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GASTROINTESTINAL, HEPATIC, PULMONARY, AND RENAL
Department of Cellular Biology and Anatomy, Center for Genomic Medicine, Medical College of Georgia, Augusta, Georgia (G.D., L.W., C.-Y.W., Z.D.); Department of Internal Medicine, University of Utah, Salt Lake City, Utah (T.Y.); and Charlie Norwood Veterans Affairs Medical Center, Augusta, Georgia (M.V.K., Z.D.)
Received February 1, 2008; accepted February 27, 2008.
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Abstract |
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40% apoptosis in 18 h. The apoptosis is accompanied by notable caspase activation; however, the general caspase inhibitor VAD can only partially suppress SAHA-induced apoptosis, suggesting the involvement of both caspase-dependent and -independent mechanisms. SAHA treatment leads to cytochrome c release from mitochondria, which is suppressed by Bcl-2 but not by VAD. Bcl-2 consistently blocks SAHA-induced apoptosis. During SAHA treatment, Bcl-2 and Bcl-XL decrease, and Bid is proteolytically cleaved, whereas Bax and Bak expression remains constant. Bid cleavage, but not Bcl-2/Bcl-XL decrease, is completely suppressed by VAD. SAHA does not activate p53, and pifithrin-
(a pharmacological p53 inhibitor) does not attenuate SAHA-induced apoptosis, negating a role of p53 in SAHA-induced apoptosis. SAHA induces histone acetylation, which is not affected by VAD, Bcl-2, or pifithrin-. Trichostatin A can also induce apoptosis and histone acetylation in renal tubular cells. Together, the results have shown evidence for renal toxicity of histone deacetylase inhibitors. The toxicity may be related to protein acetylation and decrease of antiapoptotic proteins including Bcl-2 and Bcl-XL.
; Glozak and Seto, 2007; Hodawadekar and Marmorstein, 2007). Histone deacetylase inhibitors, by original definition, are chemicals that inhibit the enzymatic activity of HDACs (Marks and Breslow, 2007; Xu et al., 2007). HDAC inhibitors can be structurally different, selectively inhibitory to specific HDACs, or be more general inhibitors of a class or several HDACs. For example, the classic HDAC inhibitor SAHA, a hydroxamic acid compound, inhibits classes I, II, and IV, but not class III, HDACs (Marks and Breslow, 2007; Xu et al., 2007).
The interest in HDAC inhibitors has been intensified by the discovery of their anticancer potential. To date, various HDAC inhibitors have been shown to suppress cancer growth and induce apoptosis in vitro in cancer cell cultures and in vivo in tumor bearing animal models (Bhalla, 2005; Bolden et al., 2006; Marks and Breslow, 2007; Xu et al., 2007). It is noteworthy that several HDAC inhibitors are being actively tested in clinical trials with promising outcomes toward specific types of tumors or cancers (Marks and Breslow, 2007; Xu et al., 2007). HDAC inhibitors can induce cell cycle arrest, trigger cellular senescence, and notably, activate cell death, although the underlying mechanisms are not entirely clear (Bhalla, 2005; Bolden et al., 2006; Marks and Breslow, 2007; Xu et al., 2007).
It is remarkable that in terms of cell injury and death, normal nonmalignant cells and tissues are relatively resistant to HDAC inhibitors (Bhalla, 2005; Bolden et al., 2006; Marks and Breslow, 2007; Xu et al., 2007). In human patients, treatment with SAHA leads to side-effect syndromes including fatigue, anorexia, and dehydration, which nevertheless disappear within days of drug withdrawal (Kelly et al., 2005). Despite these observations, whether HDAC inhibitors induce renal cell injury and death has not been carefully evaluated. Cells of the kidneys, especially those of the proximal tubules, are particularly sensitive to toxic xenobiotics, due to their cellular property and their physiological function of excretion and concentration of blood filtrates including metabolic wastes and toxins. Renal toxicity has been recognized for environmental toxins, aminoglycoside antibiotics, radiocontrast medium, and chemotherapeutic agents including Adriamycin and cisplatin (Swan, 1997; Parant et al., 2001; Arany and Safirstein, 2003; Van Vleet and Schnellmann, 2003). It is thus important to determine the potential renal toxicity of HDAC inhibitors, a family of promising anticancer drugs that is currently in clinical trials.
