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Am J Physiol Heart Circ Physiol 288: H486-H496, 2005. First published September 16, 2004; doi:10.1152/ajpheart.00437.2004
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Increased expression of poly(ADP-ribose) polymerase-1 contributes to caspase-independent myocyte cell death during heart failure

Jyothish B. Pillai, Hyde M. Russell, Jai Raman, Valluvan Jeevanandam, and Mahesh P. Gupta

Department of Cardiothoracic Surgery, Committee on Molecular Medicine, University of Chicago, Chicago, Illinois

Submitted 13 May 2004 ; accepted in final form 7 September 2004


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Poly(ADP-ribose) polymerase-1 (PARP-1) plays a pivotal role in regulating genome stability, cell cycle progression, and cell survival. However, overactivation of PARP has been shown to contribute to cell death and organ failure in various stress-related disease conditions. In this study, we examined the role of PARP in the development and progression of cardiac hypertrophy. We measured the expression of PARP in mouse hearts with physiological (swimming exercise) and pathological (aortic banding) cardiac hypertrophy as well as in human heart samples taken at the time of transplantation. PARP levels were elevated both in swimming and banded mice hearts and demonstrated a linear positive correlation with the degree of cardiac hypertrophy. A dramatic increase (4-fold) of PARP occurred in 6-wk banded mice, accompanied by apparent signs of ventricular dilation and myocyte cell death. PARP levels were also elevated (2- to 3-fold) in human hearts with end-stage heart failure compared with controls. However, we found no evidence of caspase-mediated PARP cleavage in either mouse or human failing hearts. Overexpression of PARP in primary cultures of cardiac myocytes led to suppression of gene expression and robust myocyte cell death. Furthermore, data obtained from the analysis of PARP knockout mice revealed that these hearts produce an attenuated hypertrophic response to aortic banding compared with controls. Together, these results demonstrate a role for PARP in the onset and progression of cardiac hypertrophy and suggest that some events related to cardiac hypertrophy growth and progression to heart failure are mediated by a PARP-dependent mechanism.

hypertrophy; myocyte cell death

HEART FAILURE remains an important clinical and societal problem in developed nations. On the basis of animal and human studies, it has been documented that failing hearts are associated with a striking reduction of cardiac-specific gene expression, degeneration of myocytes, and increased interstitial fibrosis (1, 15, 19, 26). However, the molecular events leading to this complex clinical syndrome are still not known.

Poly(ADP-ribose) polymerase-1 (PARP-1) is a (113 kDa) prototype member of the PARP family of DNA-bound enzymes located in the nuclei and mitochondria of various cells, including cardiac myocytes (11, 34). In Drosophila, elimination of the PARP gene is lethal, but not in mice, which have at least seven different members of the PARP family (28). The NH2-terminal region of PARP-1 (subsequently referred to as PARP) contains a caspase-sensitive site. Cleavage at this site generates a ~85-kDa prominent band, which is considered a hallmark of apoptosis (6). PARP catalyzes the transfer of successive units of the ADP-ribose moiety from nicotinamide adenine dinucleotide (NAD+) to target proteins, a process called poly ADP-ribosylation (21). Under homeostatic conditions, PARP participates in regulation of many cellular processes, including DNA repair, gene transcription, cell cycle progression, cell survival, chromatin remodeling, and genome stability (6, 45, 48). However, overactivation of PARP threatens cell survival as it consumes cellular NAD+ content to add extended chains of ADP-ribose moieties on the target proteins (6, 45). Because NAD+ is essential for mitochondrial electron transport, depletion of NAD+ rapidly leads to energy deficit and eventually to cell death (17). PARP can also cause cell death by apoptosis, but without activation of caspases (46). Among the intracellular signals, oxygen- and nitrogen-derived free radicals and nuclear accumulation of Ca2+ have been shown to increase PARP activity (20, 45). Data obtained from the use of PARP inhibitors and mice lacking the PARP gene have indicated that overactivation of PARP contributes to cell dysfunction and death in various pathological conditions associated with oxidative cell stress, including reperfusion injury, inflammation, diabetes mellitus, circulatory shock, and stroke (3, 11, 30, 41, 50). However, to date, the potential role of PARP in the development of cardiac hypertrophy and failure has not been studied.

