Abstract

Despite extensive studies, the fundamental mechanisms responsible for the development and progression of cardiovascular diseases have not yet been fully elucidated. Recent experimental and clinical studies have suggested that reactive oxygen species play a major pathological role. Oxidative stress reduction induced by flavonoids has been regarded by many as the most likely mechanism in the protective effects of these compounds; however, there is an emerging view that flavonoids may also exert modulatory actions on protein kinase and lipid kinase signaling pathways. Quercetin, a major flavonoid present in the human diet, has been widely studied, and its biological properties are consistent with its protective role in the cardiovascular system. However, it remains unknown whether the cardioprotective effects of quercetin may also occur through the modulation of genes involved in cell survival. The main goal of this study was to examine the gene expression profiling of cultured rat primary cardiomyocytes treated with quercetin using DNA microarrays and to relate these data to functional effects. Results showed distinct temporal changes in gene expression induced by quercetin and a strong upregulation of phase 2 enzymes, highlighting quercetin ability to act also with an indirect antioxidant mechanism.

  • flavonoids
  • phase 2 enzymes
  • oxidative stress

heart diseases are the leading cause of morbidity and mortality in industrialized countries (55). Despite extensive studies, the fundamental mechanisms responsible for their development and progression are still under debate. Recent experimental and clinical studies have suggested that reactive oxygen species (ROS) play a major pathological role (4, 13). The cellular sources of ROS generation within the heart include cardiac myocytes, endothelial cells, and neutrophils. Potential sources of ROS in cardiac myocytes include mitochondrial electron transport, NADPH oxidase, and xantine dehydrogenase/xantine oxidase (50). One of the strongest lines of evidence for the involvement of ROS in cardiovascular diseases is the ability of a number of structurally unrelated compounds with antioxidant properties to protect against cardiovascular pathophysiology, including myocardial ischemia-reperfusion injury, cardiomyopathy, and arterial atherogenesis (14, 23, 26, 29).

Recently, much of the attention has been focused on flavonoids, naturally occurring polyphenolic compounds, as food factors that may be beneficial for cardiovascular disease prevention. Various epidemiological studies have shown an inverse correlation between the consumption of flavonoid-rich foods and cardiovascular disease risk (6, 28, 37). Oxidative stress reduction by flavonoids has been regarded as the most likely mechanism in the protective effects of these compounds; however, there is an emerging view that flavonoids may also exert modulatory effects, acting through protein kinase and lipid kinase signaling pathways (5, 59). Quercetin (Q), a major flavonoid present in the human diet, has been widely studied, and its biological properties are consistent with its protective role in the cardiovascular system (30). In a previous paper, our laboratory has demonstrated that Q is able to protect cardiac cells from oxidative stress, acting both as an antioxidant and a modulator of the signal transduction pathway related to apoptosis (5). However, it remains unknown whether the cardioprotective effects of Q may also occur through the modulation of genes involved in cell survival. Nutrigenomics is the scientific study by which some of these questions can be answered. The concept of nutrigenomics has risen, thanks to the development of new techniques, like global gene expression by microarray technology, successfully applied to study the transcriptional changes occurring in heart, vessels, and blood cells in different cardiovascular disorders. DNA microarray technology could promote an increased understanding of how Q influences gene expressions and regulates those genes responsible for cardioprotection. To our knowledge, the cardiomyocyte expression profile of genes associated with response to Q has not yet been reported. The present study was, therefore, designed to examine the gene expression profiling of cultured rat primary cardiomyocytes treated with Q using DNA microarrays and to relate these data to functional effects. The results showed distinct temporal changes in gene expression induced by Q and a strong upregulation of phase 2 enzymes. In addition, to demonstrate the physiological relevance of the gene expression analysis, cell viability and ROS production were measured in cardiomyocytes treated with Q for selected time periods in the presence of oxidative stress.

MATERIALS AND METHODS

Chemicals.