In the present study, we show that HDAC inhibitors including SAHA and Trichostatin A (TSA) can induce apoptosis in renal proximal tubular cells at micromolar concentrations. SAHA-induced apoptosis occurs via caspase-dependent and -independent mechanisms. The apoptosis involves decreases of antiapoptotic Bcl-2 proteins and consequent mitochondrial injury but does not seem to involve p53 activation.
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Materials and Methods |
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). The cells were cultured for experiments as described recently (Jiang et al., 2007; Pabla et al., 2008). Sources of antibodies used in this study were as follows: polyclonal antibodies specific for histone H3, acetylated histone H3 (Lys9), histone H4, acetylated H4 (Lys8), p53, and phospho-p53 (Ser15) from Cell Signaling Technology Inc. (Danvers, MA); monoclonal anti-cytochrome c from BD Pharmingen (San Diego, CA); polyclonal anti-Bcl-2 from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA); monoclonal anti-Bax and anti-Bcl-xL from NeoMarkers (Fremont, CA); polyclonal anti-Bak from Upstate Biotechnology (Lake Placid, NY); polyclonal anti-Bid from Dr. Xiao-Ming Yin (University of Pittsburgh, Pittsburgh, PA); monoclonal anti-β-actin from Sigma-Aldrich (St. Louis, MO); and all secondary antibodies from Jackson ImmunoResearch Laboratories Inc. (West Grove, PA). SAHA and TSA were obtained from BIOMOL Research Laboratories (Plymouth Meeting, PA). The caspase inhibitor (VAD) and fluorogenic substrate DEVD.AFC were purchased from Enzyme Systems Products (Dublin, CA). Other reagents including cisplatin and pifithrin- were purchased from Sigma-Aldrich.
Treatment of RPTCs. RPTCs were plated to reach 90% confluence the next day. For experiment, the cells were incubated with indicated concentrations of SAHA, TSA, or cisplatin in full culture medium for various durations. To test the effects of specific inhibitors, the inhibitors were given during the incubation period. At the end of incubation, cells were evaluated for apoptotic morphology or lysed to collect cell lysates for analysis of different proteins.
Analysis of Apoptosis. Apoptosis was analyzed by morphological methods and also indicated by the measurement of caspase activity (Dong and Wang, 2004; Jiang et al., 2004, 2007; Pabla et al., 2008). For morphological assessment, cells were fixed with 4% paraformaldehyde and stained with Hoechst 33342. Cellular and nuclear morphologies were examined by phase contrast and fluorescence microscopy, respectively. Typical apoptotic morphology examined included cellular shrinkage, formation of apoptotic bodies, and nuclear condensation and fragmentation. In each dish, four fields with 200 cells per field were examined to estimate the percentage of apoptosis. Images of representative fields were also recorded by microscopy. For caspase measurement, a standard enzymatic assay was conducted. In brief, cell lysate collected with 1% Triton X-100 was added to an enzymatic reaction containing 50 µM DEVD.AFC, a fluorogenic substrate for caspases. Fluorescence generated during the reaction was measured to calculate the caspase activity.
Analysis of Cytochrome c Release from Mitochondria. The release of cytochrome c from mitochondria into cytosol was monitored by immunofluorescence localization and by cellular fractionation (Dong et al., 2003; Dong and Wang, 2004). For immunofluorescence, cells were grown on collagen-coated glass coverslips. After experimental incubation, the cells were fixed in a modified Zamboni's fixative and permeabilized with 0.1% SDS. The cells were then incubated sequentially to blocking buffer with 5% normal goat serum, primary anti-cytochrome c antibody, and Cy-3-labeled goat anti-mouse secondary antibody. Immunofluorescence was captured by fluorescence microscopy. For cellular fractionation, cells were exposed to 0.05% digitonin in an isotonic buffer (250 mM sucrose, 10 mM HEPES, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, and 1 mM EGTA, pH 7.1) for approximately 2 min at room temperature. The solubilized fraction was collected as cytosolic fraction and insoluble part collected as membrane-bound organellar fractions. This fractionation method for analysis of cytochrome c release has been successfully used and verified in previous studies. The cytosolic and organellar fractions were analyzed for cytochrome c by immunoblot analysis.