Here, we report that PARP is not cleaved but rather that a robust expression of PARP occurs in failing hearts of both animals and humans. Overexpression of PARP markedly represses cardiac gene transcription and eventually leads to cell death. By analyzing PARP–/– mice, we further present evidence showing that PARP contributes to the pathogenesis of heart failure. These data demonstrate that PARP-mediated cell death could be a novel (caspase independent) mechanism contributing to myocyte cell loss during the progression of cardiac hypertrophy to heart failure.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
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Aortic banding. Adult mice (CD1, n = 40) weighing 30–40 g were anesthetized and ventilated. The chest was opened at the second intercostal space. The aorta was isolated from the adjacent tissue and banded between carotid arteries over a 27-gauge needle, which was immediately removed (31). Animals that had sham surgery underwent an identical procedure with the exception of band placement. Animals were studied from day 1 to 6 wk postsurgery. The PARP–/– mice were purchased from The Jackson Laboratories (www.jax.org).

Swimming protocol. Adult mice (CD1, n = 20) were subjected to a swimming exercise program (25). Briefly, animals were initially exercised for 30 min twice daily, with increments of 10 min daily. The final duration of exercise was 90 min, twice daily, for 8 wk. Development of cardiac hypertrophy was monitored by obtaining echocardiographic measurements. The Institutional Animal Care and Use Committee of the University of Chicago approved all the animal protocols.

Human failing heart samples. Left ventricular (LV) samples were obtained from 14 patients with end-stage heart failure at the time of heart transplantation. Patients studied had either ischemic cardiomyopathy (n = 7), idiopathic dilated cardiomyopathy (n = 5), or restrictive cardiomyopathy (n = 2). Control myocardial specimens were obtained from five patients with nonfailing hearts who underwent valve surgery. All samples were immediately frozen in liquid nitrogen and stored at –80°C until analyzed. Patients received different combinations of the following pharmacological agents: digoxin, dobutamine, angiotensin-converting enzyme (ACE) inhibitors, Ca2+ channel blockers, {beta}-adrenergic blockers, and diuretics. The University of Chicago Institutional Review Board approved all procedures involving the use of human tissue.

Western blot analysis. Whole cell extracts of mouse and human LVs were made in a urea extraction buffer [62.5 mM Tris (pH 6.8), 6 M urea, 10% glycerol, 2% SDS, 0.003% bromophenol blue, 5% 2-mercaptoethanol, and a cocktail of protein inhibitors]. The tissue was crushed to form powder using a precooled pestle and mortar, followed by sonication for 20 s. The protein concentration of the samples was measured using Bio-Rad protein assay dye reagent. The Westen blot analysis of proteins was carried out essentially as described previously (9). Anti-PARP-1, anti-caspase-3, and anti-apoptosis inducible factor (AIF) antibodies were purchased from Santa Cruz Biotechnology. The monoclonal anti-poly(ADP-ribose) antibody was purchased from Alexis Biochemicals.

Immunohistochemistry. Thin cryosections of heart tissue were fixed in cold acetone for 15 min. Sections were repeatedly washed in PBS wash buffer (PBS and 1% NP-40), blocked with 10% goat serum for 1 h at room temperature, and then incubated with the anti-poly(ADP-ribose) (1:100) antibody for 2 h. After the sections were washed extensively in wash buffer, the immunoreactivity was detected using horseradish peroxidase-conjugated goat anti-mouse IgG (1:500). Color was developed using diaminobenzidine as a substrate. Sections were counterstained with nuclear fast red to visualize nuclei. Some heart sections were also stained using a standard hematoxylin and eosin (H&E) and/or Masson trichrome staining protocol (Poly Scientific; Bay Shore, NY). Heart sections were viewed under a Zeiss Axioskop microscope equipped with x10 Acroplan (0.25 numerical aperture) objective lens with an AxioCam color camera and the ocular power of x10 (i.e., x100 magnification).

Terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling staining. Myocyte cell death in heart sections was detected using the terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick-end labeling (TUNEL) technique with Cardio-TACS (Trevigen; Gaithersburg, MD) according to the manufacturer's instructions. In this procedure, nuclei undergoing apoptosis stained blue (TUNEL positive). Nuclear fast red was used as a counterstain to visualize nuclei. Positive controls included samples treated with nuclease-1, and negative controls lacked the TdT enzyme in the labeling reaction. Sections were also stained with H&E for comparison. TUNEL-positive myocytes were determined by randomly counting 500 cells in 10 fields. The index of apoptosis was calculated as the number of apoptotic myocytes per total number of myocytes x 100.

Primary cultures of cardiac myocytes and the transient transfection assay. Primary cultures of cardiac myocytes were prepared from 2-day-old neonatal rat hearts as previously described (9). Myocytes were grown in DMEM supplemented with 10% fetal bovine serum and a mixture of 5 mg/ml penicillin and streptomycin (Life Technologies). Cells were transfected 48 h after being plated using Tfx-TM20 reagent (Promega) according to the manufacturer's protocol. After 48 h of transfection, cells were harvested, and cell lysate was prepared and assayed for luciferase (Luc), {beta}-galactosidase ({beta}-gal), and protein content.

