Accelerated cardiomyocyte senescence contributes to late-onset doxorubicin-induced cardiotoxicity

Abstract

Children surviving cancer and chemotherapy are at risk for adverse health events including heart failure that may be delayed by years. Although the early effects of doxorubicin-induced cardiotoxicity may be attributed to a direct effect on the cardiomyocytes, the mechanisms underlying the delayed or late effects (8–20 yr) are unknown. The goal of this project was to develop a model of late-onset doxorubicin-induced cardiotoxicity to better delineate the underlying pathophysiology responsible. The underlying hypothesis was that doxorubicin-induced “late-onset cardiotoxicity” was the result of mitochondrial dysfunction leading to cell failure and death. Wistar rats, 3–4 wk of age, were randomly assigned to vehicle or doxorubicin injection groups (1–45 mg/kg). Cardiovascular function was unaltered at the lower dosages (1–15 kg/mg), but beginning at 6 mo after injection significant cardiac degradation was observed in the 45 mg/kg group. Doxorubicin significantly increased myocardial mitochondrial DNA (mtDNA) damage. In contrast, in isolated c-kit left ventricular (LV) cells, doxorubicin treatment did not increase mtDNA damage. Biomarkers of senescence within the LV were significantly increased, suggesting accelerated aging of the LV. Doxorubicin also significantly increased LV histamine content suggestive of mast cell activation. With the use of flow cytometry, a significant expansion of the c-kit and stage-specific embryonic antigen 1 cell populations within the LV were concomitant with significant decreases in the circulating peripheral blood population of these cells. These results are consistent with the concept that doxorubicin induced significant damage to the cardiomyocyte population and that although the heart attempted to compensate it eventually succumbed to an inability for self-repair.

INTRODUCTION

Long-term survival of childhood cancers is now >70% (9). The cytotoxic classes of drugs known as anthracyclines are some of the most efficacious anticancer drugs available. Unfortunately, these survivors are subject to several adverse health events including heart failure. Other complications include abnormal pulmonary function, auditory impairment, endocrine conditions, and neurocognitive impairment, whereas abnormalities involving hepatic dysfunction, osteoporosis, and kidney dysfunction are less frequently seen (29). Doxorubicin also impacts on metabolism, and follow-up studies of survivors observed increased obesity, insulin resistance, hyperlipidemia, and other markers of diabetes (56). In patients, the short-term effects of doxorubicin-induced cardiotoxicity (2–3 mo) are thought to directly involve the cardiomyocytes. The early effects of doxorubicin-induced cardiotoxicity may be attributed to altered Ca²⁺ dynamics, increased oxidative stress, and altered myocardial energetics (10, 13). Unfortunately, in children surviving cancer, a significant delayed form of heart failure has been also observed (41, 54, 55). The mechanisms underlying the delayed or late effects (8–20 yr) are unknown, but cardiac dysfunction leading to heart failure remains a major problem and accounts for 2–3% of heart transplants in young adults (4).

Doxorubicin’s usefulness as a neoplastic reagent lies in its ability to stabilize the topoisomerase-DNA complex to increase double-strand DNA breakage in proliferating cancer cells. Topoisomerases are highly conserved proteins, are present in all cells, and serve to resolve the topological difficulties of DNA replication by allowing the helix to pass through itself. Doxorubicin is known to accumulate in both the nucleus and the mitochondria and makes no distinction between cancerous and healthy cells (45). Three unique isoforms of topoisomerase have been demonstrated in the mitochondria (57). It has been suggested that the long-term effects of doxorubicin effects may be mediated by a persistent increase in cellular oxidant stress leading to mitochondrial dysfunction (49, 63). Although the linkage between doxorubicin and nuclear topoisomerases is reasonably understood, very little is known about its impact on myocardial mitochondrial topoisomerases.

Mitochondrial dysfunction has a significant role in the development and complications of different cardiomyopathies (18, 21, 25). Mitochondrial DNA (mtDNA) is particularly susceptible to damage. At autopsy, patients receiving doxorubicin presented with myocardial mtDNA alterations and deletions (36, 37). We (42) have reported that a shift in mitochondrial topoisomerase function underlies diabetic cardiomyopathy. Others have observed that that any increase in mtDNA mutations may initiate apoptosis (61). Here, failure was not just dysfunctional cardiomyocytes, but also an inability of the heart to replace the rapid dropout of cardiomyocytes. It may not be relevant that a cell dies but that the balance of cell death and cell replacement is upset. This conclusion is applicable to doxorubicin-induced cardiomyopathies.

The goal of this study was to develop and investigate a rodent model of heart failure that mimicked the delayed cardiotoxicity observed in survivors of chemotherapy. The underlying hypothesis was that late-onset doxorubicin-induced cardiotoxicity is the result of a progressive cellular failure. This cellular failure is the result of doxorubicin-induced mitochondrial dysfunction that ultimately leads to activation of apoptotic pathways and accelerated cell loss. Although it is evident that compensatory mechanisms were activated, heart failure results from an inability to maintain the balance of repair/replacement of cells.