Immunoblot Analysis. Immunoblotting was conducted using the NuPAGE Gel Systems by standard procedures. Protein concentration was determined with the bicinchoninic acid reagent (Pierce Chemical, Rockford, IL). The same amounts (usually 25 µg) of protein were loaded in each lane for electrophoresis under reducing condition. The resolved proteins were electroblotted onto polyvinylidene difluoride membranes. The membranes were incubated in 1% bovine serum albumin and 2% fat-free milk for blocking and then exposed to the primary antibodies overnight at 4°C. After extensive washing and blocking, the blot membranes were incubated with the horseradish peroxidase-conjugated secondary antibody, and antigens on the blots were revealed using the enhanced chemiluminescence kit from Pierce Chemical. All blots shown in this study are representatives of at least three separate analyses.
Statistical Analysis. Quantitative data were expressed as mean ± S.D. (n 
4). Statistical analysis was conducted using the GraphPad Prism software (GraphPad Software Inc., San Diego, CA). Statistical differences in multiple groups were determined by multiple comparisons with Tukey's post-tests following analysis of variance. Statistical differences between two groups were determined by Student's t test. P < 0.05 was considered statistically significant.
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; Jiang et al., 2004, 2007; Wei et al., 2004). When RPTCs were incubated in full culture medium containing 5 µM SAHA, apoptosis was initially noticed at 12 h and increased thereafter. By the end of 18 h, 30 to 40% cells underwent apoptosis. These cells displayed a typical apoptotic morphology, showing cellular shrinkage and formation of apoptotic bodies (Fig. 1A). Hoechst staining also revealed condensed and fragmented nuclei in SAHA-treated cells (Fig. 1A). Propidium iodide staining, on the other hand, was minimal in these cells (data not shown), suggesting that SAHA mainly induced apoptosis and not necrosis under the experimental condition. As expected, SAHA induced RPTC apoptosis does-dependently. As shown in Fig. 1B, SAHA did not induce significant apoptosis at 1 to 2 µM; however, 5 and 10 µM SAHA induced 35 and 60% apoptosis, respectively.
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SAHA-Induced Cytochrome c Release from Mitochondria. Apoptosis due to chemical stress or toxicity frequently involves the intrinsic pathway, which is characterized by mitochondrial outer membrane permeabilization, releasing apoptotic factors such as cytochrome c (Wang, 2001; Green and Kroemer, 2004). To test this during SAHA-induced RPTC apoptosis, we initially examined cytochrome c release from mitochondria into cytosol by immunofluorescence. As shown in Fig. 3A, in control cells, cytochrome c showed a punctate mitochondrial staining, which was perinuclear. In SAHA-treated dishes, there were a group of cells that had a diffuse cytosolic staining (marked by asterisks), indicating the release of mitochondrial cytochrome c into the cytosol. To confirm the results, we fractionated the cells into cytosolic and mitochondrial fractions for immunoblot analysis of cytochrome c. As shown in Fig. 3B, without SAHA treatment (lane 1), cytochrome c was detected only in the mitochondrial fraction. After 5 µM SAHA treatment, a significant amount of cytochrome c appeared in the cytosolic fraction, which was accompanied by a reduction of mitochondrial cytochrome c (lane 2). Cytochrome c release during SAHA treatment was not affected by VAD (lane 3) but was attenuated in cells stably transfected with Bcl-2 (lane 4). Consistently, SAHA-induced apoptosis was suppressed from 36 to 12% in Bcl-2-transfected cells (Fig. 3C). Overexpression of Bcl-2 in the transfected cells was verified by immunoblot analysis, and, interestingly, SAHA induced a decrease of Bcl-2 in both wild-type and Bcl-2-transfected cells (Fig. 3C). The results suggest a role for the intrinsic apoptotic pathway in SAHA-induced renal cell toxicity. In this pathway, Bcl-2 family proteins may control mitochondrial integrity and therefore apoptosis.
