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Am J Physiol Heart Circ Physiol 291: H2680-H2691, 2006. First published June 16, 2006; doi:10.1152/ajpheart.00395.2006
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Heat shock protects cardiac cells from doxorubicin-induced toxicity by activating p38 MAPK and phosphorylation of small heat shock protein 27

C. D. Venkatakrishnan,1 Arun K. Tewari,2 Leni Moldovan,1 Arturo J. Cardounel,3 Jay L. Zweier,1 Periannan Kuppusamy,1 and Govindasamy Ilangovan1

1Center for Biomedical EPR Spectroscopy and Imaging and Division of Cardiovascular Medicine, Department of Internal Medicine, 2Proteomics Core Facility, and 3Department of Pharmacology, Davis Heart and Lung Research Institute, The Ohio State University, Columbus, Ohio

Submitted 14 April 2006 ; accepted in final form 8 June 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Doxorubicin (DOX) and its derivatives are used as chemotherapeutic drugs to treat cancer patients. However, production of DOX-mediated reactive oxygen species (ROS) by prolonged use of these drugs has been found to cause dilative cardiomyopathy and congestive heart failure. Thus various preventive modalities have been developed to avoid this side effect. We have found that the DOX-mediated oxidant-induced toxicity in cardiac cells could be minimized by hyperthermia-induced small heat shock protein 27 (HSP27); that is, this protein acts as an endogenous antioxidant against DOX-derived oxidants such as H2O2. Heat shock-induced HSP27 was found to act as an antiapoptotic protein (reducing ROS and Bax-to-Bcl2 ratio) against DOX, and its phosphorylated isoforms stabilized F-actin remodeling in DOX-treated cardiac cells and, hence, attenuated the toxicity. Protein kinase assays and proteomic analyses suggested that higher expression of HSP27 and its phosphorylation are responsible for the protection in heat-shocked cells. Two-dimensional gel electrophoresis showed six isoforms (nonphosphorylated and phosphorylated) of HSP27. Matrix-assisted laser desorption/ionization time of flight analyses showed {alpha}- and beta-isoforms of HSP27, which are phosphorylated by various protein kinases. Ser15 and Ser85 phosphorylation of HSP27 by MAPK-assisted protein kinase 2 was found to be the key mechanism in reduction of apoptosis and facilitation of F-actin remodeling. The present study illustrates that hyperthermia protects cells from DOX-induced death through induction and phosphorylation of HSP27 and its antiapoptotic and actin-remodeling activities.

apoptosis; cardiomyopathy; hyperthermia

ADRIAMYCIN DERIVATIVES, such as doxorubicin (DOX), are widely used as anticancer drugs. Unfortunately, prolonged use of these drugs to treat cancer patients has resulted in the development of dilative cardiomyopathy (DCM) and congestive heart failure (CHF) within a few years after cessation of treatment, irrespective of age and race (36). To avoid this side effect, new anthracyclin derivatives with less toxicity to the heart, new formulations in the form of encapsulation into delivery vehicles such as microemulsions, and new targeted delivery modalities to selectively deliver the drug to the desired site (preventing accumulation in the heart) have emerged (39). However, there has been no significant improvement in our ability to avoid this problem, and this effect has not been completely eliminated. At the same time, attempts to understand the actual mechanism of DOX-induced DCM and CHF have been appearing with increasing frequency in recent literature. Fortunately, the mechanisms whereby DOX and its analogs kill tumor cells (intended effect) and cause DCM and CHF (unwanted effects) are different, indicating that the mode of action of these drugs depends on the nature of the tissues. This difference in action is reason for hope that selective targeting of some of the pathways through new approaches could prevent the inevitable cardiomyopathy and CHF among DOX-treated cancer patients. Thus there is renewed interest in identifying and selectively targeting these pathways.

The anthracyclins, particularly DOX, kill cancer cells primarily by DNA intercalation and damage and inhibition of topoisomerase II (36). On the other hand, cardiotoxicity is caused by mitochondria-triggered apoptosis, which is mediated by reactive oxygen species (ROS) generated in the redox reactions of DOX (11, 17). Cardiomyocytes are more susceptible to ROS-induced apoptosis, because they exhibit low levels of catalase (which detoxifies the H2O2) and readily undergo inactivation of selenium-dependent GSH peroxidase-1, which reduces NF-{kappa}B activation (46). Thus the DOX-induced apoptotic pathway could be successfully suppressed by external supplementation of antioxidants or induction of endogenous antioxidants such as cardiac-specific overexpression of cysteine-rich metallothionines (48). The ROS generated in the redox cycling of DOX are aggravated by many factors in cells, such as iron (2, 4, 27, 36). It has also been reported that DOX-generated ROS induce apoptosis by activating p38 MAPKs subunits {alpha} and beta. Interestingly, heat shock proteins (HSPs), which are induced in response to stress, have been found to be phosphorylated by the MAPK-assisted protein kinase 2 (MAPKAP-2, downstream of p38 MAPK), and the phosphorylated HSPs can act as negative regulators of apoptosis (5, 6), suggesting that induction of HSPs should regulate the p38 MAPK and, possibly, inhibit DOX-induced apoptosis, although this novel hypothesis remains untested.