Annexin V, Hoechst 33342, and propidium iodide staining. Primary cultures of cardiac myocytes were transfected with a PARP-expressing vector. On the third day after transfection, cells were stained with Annexin V-FITC, Hoechst dye, and/or propidium iodide (PI) to detect cell viability according to the manufacturer's protocol (Santa Cruz Biotechnology and Molecular Probes). For a positive control, cells were treated with camptothecin (2 µM) for 5 h. Cells transfected with an empty vector lacking PARP cDNA were used as negative controls. After cells were incubated for 15 min with the dye reagent, they were washed with PBS and visualized under the fluorescence microscope.

RNA extraction and Northern blot analysis. Total RNA was extracted from control and banded mouse hearts with TRIzol reagent (Life Technologies) according to methods provided by the manufacturer. Northern blot analysis was performed using mouse atrial natriuretic factor (ANF) and GAPDH cDNA probes and synthetic oligonucleotide probes for mouse {alpha}-myosin heavy chain (MHC), {beta}-MHC, and brain natriuretic peptide (BNP). The sequences of oligonucleotide probes were as follows: {alpha}-MHC, 5'-CGA ACG TTTA TGT TTAT TGT GGA TTG GCC ACA GCG AGG GTC TGC TGG AGA GGT TAT TCC TCG TC-3'; {beta}-MHC, 5'-GAG GGC TTC ACG GGC ACC CTT AGA GCT GGG TAG CAC AAG ATC TAC TCC TCA TTC AGG CC-3'; and BNP, 5'-CAG CTT GAG ATA TGT GTC ACC TTG GAA TTT TGA GGT CTC TGC TGG ACC CGG AGG GTG CTG-3'.

Scanning densitometry and statistical analysis. Autoradiograms were scanned using Scion Image for Windows analysis software, based on NIH Image for the Macintosh by Wayne Rasband (National Institutes of Health; Bethesda, MD). Signal intensity was adjusted for background density of the blot. Student's paired t-test was utilized to determine the statistical significance between two groups.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Induction of PARP expression during cardiac hypertrophy. To understand the mechanisms of cell growth during hypertrophy, we analyzed the expression of PARP in settings of both physiological and pathological hypertrophy. Mice were subjected to forced swimming for 8 wk to induce physiological hypertrophy and to aortic banding of different time periods to produce pressure overload (pathological) hypertrophy. Hearts with pressure overload hypertrophy displayed marked changes in cardiac gene expression. Expression of atrial natriuretic peptide (ANP), BNP, and {beta}-MHC genes were all elevated in pressure overload hypertrophy. In contrast, expression of these genes was not altered in the hearts of swimming mice. In hearts of banded mice, changes in ventricular thickness and chamber dilatation could be detected as early as 1 wk after mice were banded, but it was more pronounced at 6 wk, with the latter showing clear evidence of LV dilatation consistent with symptoms of clinical heart failure. The heart weight-to-tibia length ratio increased by 70% in 6-wk banded mice, whereas it was increased only by 15% in swimming mice.

Nuclear extracts of these hearts were subjected to Western blot analysis to detect expression level of PARP and poly-ADP-ribosylation of nuclear proteins using anti-PARP and anti-ADP-ribose antibodies, respectively. As shown in Fig. 1, PARP expression was elevated in hearts of both swimming mice and banded mice compared with their respective controls. A dramatic increase (4-fold) of PARP occurred in mice banded for 6 wk; however, no cleavage of PARP was observed in either of these hearts. The increase in PARP at different time points after aortic banding showed a linear correlation with the degree of cardiac hypertrophy (Fig. 1D). To demonstrate a change in the catalytic activity of PARP during hypertrophy, we analyzed poly-ADP-ribosylation of nuclear proteins. Results showed enhanced protein ADP-ribosylation as early as 1 day after mice were aortic banded, which increased gradually with the time of aortic banding. At 4 and 6 wk of aortic banding, when signs of LV dilatation became apparent, a massive increase in protein ADP-ribosylation was observed in every heart we analyzed. A mild increase of protein poly-ADP-ribosylation was also detected in the hearts of swimming mice (Fig 1C). To further confirm these results, we performed immunohistochemical staining of heart sections with an anti-ADP-ribose antibody. As shown in Fig 1F, an increased nuclear staining for protein poly-ADP-ribosylation was detected throughout the muscle sections of 6-wk banded mice compared with controls (Fig 1E).