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Bcl-2 Family Proteins during SAHA Treatment of RPTCs. We went on to examine the expression level of several critical Bcl-2 family proteins. As shown in Fig. 4A, the level of Bcl-2 decreased after 5 µM SAHA treatment (lane 4), and the decrease was even more at 10 or 20 µM SAHA (lanes 5 and 6). VAD did not prevent the Bcl-2 decrease induced by 5 µM SAHA (lane 7 versus lane 4). Likewise, Bcl-XL also showed a decrease, which started from 1 µM SAHA and became obvious at 10 to 20 µM SAHA. Bcl-XL decrease induced by SAHA was not prevented by VAD either (lane 7 versus lane 4), suggesting that the Bcl-2/Bcl-XL changes were not mediated by caspases. This conclusion was supported by analysis of the time course of Bcl-XL decrease. As shown in Fig. 4B, 5 µM SAHA induced an obvious Bcl-XL decrease within 6 h (Fig. 4B, lanes 5 versus 6), a time point that did not show caspase activation or apoptosis (data described above). Thus, the Bcl-2/Bcl-XL decrease induced by SAHA was not due to proteolysis by caspases; rather, it might be upstream of mitochondrial injury and caspase activation. The expression levels of Bax and Bak, two multidomain proapoptotic proteins, remained relatively constant during SAHA treatment (Fig. 4A). Bid, a BH3-only proapoptotic protein, was proteolytically cleaved in SAHA-treated cells, releasing truncated Bid or tBid. Bid cleavage started at 5 µM SAHA and became more at 10 to 20 µM (Fig. 4A). Bid cleavage was completely blocked by VAD (Fig. 4A), indicating that caspases were responsible for Bid cleavage under the experimental condition.
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No Critical role for p53 in SAHA-Induced RPTC Apoptosis. p53 has been implicated in SAHA-induced apoptosis in cancer cells (Henderson et al., 2003; Shen et al., 2007). In renal cells, p53 has a role in acute kidney injury induced by ischemia as well as cisplatin (Cummings and Schnellmann, 2002; Kelly et al., 2003; Jiang et al., 2004). To determine whether p53 is involved in SAHA-induced RPTC apoptosis, we first examined p53 phosphorylation and protein stabilization or accumulation. As shown in Fig. 5, SAHA did not induce p53 phosphorylation at the common Ser15 site. Consistently, p53 did not show protein accumulation during 1 to 20 µM SAHA treatment (lane 1 versus lanes 2–6). In contrast, cisplatin induced both p53 phosphorylation and accumulation in the same experiments (lane 9). We further showed that pifithrin-, a commonly used pharmacological inhibitor of p53, did not have significant inhibitory effects on SAHA-induced apoptosis in RPTCs (Fig. 5B). As a positive control, pifithrin- partially inhibited cisplatin-induced apoptosis (data not shown; Cummings and Schnellmann, 2002; Jiang et al., 2004). Together, the results suggest that p53 may not play a critical role in SAHA-induced apoptosis in renal tubular cells.
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SAHA-Induced Histone Acetylation in RPTCs. As a histone deacetylase inhibitor, SAHA was expected to increase histone acetylation. To determine the correlation between histone acetylation and apoptosis during SAHA treatment, we first analyzed the dose dependence of SAHA-induced acetylation of histone H3 and H4. As shown in Fig. 6A, SAHA at 1 and 2 µM induced histone acetylation (lanes 2 and 3), which was increased significantly by 5 µMor higher concentrations of SAHA (lanes 4–6), whereas the level of total histone did not change significantly during SAHA treatment (Fig. 6A). We further examined the time course of histone acetylation during 5 µM SAHA incubation. It was shown that histone acetylation was induced within hours of SAHA treatment (Fig. 6B), whereas apoptosis was not obvious until 12 h of SAHA incubation. The dose dependence and time course experiments suggest that excessive acetylation of histones and related proteins could trigger cell injury and apoptosis during SAHA treatment of RPTCs.