Cardioprotection by small HSPs against ischemia-reperfusion injury and other oxidative stresses has been investigated recently (23, 33, 37, 40). Specifically, HSP27 and its murine homolog HSP25 have been shown to play important roles in the protection of cardiomyocytes under various stresses. Recently, it was reported that phosphorylation of Ser15 and Ser82 of human HSP27 was not required to protect the heart from ischemic injury (23). Attenuation of ROS and lipid peroxidation has also been shown by induction and phosphorylation of the small HSPs (1, 3). As described above, because the DOX-induced cardiomyopathy is caused by DOX redox-generated ROS and its mediation in apoptosis, it is logical to expect that overexpression of HSP27 protects against DOX-induced cardiotoxicity. In the present work, we evaluated the hypothesis that HSP27, induced by hyperthermia, can minimize DOX-induced toxicity in cardiac cells. Although various transgenic approaches have been reported to overexpress HSPs (31), we used hyperthermia as a mode of HSP overexpression because of its clinical relevance.

Cultured cardiac H9c2 cells, a clonal heart muscle cell line that is derived from embryonic rat hearts and retains many cardiomyocyte phenotypes, were used in the present study (16, 21). Even though the DOX-induced cardiotoxicity is common in adults, this cell line has been used to investigate the cardiotoxic effect caused by DOX (12, 31). Thus we chose cardiac H9c2 cells, instead of freshly isolated adult cardiomyocytes, for this study. The results obtained in the present study demonstrate that phosphorylation of HSP27 by MAPKAP-2, a downstream product of p38 MAPK, protects cardiac cells from DOX-induced ROS, revealing a novel property of small HSPs, i.e., these proteins can act as endogenous antioxidants against DOX-derived oxidants.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture and viability measurements. Cardiac H9c2 cells [catalog no. CRL 1446, American Type Culture Collection (ATCC), Rockville, MD] were cultured in DMEM (American Type Culture Collection) supplemented with 10% FBS (ATCC) and 1% antibiotic-antimycotic solution (100x; Sigma, St. Louis, MO). Jurkat cells, treated with campothecin, were used as internal standard in flow cytometry experiments. Jurkat cells were maintained in RPMI 1640-GlutaMax supplemented with 10% FBS, 1% antibiotic-antimycotic solution (100x), Na2HCO3 (1.5 g/l), glucose (4.5 g/l), 10 mM HEPES, and 1.0 mM sodium pyruvate. Cells were treated with DOX at 0–10 µM, which is the concentration range used in clinical settings (36), and cell viability was measured by cell counting and 3-(4,5 dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. The viable cell count was carried out in an automated cell counter (Nucleo, New Brunswick Scientific, Edison, NJ) according to the manufacturer's instructions. MTT assays were performed as described previously (4).

Hyperthermia (heat-shock) treatment. Cells grown in 150-cm2 flasks at 40–50% confluency were transferred to an incubator preset at 42°C in 95% air-5% CO2 and incubated for 2 or 4 h. Heat-shock temperature of 42°C was chosen with reference to previous studies reported in the literature (65). After heat-shock treatment, the flasks were reincubated at 37°C for ≥18–24 h to express HSPs.

Western blots. After appropriate treatment, the cells were lysed in RIPA buffer (1x PBS, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, 10 µl of 1x protease inhibitor cocktail, 1 mM PMSF, and 1 mM NaVO3). Normalized lysates were used for SDS gel electrophoresis or immunoprecipitated with appropriate antibodies. Total protein (20–30 µg) or the immunoprecipitates (30 µl) were resolved on a precasted 4–12% Bis-Tris acrylamide gel (Invitrogen) and transferred to a polyvinylidene difluoride membrane. After the membranes were blocked in 5% fat-free milk in TBS + Tween 20, they were incubated with respective primary antibody at appropriate dilution overnight at 4°C and then with horseradish peroxidase-linked secondary antibody (1:3,000 dilution).

p38 MAPK activity assay. A nonradioactive p38 MAPK assay was carried out to determine p38 MAPK activity in cells treated with different concentrations of DOX. This assay measures the activity of p38 MAPK through phosphorylation of activating transcription factor 2 (ATF-2), a substrate of p38 MAPK, by Western blot. The assay was carried out according to the manufacturer's protocol (p38 MAPK assay kit, Cell Signaling). Control and heat-shocked cells were treated with various concentrations of DOX as described above, and after 24 h of incubation, the cells were lysed and normalized for protein concentration as described above. Immobilized phosphorylated p38 MAPK (Thr180/Tyr184) antibody beads (20 µl) were added to the normalized samples, which were incubated at 4°C overnight. The beads were washed twice with 500 µl of kinase buffer and suspended in 50 µl of 1x kinase buffer supplemented with 200 µM cold ATP and 1 µg of ATF-2 (substrate) per tube and incubated at 30°C for 40 min. The reaction was terminated by addition of 20 µl of 3x SDS sample buffer and boiling at 100°C for 5 min. Equal amounts of samples were loaded on a precasted 4–12% Bis-Tris gel and subjected to electrophoresis and Western blotting. Phosphorylated ATF-2 (1:1,000 dilution) was detected by chemiluminescence.

Flow cytometry. DOX-induced apoptosis or necrosis was studied using the annexin V-propidium iodide (PI) apoptosis kit (Sigma). ROS generation was measured in cardiac H9c2 cells stained with 5,6-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate. In the dark, 5 µl of annexin V-FITC and then 10 µl of PI were added to the suspension. For dual-color analysis (FL1-H filter for annexin V-FITC and FL2-H filter for PI), 10,000 cells were subjected to fluorescence-activated cell sorting (FACS)-SCAN flow cytometry. As a positive control, Jurkat cells cultured as described above were treated with 10 µM campothecin for 4 h, pelletized, and treated with annexin V-FITC and PI, as described above. For dichlorodihydrofluorescein diacetate (DCFDA) staining, cells treated with DOX under appropriate conditions were washed with ice-cold PBS and treated with 10 µM 5,6-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate (Molecular Probes and Invitrogen) in serum-free medium for 45 min. Then the cells were trypsinized and suspended in PBS, and fluorescence was studied at 535 nm.

