Toxicology in Vitro 28 (2014) 722–731 Gahide, Finet, G., Andre-Fouet, X., Revel, D., Kirkorian, G., Monassier, J.P., Derumeaux, G., Ovize, M., . Effect of cyclosporine on reperfusion injury in acute myocardial infarction. New Engl. J. Med. 359, 473–481. Semafuko, W.E., Bowie, W.C., 1975. Papillary muscle dynamics: in situ function and responses of the papillary muscle. Am. J. Physiol. 228, 1800–1807. Shiga, Y., Onodera, H., Matsuo, Y., Kogure, K., 1992. Cyclosporin A protects against ischemia-reperfusion injury in the brain. Brain Res. 595, 145–148. Singal, P.K., Iliskovic, N., 1998. Doxorubicin-induced cardiomyopathy. New Engl. J. Med. 339, 900–905. Sivaraman, V., Mudalagiri, N.R., Di Salvo, C., Kolvekar, S., Hayward, M., Yap, J., Keogh, B., Hausenloy, D.J., Yellon, D.M., 2007. Postconditioning protects human atrial muscle through the activation of the RISK pathway. Basic Res. Cardiol. 102, 453–459. Sperelakis, N., 2001. Heart Physiology and Pathophysiology, fourth ed. Academic Press, San Diego, California. Stehle, R., Iorga, B., 2010. Kinetics of cardiac sarcomeric processes and rate-limiting steps in contraction and relaxation. J. Mol. Cell. Cardiol. 48, 843–850. Syme, D.A., Josephson, R.K., 1995. Influence of muscle length on work from trabecular muscle of frog atrium and ventricle. J. Exp. Biol. 198, 2221–2227. Tanveer, A., Virji, S., Andreeva, L., Totty, N.F., Hsuan, J.J., Ward, J.M., Crompton, M., 1996. Involvement of cyclophilin D in the activation of a mitochondrial pore by Ca2+ and oxidant stress. Eur. J. Biochem. 238, 166–172.

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Vinnakota, K.C., Bassingthwaighte, J.B., 2004. Myocardial density and composition: a basis for calculating intracellular metabolite concentrations. Am. J. Physiol. Heart Circ. Physiol. 286, H1742–H1749. Wang, Y.X., Korth, M., 1995. Effects of doxorubicin on excitation–contraction coupling in guinea pig ventricular myocardium. Circ. Res. 76, 645–653. Wang, G.X., Wang, Y.X., Zhou, X.B., Korth, M., 2001. Effects of doxorubicinol on excitation–contraction coupling in guinea pig ventricular myocytes. Eur. J. Pharmacol. 423, 99–107. Yang, C.W., Ahn, H.J., Han, H.J., Kim, W.Y., Li, C., Shin, M.J., Kim, S.K., Park, J.H., Kim, Y.S., Moon, I.S., Bang, B.K., 2001. Pharmacological preconditioning with lowdose cyclosporine or FK506 reduces subsequent ischemia/reperfusion injury in rat kidney. Transplantation 72, 1753–1759. Yellon, D.M., Hausenloy, D.J., 2007. Myocardial reperfusion injury. New Engl. J. Med. 357, 1121–1135. Zhang, Y.W., Shi, J., Li, Y.J., Wei, L., 2009. Cardiomyocyte death in doxorubicininduced cardiotoxicity. Arch. Immunol. Ther. Exp. (Warsz) 57, 435–445. Zimmer, H.G., 2002. Who discovered the Frank–Starling mechanism? News Physiol. Sci. 17, 181–184.