MATERIALS AND METHODS

These studies were performed across a number of experimental platforms that included cultured cells and animals. A total of 80 rats were used in these experiments. Wistar rats (3–4 wk of age) were randomly assigned to saline or doxorubicin injection groups. Doxorubicin (Enzo, Farmingdale, NY) injections were made on alternate days and included a cumulative total of 1, 5, or 15 mg/kg ip over 3 injections or 45 mg/kg ip over 9 injections to use the same concentration at each injection. Our preliminary studies as well as others suggested that the animals did not tolerate injection of higher concentrations of doxorubicin (24). During the injection period and for 72 h afterward, the animals were housed on mesh-bottom cages to allow urine and fecal matter to fall through to prevent secondary exposure to doxorubicin. Following this, the animals were returned to normal caging. Experimental protocols had institutional approval, and animals were maintained in accordance with the American Physiological Society’s Guiding Principles for the Care and Use of Animals in Research and Teaching and the Guide for the Care and Use of Laboratory Animals.

Cardiovascular function.

Echocardiography measurement of cardiac function was initiated 2 mo following the final doxorubicin injection. Echocardiography was assessed using an Acuson Sequoia C256 system as described previously (35). In brief, animals were anesthetized using 1–1.5% isoflurane-100% O₂ metered through an Isotec 4 vaporizer (VetEquip, Pleasanton, CA) and maintained on a heated pad throughout the protocol. Once asleep, animals were allowed to stabilize for 10 min before baseline measurements were made. Following this, dobutamine was injected (50 µg/kg ip) and measurements were made after a 5-min stabilization period.

Cell culture.

Cell culture experiments used rat neonatal cardiomyocytes, bone marrow cells, or H9c2 cells [CRL-1446; Research Resource Identifier (RRID): CVCL_0286; American Type Culture Collection] that were originally derived from rat heart. H9c2 cells were maintained in DMEM + 10% FBS + 1× penicillin-streptomycin (pen/strep) as previously described (42). Bone marrow cells were maintained in Iscove’s + 10% FBS + 1× pen/strep as previously described (30). Neonatal cardiomyocytes from 1- to 3-day-old rats were prepared using collagenase IV, as previously described (15, 16). Following preparation, the cardiomyocytes were plated overnight in low-glucose (LG) DMEM + 10% FBS + 0.1 mM BrdU + 1.0 mM d-valine before switching to the experimental media (LG-DMEM, 1% FBS, 1× nonessential amino acids, 2 mM glutamine, 1× pen/strep).

Cellular and mitochondrial function.

Cytochrome oxidase (complex IV) was measured by following the oxidation of reduced cytochrome c at 550-nm absorbance, as previously described (42). Cellular senescence was determined by a β-galactosidase assay using o-nitrophenyl-β-d-galactoside (ONPG). In brief, left ventricular (LV) extracts were incubated for 8 h at 37°C in ONPG buffer (13 mM ONPG, 40 mM citric/NaPO₄, pH 6.0, 150 mM NaCl₂, 2 mM MgCl₂). Absorbance was measured at 420 nm, using 520 nm as a reference. Histamine analysis was performed on the LV using the fluorescence protocol as described by Shore (52). Measurement of chymase activity was performed on the LV using the substrate 0.2 M Suc-Ala-Ala-Pro-Phe-pNA as described by Kivinen et al. (31).

DNA cleavage assay.

DNA cleavage was determined by degradation of linear DNA using a Cy5-fluorescent probe (35, 42). In brief, a linear mtDNA was amplified by a PCR using an internally labeled Cy5-labeled primer: forward, 5′-AAATTTCCCGACACAAAATCTTTCC^(Cy5)TCCTAACTAA ACCCTCTTTACTTGC-3′, and reverse, 5′-CTCTTGGTAAGTAAATTTCTTTCTCC-3′, using mtDNA as the template to generate a 1,274-bp probe. To perform the cleavage assay, isolated mitochondria (1–10 µg of protein) were incubated in buffer (50 mmol/L Tris·HCl, pH 7.5, 100 mmol/L KCl, 0.5 mM EDTA, 0.5 mmol/L DTT, 30 µg/mL BSA) to a final volume of 20 µL. Tubes incubated without mitochondrial extracts controlled for nonspecific degradation of DNA. Three unique topoisomerase inhibitors were studied: doxorubicin (0.5 mM), dexrazoxane (5 µM), and hydroxycamptothecin (100 µM). The mitochondrial extracts were incubated in the absence or presence of topoisomerase inhibitors for 60 min at 37°C before the DNA was added. The reaction was then incubated at for a further 30 min at 37°C, to which 6-µL loading buffer was then added. Seven microliters was electrophoresed on a 4% polyacrylamide gel and visualized using a Storm 840 PhosphorImager (Molecular Dynamics). Band density was quantified using ImageJ. Where indicated, mitochondria were isolated using a differential centrifugation protocol that was nuclei-free, as previously described (25, 42).

Flow cytometry.