Electron paramagnetic resonance spin trapping. Cells in culture (75 cm2) were treated with DOX for 6 h, and the DOX-containing medium was exchanged with regular medium. Then the cells were collected from culture dishes by trypsinization, and viability was evaluated using the trypan blue exclusion method. The cells were resuspended in HEPES-buffered saline containing glucose (1 mg/ml), BSA (1 mg/ml), and diethylenetriaminopentaacetic acid (0.1 mM) and stored on ice. The required amount of sample was taken from the culture, incubated at 37°C in a water bath for 15 min, added to 50 mM spin trap [5',5-dimethyl-1-pyroline-N-oxide (DMPO) or 5-diethoxyphosphoryl-5-methyl-1-pyrroline-N-oxide], and transferred immediately to an electron paramagnetic resonance (EPR) flat cell and used in the measurements.

Microscopic fluorescence coimaging of HSP27 and F-actin. The effects of heat shock on the localization of HSP27 and stability of the F-actin cytoskeleton network in DOX-treated cells were studied by double-staining (phalloidin conjugated with FITC for F-actin and HSP27 antibody) fluorescence microscopy. Cardiac H9c2 cells grown in four-well chambers (fixed on a glass slide; LabTek, Fisher) were subjected to heat shock and then treated with DOX, as described above. After 12, 24, and 36 h of DOX treatment, the cells were fixed and permeabilized with 0.2% Triton X-100 in PBS for 10 min. Then the cells were washed with ice-cold PBS and immunostained with rabbit anti-HSP27 antibody (Stressgen, Victoria, BC, Canada) and incubated for 1 h. The excess antibody was washed off, and the cells were incubated with secondary anti-rabbit IgG antibody conjugated with Texas red (Stressgen) for 30 min at room temperature in the dark. The same cells were stained with phalloidin conjugated with FITC for visualization of the F-actin filaments. The slides were mounted using mounting medium (Molbiol-Calbiochem) and imaged using a fluorescence microscope (Nikon Eclipse 800). The images were analyzed using MetaMorph software (Universal Imaging).

Two-dimensional gel electrophoresis and matrix-assisted laser desorption/ionization time-of-flight analyses. DOX-treated control and heat-shocked cells were lysed and subjected to two-dimensional (2D) gel electrophoresis. Proteins in the 2D gels were identified by Western transfer or proteomics analysis using matrix-assisted laser desorption/ionization (MALDI) time of flight (MALDI-TOF). SDS was removed from the protein samples with a 2D clean-up kit (Bio-Rad), and the proteins were dissolved in the sample buffer (7 M urea, 2 M thiourea, 4% CHAPS, 0.5% ampholytes, and 20 mM DTT). An equal amount of protein was loaded on immobilized pH gradient (IPG) strips and focused with a Protean isoelectric focusing cell (Bio-Rad) using ampholines for separation of proteins by isoelectric focusing (isoelectric point 4–7) in one dimension. For 2D electrophoresis, the IPG strips were equilibrated for 10 min with 2% (wt/vol) DTT in 4.2 ml of an equilibration solution [0.38 M Tris base, pH 8.8, 6 M urea, 2% (wt/vol) SDS, and 20% (vol/vol) glycerol] per IPG strip and for 10 min with 2.5% (wt/vol) iodoacetamide in the equilibration solution. Each IPG strip was loaded onto a gel of the appropriate percentage of acrylamide, sealed with 1% agarose in buffer, and electrophoresed in a buffer that contained 25 mM Tris, 190 mM glycine, and 0.1% SDS. After electrophoresis, the proteins were fixed and visualized using colloidal Coomassie staining.

Stained gels were scanned with a computer-assisted densitometer (Versadoc 3000, Bio-Rad). 2D image analysis was performed using PDQuest software, and the proteins of interest were marked for excision. Spots of interest in the gels were marked, selected, and quantified. The density of the selected spots in each gel was normalized with respect to the background density. Selected gel spots from the PDQuest analysis were excised using a ProteomeWorks spot cutter (Bio-Rad). The gel plugs were transferred directly to a 96-well plate by the spot cutter. The 96-well plate was transferred to an autodigester and spotter (ProteomeWorks System, Perkin Elmer). The proteins were enzymatically digested, and the tryptic peptides were ZipTip purified. After ZipTip purification, the tryptic peptides were eluted from the ZipTip with 2 mg/ml cyano-4-hydroxycinnamic acid solution in 60% acetonitrile-0.2% formic acid and spotted directly onto a MALDI target. The tryptic peptides on the MALDI target plate were analyzed with a MALDI-TOF mass spectrometer (Waters, Milford, MA). Mass spectra were acquired in the positive-ion, delayed-extraction mode. All spectra were acquired with 20-kV accelerating voltage. Accuracy of masses for the trypsin autodigestion peptides and matrix peaks was checked for all spectra. If an inaccuracy was detected, the spectra were internally mass calibrated with the protonated molecular ions (M + H)+ of trypsin autodigestion peptides [mass-to-charge ratio (m/z) 515.33, 842.51, and 2211.10] and matrix peak (m/z 568.12). Data were analyzed using the web-based MS-Fit program.

Data analysis. Values are means ± SE. Statistical analysis was performed using Student's t-test and one-way ANOVA. The general acceptance level of significance was P < 0.05.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Attenuation of acute DOX toxicity in heat-shocked cardiac H9c2 cells. After control and heat-shocked cells were treated with 0–10 µM DOX for 6 h, the medium was replaced with regular medium. Cell viability was measured by MTT assay in DOX-treated cells at 6, 12, 24, and 36 h. Viability of DOX-treated control and heat-shocked cells after 24 h is shown in Fig. 1A. At all DOX concentrations, viability was significantly higher in heat-shocked than in control cells. These results indicate that hyperthermia protects cardiac H9c2 cells against DOX-induced toxicity.