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Effects of Bcl-2, VAD, and Pifithrin- on RPTC Apoptosis Induced by SAHA. We further compared SAHA-induced histone acetylation in wild-type and Bcl-2-transfected RPTCs (Fig. 7A). It is clear that Bcl-2 did not attenuate SAHA-induced histone acetylation (lane 2 versus 4), although it blocked cytochrome c release and apoptosis in this experimental model (Fig. 3). In addition, SAHA-induced histone acetylation was not affected by VAD and pifithrin-, respective inhibitors of caspases and p53 (Fig. 7B). The results further suggest that acetylation of histones and related proteins may be upstream of mitochondrial injury and caspase activation during SAHA-induced apoptosis.
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Trichostatin A-Induced Apoptosis and Histone Acetylation in RPTCs. Our results have suggested a role for excessive protein acetylation in SAHA-induced apoptosis in RPTCs (Figs. 6 and 7). To substantiate these observations, we examined the effects of TSA, another commonly tested histone deacetylase inhibitor. We showed that TSA could induce apoptosis in these cells in a dose-dependent manner. At 5 µM, TSA induced 25% apoptosis within 18 h. At 10 µM, TSA induced 60% apoptosis (Fig. 8A). We further analyzed histone acetylation during TSA incubation. As shown in Fig. 8B, acetylation of both histone H3 and H4 was induced by TSA in a dose-dependent manner, whereas total histones remained relatively constant.
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Discussion |
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Epigenetic regulation in renal cells and tissues is largely unknown. The latest work by Arany et al. (2007) demonstrated an interesting observation that TSA can protect mouse proximal tubular cells against cisplatin injury, and the protection is mechanistically related to the maintenance of CREB activity. The observed cytoprotective effects of TSA seem to contradict with our current results of renal cell toxicity of SAHA and TSA. However, these two studies have used different proximal tubular cell lines. In addition, a much lower concentration of 50 nM TSA was used by Arany et al. In our study, significant apoptosis is induced by 5 µM or higher concentrations of SAHA and TSA, but not by 1 to 2 µM (Fig. 1). Thus, whether HDAC inhibitors are protective or injurious depends on the experimental conditions including cell types used and drug concentrations tested. An important consideration is the pharmacokinetics of the HDAC inhibitors after being delivered into patients. Due to the renal function of excretion and reabsorption, drugs are frequently concentrated in kidney tissues, leading to the exposure of renal tubular cells to relatively high concentrations of drugs. These possibilities again call for the need of vigorous renal monitoring when the HDAC inhibitors are used clinically.
In a mechanistic manner, we show that SAHA-induced tubular cell apoptosis occurs via both caspase-dependent and -independent mechanisms. By estimation, about half of the induced apoptosis can be inhibited by VAD and thus is mediated by caspases, whereas the other half is executed in caspase-independent manner. Likewise, caspase-dependent and -independent apoptosis has been demonstrated in renal proximal tubular cells treated with cisplatin, a widely used chemotherapy drug (Cummings and Schnellmann, 2002; Jiang et al., 2004). In cancer cells, HDAC inhibitors including SAHA can induce typical caspase-dependent apoptosis, atypical apoptosis that is independent of caspases, mitotic cell death, and autophagic cell death (Xu et al., 2007). Although the different types of cell death are induced by HDAC inhibitors in cancers cells, much less is known about the distinctive underlying mechanisms (Xu et al., 2007). In the present study, we show that Bcl-2 is particularly effective in antagonizing SAHA-induced apoptosis in renal tubular cells. In fact, Bcl-2 seems to be more effective than VAD, suggesting that Bcl-2 is cytoprotective against both caspase-dependent and -independent apoptosis.
The potent antiapoptotic effects of Bcl-2 shown in our study also suggest a role for mitochondria injury in SAHA-induced tubular cell apoptosis, because, within these cells, Bcl-2 has a main localization in mitochondria (Bhatt et al., 2007). Indeed, we show that cytochrome c is released from mitochondria into cytosol during SAHA treatment of tubular cells (Fig. 3). It is noteworthy that cytochrome c release is attenuated by Bcl-2 but not VAD. These observations are consistent with the scenario that mitochondrial injury induced by SAHA leads to the release of cytochrome c into cytosol, which in turn activates caspases, resulting in apoptosis.