Flow cytometry was used to determine 1) cell apoptosis/cell death using annexin V and propidium iodide, 2) cellular senescence using p16^INK4a as a biomarker, 3) cell doubling time, 4) mitochondrial superoxide levels using MitoSox (25), and 5) cell identities, and relative proportions were measured using primary antibodies for specific targets and when necessary secondary antibodies conjugated to fluorochromes. When used, H9c2 or neonatal cardiomyocytes cells were harvested by brief exposure to a trypsin-EDTA transferred to a tube containing horse serum and collected by centrifugation (300 g; 5 min). Cells were suspended in DMEM, counted, and maintained on ice. When used, adult heart cells were isolated by collagenase II dissociation. In brief, a portion of the heart was minced using fine scissors, and the mince was added to a collagenase II solution (1% collagenase type II, Joklik medium, 60 mM taurine, 20 mM creatine, 5 mM HEPES, 0.1% BSA, 10 mM butanedione monoxime, pH 7.4) and shaken gently at 37°C for 5 min; the cells were allowed to settle for 30 s, and the supernatant was transferred to a new tube containing horse serum. This process was repeated three times, and the cells were collected by centrifugation (300 g; 5 min). Cells were suspended in DMEM, counted, and maintained on ice. One million cells per sample suspended in cell staining buffer (cat. no. 420201; BioLegend) were used for flow cytometry as described previously (35). For antibodies to internalized proteins, the cells were permeabilized in ice-cold acetone for 5 min before being collected by centrifugation (300 g; 5 min) and suspended in cell staining solution. Cells were then treated with select antibodies for 60 min at 4°C and washed twice in filtered (0.2 µm) PBS. Flow cytometry was performed using a Guava easyCyte mini flow cytometer, and cells were gated to exclude those <2 µm. The primary antibodies used included c-kit (cat. no. 105805; RRID: AB313214; BioLegend), p16^INK4a (cat. no. sc55600; RRID: AB112694; Santa Cruz Biotechnology), stage-specific embryonic antigen 1 (SSEA1; cat. no. 125608; RRID: AB1089188; BioLegend), Nkx-2.5 (cat. no. sc14033; RRID: AB650281; Santa Cruz Biotechnology), myosin (cat. no. sc101334; RRID: AB1126445; Santa Cruz Biotechnology), N-cadherin (cat. no. sc31029; RRID: AB2077523; Santa Cruz Biotechnology), and Sca-1(cat. no. 122505; RRID: AB756190; BioLegend). Cell doubling time was used as a determination of long-term consequences of doxorubicin treatment following the protocol of Wolfs et al. (59). H9c2 cells underwent a single 24-h doxorubicin challenge (5, 50, or 500 nM), after which the cells were washed twice with media. The cells were maintained in culture changing the media on alternate days and later used to measure cell doubling times at 2 days, 2 wk, and 10 wk after doxorubicin exposure. To determine cell doubling times, 10⁵ H9c2 cells were plated onto a 24-well plate and then recounted after 72 h; the differences in the initial and final cell counts were used to calculate the doubling time following the protocol of Wolfs et al. (59).

mtDNA damage.

Damage of mitochondrial DNA was assessed by long-range PCR (LRPCR; Ref. 14). In brief, the rationale is that any lesion (strand breaks, base modifications, and apurinic sites) will stop a thermostable DNA polymerase capable of generating a long DNA product. This decrease in DNA yield reflects an increase in DNA damage. The amount of damage was derived from the LRPCR amplification compared with the amplification of a short-range PCR (SRPCR) product (150–250 bp) by the comparative cycle threshold (2ΔCt) method. The SRPCR primers amplified the mtDNA in the 12S rRNA coding region. Mitochondrial copy number was derived from the SRPCR-to-β-actin ratio by the 2ΔCt method. mtDNA damage and mitochondrial copy number are expressed as means ± SE and normalized to the respective control group. With the use of dissociated LV cells as described above, an immunoprecipitation of c-kit⁺ cells was performed as described by Goswami et al. (20). In brief, freshly dissociated cells were maintained on ice in PBS to which 2–5 µg of antibody was added maintained on ice for 30 min before 50 µL of Dynabeads Protein G (Invitrogen, Carlsbad, CA) was added and the mixture was incubated for an additional 30 min. The supernatant was cleared by magnetic separation, and the cells were washed twice in cold PBS before they were collected for DNA isolation using the Extract-N-Amp kit from Sigma-Aldrich (St. Louis, MO).

DNA methylation status.

Methylation status was determined using two protocols. Global DNA methylation status of left ventricle DNA was determined using the MethylFlash Global DNA Methylation ELISA protocol as described by the manufacturer (EpiGentek, Farmingdale, NY). The second approach used a MethylScreen protocol as described by Holemon et al. (26). In brief, genomic DNA was cut using the HhaI restriction endonuclease that only cuts unmethylated DNA. An aliquot of the digest was then used in a quantitative PCR protocol with primers to amplify regions containing known methylation-sensitive restriction sites. An increase in the crossing point compared with an uncut control indicates a lower DNA yield, which reflects a decrease in DNA methylation. The primers used were atrial natriuretic factor (ANF) sense 5′-CTCTCTCCTCCCGCCC TTATTTG-3′, ANF antisense 5′-CTCGAGTGATGTTTGCTGTCTCGGCT-3′, mtDNA sense 5′-GGACTAGCCCCATT CCACTAC-3′, and mtDNA antisense 5′-GATAGGAGGATGATGGATGC-3′.

Statistical analysis.

Statistical analyses were performed using NCSS software (Kaysville, UT). Where appropriate, Student’s t-test or ANOVA was used: post hoc analysis was done using a Fisher least significant difference analysis. For cardiac functional analyses, doxorubicin treatment, dobutamine treatment, and time were used as main effects in a three-way ANOVA [total degrees of freedom (df) = 138]. We also used a repeated-measures ANOVA (total df = 104) including only those animals that survived out to 8 mo. In some cases, where the equality of variance or normality was not achieved, a Mann–Whitney U test was used. Values presented are means ± SE, and statistical significance was set at P < 0.05 unless otherwise indicated.