Figure 1
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  		Fig. 1. Toxicity and reactive oxygen species (ROS) measurements in doxorubicin (DOX)-treated cardiac H9c2 cells. A: 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. Control and heat-shocked cells, at 70–80% confluency, were treated with 0.25–10 µM DOX for 6 h, incubated in drug-free medium for 24 h, and evaluated by MTT assay. Values (means ± SE from 3 sets of experiments) are presented as percentage of cell viability relative to non-DOX-treated control cells. At 0.25–10.0 µM DOX, significant protection was observed in heat-shocked cells (P < 0.01). B: fluorescence-activated cell sorting (FACS) analysis. Control and heat-shocked (HS) cells were treated with 0.25 µM DOX for 6 h, and DOX medium was exchanged with regular medium. After 24 h, cells were stained with 5,6-chloromethyl-2',7'-dichlorodihydrofluorescein (DCF) diacetate (CM-H2DCFDA). C: spin trapping of radicals. Electron paramagnetic resonance spectra of 5',5-dimethyl-1-pyroline-N-oxide (DMPO) spin adducts were obtained in control cells treated with 0.25 µM DOX. D: quantitative measures of DMPO spin adduct spectra in control and heat-shocked cells treated with 0.25 µM DOX. *P < 0.05 vs. control.

 
Heat shock attenuates DOX-induced ROS. ROS generation in DOX-treated heat-shocked and control cells was measured as an index of oxidative stress by flow cytometry using DCFDA staining under the same conditions used for viability measurements in Fig. 1A. Data from control and heat-shocked cells after 24 h of treatment with 0.25 µM DOX are shown in Fig. 1B. Similar results were obtained at all concentrations of DOX (0.5–10 µM). In the case of control cells, the fluorescent cell population shifts to the right (higher fluorescence), indicating that more ROS is generated. In the case of heat-shocked cells, the histogram shifts to the left, indicating that ROS generation is decreased in heat-shocked cells (Fig. 1B). These results are consistent with the finding from the MTT assay of viability that hyperthermia offers protection. Thus it appears that the increased viability in heat-shocked cells after 24 h of DOX treatment is likely due to the increased attenuation of DOX-mediated ROS generation.

DCFDA staining is nonspecific and reacts with free radicals, as well as H2O2, to yield fluorescence. We used EPR spin trapping to estimate the free radicals generated in DOX-treated control and heat-shocked cells. EPR spectra obtained from control cells that were treated with 0.25 µM DOX and spin trapped with 50 mM DMPO are shown in Fig. 1C. For non-DOX-treated cells, the amplitude of the EPR spectrum was very low. However, DOX increased the signal amplitude, indicating that free radicals were generated from redox cycling of DOX in the cells, as described previously (25). The DMPO spin-adduct spectrum shown in Fig. 1C is illustrative of an ·OH adduct. Although DMPO spin strap is not sensitive enough to distinguish whether ·OH or O2–· is generated in the DOX redox reaction, previous studies with a different spin trap, namely, 5-butyl-5-methyl-1-pyrroline 1-oxide (BMPO), showed that ·OH is indeed generated in endothelial cells by the redox reactions of DOX (25). For the heat-shocked cells, the magnitude of the observed EPR spin adduct spectrum significantly decreased (Fig. 1D), indicating that heat shock reduces free radical generation by DOX.

Increased expression of HSPs in the presence of DOX. DOX treatment is known to induce HSP expression. For determination of the level of DOX-induced HSP expression, control and heat-shocked cells were treated with different concentrations of DOX. The representative Western blots and quantitative results for HSP27 are illustrated in Fig. 2. Without DOX treatment (basal), HSP27 expression was higher in heat-shocked than in control cells. In control and heat-shocked cells, DOX increased HSP27 content. The increase was higher in the case of heat-shocked cells, indicating that these proteins are expressed in response to DOX treatment. A similar increase was observed with other major HSPs, such as HSP60, HSP70, and HSP90 (data not shown).


Figure 2
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  		Fig. 2. Top: representative Western blots of heat shock protein (HSP) 27 (HSP27) expression in DOX-treated control and heat-shocked cells. Control and heat-shocked cells were treated with 0.25–10 µM DOX for 6 h, and DOX medium was exchanged with regular medium. After 24 h, cells were lysed and used (20 µg of protein) for Western blots. The same membrane was reprobed for beta-actin to ensure equal loading of protein samples. Bottom: quantitative data (normalized with respect to non-DOX-treated control). Values are means ± SE from 3 sets of experiments. Open bars, control cells; solid bars, heat-shocked cells. *P < 0.05 vs. respective control.

 
Protection of F-actin integrity in heat-shocked cardiac cells. Control and heat-shocked cells treated with various concentrations of DOX were double-stained with FITC-tagged phalloidin for F-actin and Texas red-tagged IgG secondary antibody for HSP27, as described in MATERIALS AND METHODS. Fluorescence images of F-actin and HSP27 at two different excitation/emission wavelengths at 24 h of DOX treatment are superimposed in Fig. 3A. The high-resolution F-actin networks were observed in non-DOX-treated control cells (Fig. 3A, top). The number of F-actin filaments was diminished by DOX, especially at higher concentrations. These results are consistent with previous observations in DOX-treated Chinese hamster ovary AA8 (19) and HL60 (18) cells, which were rounded, had no stress fibers and a weak network of F-actin, and were rather disrupted (19). In HSP27 stained with Texas red-tagged secondary antibody, the cytosolic oligomeric form was translocated onto the nucleus, as shown by red nuclei in the images (Fig. 3A). Integrity of F-actin filaments was somewhat retained in heat-shocked cells, even at >10 µM DOX (Fig. 3A, bottom). Similarly, red fluorescence intensity for HSP27 was higher in heat-shocked than in control cells, as seen in the background (Fig. 3A). Phosphorylation of HSP27 in heat-shocked cells was evaluated by Western blotting in Fig. 3B. Because phosphorylation of Ser15 and Ser85 has been observed to cause deoligomerization of HSP27, specific antibodies for phosphorylation at Ser15 and Ser85 were used. In heat-shocked cells, phosphorylation of Ser15 and Ser85 increased about sevenfold (Fig. 3B).