It is unclear as to how mitochondrial injury is induced by HDAC inhibitors, including SAHA. In acute T cell leukemia CEM-CCRF cells, SAHA induces Bid cleavage before mitochondrial damage, suggesting that the resultant truncated Bid may become active and target mitochondria for injury (Ruefli et al., 2001). We also show Bid cleavage during SAHA treatment of RPTCs (Fig. 4). Note that, in our study, the observed Bid cleavage is completely abolished by the general caspase inhibitor VAD, which nevertheless does not attenuate SAHA-induced cytochrome c release from mitochondria. These results therefore do not support an important role for Bid cleavage or activation in mitochondrial injury in our experimental model; otherwise, by preventing Bid activation, VAD should suppress cytochrome c release. The expression of proapoptotic Bax and Bak does not change significantly during SAHA-induced apoptosis. However, the levels of the antiapoptotic Bcl-2 and Bcl-XL decrease (Fig. 4). For Bcl-2, the decrease is obvious at 5 µM SAHA and becomes even more at 10 or 20 µM SAHA. VAD cannot prevent Bcl-2 decrease induced by 5 µM SAHA. Bcl-XL decrease starts from 1 µM SAHA and becomes obvious at 10 to 20 µM SAHA. Again, VAD cannot prevent SAHA-induced Bcl-XL decrease. Based on these results, it is concluded that Bcl-2 and Bcl-XL decrease during SAHA treatment is not mediated by caspases. This inference is further supported by the results of time course experiments. For example, Bcl-XL decrease occurs within hours of SAHA (5 µM) treatment, clearly preceding caspase activation and apoptosis (Fig. 4B). The results suggest a role for Bcl-2/Bcl-XL decrease in SAHA-induced tubular cell apoptosis. It is conceivable that Bcl-2/Bcl-XL decrease can sensitize the cells to apoptotic injury or liberate the proapoptotic molecules (e.g., Bax and Bak) for activation. The mechanism(s) of SAHA-induced Bcl-2/Bcl-XL decrease is currently unknown. It can be a result of transcriptional or translational inhibition, protein degradation, or both.
Whether p53 is involved in cell death induced by HDAC inhibitors and their anticancer effects has been quite controversial (Vrana et al., 1999; Henderson et al., 2003; Shen et al., 2007). For example, Vrana et al. (1999) showed that SAHA-induced apoptosis in U937 human leukemia cells is independent of p53, whereas Henderson et al. (2003) suggested that BJ cell apoptosis induced by SAHA or TSA is regulated by p53. In renal tubular cells, we show that p53 is not activated by SAHA at concentrations that induce massive apoptosis (Fig. 5). In addition, pifithrin-, a pharmacological inhibitor of p53, does not diminish SAHA-induced apoptosis, although it suppresses apoptosis induced by cisplatin. These data suggest that p53 may not play a critical role in SAHA-induced apoptosis in renal cells.
As expected, both SAHA and TSA induce histone acetylation in renal tubular cells (Figs. 6, 7, 8). It is noteworthy that the acetylation is not blocked by either VAD or Bcl-2, suggesting that protein acetylation is not a consequence, rather a trigger of cell injury under the experimental conditions. Protein acetylation may initiate a signaling cascade, resulting in mitochondrial injury, caspase activation, and consequent apoptosis. However, it is noteworthy that HDACs have substrates other than histones and thus are more properly called "lysine deacetylase" (Glozak and Seto, 2007; Xu et al., 2007). Whether apoptosis is triggered by acetylation of histone or other proteins remains to be investigated.
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Footnotes |
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: HDAC, histone deacetylase; SAHA, suberoylanilide hydroxamic acid; TSA, Trichostatin A; RPTC, rat renal proximal tubular cell.
Address correspondence to: Dr. Zheng Dong, Department of Cellular Biology and Anatomy, Medical College of Georgia, 1459 Laney Walker Blvd., Augusta, GA 30912. E-mail: zdong{at}mail.mcg.edu
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