RESULTS

A goal of this investigation was to replicate the delayed onset of heart failure observed in many survivors of childhood cancer. Cardiac performance was determined by echocardiography. In animals that received a total cumulative dose of 1–15 mg/kg and followed out to 16 mo, no degradation of cardiovascular function was evident (Fig. 1 and supplement data; all supplemental material is available at https://doi.org/10.6084/m9.figshare.9924935.v1). In contrast, at a total cumulative dose of 45 mg/kg, a significant degradation of cardiovascular function became evident beginning at 6 mo after injection (Fig. 1). Table 1 summarizes the three-way ANOVA that included all animals. The analysis observed a time- and treatment-dependent decline across several parameters including the short-axis ejection fraction, with a significant interaction for treatment and time but not the response to dobutamine. Dobutamine is a β₁-adrenergic agonist and used clinically to stress the heart to reveal hidden dysfunction. As shown in Fig. 1, dobutamine significantly increased the ejection fraction at all time points, but there was no suggestion of hidden dysfunction before overt changes in ejection fraction. Not all animals survived out to 8 mo. A repeated-measures ANOVA that included only surviving animals yielded a similar outcome to observe a time- and treatment-dependent decline in ejection fraction.

Fig. 1.

Doxorubicin-induced degradation of cardiac function was delayed. Echocardiography was performed to measure short-axis ejection fraction 2–8 mo (M) after injection (45 mg/kg). Determinations were made under isoflurane anesthesia for both basal and following dobutamine injection (50 µg/kg ip) as described in materials and methods. Three-way ANOVA with doxorubicin, dobutamine, and time as main effects was used, with post hoc analysis using a Fisher least significant difference protocol. Values are means ± SE. *P < 0.05 compared with respective time-matched control, #P < 0.10 compared with respective time-matched control.

Table 1.

Three-way ANOVA of cardiovascular parameters

	Group	Dobutamine	Time	Group × Time
M-mode
IVSd, cm	ns	†	ns	ns
IVSs, cm	ns	ns	ns	ns
LVd, cm	†	*	†	ns
LVs, cm	ns	†	†	ns
FS	*	†	*	ns
Diastolic volume, mL	†	†	†	ns
Systolic volume, mL	ns	†	†	ns
Stroke volume, mL	†	ns	†	ns
Ejection fraction	†	†	*	ns
Short-axis
Left ventricular area/diastole, cm2	†	*	†	ns
Left ventricular area/systole, cm2	ns	†	†	*
Ejection fraction	†	†	†	*
Long-axis
Left ventricular area/diastole, cm2	ns	ns	ns	ns
Left ventricular area/systole, cm2	ns	†	†	ns
Ejection fraction	ns	†	†	ns

FS, fractional shortening; IVSd and IVSs, interventricular septal wall thickness in diastole and systole; LVd and LVs, left ventricular diameter in diastole and systole.

^*P < 0.05;

^†P < 0.001;

ns, not significant.

A known complication of chemotherapy is altered growth patterns in children. Across different concentrations of doxorubicin treatments, we observed that a total cumulative dose of 45 mg/kg significantly retarded growth of the female rats, whereas 15 mg/kg was without effect (Fig. 2A). In male rats, the 45 mg/kg injections protocol also significantly decreased body weights (control 600 ± 25 g, doxorubicin 432 ± 30 g; P < 0.05) as well as heart weight (control 1.16 ± 0.03 g, doxorubicin 1.04 ± 0.03 g; P < 0.05). The depressed growth was, in part, responsible for some alterations in cardiac volumes/dimensions. With the use of echocardiography, end-diastolic volumes were found to be significantly depressed by doxorubicin treatment (Fig. 2B). Another complication of chemotherapy is altered endocrine function. Nonfasting blood glucose levels were significantly elevated in the 45 mg/kg doxorubicin treatment compared with control animals (control 131 ± 6 mg%, doxorubicin 219 ± 15 mg%; P < 0.05). These results are consistent with clinical observations that doxorubicin chronically alters physiological function far beyond the time that it is present in the body.

Fig. 2.

Doxorubicin depressed growth. A: body weights in female rats. Animals were injected with doxorubicin (15 or 45 mg/kg) at 3–4 wk of age. Two-way ANOVA was performed using drug and time as main effects. Values are means ± SE and represent three observations per group. *P < 0.05 compared with respective control. B: end-diastolic volume (EDV) of vehicle or doxorubicin (45 mg/kg)-injected male rats. EDV was determined under basal conditions as described in materials and methods. For each time point, an unpaired t-test was performed: *P < 0.05 compared with respective control, #P < 0.10 compared with respective control. M or Mon, months.

Doxorubicin partitions to both the nucleus and the mitochondria, leading to mtDNA depletions and mtDNA rearrangements (37, 45). With the use of isolated myocardial mitochondria, hydroxycamptothecin or doxorubicin treatment significantly increased in vitro DNA cleavage activity demonstrating the presence of both type 1 and type 2 topoisomerases within the mitochondria. In contrast, dexrazoxane blunted basal DNA cleavage activity in the mitochondria (Fig. 3). In cardiomyocytes, 7 days following a transient (3 h) doxorubicin exposure, we observed a significant increase in mtDNA damage (Fig. 4A). Importantly, mtDNA damage was evident at doxorubicin concentrations (50 nM) that did not significantly increase mitochondrial superoxide levels (Fig. 4B). Consequence to the mtDNA damage was a significant increase in cell death following doxorubicin exposure (Fig. 4C). These findings demonstrated that doxorubicin-induced mitochondrial topoisomerase dysfunction resulted in significant mtDNA damage that contributed to increased cell death.

Fig. 3.

