AJP - Heart Calcium Transients and Cell-Sarcomere
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 274: H965-H973, 1998;
0363-6135/98 $5.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Borger, D. R.
Right arrow Articles by Essig, D. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Borger, D. R.
Right arrow Articles by Essig, D. A.
Vol. 274, Issue 3, H965-H973, March 1998

Induction of HSP 32 gene in hypoxic cardiomyocytes is attenuated by treatment with N-acetyl-L-cysteine

D. R. Borger and D. A. Essig

Department of Exercise Science, Laboratory of Exercise Molecular Biology, University of South Carolina, Columbia, South Carolina 29208

    ABSTRACT
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Increased synthesis of stress proteins may enhance myocardial viability during periods of low oxygen delivery. Our purpose was to determine if the oxidative stress protein heme oxygenase-1 [heat stress protein 32 (HSP 32)] was induced in hypoxic cardiomyocytes and whether this induction might be mediated by a redox-sensitive mechanism. Primary rat neonatal cardiomyocytes, cultured to express a tissuelike phenotype, responded to 12 h of hypoxia (<0.5% ambient oxygen) with an approximately fivefold (range 3- to 7.5-fold; P < 0.05) increase in HSP 32 mRNA and a threefold (P < 0.05) increase in HSP 32 protein content. Exposure to 80 µM H2O2 for 3 h increased HSP 32 mRNA content to a similar extent. Expression of heme oxygenase-2 mRNA was unaffected by H2O2 or hypoxic treatments. Inclusion of 20 mM N-acetyl-L-cysteine in the medium during hypoxia reduced the increase in HSP 32 mRNA and protein expression by 25-50% compared with hypoxia alone. The data suggest that induction of HSP 32 protein may lead to an improved antioxidant defense in cardiomyocytes during hypoxia and that a redox-sensitive pathway mediates at least a portion of the hypoxic induction of the HSP 32 gene.

hypoxia; ischemia; oxidative stress; heme oxygenase; heat stress protein 32; HSP 32

    INTRODUCTION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

MYOCARDIAL HYPOXIA RESULTS from reduced blood flow due to either an abrupt arterial thrombosis or gradual atherosclerotic narrowing. There are a variety of metabolic responses associated with the onset and progression of hypoxia, including a decrease in cellular ATP content, loss of contractile activity, acidosis, loss of membrane integrity, and increased production of reactive oxygen species (ROS) (13, 22). Heat stress proteins (HSPs) are a family of proteins whose expression constitutes a ubiquitous intracellular protective response to stress. Expression of the inducible isoform of the gene HSP 70 (HSP 70i), for example, has been shown to increase in myocardial cells after 30-min myocardial ischemia before reperfusion (23) or in cultured cardiomyocytes after several hours of hypoxia (13). Elevated levels of HSP 70i caused by overexpression of the HSP 70i gene in transgenic mice have been shown to improve the recovery of contractile force after ischemia-reperfusion (17, 24). However, the role that other stress proteins may have in the response to myocardial hypoxia is less characterized.

Recently, the expression of the stress protein gene, HSP 32, has been shown to be increased in cultures of isolated neonatal rat cardiomyocytes after prolonged (13 h) severe hypoxia (8). This is similar to what has been shown in several other cell types, including vascular smooth muscle (16, 19) and Chinese hamster ovary cells (20). In addition, exposure of rats to hypoxia (7-10% O2) markedly increased levels of the HSP 32 transcript in several tissues, including heart (14, 16). HSP 32 is the inducible isoform of heme oxygenase (HO-1) and, along with the constitutive isoform HO-2, catalyzes the first and rate-limiting step in heme degradation. HSP 32 functions as a stress protein by at least two mechanisms. Increased levels of HSP 32 have been shown to result in an improved antioxidant defense (1), presumably by accelerating conversion of the pro-oxidant heme substrate into the potent anti-oxidant bilirubin (28). The heme oxygenase reaction also results in the production of CO, which has been shown to stimulate guanyl cyclase and production of guanosine 3',5'-cyclic monophosphate. CO from vascular smooth muscle cells has been hypothesized to serve a physiological role in regulating vascular tone during hypoxia (19). Hence, induction of HSP 32 during hypoxia in cardiomyocytes could lead to an adaptive response characterized by an improved antioxidant defense and the release of a potential vasodilatory factor (CO).

The signals and pathways that mediate the hypoxic induction of HSP 32 in cardiomyocytes has not been well characterized. In other cell types it is clear that hypoxia leads to an increase in HSP 32 protein by increasing the rate of transcription of the HSP 32 gene (16, 19). One major signaling pathway that directly senses the decrease in cellular PO2 is linked to the synthesis and binding of the hypoxia-inducible factor 1 (HIF-1) transcription factor. HIF-1 has been shown to mediate the transcriptional induction of multiple genes during hypoxia, including HSP 32 (5, 16). A second or additional pathway has also been proposed to be involved in the induction of the HSP 32 gene during hypoxia. Murphy et al. (20) have suggested on the basis of experiments in Chinese hamster ovary cells that decreases in levels of reduced glutathione during hypoxia (15) and the associated oxidant stress may be linked to induction of the HSP 32 gene. However, to date the potential contribution of this pathway has not been evaluated experimentally in any cell type. Such a pathway may also be important in cardiomyocytes as there is evidence that during hypoxia glutathione content decreases and leads to oxidative stress in ischemic myocardium before reperfusion (22).

Given the potentially important cardioprotective effects of HSP 32 induction during hypoxia, it is imperative to further examine several features associated with induction of HSP 32 observed in myocardial cells. First, induction of HSP 32 during hypoxia, as measured in primary cultures of neonatal cardiomyocytes, has only been assessed at the mRNA level. To better understand the potential physiological significance of HSP 32 induction, the changes in protein expression need to be measured. Second, studies to date have not yet assessed HO-2 expression during hypoxia. Though HO-2 is typically not stress inducible, the extent and timing of HSP 32 induction may be related to the levels of this constitutively expressed isoform of heme oxygenase (3). Finally, given the antioxidant role of HSP 32, it is of interest to determine if oxidant levels play an important role in the signaling pathways that mediate the potentially beneficial increases in HSP 32 expression in hypoxic cardiac cells.

In the present study, we performed a series of experiments to further characterize the expression of heme oxygenase isoforms HSP 32 and HO-2 in primary rat cardiomyocytes after hypoxia. To delineate the specific effects of hypoxia, we have used a culture model similar to that of Webster et al. (31, 32), in which oxygen content was <0.5% but other potential stressors associated with ischemia such as low glucose and serum were minimized. Initially, we established the expression pattern of both heme oxygenase mRNA isoforms subjected to 12 h of hypoxia or H2O2. We then tested the hypothesis that the hypoxic induction of HSP 32 was potentially mediated by a redox-sensitive pathway. Cardiomyocytes were pretreated with the antioxidant and reduced glutathione precursor N-acetyl-L-cysteine (NAC), shown previously (2) to blunt the H2O2-induced transcription of the HSP 32 gene. Our results indicated that hypoxia and H2O2 both induced the expression of HSP 32 mRNA, whereas the expression of HO-2 mRNA was not changed from baseline. Moreover, the hypoxic induction of HSP 32 mRNA and protein was significantly diminished after pretreatment with NAC, suggesting that a component of the signaling pathway in cardiomyocytes may involve a redox mechanism.