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  		Fig. 1. Increased activity of poly(ADP-ribose) polymerase-1 (PARP-1) during physiological and pathological cardiac hypertrophy. Cardiac nuclear extract of mice subjected to swimming or aortic banding was analyzed by Western blot analysis using anti-PARP-1 (A and B) or anti-ADP-ribose antibodies (C). Anti-GAPDH antibody was used as a reference control. D: linear regression analysis of PARP activity with degree of cardiac hypertrophy in mice with aortic banding. Heart sections of control (E) and 6-wk banded (F) mice were immunostained for poly-ADP-ribosylation of nuclear proteins. Note the increased brown color in F (arrow) showing poly-ADP-ribosylation of nuclear proteins.

 
Increased expression of PARP in human failing hearts. To examine the role of PARP in clinical heart failure, we analyzed human failing heart samples. Of 14 failing hearts analyzed, 7 were diagnosed with ischemic cardiomyopathy, 5 had dilated cardiomyopathy, and 2 had restrictive cardiomyopathy. Control LV samples (n = 5) were obtained from patients undergoing valvuloplasty who showed no clinical sign of heart failure. All samples were minced in a homogenization buffer and subjected to Western blot analysis. A full-length 113-kDa PARP band was detected in all human samples analyzed, and it was consistently increased from two- to threefold in failing hearts compared with controls (Fig. 2A). However, we did not detect a cleaved fragment of PARP in either of these hearts, consistent with the results of mouse heart studies. To examine whether PARP of human cardiac samples was indeed sensitive to caspase cleavage, we digested one sample with active caspase-3 and analyzed protein cleavage by Western blot analysis. As shown in Fig. 2 (lane 7), an ~85-kDa band was observed, which was associated with a proportionate decrease of 113-kDa full-length PARP, indicating that PARP present in these samples was indeed sensitive to caspase cleavage. Experiments carried out to test the enzymatic activity of PARP also revealed increased poly-ADP-ribosylation of nuclear proteins in failing hearts compared with controls (Fig. 2B). Together, these results indicated that PARP expression and enzymatic activity are elevated in failing hearts.



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  		Fig. 2. Increased activity of PARP in human failing hearts. A: Western blot analysis of whole cell extract with anti-PARP antibody. Lanes 1 and 2 show nonfailing (NF) controls; lanes 3–6 show end-stage failing hearts. Lane 7 shows a cleaved fragment of PARP (~85 kDa) after incubation of the extract with caspase-3. B: Western blot analysis of extracts with anti-ADP-ribose antibody. C: loading control. D: quantification of PARP activity (means ± SE) in nonfailing (n = 5) and failing (n = 14) hearts. *P < 0.01.

 
Increased expression of PARP caused myocyte cell death. PARP has been shown to play a pivotal role in cell survival and death (6, 21). To test whether hearts with increased activity of PARP were associated with cell degeneration, we examined myocyte cell death by TUNEL staining of heart sections. For positive controls, heart sections were incubated with nuclease-1, which induced massive DNA damage, and cells became heavily positive to TUNEL staining (Fig. 3C). In 6-wk banded hearts, TUNEL-positive myocytes were seen scattered throughout the heart section (Fig. 3B). Although few TUNEL-positive cells were also detected in hearts of swimming mice, they were not different from control mice, consistent with previous reports (27), whereas in the 6-wk banded mice with dilated hearts, the TUNEL-positive cells were far greater in number (>20-fold) compared with controls. In these heart samples, we also analyzed markers of apoptotic cell death, viz., AIF and caspase-3. However, no change was observed in the expression levels of these enzymes among different hearts (Fig. 3D). Although procaspase-3 (32 kDa), the precursor of caspase-3, was detected, no active form of caspase-3 (20 kDa) was found in these samples, thus suggesting the involvement of a caspase-independent mechanism of myocyte cell death in our models of heart failure.



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  		Fig. 3. Myocyte cell death in hearts expressing high levels of PARP. A–C: terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) staining of heart sections of control (A), 6-wk banded mouse (B), and nuclease-1-treated sections (C). Blue nuclei are positive for TUNEL staining. D: Western blot analysis of different mouse heart samples for apoptosis inducible factor (AIF) and caspase-3 using protein-specific antibodies. Arrows in B show TUNEL-positive cells in a hypertrophic heart.