Figure 3
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  		Fig. 3. A: fluorescence microscopic coimaging of F-actin filaments and HSP27 in control and heat-shocked cells. Control and heat-shocked cells were treated with DOX for 4 h and then incubated for 24 h in drug-free medium. Cells were treated with FITC-tagged antibody for F-actin and Texas red-tagged IgG secondary antibody, and fluorescence was measured. F-actin filaments are degraded by DOX, which is inhibited by heat shock. B: Western blots for phosphorylated Ser15 and Ser85 of HSP27 in control and heat-shocked cells treated with 0.25–10 µM DOX (top). Bottom: quantitative results from Western blot. Values are means ± SE from 3 independent experiments. Open bars, control, Phospho Ser15; gray hatched bars, control, Phospho Ser85; solid bars, heat-shocked, Phospho Ser15; black hatched bars, heat-shocked, Phospho Ser85. C: schematic illustration of inhibition of actin repolymerization by HSP27 and phosphorylated HSP27-assisted repolymerization in control and heat-shocked cells.

 
Reduced Bax-to-Bcl2 ratio in heat-shocked cells. The execution of apoptosis is regulated by specific apoptotic signaling molecules. To understand how heat shock-induced HSP27 interfered with DOX-induced apoptosis, we estimated the Bax and Bcl2 levels in DOX-treated control and heat-shocked cells. These pro- and antiapoptotic proteins were estimated using immunoblots of lysates of DOX-treated heat-shocked cells (Fig. 4A). Addition of DOX induced Bax, which increased with increasing DOX concentration, suggesting that Bax is involved in DOX-induced apoptosis signaling. The Bax-to-Bcl2 ratio, a marker of apoptosis, was consistently low in heat-shocked cells (Fig. 4B), indicating that heat shock had an antiapoptotic effect. This result is consistent with FACS analysis of DCFDA-stained cells (Fig. 1B), which showed a much greater reduction of fluorescence intensity after 24 h of drug treatment.


Figure 4
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  		Fig. 4. Bax-to-Bcl2 ratios. A: Bcl2 and Bax immunoblotted from heat-shocked cells (HS) treated with 0.25–10 µM DOX and blotted after 24 h of postdrug treatment. B: quantitative measures (normalized with respect to 0 µM DOX) of Bax-to-Bcl2 ratio for heat-shocked (solid bars) and control (open bars) cells.

 
Attenuation of DOX-induced apoptosis in heat-shocked cardiac H9c2 cells. DOX-treated control and heat-shocked cells were double-stained with annexin V and PI and subjected to FACS analysis for further evaluation of the role of apoptosis in cell death. The cells were treated with 0.25, 5, and 10 µM DOX and subjected to FACS analysis. The FACS measurements at higher concentrations were not reliable because of background red fluorescence from DOX, which interfered with PI fluorescence. Thus the FACS analysis of 0.25 µM DOX, where no significant background DOX fluorescence was observed, is presented in Fig. 5. The viable cell population was greater for heat-shocked than for control cells: 79.8 vs. 33.2%. Also, PI- and annexin V-positive staining revealed a significant difference in the cell population: 21.8 and 4.6% in control and heat-shocked cells, respectively (top right quadrants in Fig. 5, C and D). This indicates that heat shock protects cells from necrosis and that the apoptotic pathway is attenuated.


Figure 5
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  		Fig. 5. Flow cytometric analyses of control and heat-shocked cells double-stained with annexin-FITC and propidium iodide (PI). Cells were treated with DOX and double stained as described in MATERIALS AND METHODS. A: non-DOX-treated control cells. B: non-DOX-treated heat-shocked cells. There was no difference in the percentage of viability. C: control cells treated with 0.25 µM DOX for 6 h and assessed after 24 h. D: heat-shocked cells treated as described in C. Viability is increased in heat-shocked cells.

 
Increased p38 MAPK activity in heat-shocked cells. DOX activates various stress-induced kinases (14). Because attenuation of apoptosis and increased protection against DOX-induced toxicity are mainly due to increased expression of HSP27, we assessed the activity of p38 MAPK. Phosphorylation of Ser85 in HSP27 is the key factor in deoligomerization and relocation onto the F-actin networks (49). Immunoblots were carried out for total p38 MAPK, and p38 MAPK activity was measured by ATF-2 phosphorylation. Total p38 MAPK protein content was the same in control and heat-shocked cells at 0.25, 5, and 10 µM DOX (data not shown). However, p38 MAPK activity was proportional to the DOX concentration (Fig. 6). Activity of p38 MAPK was higher in heat-shocked cells at all DOX concentrations (Fig. 6). The quantitative measurements showed an almost fourfold increase in p38 MAPK activity in heat-shocked cells (Fig. 6B). It has been reported recently that the p38 MAPK {alpha}-isoform is proapoptotic, whereas the beta-isoform is prosurvival (26). To evaluate whether the beta-isoform is selectively activated in heat-shocked cells, we carried out studies with a selective inhibitor of the {alpha}-isoform, SB-203580. Addition of this inhibitor reduced the p38 MAPK activity in control and heat-shocked cells. However, even in the presence of this inhibitor, p38 MAPK activity was higher in heat-shocked cells (Fig. 6, A and B), indicating that, indeed, p38 MAPK beta-isoform activity is higher in heat-shocked than in control cells.