    MATERIALS AND METHODS
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Cardiomyocyte isolation. Cardiomyocytes were isolated from 3- to 4-day-old Sprague-Dawley rats using modifications to the methods of Borg et al. (4). After rats were decapitated, hearts from 25 to 30 neonatal pups were rapidly excised and pooled in ice-cold Moscona's saline [containing (in mM) 28.6 KCl, 136.8 NaCl, 9.4 glucose, 11.9 NaHCO3, and 0.08 NaH2PO4, pH 7.4]. Excess connective tissue was removed, and the hearts were washed and then minced into 1-mm3 pieces in ice-cold Krebs-Ringer buffer [KRB; containing (in mM) 4.7 KCl, 118.4 NaCl, 23.8 NaHCO3, 2.4 MgSO4, and 1.5 KH2PO4, gassed with 5% CO2] supplemented with 2 mg/ml glucose, 1 mg/ml bovine serum albumin (BSA) fraction V, 100 U/ml penicillin, and 100 µg/ml streptomycin (pH 7.4). The minced tissue was transferred to a 100-ml flask containing 10 ml of fresh KRB II-collagenase [KRB gassed with 5% CO2 and supplemented with 2 mg/ml glucose, 20 mg/ml BSA fraction V, 100 U/ml penicillin, 100 µg/ml streptomycin, 1% deoxyribonuclease (DNase), and 100 U/ml collagenase (pH 7.4)] and digested at 37°C for 10 min in a shaking water bath. The tissue was allowed to settle and the supernatant was removed and replaced with fresh KRB II-collagenase. This digestion procedure was repeated seven times. The supernatant obtained from the first enzymatic digestion was discarded. During each subsequent enzymatic digestion, before removal of the supernatant, loosened cells were mechanically dissociated from the tissue through repeated pipetting of the digestion solution. The dissociated cells in supernatants 2-7 were washed of collagenase by centrifugation at 800 g for 8 min in a 2× vol of KRB II without collagenase and DNase. The pelleted cells were resuspended in Dulbecco's modified Eagle's medium (GIBCO BRL, Life Technologies, Gaithersburg, MD) supplemented with 10% newborn bovine serum (GIBCO), 5% horse serum (Flow Laboratories, McLean, VA), and 4 µg/ml cytosine arabinoside (hereafter referred to as standard medium) and were pooled together. The suspended cells were filtered through a 20-µm nylon mesh, collected by centrifugation at 800 g for 8 min, and resuspended in standard medium. Cardiomyocytes were enriched through a 30-min interval of differential adhesion. A small aliquot was stained with trypan blue, and cell number was determined using a hemocytometer. This process routinely yielded ~2 × 106 cells/heart. On the basis of immunocytochemical analysis using cell-specific antibodies, cardiomyocytes represented 93-96% of the cell population.

Cell plating. Cardiomyocytes were plated on sterile 100-mm culture dishes coated with an aligned collagen matrix after the methods of Simpson et al. (27) to ensure a more "in vivo tissue"-like morphology. In brief, a 3.5-ml ice-cold layer of 3.0 mg/ml collagen type I (Collagen, Palo Alto, CA) was applied over a 1-ml ice-cold mix (1:1) of 10× minimal essential medium (MEM; GIBCO) and 0.2 M N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), pH 9, in a 50-ml centrifuge tube. The collagen solution was gently mixed and diluted to 10 ml with ice-cold Moscona's saline. A 3-ml volume of this solution was pipetted onto the lowered peripheral edge of a culture dish tilted at a 45° angle. After the lowered edge was rotated upward in the opposite direction, the solution was drawn across the bottom with a continuous and downward stroke with a cell scraper until the entire dish bottom was covered. The dish was propped at a 45° angle, and the excess collagen was carefully aspirated off the lowered edge. Collagen-coated dishes were allowed to polymerize on a flat surface at 37°C for at least 1 h before use. Cardiomyocyte cells were plated at a density of 6 × 106 cells/100-mm dish in standard medium and allowed to adhere for 24 h. Cultures were maintained at 37°C in a humidified, 5% CO2 supplemented air environment and were fed fresh standard medium (prewarmed to 37°C) 24 h after plating and thereafter at 2-day intervals. Cytosine arabinoside was included as a component of the standard medium to inhibit the proliferation of any remaining fibroblast cells.

Experimental designs. Confluent cardiomyocytes (7-8 days after isolation) were utilized in a series of three experiments. Experiment 1 was designed to evaluate cardiocyte membrane integrity (medium creatine kinase), metabolic response (medium glucose and lactic acid), and heme oxygenase mRNA isoform expression after 12 h of hypoxia (n = 3 plates) or 12 h of normoxia (n = 3 plates). Previous studies (19, 20) had shown that 12 h of hypoxia produced the greatest increase in HSP 32 expression. Additional plates were also included to confirm the ability of the cardiomyocyte HSP 32 gene to respond to oxidative stress as shown earlier by Hoshida et al. (10). Plates with fresh standard media were supplemented with H2O2 to achieve a final concentration of 80, 160, or 300 µM H2O2 (n = 1 plate per concentration) and were incubated for 3 h before harvest. In experiment 2, two control studies were performed. To rule out medium replacement as a form of culture stress that might influence HSP 32 expression (10), we determined HSP 32 and HO-2 mRNA content 0, 2, 6, or 12 h (n = 4 plates/time point) after replacement of medium under normoxic conditions. A similar study was performed to examine the effects of inclusion of NAC on basal expression of HSP 32 and HO-2 mRNA. Cells were harvested 0, 2, 6, or 12 h (n = 4 plates/time point) after replacement of standard medium with medium supplemented with 20 mM NAC (Sigma, St. Louis, MO). In experiment 3, plates of cells were divided into three groups (n = 4 plates/group). Cells were preincubated for 1 h with either fresh standard medium (Hyp group) or standard medium supplemented with 20 mM NAC (Hyp + NAC group) and harvested after 12 h of hypoxia. Cells incubated in a normoxic environment for 12 h served as control (NC group). After cells were harvested, the contents of HSP 32 mRNA and HSP 32 protein in the cardiomyocytes and medium concentrations of glucose and lactic acid were measured.

Hypoxia was produced in a humidified and airtight controlled atmosphere culture chamber (Bellco Glass, Vineland, NJ) placed in a standard 37°C incubator and connected to a vacuum and gas manifold. The chamber and manifold were flushed with preanalyzed hypoxic gas (5% CO2-95% N2) for 5 min before cells were sealed. An evacuation process, involving the replacement of vacuum chamber gases with the hypoxic gas mixture, was performed initially three times in succession and then once every 10 min over a 1-h duration after the cells were sealed in the chamber. This procedure produced an environment of <0.5% O2, as visually confirmed by the complete reduction (white) of a prewetted methylene blue dry anaerobic indicator strip (BBL Microbiology Systems, Becton Dickinson, Cockeysville, MD). Slight positive pressure of hypoxic gas within the chamber was maintained for the duration of the hypoxic period to prevent ambient air contamination. Normoxic cells were maintained at 37°C in a separate incubator supplemented with 5% CO2 air (~21% O2). All cells were immediately processed on completion of their respective treatments (within 2 min), and the samples were stored at -75°C for subsequent analysis. Cultures that constituted each series of experiments were obtained from 1 to 2 separate cell preparations.