Mitochondrial DNA cleavage is brought about by types I and II mitochondrial topoisomerases. Dexrazoxane (5 µM) and doxorubicin (0.5 mM) are topoisomerase type 2 inhibitors. Hydroxycamptothecin (100 µM) is a topoisomerase type 1 inhibitor. Isolated mitochondria were incubated in the absence or presence of inhibitors for 60 min before the addition of linear DNA to perform a DNA cleavage assay as described in materials and methods. Tubes containing no mitochondria (No Mito) served as a negative control. Values are means ± SE and are normalized to the average of the No Mito group. One-way ANOVA was performed using drug concentration as a main effect with Fisher least significant difference for post hoc analysis. *P < 0.05 compared with No Mito group, #P < 0.05 compared with +mito group.

Fig. 4.

Doxorubicin induced mitochondrial DNA (mtDNA) damage and apoptosis in cardiomyocytes. A: doxorubicin increased in mtDNA damage. Damage of mtDNA was assessed by long-range quantitative PCR as described in materials and methods. Values are means ± SE and normalized to the mean of the control value (100%). B: higher concentrations of doxorubicin increased mitochondrial superoxide levels. Cardiomyocytes were treated with doxorubicin for 3 h before being washed twice with fresh media and incubated for 24 h before harvest. Values are means ± SE and normalized to the mean of the vehicle control (100%). C: doxorubicin induced cell death. Cells were analyzed by flow cytometry using annexin V (AnV) and propidium iodide (PI) as described in materials and methods. Values presented are means ± SE of the relative proportion of all cells counted. D: representative cytograms of flow cytometry. Values are means ± SE and represent twelve observations per group. One-way ANOVA was performed using drug concentration as a main effect with Fisher least significant difference for post hoc analysis. *P < 0.05 compared with respective control.

Myocardial mtDNA alterations have been demonstrated in patients previously treated with doxorubicin (36). Mitochondrial functional was compromised in doxorubicin-treated animals as evidenced by a significant decrease in LV cytochrome oxidase activity (control 1.31 ± 0.17 nmol·min⁻¹·mg⁻¹ protein, doxorubicin 0.72 ± 0.21 nmol·min⁻¹·mg⁻¹ protein; P < 0.05; n = 8). In both the nucleus and the mitochondria, doxorubicin binds to topoisomerase 2β to promote double-strand breaks. In cultured cells, doxorubicin exposure caused a significant increase in mtDNA damage (Fig. 4A). Thus we examine mtDNA damage in the LV from control and doxorubicin-treated animals. At 8 mo after injection, a significant increase in mtDNA damage as well as decreased mtDNA copy number was evident in the LV mitochondria of doxorubicin-treated animals (Fig. 5). Extending those observations, the LV was disassociated to single cells to allow for immunoprecipitation of LV c-kit⁺ cells. In contrast, doxorubicin treatment did not alter mtDNA damage or mitochondrial copy number of the isolated LV c-kit⁺ cells (Fig. 5). These results indicate a persistent effect of doxorubicin on the myocardial mitochondria that may be compartmentalized.

Fig. 5.

Past exposure to doxorubicin promoted degradation of left ventricular (LV) mitochondrial DNA (mtDNA) but not c-kit⁺/LV resident mtDNA of doxorubicin-injected animals. Long-range PCR analysis was used to determine mtDNA damage and mitochondrial cell numbers (Copy#) in LV or LV/c-kit⁺ cells as described in materials and methods. c-kit⁺ Cells were isolated by immunoprecipitation before mtDNA damage analysis. LV samples were obtained 8 mo following injection with saline or doxorubicin (45 mg/kg). Values are means ± SE and normalized to respective control samples. Unpaired t-test was performed: *P < 0.05 compared with respective control.

Doxorubicin has a half-life in the plasma of ~18 h, and similar intracellular values have been reported (34). H9c2 cells were treated with doxorubicin for 24 h before the cells were washed twice with media. The cells were then maintained in culture with media changes on alternate days and used to measure cell doubling times after 2 days, 2 wk, or 10 wk’ doxorubicin exposure. In comparison with time-matched controls, 50 and 500 nM doxorubicin treatments significantly increased doubling time for all time points (Fig. 6). The cell cycle distribution was determined in bone marrow cells (BMC) and neonatal cardiomyocytes 2 wk following a transient (24 h) doxorubicin treatment. In the BMC, we observed a significant increase in cells in the G₂/M phase (Fig. 7, region 3). Concomitant with this was a significant decrease in the proportion of cells in the G₁ phase (Fig. 7, region 1). Doxorubicin treatment also significantly increased the proportion of cells exhibiting polyploidy (Fig. 7, region 4). The response in neonatal cardiomyocytes was very different. Following a similar transient treatment, doxorubicin treatment produced a significant decrease of the number of cells in the G₂/M region (control 14.1 ± 1.2%, 5 nM doxorubicin 1.8 ± 0.4%; P < 0.05). Doxorubicin treatment of the cardiomyocytes also significantly increased the relative proportion of cells undergoing apoptosis (control 16.1 ± 1.9%, 5 nM doxorubicin 49.3 ± 4.2%; P < 0.05). These results indicate a persistent effect of doxorubicin and suggest distinct differences in the response by proliferating cells (BMC) versus cells that are predominantly terminally differentiated (neonatal cardiomyocytes).

Fig. 6.

Doxorubicin increased cell doubling time. Following a single 24-h doxorubicin exposure, H9c2 cells were maintained in culture for the time indicated before 10⁵ cells were replated and then counted after 72 h as described in materials and methods. Values are means ± SE. For each time point, a 1-way ANOVA was performed using drug concentration as a main effect with Fisher least significant difference for post hoc analysis. *P < 0.05 compared with respective time-matched vehicle control.