 
To confirm the role of PARP in myocyte cell death, we overexpressed PARP in primary cultures of cardiac myocytes. As a positive control, cells were treated with camptothecin, a known apoptosis-inducing agent. Cell death was examined by Annexin V-FITC, Hoechst 33342, and PI labeling of cells. Annexin V binds to negatively charged phospholipids, including phospholipid serine (PS), which is located in the cytoplasmic interface of the cell membrane. During early events of cell death, the loss of membrane asymmetry exposes PS to the extracellular space and makes it accessible to Annexin V labeling, which gives the intense green color of the FITC tag. Hoechst and PI are two DNA-binding dyes that give a blue and red color, respectively. Hoechst readily penetrates the cell membrane and stains the nuclei of both live and dead cells. PI, which does not penetrate the cell membrane, stains nuclei of dead cells red, whereas living viable cells remain unstained. As shown in Fig. 4A, cells transfected with the PARP (5 µg) expression plasmid shrank, became round, and stained positive to Annexin V as early as 12 h after transfection, but not in the negative controls transfected with the mock plasmid. Camptothecin-treated cells stained positive to Annexin V as early as 45 min after stimulation. To confirm that PARP was indeed overexpressed in these transfections, we also stained a few plates with anti-PARP antibody using phycoerythrin-conjugated IgG as a secondary antibody, which gives an intense red color of the immune complex. As shown in Fig. 4B, PARP is highly expressed in transfected cells and localized primarily to cell nuclei. To quantify PARP-mediated cell death, we overexpressed different amounts of PARP (1–5 µg) in cardiac myocytes, and cells were double stained with Hoechst and PI dyes 48 h after transfection. Hoechst dye (blue) stains all cells (both live and dead) attached to the bottom of the plate, whereas PI (red) stains only dead cells; therefore, the subtraction of red cells from blue cells gives the number of viable cells in a given field. As shown in Fig. 5, PARP overexpression resulted in concentration-dependent cell death of myocytes. In plates transfected with 1 µg of PARP expression plasmid, ~20% myocyte stained positive to PI stain, whereas at 2 µg of the plasmid, the number of PI-positive cells rose to >50%. At higher amounts of PARP (5 µg), >50% cells detached from the bottom of the plate and almost all remaining cells became round and stained positive to both Hoechst and PI dyes, indicating massive myocyte cell death by PARP overexpression. Again, in the negative controls, no cell death could be detected in cultures transfected with the empty vector lacking PARP cDNA. These results thus demonstrate that PARP overexpression induces robust myocyte cell death in cultures.



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  		Fig. 4. PARP overexpression induces myocyte cell death. A: control cardiac myocytes (top), camptothecin-treated cells stained 45 min after stimulation (middle), and cells overexpressing PARP stained 12 h after transfection (bottom). Cells were stained with Annexin V-FITC and visualized under a fluorescence microscope. B: cardiac myocytes overexpressing PARP were immunostained with anti-PARP primary antibody and phycoerythrin-conjugated IgG (red) as secondary antibody. Arrows indicate the position of nuclei in the cell where PARP expression is detected most.

 


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  		Fig. 5. A: cardiac myocytes were transfected with different amounts of PARP expression plasmid or with a control vector lacking PARP cDNA (p-dp or pCMV-dp). Forty-eight hours after transfection, cells were doubly stained with Hoechst dye (stains both live and dead cells) and propidium iodide (PI), a dead cell stain. Note that at the 5-µg amount of PARP DNA, almost all cells became positive to both stains, showing massive cell death. B: quantification of cell death after PARP overexpression. Cells were counted in at least 3 different fields in each plate. Values are means ± SE of 4 different transfections.

 
To examine the mechanism of cell death by PARP, we analyzed the effect of PARP overexpression on the transcription activity of different cardiac-specific and -nonspecific gene promoters. Cells were transfected with and without PARP expression vectors (0.5–1.0 µg) together with promoter/reporter plasmids, viz., {alpha}-MHC/Luc, skeletal {alpha}-actin/Luc, 5xCArG-heat shock protein (HSP)/{beta}-gal, and HSP/{beta}-gal reporter plasmids (9). All transfections were repeated at least three times with different preparations of DNA. The reporter gene activity normalized for the protein content is shown in Fig. 6. PARP markedly repressed (40–80%) transcription activity of both cardiac-specific (e.g., {alpha}-MHC and skeletal {alpha}-actin) and -nonspecific (5xCArG-HSP/{beta}-gal and HSP/{beta}-gal) promoters in a concentration-dependent manner from 0.5 to 1.0 µg of PARP plasmid, indicating that full-length PARP has a profound negative gene-regulatory effect in cardiac myocytes. To examine whether PARP-mediated repression of gene activity was a result of energy deficit, we measured ATP content of PARP-transfected cardiac myocytes. The results indicated that the amount of PARP that repressed gene transcription by 80% had no noticeable effect on the cell ATP contents, thus excluding the possibility of energy depletion as a cause of gene repression by PARP. From these results, we deduce that the PARP-mediated myocyte cell death could be, in part, due to a loss of basic transcriptional support of myocytes.