RNA isolation. After the medium was removed, cardiomyocytes plated on 100-mm dishes were immediately lysed in 1 ml of Ultraspec (Biotecx Laboratories, Houston, TX). The lysate was collected with the aid of a cell scraper and transferred to a sterile 2-ml Eppendorf tube. After a 5-min incubation on ice, 0.2 ml of chloroform was added and the sample was vortexed vigorously for 30 s. The RNA containing supernatant obtained from a 20-min centrifugation at 12,000 g (4°C) was transferred to a fresh prechilled Eppendorf tube, isopropanol precipitated (1:1 vol/vol) overnight at -75°C, and collected by centrifugation at 14,000 g (4°C) for 15 min. Pellets were washed twice in 1 ml of 75% ethanol through vortexing and a 10-min centrifugation at 14,000 g (4°C). After a final wash in 0.5 ml of 100% ethanol, the ethanol was aspirated off, and air-dried pellets were resolubilized in 15 µl of RNase-free deionized H2O (dH2O) with the aid of a 5-min incubation at 60°C. A small aliquot (1 µl) was diluted 1:1,000 with dH2O, and the concentration was determined. Only total RNA samples with an absorbance ratio (A260/280) of 1.8-2.0 were further analyzed.

Northern blot analysis and probes. Equal amounts of total RNA (as designated in legends of Figs. 1 and 2) were denatured at 65°C for 5 min in 15 µl of sample buffer (13.4 mM NaH2PO4, pH 7.4, 6.7 mM EDTA, pH 8.0, 8.83% formaldehyde, 66.8% formamide, and 0.067 mg/ml ethidium bromide), electrophoresed in a 1% agarose-1.12 M formaldehyde denaturing gel, and capillary diffused overnight onto a Zeta-Probe membrane (Bio-Rad Life Science Research Products, Hercules, CA) using standard procedures (25). RNA was covalently bonded to the membranes with ultraviolet (UV) crosslinking (UV Stratalinker 1800; Stratagene Cloning Systems, La Jolla, CA).

Plasmid containing a 1,341-base pair (bp) rat HSP 32 cDNA insert (generously provided by Dr. Richard Levine, Department of Medicine, New York Medical School) was denatured at 95°C for 5 min and then labeled in the presence of [alpha -32P]dATP (3,000 Ci/mmol; NEN Research Products, DuPont, Boston, MA) by the random priming method (Prime-It II Random Primer Labeling Kit; Stratagene) and unincorporated nucleotides were removed (NucTrap Push Columns; Stratagene). The blots were prehybridized for 1 h and then hybridized with ~3 × 106 counts/min (cpm)/ml of denatured probe for 16 h at 42°C in a solution containing 200 µg/ml denatured salmon sperm DNA (Stratagene), 50% formamide, 5× Denhardt's buffer, 5× SSPE (sodium chloride-sodium phosphate-EDTA), and 0.1% sodium dodecyl sulfate (SDS). Membranes were washed four times at room temperature in 2× standard saline citrate (SSC), 0.1% SDS for 15 min and three times at 56°C in 0.1× SSC, 0.1% SDS for 30 min. Air-dried membranes were exposed to X-ray film (X-O-MAT AR; Eastman Kodak, Rochester, NY) at -75°C with intensifying screens. The developed autoradiograms were scanned and quantified with an IS-1000 digital imaging system densitometer (Alpha Innotech, San Leandro, CA). The linearity of the autoradiographic signal was confirmed using several exposure times for each blot. Comparable loading and integrity of the RNA was assessed by ethidium bromide staining of the 28S and 18S rRNA on the original gel.

Membranes were subsequently stripped by boiling in 0.1× SSC, 0.1% SDS for 10 min, then prehybridized and reprobed using identical procedures but with a 190-bp HO-2 cDNA fragment. This probe was generated by reverse transcriptase-rapid polymerase chain reaction (RT-RPCR) using a 1605 Air Thermo-Cycler (Idaho Technologies, Idaho Falls, ID) after the methods of Tan and Weis (29) with further modifications described by Essig et al. (6). Polyadenylated RNA was isolated from 0.5-1.0 µg of rat liver total RNA using the PolyATtract mRNA isolation system (Promega, Madison, WI) according to the instructions supplied. Complementary DNA was synthesized during a 1-h incubation at 37°C using 2 µg of liver polyadenylated RNA in 50 µl of reverse transcriptase (RT) reaction buffer [1× RT buffer (GIBCO BRL, Life Technologies, Gaithersburg, MD), 10 mM dithiothreitol (DTT), 0.5-ng random primers (New England Biolabs, Beverly, MA), 400 U M-MLV RT (GIBCO BRL), and 125 mM each of dATP, dGTP, dCTP, and dTTP (New England Biolabs)]. The RNA template was subsequently removed by a 3-min incubation on ice with 2 µg of DNase-free RNase A (Promega). The reaction solution was adjusted to 220 µl with 0.4 M NaCl and chloroform:phenol and then chloroform extracted (1:1 vol/vol). The DNA contained in the supernatant was precipitated in 2.5 vol of 100% ethanol at -75°C for 1 h and collected by centrifugation at 14,000 g (4°C) for 20 min. Pellets washed in 1 ml of 70% ethanol, then 0.5 ml 100% ethanol at 14,000 g (4°C) for 10 min, were allowed to air dry and then were solubilized in 10 µl of 10 mM tris(hydroxymethyl)aminomethane (Tris)-1 mM EDTA, pH 8.0. The concentration was determined as described in RNA isolation.

RPCR amplification of HO-2 cDNA was performed on a 10-µl reaction mix containing 100 ng rat liver cDNA, 1× high-Mg2+ buffer (Idaho Technologies), 1× deoxynucleotide triphosphates (Idaho Technologies), 7 µM forward primer (5'-GCTTACACTCGTTACATGGGGGG-3'), 7 µM backward primer (5'-TGGTCTTCA-TACTCAGGTCCAAGG-3'), and 0.83 U Taq DNA polymerase [5 U/µl Taq (Promega) diluted 1:6 in enzyme diluent (Idaho Technologies)] (6). This sample reaction mix (10 µl) was sealed in a glass microcapillary tube (Idaho Technologies) and subjected to 30 cycles of PCR amplification at 94°C, 0-s denaturation; 60°C, 0-s annealing; and 72°C, 5-s elongation in a 1605 Air Thermo-Cycler (Idaho Technologies). The amplified product was gel purified in a 2% low-melt agarose gel. The 190-bp fragment corresponding to the amplified HO-2 cDNA was excised from the gel and eluted in 5 vol of 20 mM Tris · HCl, 1 mM EDTA, pH 8.0, at 65°C for 5 min. The agarose was removed from the sample (after cooling to room temperature) by phenol, phenol:chloroform, and then chloroform extraction (1:1 vol/vol) at 4,000 g for 10 min at room temperature. The fragment contained in the resulting supernatant was precipitated in 2 vol of ice-cold 100% ethanol and 0.2 vol of 10 M ammonium acetate. The pellet was collected by a 20-min centrifugation at 14,000 g (4°C). Ethanol washing and spectrophotometry were performed as per above. The purified HO-2 DNA fragment was radiolabeled by the random priming method described above.