Fig. 7.

Doxorubicin exposure altered cell cycle distribution. A: following a single 24-h doxorubicin (Doxo) exposure, bone marrow cells were washed twice with media and then maintained in media for 2 wk before being analyzed for cell cycle distribution. B: representative cytograms of flow cytometry for control and 5 nM doxorubicin treatments using bone marrow cells (BMC) or neonatal cardiomyocytes. Cells were stained with propidium iodide to determine G₁ (region 1), S phase (region 2), G₂/M (region 3), and polyploidy (region 4) and analyzed using Guava FC Cell Cycle Software. Values are means ± SE and represent three observations per group. One-way ANOVA was performed using drug concentration as a main effect with Fisher least significant difference for post hoc analysis. *P < 0.05 compared with respective control.

Doxorubicin appeared to accelerate the aging process. With the use of H9c2 cells in culture, 7 days following a single transient doxorubicin treatment, a significant increase in senescence β-galactosidase activity was observed (Fig. 8A). Similarly in vivo, but with a much longer timeframe, 8 mo following doxorubicin treatment, senescence β-galactosidase activity was significantly increased in the left ventricle (LV) of doxorubicin-treated animals compared with controls (Fig. 8B). As previously reported, p16^INK4a is a recognized marker of cellular senescence (1, 2, 51). Compared with age-matched controls, 8 mo following doxorubicin treatment, there was a significant increase in p16^INK4a+/myosin⁺ cells in the left ventricle that was concomitant with a significant decrease in p16^INK4a-/myosin⁺ cells (Fig. 8D). These results indicate a significant shift in the cardiomyocytes toward an aged phenotype.

Fig. 8.

Doxorubicin (Doxo) increased markers of cellular senescence. A: senescence β-galactosidase activity in H9c2 cells was increased 7 days after transient doxorubicin treatment. Values are means ± SE and are normalized to the control group. One-way ANOVA was performed using drug concentration as a main effect with Fisher least significant difference for post hoc analysis. B: senescence β-galactosidase activity in left ventricle obtained 8 mo following saline- or doxorubicin (45 mg/kg)-treated animals. Values are means ± SE and normalized to respective control, and a Mann–Whitney U test was performed: *P < 0.05 compared with control. C: representative flow cytometry cytograms illustrating the relative myosin (green, GRN; x-axis) and p16^INK4a (red: y-axis) fluorescent distributions from doxorubicin-treated (45 mg/kg) animals. D: summary of myosin heavy chain-positive (MHC⁺)/p16^INK4a-positive (p16⁺) cells from the left ventricle of control or doxorubicin-treated animals shown in the upper right quadrant and the MHC⁺/p16⁻ cells in the lower right quadrant. Values are means ± SE, and a Mann–Whitney U test was performed to analyze the MHC⁺/p16⁺ cells: *P < 0.05 compared with respective control.

The identity and source of the stem/progenitor cells that may serve to regenerate the heart remains highly controversial. However, several investigations have demonstrated the importance of c-kit⁺ cells as a biomarker of progenitor cells that contribute to the maintenance of the heart (3, 17, 53). With the use of flow cytometry, we determined the relative presence of select biomarkers for progenitor cells. In cells isolated from the bone marrow of control or doxorubicin-treated rats, no differences were observed in the relative proportion of c-kit⁺ or SSEA1⁺ cells (data not shown). In contrast, 8 mo after doxorubicin treatment resulted in a significant decrease in blood-borne circulating c-kit⁺ and SSEA1⁺ cells (Fig. 9). Within the heart, some markers of pluripotency including c-kit⁺ and SSEA1⁺ were significantly increased in the left ventricle 8 mo following doxorubicin injection, whereas Sca-1 levels remained unaltered (Fig. 10). Both Nkx2.5 and N-cadherin are thought to be markers for commitment to a cardiac progenitor cell lineage (27). Although doxorubicin significantly increased LV Nkx-2.5 levels, N-cadherin levels were unaltered. Additionally, we observed a significant increase in the number of Ki67⁺ cells suggesting active proliferation within the LV. In parallel, we observed significant changes in DNA methylation. With the use of both mtDNA and the ANF promoter as a genomic target, doxorubicin significantly decreased DNA methylation in the LV DNA from doxorubicin-treated animals compared with controls (Fig. 11A). With the use of a protocol to measure global DNA methylation status, LV DNA methylation was also significantly decreased in the doxorubicin-treated animals (Fig. 11B). Mast cell content was examined using flow cytometry. With the use of FcεR1 receptor as a biomarker of mast cells, we were able to differentiate mature (high side scatter) mast cells from immature (low side scatter; Fig. 12A). Doxorubicin treatment did not alter the numbers of either subpopulation or the total number of mast cells (control 3.3 ± 0.6%, doxorubicin 2.7 ± 0.5%) within the LV (Fig. 12B). In contrast, we observed a significant increase in LV histamine content (Fig. 12C), indicative of mast cell activation. Additionally, chymase activity was more than doubled in the LV doxorubicin-treated animals compared with controls (control 0.91 ± 0.14 µmol·min⁻¹·mg⁻¹, doxorubicin 1.84 ± 0.39 µmol·min⁻¹·mg⁻¹; P < 0.05; n = 10).

Fig. 9.