CelLytic M, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-tetrazolium bromide (MTT), 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA), ferrozine, Q, H2O2, digitonin, glucose-6-phosphate, glucose-6-phosphate dehydrogenase, bovine serum albumin, NADP, FAD, DMSO, FeCl3, menadione, monochlorobimane (MCB), 1-chloro-2,4-dinitrobenzene (CDNB), 5,5′-dithiobis(2-nitrobenzoic) acid (DTNB), glutathione (GSH), and all other chemicals of the highest analytical grade were purchased from Sigma Chemical (St. Louis, MO), unless otherwise stated. Q was dissolved in DMSO at a concentration of 30 mM and kept at −20°C until use.

Cell culture and treatments.

Ventricular cardiomyocytes were derived from the heart of 2- to 4-day-old Wistar rats, as previously reported (7). All experiments were conducted under the “Guidelines for the Care and Use of Laboratory Animals,” published by the Office of Science and Health Reports, National Institutes of Health, and approved by the Ethics Committee of our Institution. Cells were seeded at a density of 1.2 × 106 cells/ml and were grown in DMEM F-12 culture medium treated with 10% fetal calf serum, 10% horse serum (complete medium), and 1% sodium pyruvate. Cells were routinely grown in a humidified incubator (95% humidity) with 5% CO2 at 37°C until complete confluence. In some dishes, 30 μM Q were added to the culture medium for 6, 12, and 24 h. The low micromolar concentration of Q employed in this study is similar to that used by other investigators (30, 32, 53).

RNA extraction.

Total RNA was extracted using Total RNA Isolation Mini Kit (Agilent Technologies, Palo Alto, CA), following the manufacturer's protocol. The yield and purity of the RNA were measured using NanoDrop ND-1000 Spectrophotometer (NanoDrop Technologies, Rockland, DE). The 28s and 18s ribosomal bands were checked using Agilent 2100 bioanalyzer with an RNA 6000 Nano LabChip Kit.

mRNA amplification and labeling.

For each sample, mRNA was amplified starting from 1.5 μg of total RNA by Amino Allyl MessageAmp I aRNA Kit (Ambion, Woodward Austin, TX) to obtain amino allyl antisense RNA (aaRNA), following the method developed by Van Gelder et al. (54). Only one round of amplification was performed, according to the manufacturer's protocol, with minor modification. Briefly, mRNA was reverse transcribed into cDNA single strand; after the second-strand synthesis, cDNA was in vitro transcribed in aaRNA, including an amino allyl modified nucleotide (aaUTP). Both double-stranded DNA and aaRNA underwent a purification step using columns provided with the kit. Labeling was performed using NHS ester Cy3 or Cy5 dies (Amersham Biosciences, Piscataway, NJ), which are able to react with the modified RNA. At least 5 μg of mRNA, for each sample, were labeled and purified with columns. mRNA quality, concentration, and labeling were checked by RNA 6000 Nano LabChip assay (Agilent Technologies) and Agilent 2100 Bioanalyzer. Concentrations were also checked by NanoDrop ND-1000 Spectrophotometer.

Microarray hybridization and scanning.

One hybridization with dye-swap duplication was performed to compare samples vs. reference. Reverse labeling was performed to reduce dye-specific biases in signal intensity. The same quantity of differentially labeled sample and reference (0.75 μg) was put together, fragmented, and hybridized to high-density rat arrays containing 22,575 (60-mer) oligo-nucleotide probes representing over 20,000 well-characterized rat genes, expressed sequence tags, and expressed sequence tag cluster. All steps were performed using the In Situ Hybridization kit-plus (Agilent Technologies) and following the 60-mer oligo microarray processing protocol (Agilent Technologies). Then slides were washed with the saline sodium citrate wash procedure (Agilent Technologies) and scanned with the dual-laser microarray scanner Agilent B (Agilent Technologies) at 5-μm resolution. Feature extraction and data normalization were performed with the Agilent Feature Extraction software.

Data analysis.

Raw result files were then loaded into the Resolver SE System (Rosetta Biosoftware, Seattle, WA) for data processing and normalization using the Agilent platform-specific error model. Replicated expression profiles were combined to form ratio experiments where each gene is associated to an expression fold change and a P value that assesses the statistical significance of its modulation in the treated sample compared with the control reference. Sequences with absolute fold change ≥2 and P value ≤0.001 were considered as differentially expressed.