To control for variability in gel loading, blots used to determine HSP 32 and HO-2 content were rehybridized with a 40-bp rat glyceraldehyde-3-phosphate dehydrogenase (GAPDH) oligonucleotide probe (Oncogene Research Products, Calbiochem, La Jolla, CA). The probe was labeled and hybridized according to the manufacturer's procedures. Briefly, a 10-µl reaction mix containing 0.1 M Tris · HCl, pH 7.5, 10 mM MgCl2, 10 mM DTT, 10 U polynucleotide kinase, 40 µCi [gamma -32P]ATP, and 32.5 ng GAPDH oligonucleotide was incubated at 37°C for 30 min. This labeling reaction was subsequently terminated with the addition of 40 µl 0.1 M EDTA, pH 8.0. Unincorporated nucleotides were removed using NucTrap push columns (Stratagene). The blot was prehybridized for 1 h and then hybridized with the GAPDH probe for 16 h at 65°C in 5 ml of hybridization solution (1.0 M NaCl, 50 mM Tris · HCl, pH 7.5, 10% dextran sulfate, 1% SDS, and 100 µg/ml denatured sonicated salmon sperm DNA). The blot was washed four times briefly at room temperature, 30 min at 65°C, and 5 min at room temperature in 2× SSC, 0.1% SDS. A final rinse was performed at room temperature in 2× SSC. The blot was exposed to X-ray film, and the autoradiogram was quantified densitometrically.

Western blot analysis. Cardiomyocytes were washed five times in ice-cold phosphate-buffered saline (PBS) and drained. Cells were lysed in 1 ml of ice-cold 1× SDS loading buffer (62.5 mM Tris · HCl, pH 6.8, 1% SDS, 10% glycerol, and 5 mM DTT). Lysates were collected into a microcentrifuge tube with the aid of a cell scraper, heated at 100°C for 5 min, and subsequently centrifuged at 10,000 g for 5 min. Supernatants were transferred to a fresh microcentrifuge tube, and protein concentration was determined using a modified Lowry assay (DC Protein Assay Kit; Bio-Rad). Equal amounts of protein sample (15 µg) were resolved on a discontinuous 12% SDS-polyacrylamide gel using a Tris-glycine buffer system (25 mM Tris, 192 mM glycine, and 0.1% SDS). Proteins were transferred onto Protean nitrocellulose (Schleicher & Schuell, Keene, NH) by electroblotting for 5 h (120 V, constant voltage) at 4°C in 25 mM Tris, 192 mM glycine, and 20% methanol. Air-dried membranes rinsed briefly in several changes of PBS were blocked overnight at room temperature in 10 ml of blocking solution [5% Carnation skim milk powder and 0.02% sodium azide dissolved in 1× PBS, 0.05% Tween-20 (Sigma)]. Polyclonal rabbit anti-rat HSP 32 primary antibody (StressGen, Victoria, BC, Canada) was diluted 1:1,000 in fresh blocking solution (10 ml) and immunoreacted to the blot for 1 h at room temperature with gentle agitation. After extensive washings in 1× PBS, 0.05% Tween-20, blots were subsequently immunoreacted with a 1:3,000 dilution of goat anti-rabbit horseradish peroxidase-conjugated secondary antibody (Bio-Rad) in fresh blocking solution (10 ml) also for 1 h at room temperature. A second series of washes in 1× PBS, 0.5% Tween was followed by enzymatic chemiluminescense detection using a commercially available kit (ECL Western blotting detection reagents; Amersham, Arlington Heights, IL). Membranes were exposed to X-OMAT AR film (Eastman Kodak) at room temperature for 30-120 min, and autoradiograms were quantified densitometrically. Purified recombinant rat HSP 32 (10 ng) and HO-2 (50 ng) antigen samples (StressGen) were used to demonstrate the specificity of the immunoreaction, and parallel gels stained with Coomassie brilliant blue R-250 were used to confirm relative sample loading.

Metabolite and enzyme assays. The concentration of lactic acid and glucose was determined on aliquots of medium obtained from normoxic and hypoxic treated cultures. An additional aliquot of the medium was saved for subsequent analysis of creatine kinase activity. For lactic acid determination, 0.5 ml of medium sample was quickly deproteinized in 1 ml of 8% perchloric acid and centrifuged at 5,000 g for 5 min at 4°C. An aliquot of the supernatant (10 µl) was added to 0.3 ml of glycine-hydrazine buffer (0.33 M glycine and 0.27 M hydrazine, pH 9.2) supplemented with 1.5 U lactate dehydrogenase (Sigma) and 0.363 mM beta -nicotinamide adenine dinucleotide (Sigma). Sample mixtures (in triplicate) were incubated at 37°C for 45 min on a microplate reader (MR 5000, Dynatech) read at 340 nm, and lactic acid concentration was determined against a standard curve. Medium glucose concentration was assayed in duplicate using the o-toluidine procedure (12). The activity of creatine kinase in the medium was determined using a kit (Sigma).

Statistical analysis. Data from individual experiments are reported as means ± SE. Intergroup comparisons were performed with an analysis of variance with repeated measures. The Scheffé's post hoc test was used to determine specific differences between treatment groups when a significant F value was found. Differences were considered statistically significant at P < 0.05.

    RESULTS
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Metabolic characteristics of hypoxic aligned cardiomyocytes. Primary rat neonatal cardiomyocytes were plated on an aligned collagen matrix to produce cultures with an elongated rod shape and an organized myofibrillar structure more similar to the intact myocardial phenotype (27). Aligned cardiomyocytes (7-8 days postplating) with synchronous beating rates were subjected to 12 h of hypoxia in an airtight chamber. The reversible reduction reaction of the methylene blue-anaerobic indicators within the chamber verified an environment of <0.5% O2 throughout the duration of hypoxia. Standard culture medium containing both serum and glucose was replenished before the cardiomyocyte cultures were subjected to hypoxic (<0.5% O2) or control normoxic (21% O2) conditions at 37°C. In an initial experiment, the activity of creatine kinase and concentration of glucose and lactic acid were determined in the medium after 12 h of hypoxia. The activity of creatine kinase in medium from both control (n = 3) and hypoxic cells (n = 3) was below the limit of detection (data not shown). The concentration of glucose in the medium of hypoxic cells was 16.6 ± 0.5 mM (n = 3) and was 17% lower compared with medium of normoxic controls (19.9 ± 0.5 mM; P < 0.01). Lactic acid concentrations in the medium of the cardiomyocytes exposed to the hypoxic environment averaged 14.7 ± 0.4 mM (n = 3) and were ~1.8-fold higher than in medium from cardiomyocytes cultured under normoxic conditions (8.0 ± 1.7 mM, n = 3; P < 0.005).

Heme oxygenase isoform mRNA expression during hypoxic and oxidative stress in aligned cardiomyocytes. Potential modulations in heme oxygenase gene expression in response to a 12-h exposure to hypoxic stress were evaluated by measuring steady-state levels of both the HSP 32 and HO-2 transcripts via Northern blot analysis. The content of GAPDH mRNA remained unchanged during hypoxia (Fig. 1A) and was used to normalize HSP 32 and HO-2 mRNA data (summarized in Fig. 1C) to correct for variability in sample loading. In this experiment, hypoxia induced HSP 32 mRNA levels by 7.5-fold above the normoxic counterpart (P < 0.05). To confirm inducibility of the HSP 32 gene to an oxidative challenge in the aligned cardiomyocyte cultures, we incubated cells with 80, 160, or 300 µM H2O2 for 3 h under normoxic conditions. The 80 µM H2O2 treatment induced HSP 32 mRNA to a level similar to that seen with 12 h of hypoxia (Fig. 1C) and was higher than the two- to threefold induction we have observed in other experiments with 100 µM H2O2 [in densitometric units: H2O2 treated, 23.5 ± 0.6 (n = 2); control, 8.5 ± 0.5 (n = 3)] and recently reported by Hoshida et al. (10). It should be noted that although total RNA from cells treated with 80 or 100 µM H2O2 appeared intact, RNA isolated from cardiomyocytes exposed to 160 and 300 µM H2O2 consistently showed evidence of extensive degradation (Fig. 1B). Thus the lower level induction observed at H2O2 concentrations >100 µM may occur because of the competing influence of RNA degradation on HSP 32 mRNA concentration. These results are consistent with the findings of Nag et al. (21), which indicated that myofibrillar proteins in neonatal cardiomyocytes were degraded at concentrations between 30 and 100 µM H2O2.