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  		Fig. 6. Overexpression of PARP represses gene transcription. Primary cultures of cardiac myocytes were transfected with increasing amounts of PARP expression vector (50 ng–1.0 µg) together with 2.0 µg of reporter plasmids, viz., {alpha}-myosin heavy chain (MHC)-luciferase (Luc), skeletal {alpha}-actin-Luc, 5xC(Ar)G heat shock protein (HSP)-{beta}-galactosidase ({beta}-gal), or HSP-{beta}-gal. Bars show normalized Luc activity to protein content (means ± SE) derived from 5 different transfections.

 
Attenuation of cardiac hypertrophy in PARP knockout mice. To further substantiate a role of PARP in the development of cardiac hypertrophy, we studied mice lacking the PARP gene (PARP–/– mice). In general appearance, PARP–/– mice are smaller in size and sluggish compared with PARP+/+ mice of the same age. They are less fertile compared with their wild-type counterparts. Weight-matched PARP–/– and PARP+/+ mice were subjected to 6 wk of aortic banding, and the development of cardiac hypertrophy was analyzed using different histological and biochemical markers. As shown in Fig. 7, PARP–/– mice did produce cardiac hypertrophy (20%); however, it was far less than that which occurred in PARP+/+ mice (70%). On the basis of the heart weight-to-tibia length ratio, there was an ~60% reduction of cardiac hypertrophy in PARP–/– mice compared with PARP+/+ mice. The rate of animal survival after aortic banding was also significantly different between the two groups of mice. Most of the PARP–/– animals (>75%) that survived the first 2 days of aortic banding remained alive beyond 6 wk, whereas >50% of the PARP+/+ mice died within the 6 wk of study (Fig. 8). PARP+/+ mice that survived 6 wk of aortic banding had a significant amount of LV dilatation, whereas no apparent sign of ventricular dilation was seen in PARP–/– mice (Fig. 8). The histological analysis of mouse heart sections showed disorganization of myofibers, scattered vacuoles, and intense Masson's trichrome blue staining in PARP+/+ mice, whereas these changes were either undetectable or of markedly less magnitude in PARP–/– mice, suggesting increased interstitial fibrosis in the wild-type but not in PARP-negative mice during the course of pressure overload hypertrophy (Fig. 8). Attenuation of cardiac hypertrophy in PARP–/– mice was also evident from the analysis of hypertrophic marker genes. In PARP+/+ banded mice, the levels of ANF, BNP, and {beta}-MHC mRNA were highly elevated, whereas {alpha}-MHC levels were repressed, as expected. On the contrary, in PARP–/– banded mice, no repression of {alpha}-MHC levels was observed, and the levels of ANF, BNP, and {beta}-MHC were increased to a much lesser extent (Fig. 7). From these results, we conclude that PARP participates in certain events of cardiac hypertrophy, particularly in myocyte cell degeneration, interstitial fibrosis, and subsequent ventricular dilation, which are signs of propensity toward heart failure.



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  		Fig. 7. Attenuation of cardiac hypertrophy in PARP–/– mice. A: heart weight-to-tibia length ratio (HW/TL) in PARP–/– and PARP+/+ mice. Values are means ± SE of 7–10 different animals. B and C: RNA analysis of hypertrophy marker genes from control (C) and banded (B) mice of PARP+/+ and PARP–/– groups. The change in RNA expression was quantified from 3 different blots. BNP, brain natriuretic peptide; ANF, atrial natriuretic factor.

 


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  		Fig. 8. Prevention of fibrosis in PARP–/– mice after aortic banding. A: heart sections of 6-wk banded mice stained with Masson trichrome stain, which includes hematoxylin and eosin (cyan) as nuclear stain, scarlet acid fuchin (red) as cytoplasm and muscle stain, and aniline blue as collagen stain. Note the degeneration of myocytes and intense interstitial fibrosis (blue) in PARP+/+ 6-wk banded mice. B: animal survival in PARP–/– and PARP+/+ groups of mice after 6 wk of aortic banding. Bars represent means ± SE of 5–25 animals in different groups. C: change in left ventricular (LV) dimension in 2 groups of mice as measured by an echocardiogram. Values are means ± SE obtained from 5–7 different animals. *Significantly different (P < 0.01) from PARP–/– animals.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
PARP cleavage is considered a hallmark of apoptosis being mediated by caspases (6). In this study, however, we showed that during the progression of hypertrophy to heart failure, no PARP cleavage could be detected. Instead, its expression and enzymatic activity was gradually increased with the degree of cardiac hypertrophy. These changes were the same between mouse and human failing hearts. Overactivation of PARP suppressed cardiac gene expression and contributed to myocyte cell death. Interestingly, we found that increased expression of PARP is not only limited to pathological hypertrophy but also occurred to a mild extent in physiological hypertrophy. These data, for the first time, demonstrate a role of PARP in the evolution and progression of cardiac hypertrophy and suggests that some events (if not all) related to cardiac growth, muscle gene regulation, and cell death might be mediated by PARP-dependent signaling pathways.