Past exposure to doxorubicin treatment decreased circulating levels of progenitor cells. A: quantification of the upper left (UL) region, which contained positive stained cells in the 2- to 10-µm range. B: representative cytographs; UL region, 2–10 µm; upper right (UR), >10-µm diameter. Blood was obtained 8 mo after injection, and nucleated cells were prepared by standard methods. One million cells were stained with anti-c-kit-peridinin-chlorophyll-protein complex (anti-c-kit-PerCP) or anti-stage-specific embryonic antigen 1 (SSEA1)-Alexa Fluor 488 for 60 min before counted in a Guava Flow Cytometer. Samples are from control and doxorubicin (45 mg/kg) groups. Values are means ± SE with sample sizes indicated in each bar, and an unpaired t-test was performed: *P < 0.05 compared with respective control cells. FSC, forward scatter; GRN, green.

Fig. 10.

Doxorubicin (Doxo) treatment increased some markers of cardiac progenitor cells in the heart. One million left ventricular cells were incubated with indicated antibodies before analyzed by flow cytometry. Samples are from control and doxorubicin (45 mg/kg) groups. Values are the upper left (UL) region as illustrated in Fig. 9 and are the means ± SE, and unpaired t-tests were performed, with the exception of Nkx-2.5 data where a Mann–Whitney U test was performed: *P < 0.05 compared with respective control cells. SSEA1, stage-specific embryonic antigen 1.

Fig. 11.

Past exposure to doxorubicin decreased left ventricle (LV) DNA methylation. A: DNA methylation analysis of the mitochondrial DNA (mtDNA) and atrial natriuretic factor (ANF) promoter regions by quantitative PCR (QPCR). DNA from LV of control or doxorubicin-treated animals was isolated and cut with HhaI before QPCR. Values are relative to uncut DNA and are means ± SE. B: global LV DNA methylation was determined by an ELISA protocol as described in materials and methods. Values are means ± SE, and an unpaired t-test was performed: *P < 0.05 compared with respective control cells.

Fig. 12.

Doxorubicin exposure activated myocardial mast cells. Dissociated left ventricle cells were incubated with anti-FcεR1-FITC and anti-c-kit-peridinin-chlorophyll-protein complex (anti-c-kit-PerCP) as described in materials and methods. A: cells were gated to differentiate immature (lower) and mature (upper) mast cells. B: representative cytograms illustrating flow cytometry analysis. No differences in mast cell numbers were observed between the groups for all cells or cells gated by side scatter. C: histamine content of the left ventricle. Samples are from control and doxorubicin (45 mg/kg) groups. Values are means ± SE, and unpaired t-test was performed: *P < 0.05 compared with respective control. FSC, forward scatter; GRN, green; LV, left ventricle; SSC, side scatter.

DISCUSSION

These findings indicate a dose and time dependency for the onset of doxorubicin-induced cardiac dysfunction that replicated the delayed or late-onset heart failure observed in survivors of childhood cancer. Observation of a threshold for the development of doxorubicin-induced cardiotoxicity is also consistent with clinical reports. The model also recapitulates the depressed growth and markers of diabetes that are an associated outcome of chemotherapy. The data suggest that the heart attempts to recover from doxorubicin-induced stress as suggested by the increases in biomarkers for progenitor cells and proliferation. Unfortunately, the response appears to be insufficient as evidenced by chronic mitochondrial dysfunction and progressive degradation of cardiovascular function.

Myocardial mtDNA damage was an early (Fig. 2A) and persistent (Fig. 8) consequence of doxorubicin exposure. Doxorubicin accumulates in both the nucleus and the mitochondria where it binds to type 2 topoisomerase and promotes double-strand DNA breaks (45, 46). Doxorubicin exposure impacts heavily on the mitochondria as significant mtDNA depletion and mtDNA rearrangements have been reported in the heart and kidney (37, 38). The early cardiac crisis often seen clinically may be doxorubicin-induced cardiotoxicity manifested as apoptosis, resulting of the removal of cells with severely damaged mitochondria as we (Fig. 2) and others have reported (10, 13). At autopsy, myocardial mtDNA alterations were present in doxorubicin-treated patients indicating the persistence of doxorubicin’s effects, an effect similar to the present findings (Fig. 8; Ref. 36). Generation of mutant mtDNA is known to activate autophagy; however, this alone is not sufficient to completely eliminate all mutant mtDNA (11, 19). The mitochondrial genome continues to replicate about once a month (8). It has been suggested that the metabolic inefficiency of mutated mitochondria elicits a compensatory response driving increased mitochondrial biogenesis and the mutated mtDNA is dragged along as a hitchhiker (7). The lack of damage in the c-kit⁺ fraction suggests an expansion of only the healthier progenitor cells. Our findings of distinct differences in cell cycle distribution between proliferating and terminally differentiated cells support this concept. It is unknown whether the transformation of primitive/progenitor cells to cardiomyocytes is altered by mtDNA damage. However, any compensatory response appears to be insufficient as evidenced by the significant decreases in LV mitochondrial function. The near-complete reliance of the myocardium on aerobic metabolism to meet energy demand engenders its sensitivity to mitochondrial function.

Within any organ even under normal conditions, there is heterogeneity of cellular ages. Aguayo-Mazzucato et al. (1) observed individual pancreas β-cells of different cell ages. They reported that p16^INK4a as well as Igf1r and Bambi positively correlated with senescence galactosidase activity (1). Rota et al. (51) demonstrated that within 3-mo-old animals, there was heterogeneity of heart cell age, an observation that is consistent with the concept that the heart has a continual turnover of cells. In that study, the aged cells were positive for p16^INK4a and apoptosis was evident in some cells. Baker et al. (2) reported that elimination of p16^INK4a+ cells delayed the onset of age-related pathologies in adipose tissue and skeletal muscle. In the present study, doxorubicin increased the relative proportion of p16^INK4a+/myosin⁺ cells at the expense of the p16^INK4a−/myosin⁺ as well as increased senescent β-galactosidase activity in the left ventricle. In combination with the changes in mitochondrial function, these findings are all indicative of an accelerated aged cardiomyocyte profile that underlies degradation of cardiovascular function.