For all selected genes, information was obtained from the National Center for Biotechnology Information (NCBI) websites UniGene, OMIM, and PubMed (www.ncbi.nlm.nih.gov). All of the details about the microarrays and data on gene expressions have been deposited in NCBIs Gene Expression Omnibus and are accessible through GEO Series accession number GSE7222 (www.ncbi.nlm.nih.gov/geo/).

RT-PCR analysis of mRNA expression.

NAD(P)H:quinone oxidoreductase (NQO1), heme oxygenase 1 (HO-1), thioredoxin (TRX) reductase 1 (TR), and glutathione S-transferase (GST) genes were validated using the End-Point RT-PCR Assay, designed for conventional gel-based detection (Superarray, Frederick, MD). Total RNA was extracted as previously reported. cDNA was synthesized from 1 μg of total RNA using the ReactionReady First-Strand cDNA Synthesis Kit, according to the manufacturer's directions (Superarray). cDNA was reverse transcribed at 37°C for 60 min; finally the reaction was stopped by heating at 95°C for 5 min. To ensure optimal results, two different dilutions (1:5 and 1:10) of cDNA were prepared from each samples. PCR reaction was carried out in 25-μl volume containing ReactionReady HotStart Sweet PCR master mix (10 mM Tris·HCl, 50 mM KCl, 1.5 mM MgCl2, 0.2 mM dNTPs, and Taq DNA polymerase), 0.4 μM of each primer (Superarray), and 1 μl diluted cDNA. The cDNA amplification was started by denaturating the samples for 10 min at 95°C, followed by 35 cycles of 15 s at 95°C, 30 s at 55°C, and 30 s at 72°C. Finally, samples were held at 72°C for 7 min to ensure the complete extension of the PCR products. Fragment sizes were predicted on the basis of the mRNA sequence, as reported in Table 1.

View this table:
Table 1.

Primers used in RT-PCR

Six microliters of the amplified products were separated by electrophoresis on 10% polyacrylamide gel in Tris-borate-EDTA buffer (precast gel, Bio-Rad). The bands were stained with ethidium bromide and visualized under UV light using a Versa-Doc 4000 Imaging system (Bio-Rad).

NQO1 enzymatic activity assay.

NQO1 enzymatic activity was measured as described previously (40). Briefly, cardiomyocytes were plated in 96-well plates (2.5 × 103 cells/well) and grown under normal conditions until confluence. The plates were treated with Q for 6, 12, and 24 h, then lysated with a solution containing 0.8% digitonin. Two hundred microliters of reaction mix (0.025 mM Tris·HCl, 0.67 mg/ml bovine serum albumin, 0.01% Tween 20, 5 μM FAD, 1 mM glucose-6-phosphate, 30 μM NADP, 2 U/ml yeast glucose-6-phosphate dehydrogenase, 0.3 mg/ml MTT, 50 μM menadione) were added to each well, and the reaction was arrested after 5 min by the addition of a solution containing 0.3 mM dicoumarol, 0.5% DMSO, and 5 mM potassium phosphate buffer. The plates were then scanned at 610 nm with a microplate spectrophotometer VICTOR3 V Multilabel Counter (Perkin Elmer). NOQ1 activity was expressed as nanomoles per minute per milligram protein.

Total GST enzymatic activity assay.

Total GST activity was assayed using CDNB, according to the procedure of Habig et al. (15). This assay measures the total GST activity, because all of the different GST isoforms catalyze the conjugation of GSH with CDNB (16). After Q treatment, cells were lysated with CelLytic M and centrifuged (10,000 g for 10 min). Ten microliters of supernatant were added to 990 μl of reaction mix (100 mM phosphate buffer, pH 6.5, with 1 mM EDTA, 2 mM GSH, 2 mM CDNB), and absorbance was read at 340 nm at 30-s intervals over 5 min. GST activity was expressed as nanomoles per minute per milligram protein.