Cardiac muscle gene dysregulation and cell death associated with dilated cardiomyopathy and heart failure have long been documented (1, 12, 15, 19, 26). However, the mechanism of cell death remains highly disputed (12, 19, 22). Our observation that PARP is not cleaved but rather its expression is progressively increased in relation to the degree of cardiac hypertrophy suggests that homodynamic stress endangers cardiac myocytes through a mechanism that appears different from conventional caspase-mediated apoptosis. This is in agreement with other reports showing failing heart myocytes to be negative for active caspase-3 but positive for DNA damage markers, including PARP (2, 10, 26, 27). Recently, Guerra and colleagues (15) have demonstrated that the number of necrotic cells is far greater (7-fold) than the number of apoptotic cells in patients with cardiac failure. Others have presented even stronger evidence against caspase-mediated cell death during heart failure. Knanpen et al. (26) have shown that cardiomyocytes in embryonic hearts undergo caspase-dependent cell death, whereas in the adult heart, during failure, they go through caspase-independent autophagic cell death. Similarly, DeBour et al. (10) found no sign of PARP cleavage despite evident cell death in myocardial specimens of patients with severe congestive heart failure. On the basis of these reports and data presented here, we believe that elevated levels of PARP contribute to a caspase-independent mechanism of myocyte cell death during heart failure.

Conflicting data exist concerning the influence of caspases in the development of cardiac hypertrophy and subsequent failure. Whereas some investigators have failed to detect activation of caspases or a beneficial effect of caspase inhibitors in endangered myocardium, others have found positive results (Refs. 7 and 12 and the references therein). During early stages of cardiac hypertrophy (or physiological hypertrophy), no activation of caspases has been reported; however, apoptosis has been seen accompanying the adaptive phase of pressure overload hypertrophy (8, 13). Some studies have shown that during sustained pressure overload, caspase activation peaks only during hypertrophy of the myocardium; thereafter it declines in failing hearts (23). It is therefore likely that caspase activation is an epiphenomenon occurring at a particular stage of hypertrophy, possibly during transition to heart failure. The time window between activation of caspases and their downstream effects, including PARP cleavage and finally DNA fragmentation, also may be very narrow. As a result, conflicting results could have been seen, depending on the severity of the disease and the time point when tissue was taken for the analysis. In this study, we did not detect activation of caspases in the mouse model of cardiac hypertrophy, whereas PARP was gradually increased with the intensity of hypertrophy, including during physiological hypertrophy. This suggests that PARP activation may have a much broader role in the development of cardiac hypertrophy, and its cell degeneration effects may precede the timeline of caspase activation in diseased hearts.

How does increased expression of PARP cause myocyte cell death? PARP is considered the "Cinderella" of the genome because of its pivotal role in cell survival and maintenance of genome integrity. Single-strand DNA break is the most potent signal for PARP activation (6, 21). However, recent reports have indicated that PARP also could be activated by cell stressors not necessarily associated with DNA damage. These include reactive oxygen/nitrogen species and increased intracellular levels of Ca2+, Mg2+, and polyamines (20, 29, 45). Once activated, PARP induces protein poly-ADP-ribosylation by utilizing cellular NAD+ content. Several reports have indicated that, whereas mild activation of PARP plays a physiological role in cell survival, overactivation of PARP threatens cell viability due to a drop in the intracellular NAD+/ATP pool, which leads to cell necrosis (6, 48). PARP can also induce cell death by apoptosis but without activation of caspases (46). PARP has been shown to induce translocation of AIF from the mitochondria to the nucleus, where it causes chromatin degradation and eventually cell death (46). Another mechanism for PARP-mediated cell death is considered to be dependent on the linkage between the PARP pathway and class III histone deacetylases (sirtuins; e.g., SIR2{alpha}), which are NAD+ dependent and expressed in the adult heart (37, 49). Several studies have shown that SIR2{alpha} has profound roles in chromatin remodeling, gene silencing, cell viability, and longevity (42). Overexpression of SIR2 has been shown to increase the life span of the organism by 50%, perhaps by protecting cells against oxidative stress (42). Because the enzymatic activity of SIR2 is absolutely dependent on NAD+, it is suggested that the activation of PARP downregulates SIR2 through depletion of cell NAD+ content, which then promotes cell death via activation of p53-mediated apoptosis (5, 44, 49). Although more studies are required to understand the precise mechanism of PARP-mediated myocyte cell death, our data presented here show that one way that PARP induction could threaten cardiac cell survival is by depriving cells of their basic transcriptional support.