View larger version (42K):
[in this window]
[in a new window]
  		Fig. 1.   Northern blot analysis of heme oxygenase-1 [heat stress protein 32 (HSP 32)] and heme oxygenase-2 (HO-2) mRNA levels after hypoxia and H2O2 treatments. Cardiomyocyte cultures were immediately lysed in Ultraspec after a 12-h exposure to normoxia control (NC) conditions, a 12-h exposure to hypoxia (Hyp), or a 3-h exposure to 80, 160, or 300 µM H2O2 in fresh standard medium. Total RNA (20 µg) was electrophoresed in a 1% agarose denaturing gel, blotted onto a Zeta-Probe membrane, and probed. A rat spleen total RNA sample (7 µg) was included to serve as a positive control for HSP 32 probe hybridization. A: Northern blot analysis of rat spleen (lane 1), control normoxic cardiomyocytes (lanes 2-4), hypoxic cardiomyocytes (lanes 5-7), and cardiomyocytes treated with 80 µM H2O2 (lane 8), 160 µM H2O2 (lane 9), and 300 µM H2O2 (lane 10). The blot was hybridized using a radiolabeled rat HSP 32 cDNA probe (top) and then stripped and subsequently rehybridized with a polymerase chain re action (PCR)-generated rat HO-2 cDNA probe (middle) and then a rat glyceraldehyde-3-phosphate dehydrogenase (GAPDH) cDNA probe (bottom). B: loading and integrity of total RNA samples across treatments. HSP 32 and HO-2 mRNA densitometric signals (C) were normalized against their respective GAPDH mRNA densitometric signal to correct for variability in total RNA loading. Because of the apparent stimulatory effects of the 160 and 180 µM H2O2 treatments on total RNA degradation (B), these samples were not quantified. Both NC and Hyp treatments con sisted of 3 separate cardiomyocyte cultures obtained across 2 separate batches. Each H2O2 treatment consisted of only 1 cardiomyocyte culture and was not included in the statistical analysis model. * P < 0.05 vs. NC cultures.

Levels of HO-2 mRNA were not significantly altered after either hypoxic or oxidative stress treatments (Fig. 1, A and C). Two HO-2 transcripts of 1.3 and 1.9 kilobase (kb) are expressed in the adult rat heart (7). Our PCR-generated HO-2 probe recognized both species in various adult rat organs, but only the 1.3-kb transcript was observed in the primary neonatal cultures, even when up to 20 µg of total RNA sample were used in Northern blot analysis (data not shown). Moreover, levels of HSP 32 mRNA appeared to exceed that of the HO-2 mRNA. This was evidenced by varying exposure times required for proper quantification of the autoradiograms for each probe, despite similar specific activities between them (HSP 32, 1-2 days; HO-2, 5-7 days; GAPDH, 6-12 h).

Antioxidant NAC treatment partially blocked HSP 32 induction during hypoxia. To investigate oxidative stress as a potential signal in the induction of HSP 32 during hypoxia, we supplemented cardiomyocyte cultures with the antioxidant and glutathione precursor NAC. Before these studies it was necessary to establish that HSP 32 was expressed at a constant basal level during 12 h of culture in the presence or absence of NAC. Recently, expression of HSP 32 has been shown to be increased during the apparent oxidative stress associated with the procedures used in the first 72 h after establishment of primary cardiomyocyte cultures (10). To minimize the impact of this "culture shock," we used cells 5-7 days postplating. However, we performed an experiment to determine if medium replacement could also constitute a form of culture shock. HSP 32 mRNA content (densitometric units) in cardiomyocytes (5-7 days postplating) just before a medium change (0 h; 0.79 ± 0.05) was not significantly different (P > 0.05) from that in cells 2 h (0.65 ± 0.04), 6 h (0.60 ± 0.03), or 12 h (0.86 ± 0.07) after replacement with fresh medium. Similarly, exposure to fresh standard medium supplemented with 20 mM NAC did not alter HSP 32 mRNA levels at 2 h (0.14 ± 0.03), 6 h (0.14 ± 0.02), or 12 h (0.14 ± 0.01) compared with standard medium at 0 h (0.16 ± 0.02). HO-2 mRNA levels remained unaffected by either treatments (data not shown). Thus routine maintenance of confluent aligned cardiomyocyte cultures in standard medium subsequently followed by medium replacement did not impart a significant stress to the cell cultures. Furthermore, these data indicated that a simplified experimental design could be used in subsequent experiments. First, because the 0-h and 12-h time points showed similar levels of HSP 32 mRNA, a 0-h control would not be necessary. Second, because inclusion of NAC did not change HSP 32 mRNA levels, a normoxic + NAC control group would not be necessary.

We next determined HSP 32 mRNA and protein expression in cardiomyocytes subjected to 12 h of NC, Hyp + NAC, or Hyp conditions. Cells exposed to Hyp or Hyp + NAC conditions demonstrated significant 3.4-fold and 2.6-fold increases in HSP 32 mRNA content, respectively, compared with cells exposed to NC conditions (Fig. 2, A and C). However, the extent of induction was significantly less (P < 0.05) with Hyp + NAC cardiomyocytes compared with Hyp myocytes. Similar effects of NAC pretreatment on the level of HSP 32 protein expression were also evident (Fig. 3). In the Hyp cardiomyocytes, HSP 32 protein content was increased 2.5-fold (P < 0.005) above that in NC cardiomyocytes, whereas the value in the Hyp + NAC group was only 1.5-fold higher than, and not statistically different from (P > 0.05), that in the NC cardiomyocytes. Increasing the antioxidative capacity of neonatal cardiomyocytes in culture thus significantly reduced the degree of HSP 32 mRNA and protein induction during a 12-h hypoxic exposure.


View larger version (35K):
[in this window]
[in a new window]
  		Fig. 2.   Northern blot analysis of HSP 32 mRNA induction during hypoxia with and without N-acetyl-L-cysteine (NAC). Medium from confluent cardiomyocyte cultures was replaced 1 h before a 12-h exposure to normoxic control conditions in standard medium (NC) or before a 12-h hypoxic exposure in either standard medium (Hyp) or standard medium supplemented with 20 mM NAC (Hyp + NAC). Immediately after treatment, cells were lysed with Ultraspec, and total RNA was isolated. Northern blot analysis of HSP 32 and GAPDH mRNA levels was performed using 12 µg of total RNA as described in MATERIALS AND METHODS. A: autoradiograms derived from Northern blot analysis of samples obtained from rat spleen (Spl), rat testis (Test), NC treatments, Hyp treatments, and Hyp + NAC treatments. B: total RNA loading and integrity across the treatments. Densitometric signals obtained with a radiolabeled rat HSP 32 cDNA probe were normalized against the respective signal subsequently obtained with a rat GAPDH cDNA probe (C). Each treatment consisted of a total of 4 cardiomyocyte cultures obtained across 2 separate isolations. To allow for rapid processing of only a small number of treated cell cultures at a time, one half of the cultures (the 1st isolation) was exposed to either NC, Hyp, or Hyp + NAC treatments on 1 day and the other half of the cultures (the 2nd isolation) was treated on the following day under identical conditions. P <=  0.0001 vs. * NC and ** Hyp.