TR enzymatic activity assay.

TR activity was assayed by an in vitro reduction of DTNB to 5′-thionitrobenzoic acid using a procedure adapted from Holmgren and Bjornstedt (19). Briefly, after Q treatment, cells were lysated with CelLytic M, centrifuged at 10,000 g for 10 min, and 10 μl of supernatant were added to 990 μl of reaction mix (0.25 mM DTNB, 0.24 mM NADPH, 10 mM EDTA, 100 mM phosphate buffer, pH 7.5). The conversion of DTNB to 5′-thionitrobenzoic acid was measured spectrophotometrically at 412 nm at 10-s intervals over 1 min. GST activity was expressed as milliunits per milligram protein. One unit of TR will cause an increase in absorbance at 412 nm of 1.0 per minute per milliliter (when measured in a noncoupled assay containing DTNB alone) at pH 7.0 at 25°C.

HO-1 activity assay.

HO-1 activity was determined as reported in Ref. 3, with slight modification. Briefly, cells were lysated with CelLytic M and centrifuged (10,000 g for 10 min), and 30 μl of supernatant were added to 70 μl of iron-detection reagent (6.5 mM ferrozine, 6.5 mM neocuproine, 2.5 M ammonium acetate, and 1 M ascorbic acid). The incubation was carried out for 30 min at 37°C, and the absorbance was measured at 540 nm on a microplate reader. The iron content of the sample was calculated, comparing its absorbance to that of a standard curve obtained with FeCl3 and normalized against the protein concentration.

Protein concentration.

The protein concentration of the cell lysates was determined by the Bio-Rad Bradford protein assay (Bio-Rad Laboratories, Hercules, CA).

Reduced GSH levels.

Reduced GSH levels were determined with a fluorometric assay. GSH is specifically conjugated with MCB to form a fluorescent bimane-GSH adduct, in a reaction catalyzed by GST (45). The concentration of the bimane-GSH adducts increases during the initial 10- to 12-min period of this reaction with first-order kinetic, before leveling off (66). After Q treatment, culture medium was removed, and cells were washed twice with 0.2 ml PBS and incubated for 30 min at 37°C in 0.1 ml fresh PBS containing 50 μM MCB. After incubation, fluorescence was measured at 355 nm (excitation) and 460 nm (emission) with a microplate spectrofluorometer. Reduced GSH levels were expressed as percentage of control cells (control cells = 100%).

Detection of intracellular ROS.

The formation of intracellular ROS was evaluated using a fluorescent probe, DCFH-DA, as described by Wang and Joseph (57). Briefly, cardiomyocytes were treated for 6, 12, and 24 h with 30 μM Q. The cells were washed with PBS and then incubated with 5 μM DCFH-DA in PBS for 30 min. After DCFH-DA removal and further washing, the cells were incubated with 100 μM H2O2 for 30 min. At the end of incubation, cell fluorescence from each well was measured using a microplate spectrofluorometer VICTOR3 V Multilabel Counter (Perkin Elmer, Wellesley, MA) (excitation wavelength = 485 nm and emission wavelength = 535 nm). Intracellular antioxidant activity was expressed as the percentage of inhibition of intracellular ROS produced by H2O2 exposure.

Cell viability measurement.

Cardiomyocyte viability in the presence of Q was measured using the MTT assay. After the treatment for 6, 12, and 24 h with 30 μM Q, cells were washed twice with PBS and exposed to 100 μM H2O2 for 30 min. Controls received equivalent volumes of DMSO (vehicle). Cellular damage elicited by H2O2 treatment was evaluated by measuring MTT reduction. MTT was added to the medium (final concentration 0.5 mg/ml) and incubated for 1 h at 37°C. After incubation, MTT solutions were removed, DMSO was added, and the absorbance was measured at 595 nm using a microplate spectrophotometer VICTOR3 V Multilabel Counter (Perkin Elmer). Data were expressed as percentage of viable cells with respect to controls.

Statistics.