Given the cell-damaging role of PARP in myocytes, it was surprising to note that PARP was also activated in hearts of swimming mice with physiological hypertrophy. Previous studies analyzing the effects of PARP inhibitors have demonstrated that PARP has a physiological role in cell growth and gene regulation (39, 48). In the brain, stress-mediated atrophy of neurons was shown to be associated with reduced levels of PARP, implicating a role of PARP in normal cell growth and the development of neurons (47). PARP has been shown to regulate chromatin condensation and gene expression by poly-ADP-ribosylation of nuclear target proteins (35, 43). Additionally, PARP was shown to activate gene transcription when it is catalytically inactive (18, 32). Thus it appears that PARP controls gene expression in at least two ways: 1) by its enzymatic activity to poly-ADP-ribosylate target proteins, which adds a massive negative charge on the target protein, leading to change of its function; and 2) by its nonenzymatic activities, which includes its ability to bind to other transcription factors and hence to modify the activity of the transcription complex. Recently, numerous physical and functional interactions of PARP with other transcription factors have been described. These include factors that are also known to participate in the development of hypertrophy, such as NF-{kappa}B, activator protein 2, p53, TEF1, MyoD, and YY1 (4, 18, 24, 28, 33). One of the major determinants of physiological hypertrophy is the induction of {alpha}-MHC expression. Previously, we and others (16, 40) have shown that TEF1 and YY1 regulate {alpha}-MHC expression in an opposite manner; TEF1 activates, whereas YY1 represses, {alpha}-MHC promoter activity. Interestingly, the transcription activity of TEF1 and YY1 also has been shown to be regulated in an opposite manner by their association with PARP. Whereas PARP activates TEF1 activity, it inhibits the activity of YY1 (4, 28, 33). Thus it is conceivable that the mild activation of PARP through changing the activity of TEF1 and YY1 and/or through other unidentified mechanisms [such as chromatin relaxation (35)] regulates the expression of {alpha}-MHC and other cell growth-promoting genes, which leads to the development of physiological hypertrophy. However, when PARP is overactivated by sustained hemodynamic stress, it initiates another cascade of events through its enzymatic activity, which causes poly-ADP-ribosylation of transcription factors. This kind of posttranslational modification of proteins interferes with their ability to bind to each other and to DNA, resulting in repression of gene transcription and finally to cell death. A general outcome of this process in the heart would be expected to result in gene dysregulation, myocyte cell death, and fibrosis, a series of events that are known to be associated with pathological cardiac hypertrophy and with failing hearts. A role of inflammatory cytokines and NF-{kappa}B in the induction of pathological hypertrophy is fairly well established (36). Recent reports indicate that PARP, together with other cofactors, regulates the transcriptional activity of cytokines and NF-{kappa}B; thus PARP could also initiate the process of pathological hypertrophy via changing the activity of these intermediary factors (18). From the foregoing discussion, it appears that a gradual change in PARP activity may determine the transition between physiological and pathological cardiac hypertrophy. This would be identical to the dual role of other factors, e.g., calcineurin, which has been shown to participate in both physiological and pathological forms of cardiac hypertrophy (14).

In summary, this study shows for the first time that PARP-mediated signaling may be directly involved in the evolution and progression of cardiac hypertrophy. It supports many previous studies where myocardial specimens of failing hearts were found to be negative for active caspases but had clear signs of cell death. Recent studies have strongly indicated the involvement of oxygen/nitrogen-derived free radicals in the development of cardiac hypertrophy (38). Because PARP activity is sensitive to free radical generation and PARP inhibitors are in use for different oxidative stress-mediated diseases, it is tempting to believe that PARP inhibitors may also hold potential for treating anomalies of cardiomyopathies and heart failure.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported by National Heart, Lung, and Blood Institute Grants R01 HL-68083 and R01 HL-77788, American Heart Association Grant-In-Aid N-150108, and the Department of Surgery Research Funds (to M. P. Gupta).


   ACKNOWLEDGMENTS
 
We thank Ayman Isbatan and Ursula William for expert technical assistance. All imaging was performed with the help of Shirley Bond, Digital Light Microscopy Facility, University of Chicago, Chicago, IL. PARP expression plasmid was provided by Dr. V. L. Dawson, Dept. of Neurology, Johns Hopkins University School of Medicine, Baltimore, MD.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. P. Gupta, Dept. of Surgery, Committee on Molecular Medicine, MC 5040, Univ. of Chicago, 5841 S. Maryland Ave., Chicago, IL 60637 (E-mail: mgupta{at}surgery.bsd.uchicago.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.


   REFERENCES
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