View larger version (22K):
[in this window]
[in a new window]
  		Fig. 3.   Representative Western blot of HSP 32 protein during hypoxia with and without NAC supplementation. Medium from confluent cardiomyocyte cultures was replaced 1 h before a 12-h exposure to normoxic control conditions in standard medium (NC) or before a 12-h hypoxic exposure in either standard medium (Hyp) or standard medium supplemented with 20 mM NAC (Hyp + NAC). Cells were immediately lysed in sodium dodecyl sulfate loading buffer, and proteins were separated in a 12% discontinuous acrylamide gel and blotted to Protean nitrocellulose membrane as described in detail in MATERIALS AND METHODS. A: representative autoradiogram. Purified rat recombinant HSP 32 antigen (HSP 32a; 10 ng) and rat recombinant HO-2 antigen (HO-2a; 50 ng) were used to confirm specificity of the primary and secondary antibody immunoreaction. Protein samples obtained from NC, Hyp, and Hyp + NAC treatments were immunoreacted against a primary rabbit anti-rat HSP 32 polyclonal antibody (StressGen) and detected with a horseradish peroxidase-based chemiluminescence system (Amersham). A duplicate gel was stained with Coomassie blue and confirmed relative equivalent loadings (B). Markers (M): 107, 76, 52, 36.8, 27.2, and 19 kDa. The autoradiograms were quantified densitometrically and the data are summarized as shown in C. P < 0.005 vs. * NC and ** Hyp.

Inclusion of 20 mM NAC did not appear to alter the metabolic response of the cardiomyocytes to hypoxia. Medium glucose (14.9 ± 0.6 mM) and lactic acid (12.4 ± 0.7 mM) concentrations in the Hyp group were not different (P > 0.05) than in the Hyp + NAC group (glucose = 15.2 ± 0.5 mM; lactic acid = 11.5 ± 0.8 mM). Medium glucose concentration (17.6 ± 0.4 mM) in the NC group was 15% higher (P < 0.05) compared with the Hyp (14.9 ± 0.6 mM) or Hyp + NAC (15.2 ± 0.5 mM) groups. Medium lactate concentrations in the Hyp (12.4 ± 0.7 mM) and Hyp + NAC (11.5 ± 0.8 mM) groups were both increased ~1.6-fold compared with the NC group (7.6 ± 0.6 mM)

    DISCUSSION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Hypoxia and heme oxygenase isoform expression in aligned cardiomyocytes. Primary cultures of aligned cardiomyocytes oriented on a collagen matrix gel were used to study heme oxygenase isoform expression during hypoxia. After 12 h of hypoxia, the cells did not release any measurable creatine kinase into the medium, suggesting that membrane integrity and cell viability were not significantly affected by hypoxia. These results are similar to those previously reported by Webster et al. (32) after 12 h of hypoxia using similar culture conditions. The significant decreases in medium glucose and increases in medium lactic acid after 12 h of hypoxia were similar to those noted in other cardiomyocyte cultures (13, 31) and indicated an increased reliance on anaerobic catabolism of glucose. Concomitant with the switch to anaerobic metabolism, the aligned cardiomyocytes selectively induced expression of HSP 32 mRNA but did not change expression of HO-2 mRNA. The induction of the HSP 32 isoform is similar to the recent data of Eyssen-Hernandez et al. (8) using nonaligned cardiomyocyte cell cultures. The nonresponsiveness of the HO-2 isoform to a hypoxic environment has also been observed in cultures of vascular smooth muscle cells (19). Hence, the lack of change in HO-2 isoform expression in cardiomyocytes allows the conclusion that changes in HSP 32 mRNA were not simply a compensatory response to a decrease in HO-2 mRNA. We also report that increases in HSP 32 mRNA correlated with a similar increase in HSP 32 protein. This result is consistent with pretranslational control of HSP 32 expression in cardiomyocytes during hypoxia as observed in cultured vascular smooth muscle cells (16, 19) and Chinese hamster ovary cells (20).

The elevated levels of HSP 32 protein suggest that the rate of the heme oxygenase reaction was accelerated in the cultured cardiomyocytes during hypoxia. Rat cardiomyocytes have been previously shown by immunocytochemical procedures to synthesize HSP 32 in culture (10). Acceleration of the heme oxygenase reaction as shown in other cell types leads to an increase in antioxidant protection (1). These potential adaptive consequences of an elevated HSP 32 expression are consistent with the suggestion of Hermes-Lima and Storey (9) that prolonged hypoxia may cause adaptations that are preparatory for the potentially harmful oxygen reperfusion stress. The increased expression of the HSP 32 isoform in cultured cardiomyocytes during prolonged hypoxia indicates a need for future experiments directed at studying the physiological consequences of accelerated heme catabolism in the myocardium in vivo.

Recently, it has been shown that in isolated perfused rat hearts, a short ischemic bout (5 or 20 min) failed to induce HSP 32 mRNA (18). However, after reperfusion HSP 32 mRNA and protein were both induced. Although these results could indicate that hypoxia associated with ischemia may not induce HSP 32 in the intact myocardium, it is also possible that the ischemic episode was too short to allow for an increased synthesis of HSP 32 mRNA and protein to be observed immediately after ischemia. Webster et al. (31) have suggested that the greater tolerance of the cultured neonatal cardiomyocyte to hypoxic environments may allow the investigation of underlying adaptations ordinarily missed when intact heart preparations were used.

Redox signaling during the hypoxic induction of HSP 32. Similar to Murphy et al. (20), we hypothesized that during hypoxia increased levels of unscavenged ROS may be part of the signaling pathway utilized to effect an increased expression of HSP 32. This was based on 1) the presumed function of HSP 32 induction, i.e., to increase the antioxidant capacity of the cell (1), 2) the ability of oxidants such as H2O2 to increase expression of the HSP 32 gene (10), and 3) evidence of oxidative stress during prolonged hypoxia as indicated by decreases in the level of glutathione (15) and increased lipid peroxidation (22). Our experiments in which the medium of cardiomyocytes were supplemented with the glutathione precursor NAC were designed to test this hypothesis. Hypoxia-induced HSP 32 expression was significantly reduced after pretreatment with NAC. We interpret our findings to indicate that ROS present during hypoxia were involved in one or more signaling pathways utilized to increase expression of HSP 32 in cultures of rat neonatal cardiomyocytes.

The signaling pathways utilized during adaptation to hypoxia in cultured rat neonatal cardiomyocytes have been shown to involve activation of an intracellular mitogen-activated protein kinase cascade that may culminate in the increased binding of AP-1 transcription factors and/or synthesis of the individual AP-1 subunit proteins (26, 31). In the mouse HSP 32 gene, under normoxic conditions, two inducible enhancer elements, both of which contain AP-1 factor binding sequences, are critical for mediating the induction of HSP 32 transcription after exposure to the pro-oxidant heme or oxidant H2O2 (2). Using an enhancer promoter construct stably incorporated into L929 fibroblastic cells, pretreatment of the cells with 20 mM NAC completely blocked activation of transcription by H2O2. In our experiments, 20 mM NAC partially decreased the extent of hypoxic induction of HSP 32 mRNA expression. Increasing the antioxidant status of the cardiomyocytes thus may have interrupted oxidant-sensitive pathways targeting AP-1 or perhaps other DNA binding proteins to induce the HSP 32 gene during hypoxia.