Figure 6
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  		Fig. 6. Nonradioactive p38 MAPK activity assay. p38 MAPK activity was measured in control and heat-shocked cells treated with 0.25–10 µM DOX. A: Western blots of phosphorylated activating transcription factor 2 (p-ATF-2, 37.5 kDa, a substrate for p38 MAPK) in control and heat-shocked cells. B: quantitative plots (normalized with respect control cells treated with 10 µM DOX) of p38 MAPK activity measured in the absence and presence of the selective p38 MAPK {alpha}-isoform inhibitor SB-203580 (SB) at 10 µM. High levels of phosphorylated ATF-2 indicate high p38 MAPK activity. Activity was higher in heat-shocked than in control cells at 0.25–10 µM DOX. Although addition of SB-203580 decreased activity in control and heat-shocked cells, activity was higher in heat-shocked cells, indicating higher p38 MAPK beta-isoform activity in heat-shocked cells.

 
Increased phosphorylation of HSP27 in heat-shocked cells. 2D gel electrophoresis was carried out to determine the different isoforms of HSP27. This protein has found to be phosphorylated at different sites by different protein kinases that are sensitive to hyperthermia (13). A 2D colloidal Coomassie blue-stained gel image of whole lysate of control cardiac cells treated with 0.25 µM DOX is shown in Fig. 7A. Previous 2D gel electrophoresis studies of heart tissues and cell lines reported largely diversified results for the number of HSP27 spots. Depending on the cell type, the number of isoforms with different phosphorylation status varied from 2 to 80 (10, 42, 44, 47). Especially in pathogenic tissues, increased numbers of HSP27 spots have been observed in 2D gels, likely because of disintegration of HSP27 oligomers and also different degrees of phosphorylation, such as, for example, mono-, di-, and triphosphorylation. The results in Fig. 7B are from Western transfer of 2D gels probed with HSP27-specific antibody. Once again, six spots indicate six isoforms of HSP27 in the cell lysate. All the lysates from control and heat-shocked cells treated with 0, 0.25, 5, and 10 µM DOX were subjected to similar 2D gel analysis. In all the gels, there were consistently six ~25-kDa spots. However, the intensity of each spot varied (Fig. 7C). In control cell lysate, 5.0 and 10 µM DOX increased the intensities of spots 4 and 6. Heat shock increased the intensities of all six spots at the basal level (no drug treatment). Also, DOX increased the intensities of spots 4 and 6 in heat-shocked cells (Fig. 7C).


Figure 7
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  		Fig. 7. Two-dimensional (2D) gel electrophoresis and matrix-assisted laser desorption/ionization time of flight (MALDI-TOF) analysis. A: 2D gel electrophoresis of control cells treated with 0.25 µM DOX. B: Western transfer of 2D gel probed with HSP25/27 antibody. C: quantitative measurements of all 6 isoforms of HSP27 from cells treated with 0.25–10 µM DOX. Spots in 2D gel were quantitated using PDQuest software and normalized with respect to spot of non-DOX-treated control. D: MALDI-TOF analysis of spot 1 digested from the gel. Identified peptides are shown. E: MALDI-TOF of spot 6. au, Arbitrary units.

 
The spots in the 2D gels (6 in total) were cut, digested, and subjected to MALDI-TOF for positive identification of the proteins, fragmented peptides, and phosphorylation status of each spot. The representative MALDI-TOF spectra of these six samples are presented in Fig. 7D. Analysis of the MALDI-TOF data confirmed that these spots indeed correspond to rat HSP27. In all six samples, one major peak at m/z 1149.63, corresponding to unphosphorylated peptide LFDQAFGVPR of residues 29–38, was observed with maximum intensity (Fig. 7, D and E). In spot 1, apart from this main peak, seven other peptides were positively recognized (Fig. 7D). None of these peptides was phosphorylated, indicating that this is an unphosphorylated HSP27 isoform (Table 1). The peptides analyzed from spot 2 showed phosphorylation at three sites: Ser85, Thr94, and Thr113. The peptide with m/z 1609.86 was identified as ALNRQLSphosSGVSEIR at residues 79–92. Ser85 has been identified as the site for phosphorylation by MAPKAP-2 (RxxS motif), which is downstream from p38 MAPK, indicating that p38 MAPK phosphorylates HSP27. However, another potential site of MAPKAP-2 phosphorylation, Ser15, was not phosphorylated for this sample. The other threonine phosphorylations are likely due to the other stress-induced kinases, such as JNK, inasmuch as these residues are identified to be at appropriate motifs. In the case of sample digested from spot 3, no phosphorylation was observed in the eight peptides identified. Analysis of sample 4 (spot 4) showed phosphorylation at Ser85, indicating MAPKAP-2 activity. Once again, Ser15 was not phosphorylated. However, analyses of spots 5 and 6 showed substantial phosphorylation. Spots 5 and 6 showed four phosphorylated sites: Ser9, Ser15, Ser85, and Thr116. In this case, Ser15 and Ser85 were observed to be phosphorylated by MAPKAP-2, whereas the other phosphorylations are due to the other protein kinases. From the literature, it is known that HSP27 exists as two isomers, {alpha} and beta, on 2D gel electrophoresis (9). Thus spots 1 and 3 are assigned to these two nonphosphorylated isoforms. Similarly, spots 2 and 4 are moderately phosphorylated samples (Ser85 is phosphorylated, but Ser15 is not), whereas spots 5 and 6 are due to the extensively phosphorylated isoforms (where Ser15 and Ser85 are phosphorylated by MAPKAP-2). Overall, these results indicate that the two isoforms of HSP27 are resolved as six spots because of their different phosphorylation status.