Each experiment was performed at least four times, and all values are represented as means ± SD. Student's t-test was used to analyze a statistical significance of the results (Prism 4, GraphPad Software, San Diego, CA). Values of P < 0.05 were considered as statistically significant.

RESULTS

Modulation of gene expression by Q.

Q treatment did not alter the expression of most of the ∼22,000 studied genes. A total of 91 genes (Fig. 1) were up- or downregulated, with most of the genes significantly changed after 6-h treatment. The number of the upregulated genes was greater than that of the downregulated. A total of eight genes were upregulated at any time exposure, but only two genes were upregulated following both 6- and 12-h treatment and one gene following 12- and 24-h treatment (Fig. 1A). The overlapping region of the Venn diagram of Fig. 1B shows that there were no downregulated genes shared by 6-, 12-, and 24-h Q treatment. Eleven genes were modulated in both the 12- and 24-h treatment. Figure 2 shows the hierarchical clustering display of data for treated cardiomyocytes based on the 91 filtered genes. Each gene is represented by a single row of colored bars. The genes were grouped into four main clusters. Cluster 1 shows a gene upregulation only after 6-h Q treatment, while the expression level profile was not significantly different from control cells at 12- and 24-h Q treatment. Cluster 2 evidences an induced gene expression at each treatment time. Cluster 3 shows a gene downregulation at 12 and 24 h, whereas cluster 4 shows a downregulation only at the shortest treatment time.

Fig. 1.

A: number of upregulated genes (fold change ≥2) in cardiomyocytes treated with 30 μM quercetin (Q) for 6, 12, and 24 h compared with the controls. B: number of downregulated genes (fold change ≤−2) in cardiomyocytes treated with 30 μM Q for 6, 12, and 24 h compared with the controls. The overlapping regions denote the number of common genes up- or downregulated. The non-overlapping regions denote the number of unique genes up- or downregulated.

Fig. 2.

Hierarchical clustering display of data in cardiomyocytes treated with 30 μM Q for 6, 12, and 24 h based on the significant 91 genes. The clustering display was performed using Rosetta Resolver SE software, as reported in materials and methods. Each gene is represented by a single row of colored bars. The red color indicates upregulation of the gene expression, and the green color denotes the downregulation of the gene expression compared with the controls.

Table 2 shows the list of the significant up- or downregulated genes in cardiomyocytes treated with 30 μM Q for 6, 12, and 24 h, related to their function. We focused our attention on genes codifying for antioxidant/detoxification enzymes. Interestingly, most of these genes belong to cluster 2, underlying a common functional regulation. In particular, we have chosen six genes critically involved in antioxidant/detoxification mechanisms related to cell survival: NQO1, HO-1, TR, GSTa3, Mgst1, and GSTp2.

View this table:
Table 2.

Upregulated (≥2.0-fold) and downregulated (≤[−]2.0-fold) genes in cardiomyocytes treated with quercetin for 6, 12, and 24 h

To verify the accuracy and validity of the array data obtained for the six selected genes, we performed RT-PCR in independent samples. Gene expression levels, reported in Fig. 3, are in agreement with the changes of gene expression observed in the microarray data.

Fig. 3.

Gene expression levels of NAD(P)H:quinone oxidoreductase (NQO1; A), heme oxygenase 1 (HO-1; B), thioredoxin reductase 1 (TR; C), glutathione S-transferase (GST) a3 (D), Mgst1 (E), and GSTp2 (F) in cardiomyocytes. Cardiomyocytes were treated with 30 μM Q for 6, 12, and 24 h, and gene expression was obtained by RT-PCR, as reported in materials and methods, and reported as fold change relative to the control (nontreated) cells after normalization using the GADP gene expression level. Data are means ± SD of four different cell cultures. Statistical analysis was performed by the Student's t-test comparing cardiomyocytes treated with Q with control cardiomyocytes. *At least P < 0.05.

Effects of Q on NQO1, HO-1, TR, total GST activities, and cellular GSH content.