The mechanisms or signals that trigger the hypoxic induction of genes are complex and may involve tissue-specific or gene-specific mechanisms (for review, see Ref. 5). In a recent study using cultured vascular smooth muscle cells, Lee et al. (16) reported the functional dependence of the hypoxic induction of HSP 32 transcription upon HIF-1 binding. The authors also reported that removal of the enhancer elements containing AP-1 binding sites did not interfere with the hypoxic induction. Hence, these authors hypothesized that oxidative signaling may not be important in the induction of HSP 32 gene in vascular smooth muscle cells. Although it is difficult to completely reconcile our results with that of Lee et al. (16), a number of explanations are possible. First, there may be tissue-specific differences in the way vascular smooth muscle cells and cardiomyocytes regulate the HSP 32 gene during hypoxia. This might reflect, for example, the capacity of each cell type to buffer or generate ROS. Second, the conditions used to culture the two cell types may have biased the cells to a particular response to hypoxia. For example, Hoshida et al. (10) have demonstrated that primary cultures of cardiomyocytes undergo a phase during the first several days postplating characterized by oxidative stress or culture shock. Because we performed our studies after this phase, it is not clear how adaptations during the culturing of vascular smooth muscle cells might influence the balance of oxidative stress during hypoxia. Last, because both alpha - and beta -HIF-1 subunit mRNAs are present in rat heart (33), and the inclusion of 20 mM NAC only partially attenuated HSP 32 expression during hypoxia, it is possible that the redox-sensitive pathway suggested by our data may work in conjunction with the HIF-1-dependent pathway. Interestingly, increasing the oxidant status before hypoxic treatment of HeLa cells decreases the stability of the alpha -HIF subunit and interferes with binding of HIF-1 to the erythropoietin gene (11, 30). Hence, if HIF-1 were to be solely responsible for inducing the HSP 32 gene in cardiomyocytes during hypoxia, increasing the antioxidant status might have led to an increase in HSP 32 induction rather than the decrease as observed. To determine the relative importance of signaling pathways sensitive to ROS during hypoxia, additional studies are needed to determine the DNA binding sites needed to mediate the hypoxic induction of the gene in cardiomyocytes.

Summary. The increased levels of HSP 32 protein in aligned cardiomyocyte cell cultures after hypoxia correlated with an increased expression of the HSP 32 mRNA. These data suggest that cardiomyocytes respond to hypoxic stress by producing an adaptive increase in the capacity for heme degradation. A portion of the hypoxic induction of HSP 32 expression was blocked when cells were pretreated with the glutathione precursor NAC, suggesting a component of the signaling may involve an ROS-dependent activation step. Future studies aimed at understanding the importance of ROS as a regulatory signal generated in the hypoxic myocardium during hypoxia appear to be warranted.

    ACKNOWLEDGEMENTS

We thank David Simpson for skillful technical assistance with the culturing of the aligned cardiomyocytes, Gabor Kemeny for help in the rapid harvesting of cells, and Thomas Borg for use of the culture facility.

    FOOTNOTES

Address for reprint requests: D. A. Essig, Dept. of Exercise Science, Univ. of South Carolina, Columbia, SC 29208.

Received 12 June 1997; accepted in final form 18 November 1997.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

1.   Abraham, N. G., Y. Lavorsky, M. L. Schwartzman, R. A. Stolz, R. D. Levere, M. E. Gerritsen, S. Shibahara, and A. Kappas. Transfection of the human heme oxygenase gene into rabbit coronary microvessel endothelial cells: protective effect against heme and hemoglobin toxicity. Proc. Natl. Acad. Sci. USA 92: 6798-6802, 1995[Abstract/Free Full Text].

2.   Alam, J., S. Camhi, and A. M. K. Choi. Identification of a second region upstream of the mouse heme oxygenase-1 gene that functions as a basal level and inducer-dependent transcription enhancer. J. Biol. Chem. 270: 11977-11984, 1995[Abstract/Free Full Text].

3.   Applegate, L. M., A. Noel, G. Vile, E. Frenk, and R. M. Tyrell. Two genes contribute to different extents to the heme oxygenase enzyme activity measured in cultured human skin fibroblasts and keratinocytes: implications for protection against oxidant stress. Photochem. Photobiol. 61: 285-291, 1995[Medline].

4.   Borg, T. K., K. Rubin, E. Lundgren, K. Borg, and B. Obrink. Recognition of extracellular matrix components by neonatal and adult cardiac myocytes. Dev. Biol. 104: 86-96, 1984[Medline].

5.   Bunn, H. F., and R. O. Poyton. Oxygen sensing and molecular adaptation to hypoxia. Physiol. Rev. 76: 839-885, 1996[Abstract/Free Full Text].

6.   Essig, D. A., D. R. Borger, and D. A. Jackson. Induction of heme oxygenase-1 (HSP32) mRNA in skeletal muscle following contractions. Am. J. Physiol. 272 (Cell Physiol. 41): C59-C67, 1997[Abstract/Free Full Text].

7.   Ewing, J. F., V. S. Raju, and M. D. Maines. Induction of heart heme oxygenase-1 (HSP32) by hyperthermia: possible role in stress-mediated elevation of cyclic 3',5'-guanosine monophosphate. J. Pharmacol. Exp. Ther. 271: 408-414, 1994[Abstract/Free Full Text].

8.   Eyssen-Hernandez, R., A. Ladoux, and C. Frelin. Differential regulation of cardiac heme oxygenase-1 and vascular endothelial growth factor mRNA expressions by hemin, heavy metals, heat shock and anoxia. FEBS Lett. 382: 229-233, 1996[Medline].

9.   Hermes-Lima, M., and K. B. Storey. Relationship between anoxia exposure and antioxidant status in the frog Rana pipiens. Am. J. Physiol. 271 (Regulatory Integrative Comp. Physiol. 40): R918-R925, 1996[Abstract/Free Full Text].

10.   Hoshida, S., M. Nishida, N. Yamashita, J. Igarashi, K. Aoki, M. Hori, T. Kozuya, and M. Tada. Heme oxygenase expression and its relation to oxidative stress in rat neonatal cardiomyocytes. J. Mol. Cell. Cardiol. 28: 1845-1855, 1996[Medline].

11.   Huang, L. E., Z. Arany, D. M. Livingston, and H. F. Bunn. Activation of hypoxia-inducible transcription factor depends primarily upon redox-sensitive stabilization of its alpha -subunit. J. Biol. Chem. 271: 32253-32259, 1996[Abstract/Free Full Text].

12.   Hyvarinen, A., and E. Nikkilo. Specific determination of blood glucose. Clin. Chim. Acta 7: 140-143, 1962[Medline].

13.   Iwaki, K., S.-H. Chi, W. H. Dillman, and R. Mestril. Induction of HSP 70 in cultured rat neonatal cardiomyocytes by hypoxia and metabolic stress. Circulation 87: 2023-2032, 1993[Abstract/Free Full Text].

14.   Katayose, D., S. Isoyama, H. Fujita, and S. Shibahara. Separate regulation of heme oxygenase and heat shock protein 70 mRNA expression in the rat heart by hemodynamic stress. Biochem. Biophys. Res. Commun. 191: 587-594, 1993[Medline].