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  		Table 1. Number of HSP27 spots in 2D Western blots and their phosphorylation status in control and heat-shocked cells treated with 0.25 µM DOX

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
Cardiotoxicity of DOX has been a serious limitation in its use in treatment of cancer patients. Clinical and experimental models have demonstrated that increased oxidative stress, associated with a depleted antioxidant defense system at the subcellular level, plays a critical role in the development of cardiomyopathy (36). Various approaches have been suggested to avoid the cardiac toxicity, particularly, targeting and arresting the apoptotic signaling cascades (36). In the present study, we found that hyperthermia-induced adaptation before initiation of the treatment regimen could be a novel approach to minimize this side effect. The in vitro results of the present work show that heat shock-induced HSP27 can protect against DOX-induced toxicity in cardiac H9c2 cells. Several studies have shown that small HSPs can act as negative regulators of apoptotic and necrotic cell deaths. Mehlen et al. (35) showed that HSP27 blocks Fas/Apo-1- and staurosporine-induced cell death (35). On the other hand, HSP10 and HSP60 have been found to modulate DOX-induced Bcl2 family and mitochondrial apoptosis signaling in neonatal cardiomyocytes (45). Also, these proteins have been shown to attenuate the apoptotic pathway triggered by ischemia-reperfusion injury in cardiomyocytes (22). However, in our present approach, mild hyperthermia was used to induce small HSPs. This approach is relatively simple compared with other approaches reported in the literature, because hyperthermia is a clinically relevant modality and has been used in cancer clinics for a long time. Moreover, hyperthermia not only induces HSPs; it also simultaneously activates various stress-induced protein kinases, such as p38 MAPK, JNK, and PKC (50), some of which can phosphorylate HSPs. However, activation of MAPKs and induction of HSPs by hyperthermia are time-dependent processes. MAPKs are activated as an immediate response to heat stress, whereas small HSPs are induced 12–24 h after the heat stress and survive for up to 72 h (50). As shown in Fig. 7, p38 MAPK activity is not observed after 24 h of heat shock, whereas hyperthermia significantly increases all major HSPs (including HSP27; Fig. 2). Thus the only difference between DOX-treated control and heat-shocked cells in the present work is a greater abundance of HSPs in heat-shocked than in control cells at the time of DOX treatment. However, in the present work, DOX also activated p38 MAPK in control and heat-shocked cells. Indeed, p38 MAPK activity was higher in heat-shocked than in control cells (Fig. 6).

The increased cell viability and decreased ROS generation (Fig. 1, A and B) are clearly proof of our hypothesis that the protection in the heat-shocked cells is due to attenuation of ROS. This finding is very important, because suppression of DOX-induced ROS by hyperthermia can be an effective modality for minimizing the DCM and CHF that occur under in vivo conditions. Moreover, p38 MAPK activity in the presence of DOX is increased in heat-shocked cells (Fig. 6). Despite the increased p38 MAPK activity, cell viability was higher than control, suggesting that p38 MAPK activity is not completely related to transcription of apoptotic cell signaling. Recently, Okada et al. (38) reported that p38 MAPK plays a differential role in cardiomyocytes, protecting or killing, depending on its role in signal transduction (38). A recent report (33) suggests that the p38 MAPK beta-isoform is prosurvival. In aggreement with this report, our results, with addition of specific inhibitor (Fig. 6), showed that p38 MAPK beta-isoform activity is higher in heat-shocked than in control cells. Thus it appears that the increased activity of the p38 MAPK beta-isoform quenches the DOX-induced ROS generation and subsequent apoptosis in heat-shocked cells and maintains the HSP27 phosphorylation dynamics. Such increased p38 MAPK activity and phosphorylation of HSP27 are indeed expected to increase F-actin stability and dynamics, because the p38 MAPK-effected phosphorylation of Ser15 and Ser85 (as discussed below) deoligomerizes HSP27 and translocates it onto F-actin. This interpretation is indeed supported by the increased phosphorylation of HSP27 (in 2D gel electrophoresis and subsequent MALDI-TOF analyses, Fig. 7, C–E) and the greater stability of F-actin (Fig. 3).

MAPKs are known to phosphorylate small HSPs (15). Particularly, the downstream product of p38 MAPK, namely, MAPKAP-2, has been identified to phosphorylate Ser15 and Ser85 (RxxS motif) of HSP25/27 (15, 29). Such phosphorylation of HSP25/27 has been observed to functionalize the oligomeric inactive HSP25/27 in the cytoplasm to deoligomerize and translocate onto actin filaments and sarcomeres. Previous studies have shown that phosphorylation of Ser15 alone induced only a small effect on the oligomerization, whereas phosphorylation at Ser15 and Ser85 of HSP27 led to complete dissociation of oligomers (28, 41). This deoligomerization leads to translocation and stabilization of the F-actin networks (8) and, in turn, protects the cells. The existence of various isoforms with different phosphorylation status has been confirmed by 2D gel electrophoresis. Previous studies using 2D gel electrophoresis have established that phosphorylation status of HSP27 was significantly altered in pathogenic human hearts. Scheler et al. (44) reported that, in heart tissues affected by DCM and CHF, several isoforms of phosphorylated HSP27 were detected in 2D immunoblots, suggesting some form of HSP27 degradation in these hearts. They did not consider different isoforms with different phosphorylation status. Chen et al. (9) reported that the different spots detected in the 2D immunoblots are due to different isoforms with different phosphorylation status. The results of 2D gel analysis and MALDI-TOF in the present study provide evidence for the existence of six isoforms with different phosphorylation status. Interestingly, such phosphorylation has been noticed, even in control cells treated with a higher concentration of DOX. However, the native concentration of this protein is not adequate to protect the cells; hence, p38 MAPK targets another caspase-signaling cascade, inducing apoptosis (Fig. 8).