Increased DNA methylation is thought to underlie gene silencing as a function of the aging process (28). It is then a contradiction that DNA methylation was significantly decreased in a heart that exhibited a senescence phenotype with a greater percentage of aged cardiomyocytes. The heart is a heterogeneous organ. Cardiomyocytes comprise 70% of the heart by volume but <50% by cell number, with endothelial cells, fibroblasts, mast cells, smooth muscle cells, and progenitor cells comprising the rest (5, 47). Although cardiomyocytes have minimal regenerative capacity, other cell types such as fibroblasts and smooth muscle are known to be capable of replicative function. As well, there is a low turnover of progenitor cells throughout the life of an individual, although the source of cells and rate of turnover remain highly controversial (3, 5, 6, 43, 53). In the doxorubicin-treated hearts, the greater presence of Ki67⁺, SSEA1, and c-kit⁺ cells suggests an increase in proliferating progenitor cells, and those cells would be expected to have less DNA methylation (28). The increase in Nkx2.5 suggests that the heart cells were committing to the cardiomyocyte phenotype (44). Collectively, these results suggest that the heart was attempting to compensate by the formation of new cardiomyocytes.

These contradictions may be at least partially resolved by considering the impact of myocardial mast cells. Similar to other cell types, doxorubicin is taken up by mast cells (12). Mast cell activation is proinflammatory and induces not only apoptosis, but also degradation of extracellular space that could contribute to myocardial dysfunction (33). Coculture of mast cells with cardiomyocytes promoted significant apoptosis of the cardiomyocytes (22). Additionally, it has been suggested that chymase also secreted by mast cells may induce myosin degradation within the cardiomyocytes (48). Mast cell activation with elevated histamine release has also been shown to be arrhythmogenic (58). Several investigations have reported deleterious effects of mast cell activation in response to ischemia/reperfusion injury, pressure overload, or high-fat diets (22, 23). Blockade of the H₂ receptor pathways at least partially ameliorates the pathological outcome (23, 60). Acutely, doxorubicin increased histamine release from myocardial mast cells (12). Zhang et al. (62) observed in adult animals a significant increase in cardiac mast cell number and degranulation following the doxorubicin treatment. Kondru et al. (32) reported that histamine antagonists were partially cardioprotective from doxorubicin treatment. In the present study, 8 mo following the doxorubicin injections, although there was no change in mast cell number, the increased histamine content and chymase activity suggested chronic activation of the existing population. It is not clear whether histamine alone or in concert with other proinflammatory cytokines promoting the release of senescence-associated secretory proteins enhance the aging process of other cells locally (2, 50). Collectively, these findings all are consistent with the concept of altered myocardial mast cell function contributing to an accelerated cardiomyocyte aging profile.

Conclusion.

Long-term survival of childhood cancers is now >70% (9). Unfortunately, adult survivors of childhood cancer are at risk for treatment-related adverse health outcomes. Doxorubicin-induced cardiac dysfunction remains a significant clinical problem. This is of greater concern if radiotherapy is part of the treatment protocol (39, 40). Analysis of heart transplantation patients found doxorubicin as the underlying cause in 2–3% of all cases (4).

These findings indicate a dose and time dependency for the onset of doxorubicin-induced cardiac dysfunction that replicated the late-onset heart failure observed in survivors of childhood cancer. Doxorubicin treatment chronically altered DNA structure in heart cells. A chronic shift in mast cell function may underlie many of the changes observed. The heart is heavily dependent on aerobic metabolism, and mitochondria function was compromised because of mitochondrial topoisomerase-induced mtDNA damage that appeared to accelerate cardiomyocyte senescence. Our findings suggest that the progenitor cell activity within the heart was elevated in an attempt to compensate for cell loss but was unable to match the accelerated aging and degradation of the cardiomyocyte population. The importance may be not that a cell dies but that the balance of cell death and cell replacement is upset.

GRANTS

This study was sponsored, in part, by National Institute of Child Health and Human Development (Grant R03-HD-065555101), National Aeronautics and Space Administration (Grant 80NSSC19K0436), and Touro Colleges and Universities Biomedical Fund to J. G. Edwards. B. L. Keith was supported by a Glorney-Raisbeck Medical Student Fellowship Award. We are also grateful for the support of the New York Medical College/Westchester Medical Center Stem Cell Laboratory and its Core Facility.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

D.L., C.A.E., L.M.E., and J.G.E. conceived and designed research; M.A.M., D.L., B.L.K., E.S., S.J., and J.G.E. performed experiments; M.A.M., D.L., B.L.K., E.S., C.A.E., L.M.E., S.J., S.G., and J.G.E. analyzed data; M.A.M., D.L., B.L.K., E.S., C.A.E., L.M.E., S.J., S.G., and J.G.E. interpreted results of experiments; S.J. and J.G.E. prepared figures; C.A.E. and J.G.E. drafted manuscript; C.A.E., L.M.E., S.G., and J.G.E. edited and revised manuscript; M.A.M., D.L., B.L.K., E.S., C.A.E., L.M.E., S.J., S.G., and J.G.E. approved final version of manuscript.