15.   Kwok, T. T., and R. M. Sutherland. The relationship between radiation response of human squamous carcinoma cells and specific metabolic changes induced by chronic hypoxia. Int. J. Radiat. Oncol. Biol. Phys. 16: 1301-1305, 1989[Medline].

16.   Lee, P. J., B.-H. Jiang, B. Y. Chin, N. V. Iyer, J. Alam, G. L. Semenza, and A. M. K. Choi. Hypoxia-inducible factor-1 mediates transcriptional activation of the heme oxygenase-1 gene in response to hypoxia. J. Biol. Chem. 272: 5375-5381, 1997[Abstract/Free Full Text].

17.   Marber, M. S., R. Mestril, S. H. Chi, M. R. Sayen, D. M. Yellon, and W. H. Dillman. Overexpression of the rat inducible 70-kD heat stress protein in a transgenic mouse increases the resistance of the heart to ischemic injury. J. Clin. Invest. 95: 1446-1456, 1995.

18.   Maulik, N., H. S. Sharma, and D. K. Das. Induction of the heme oxygenase gene expression during the reperfusion of ischemic rat myocardium. J. Mol. Cell. Cardiol. 28: 1261-1270, 1996[Medline].

19.   Morita, T., M. A. Perrella, M. E. Lee, and S. Kourembanas. Smooth muscle cell-derived carbon monoxide is a regulator of vascular cGMP. Proc. Natl. Acad. Sci. USA 92: 1475-1479, 1995[Abstract/Free Full Text].

20.   Murphy, B. J., K. R. Laderoute, S. M. Short, and R. M. Sutherland. The identification of heme oxygenase as a major hypoxic stress protein in Chinese hamster ovary cells. Br. J. Cancer 64: 69-73, 1991[Medline].

21.   Nag, A. C., P. Srepathi, M. L. Lee, and J. R. Reddan. Effect of oxidative insult on young and adult cardiac muscle cells in vitro. Cytobios 85: 7-27, 1996[Medline].

22.   Park, Y., S. Kanekal, and J. P. Kehrer. Oxidative changes in hypoxic rat heart tissue. Am. J. Physiol. 260 (Heart Circ. Physiol. 29): H1395-H1405, 1991[Abstract/Free Full Text].

23.   Plumier, J.-C. L., H. A. Robertson, and R. W. Currie. Differential accumulation of mRNA for immediate early genes and heat shock genes in heart after ischemic injury. J. Mol. Cell. Cardiol. 28: 1251-1260, 1996[Medline].

24.   Plumier, J.-C. L., B. M. Ross, R. W. Currie, C. E. Angelides, H. Kazlaris, G. Kollias, and G. N. Pagoulatos. Transgenic mice expressing the human heat shock protein 70 have improved post-ischemic myocardial recovery. J. Clin. Invest. 95: 1854-1860, 1995.

25.   Sambrook, J., E. F. Fritsch, and T. Maniatis. Molecular Cloning: A Laboratory Manual. Cold Springs Harbor, NY: Cold Springs Harbor Laboratory, 1989.

26.   Seko, Y., K. Tobe, K. Ueki, T. Kadowaki, and Y. Yazaki. Hypoxia and hypoxia/reoxygenation activate raf-1, mitogen-activated protein kinase kinase, mitogen-activated protein kinases and S5 kinase in cultured rat cardiac myocytes. Circ. Res. 78: 82-90, 1996[Abstract/Free Full Text].

27.   Simpson, D. G., L. Terracio, M. Terracio, R. L. Price, D. C. Turner, and T. K. Borg. Modulation of cardiac myocyte phenotype in vitro by the composition and orientation of the extracellular matrix. J. Cell. Physiol. 161: 89-105, 1994[Medline].

28.   Stocker, R., Y. Yamamoto, A. F. McDonagh, A. N. Glazer, and B. N. Ames. Bilirubin is an antioxidant of possible physiological importance. Science 235: 1043-1046, 1987[Abstract/Free Full Text].

29.   Tan, S. S., and H. Weis. Development of a sensitive reverse transcriptase PCR assay, RT-RPCR, utilizing rapid cycle times. PCR Methods Applic. 2: 137-143, 1992[Medline].

30.   Wang, G. L., B.-H. Jiang, and G. L. Semenza. Effect of altered redox states on expression and DNA-binding activity of hypoxia-inducible factor-1. Biochem. Biophys. Res. Commun. 212: 550-556, 1995[Medline].

31.   Webster, K. A., D. J. Discher, and N. H. Bishphoric. Induction and nuclear accumulation of fos and jun proto-oncogenes in hypoxic cardiac myocytes. J. Biol. Chem. 268: 14678-14681, 1993[Abstract/Free Full Text].

32.   Webster, K. A., D. J. Discher, and N. H. Bishphoric. Regulation of fos and jun immediate early genes by redox or metabolic stress in cardiac myocytes. Circ. Res. 74: 679-686, 1994[Abstract/Free Full Text].

33.   Wiener, C. M., G. Booth, and G. L. Semenza. In vivo expression of mRNAs encoding hypoxia-inducible factor 1. Biochem. Biophys. Res. Commun. 225: 485-488, 1996[Medline].

AJP Heart Circ Physiol 274(3):H965-H973
0363-6135/98 $5.00 Copyright © 1998 the American Physiological Society


This article has been cited by other articles:


Home page
 Exp. Biol. Med.Home page
T. V. Serebrovskaya, E. B. Manukhina, M. L. Smith, H. F. Downey, and R. T. Mallet
Intermittent Hypoxia: Cause of or Therapy for Systemic Hypertension?
Experimental Biology and Medicine, June 1, 2008; 233(6): 627 - 650.
[Abstract] [Full Text] [PDF]

Home page
 Physiol. Rev.Home page
S. W. Ryter, J. Alam, and A. M. K. Choi
Heme Oxygenase-1/Carbon Monoxide: From Basic Science to Therapeutic Applications
Physiol Rev, April 1, 2006; 86(2): 583 - 650.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Renal Physiol.Home page
Z.-Z. Yang and A.-P. Zou
Transcriptional regulation of heme oxygenases by HIF-1alpha in renal medullary interstitial cells
Am J Physiol Renal Physiol, November 1, 2001; 281(5): F900 - F908.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
R. B. Hunter, H. Mitchell-Felton, D. A. Essig, and S. C. Kandarian
Expression of endoplasmic reticulum stress proteins during skeletal muscle disuse atrophy
Am J Physiol Cell Physiol, October 1, 2001; 281(4): C1285 - C1290.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
R. Motterlini, R. Foresti, R. Bassi, V. Calabrese, J. E. Clark, and C. J. Green
Endothelial Heme Oxygenase-1 Induction by Hypoxia. MODULATION BY INDUCIBLE NITRIC-OXIDE SYNTHASE AND S-NITROSOTHIOLS
J. Biol. Chem., April 28, 2000; 275(18): 13613 - 13620.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
M. V. Panchenko, H. W. Farber, and J. H. Korn
Induction of heme oxygenase-1 by hypoxia and free radicals in human dermal fibroblasts
Am J Physiol Cell Physiol, January 1, 2000; 278(1): C92 - C101.
[Abstract] [Full Text] [PDF]

Home page
 Exp. Biol. Med.Home page
R. Galbraith
Heme Oxygenase: Who Needs It?
Experimental Biology and Medicine, December 1, 1999; 222(3): 299 - 305.
[Abstract] [Full Text]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Borger, D. R.
Right arrow Articles by Essig, D. A.
Right arrow