NQO1, HO1, TR, and total GST activities in cardiomyocytes treated with Q are shown in Fig. 4. The activities of all four enzymes significantly increased at any exposure time compared with control cells. The increase in activities, although different in magnitude, is in agreement with the upregulation of the corresponding genes at any treatment time.

Fig. 4.

Effect of Q treatment on NQO1 (A), HO (B), TR (C), and GST (D) activities in cardiomyocytes. Cardiomyocytes were treated with 30 μM Q for 6, 12, and 24 h, and enzyme activities were evaluated as reported in materials and methods. Data are means ± SD of four different cell cultures. Statistical analysis was performed by the Student's t-test comparing cardiomyocytes treated with Q with control cardiomyocytes. *At least P < 0.05.

Figure 5 represents GSH levels in cultured cardiomyocytes treated with Q. GSH levels significantly increased after 12- and 24-h Q treatments, with the highest increase at the longest time exposure. After 6-h treatment, no differences in GSH levels were observed with respect to control cells. This increase is in agreement with the increased expression of the gene codifying for the glutamate-cysteine ligase catalytic subunit, as obtained by microarray data analysis.

Fig. 5.

Effects of Q treatment on glutathione (GSH) levels. Cardiomyocytes were treated with 30 μM Q for 6, 12, and 24 h, and GSH level was determined using the fluorescent probe monochlorobimane, as reported in materials and methods. Data are expressed as %GSH level compared with controls. Data are means ± SD of four different cell cultures. Statistical analysis was performed by the Student's t-test comparing cardiomyocytes treated with Q with control cardiomyocytes. *At least P < 0.05.

Q reduces ROS accumulation induced by H2O2.

To verify whether antioxidant genes upregulated by Q can influence the cell's ability to counteract oxidative stress, we verified the effect of Q treatment on ROS production induced by H2O2. A significantly increased formation of ROS, as detected by DCFH-DA assay, was observed in cells exposed to H2O2 (Fig. 6). ROS levels were markedly reduced in Q-treated cells at any exposure time, with the highest inhibition after 24-h treatment.

Fig. 6.

Effect of Q treatment on intracellular reactive oxygen species (ROS) production. Cardiomyocytes were treated with 30 μM Q for 6, 12, and 24 h before the addition of H2O2 (100 μM, 30 min), and then the level of intracellular ROS was determined using the peroxide-sensitive fluorescent probe 2′,7′-dichlorodihydrofluorescein diacetate, as reported in materials and methods. Data are expressed as %inhibition of ROS produced compared with H2O2 exposure. Data are means ± SD of four different cell cultures. Statistical analysis was performed by the Student's t-test. *At least P < 0.05 compared with H2O2-exposed cells.

Q protects against H2O2-induced damage.

To verify whether the upregulation of antioxidant genes after Q treatment could lead to cell protection against oxidative damage, we determined cell viability after exposure to H2O2. Incubation of cardiomyocytes with H2O2 led to a significant decrease in cell viability, as detected by MTT reduction assay (Fig. 7). Q is responsible for a marked protection at all of the treatment times, as cell viability levels of Q-treated cardiomyocytes were comparable to controls.

Fig. 7.

Protection against peroxide-induced cell damage by Q. Cardiomyocytes were treated with 30 μM Q for 6, 12, and 24 h before the addition of H2O2 (100 μM, 30 min), and cellular damage was assessed by the MTT assay and reported as %cell viability compared with controls. Data are means ± SD of four different cell cultures. Statistical analysis was performed by the Student's t-test. *At least P < 0.05 compared with H2O2-exposed cells.