Figure 8
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  		Fig. 8. Schematic illustration of proposed mechanism of protection from DOX toxicity. DOX-induced cell death occurs through activation of stress-induced protein kinases and subsequent signal transduction for apoptosis. Hyperthermia induces similar stress-induced kinases and small HSPs, which can be phosphorylated by MAPK. Phosphorylated small HSPs protect cells. DCFDA, dichlorodihydrofluorescein diacetate.

 
The actin filament network in cardiomyocytes plays a critical role in coordination with troponin in controlling contraction (34). In vivo, loss of the F-actin network in cardiomyocytes may be responsible for loss of myofibril tension and contraction ability because of development of cardiomyopathy. In the present work, microscopic imaging of DOX-treated cells showed a reduction in the integrity of actin filaments, whereas in the case of heat-shocked cells, the F-actin network was retained to some extent at all DOX concentrations (Fig. 3). Many studies have shown an association between the phosphorylation status of HSP27 and {alpha}B crystalline (both are small HSPs) and F-actin stability (20, 32). Many earlier studies reported inhibition of actin polymerization by HSP27 and HSP25 and, thus, enhanced actin depolymerization (7). However, recent studies have reported that wild-type recombinant HSP25 does not affect actin polymerization, whereas after phosphorylation by MAPKAP-2 kinase, HSP25 starts to activate actin polymerization. Increased expression of HSP25/27 has been reported to protect actin filaments from fragmentation and preserve the contacts with the cell membranes (30, 43). Mutant HSP25/27 with replacement of Ser (which can be phosphorylated by MAPKAP-2) by Ala or Gly was unable to protect actin from different stresses (30, 43). Thus there is no consensus on the effect of HSP25 on actin polymerization. On the basis of these facts, Gusev et al. (20) supposed that, under normal conditions, the extent of HSP27 phosphorylation is low; therefore, it does not interact with membranes and remains in the cytosol. This unphosphorylated HSP27 interacts with the barbed end of actin filaments and prevents actin polymerization. On phosphorylation, its interaction with actin is reduced and, therefore, its inhibitory effect on actin polymerization is decreased. Thus retention of the F-actin network in the present work could be explained as follows (Fig. 3). In the fluorescence images obtained for actin filaments and HSP27 on treatment with DOX, the F-actin filaments vanish, and HSP27 translocates onto the nucleus; in control cells, this fragmented F-actin is stabilized by the unphosphorylated oligomeric HSP27. However, in the case of heat-shocked cells, the fragmented F-actin is reconstituted, because phosphorylated HSP27 does not interfere with actin polymerization; it simply translocates onto the nucleus, as confirmed by increased red fluorescence from the nuclei of the heat-shocked cells (Fig. 3).

Overall, the attenuation of ROS generation and phosphorylation of HSP27 in the heat-shocked cells appear to play a key role in attenuation of apoptosis and F-actin remodeling. However, we are aware of no mechanism whereby the HSPs can minimize the ROS generation from DOX. We recently reported that mitochondrial aconitase activity could be attenuated by hyperthermia-induced HSPs (24). This aconitase is known to play a significant role in mediating the DOX-induced toxicity through iron regulatory proteins (IRP1 and IRP2) (36). The 4[Fe-4S] cluster in aconitase has been found to be susceptible to O2–· (formed in the DOX redox reaction and the metabolites) attack, releasing one Fe(II), resulting in [3Fe-4S] species and a free Fe(II). This aconitase-released iron is active for redox conversion. On the other hand, the DOX metabolites have been reported to release all four Fe(II) from the aconitase, converting it to IRP-1, which binds with iron-regulating elements in the untranslated regions of transferrin receptor and ferritin mRNAs. When activated, IRPs enhance transferrin receptor mRNA stability and block ferritin mRNA translation, thereby facilitating iron uptake over sequestration. Thus we propose that hyperthermia-induced aconitase inactivation could be a mechanism of reduced DOX toxicity in the heat-shocked cells.

In conclusion, the present study has demonstrated that hyperthermia-induced HSP27 and its phosphorylation by MAPKAP-2 protects cardiac H9c2 cells, because phosphorylated HSP27 acts as an endogenous antioxidant against the DOX-derived oxidants, such as oxygen free radicals and H2O2. Perhaps, the present study has the following limitations as well: although we have followed the HSP27-involved pathway in the present work, hyperthermia also induces other HSPs; hence, their roles in the observed effect remain to be elucidated. Further studies with HSP27-overexpressed/silenced animal models and their comparison with present results may be required to confirm the proposed mechanism.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported by National Institutes of Health Grants R21 EB-004658 and R01 HL-078796-02 and American Heart Association Grant BGIA 0365203B (to G. Ilangovan).


   ACKNOWLEDGMENTS
 
The services of Davis Heart and Lung Research Institute core facilities, namely, proteomics, flow cytometry, and microscopy, are gratefully acknowledged.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. Ilangovan, The Ohio State Univ., 420 West 12th Ave, Rm. 116A, Columbus, OH 43210 (e-mail: Govindasamy.Ilangovan{at}osumc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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 GRANTS
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