DISCUSSION

Many evidences underline a critical role for oxidative stress in the development of various forms of cardiovascular disorders, including myocardial ischemia-reperfusion injury, congestive heart failure, coronary arterial atherosclerosis, and chemical-induced cardiotoxicity (21, 29, 47, 52, 58). In this context, administration of some exogenous antioxidants, such as flavonoids, has been shown to exert protective effects against oxidative stress in different “in vitro” and “in vivo” models (1, 5, 11, 23, 26, 29, 59). The ability of flavonoids to protect cardiac cells has been classically ascribed to their antioxidant activity as ROS scavengers. Recently, it has been demonstrated that flavonoids are also able to modulate important signaling pathways activated by ROS in cardiac myocytes, namely the mitogen-activated protein kinase cascades and the phosphoinositide 3-kinase/protein kinase B (Akt) pathway (5, 55, 59). One of the major representatives of flavonoids is Q, found in many edible plants. In Western populations, the intake has been estimated as 18 mg/day (41). The potential protective effects of Q have been documented in various studies (5, 36). However, the exact mechanisms underlying Q protective effects in cardiac cells remain to be elucidated. To better clarify these aspects, we have evaluated the ability of Q to modulate the cardiomyocyte global gene expression. Microarray technology was used to determine the expression levels of 22,000 genes after Q treatment. Since dietary components can also modify the translation of RNA to proteins and the posttranslational events, which can affect protein activity, we have also evaluated the activity of the enzymes coded by the selected genes. Moreover, we have evaluated the existence of a time-dependent relationship between induced enzyme activities and the ability of Q to protect cells from H2O2-elicited damage.

The results of this study demonstrated for the first time the predominant induction of antioxidant/detoxifying genes in cardiomyocytes after Q treatment. In particular, a group of cellular antioxidant/phase 2 enzymes was significantly upregulated, including GSTa3, GSTp2, Mgst1, NQO1, TR, HO-1, and glutamate-cysteine ligase. Interestingly, the hierarchical cluster analysis obtained with the microarray data revealed that all of these genes were classified in the same cluster, evidencing a similar time-dependent regulation pattern of expression and highlighting a possible common functional regulation. NQO1, HO, TR, and GST activities and GSH content are critically involved in the regulation of cellular reduction/oxidation status and have been suggested to play important roles in protecting against oxidative cardiovascular pathophysiology (16, 23, 46, 60). Surprisingly, Q treatment significantly downregulated metallothionein 1a (Mt1a) and metallothionein 2 (MT-2) gene expressions after 12 and 24 h. Other authors demonstrated that Mt1a and MT-2 mRNA levels were enhanced by Q treatment in different cell systems. These discrepancies are due to the use of DMSO as vehicle for Q. DMSO is a potent inducer of Mt1a and MT-2 mRNA transcription, as reported by Conklin et al. (10), who also demonstrated that MT levels in DMSO plus Q-treated cells were indistinguishable from levels in their respective controls. As we performed competitive microarrays in which control and treated cells are hybridized on the same microarray, the observed downregulation of Mt1a and MT-2 was only apparent. From the data reported in this paper, it is not possible to speculate about the ability of Q to modulate MT gene expression in the heart. Further studies, specifically targeted to flavonoid modulation of MT expression levels, are necessary to clarify this aspect.

Although it has been previously demonstrated that Q is capable of inducing several genes encoding for endogenous antioxidants and phase 2 enzymes in transformed cells (31, 32), the inducibility of these important antioxidant-related genes has not yet been reported in normal cells.

Regulation of the cellular reduction/oxidation (redox) balance is critically determined by several antioxidant systems, and its impairment alters multiple cell pathways. The ubiquitously expressed thiol-reducing systems include the TRX, glutaredoxin, and GSH systems (35, 39). TRX is a small and ubiquitously expressed thiol-disulfide oxidoreductase and functions as an important redox regulator in cells from Escherichia coli to mammals (18, 34). The reduction of the catalytic center in TRX is catalyzed by TR, using NADPH as a cofactor. TRX functions as a scavenger for ROS at the cellular level (2). In addition, TRX interacts with various proteins in a redox-dependent manner and modulates intracellular signaling pathways and transcription factor activity (44, 62). Turoczi et al. (51) showed that TRX-overexpressing mouse hearts had improved postischemic ventricular recovery and reduced myocardial infarct size compared with normal hearts. It has been demonstrated that TRX overexpression in adult rat cardiomyocytes prevents α-adrenergic receptor-stimulated hypertrophy (22), suggesting a protective role of endogenous TRX in failing heart. Our results evidenced a marked upregulation of TR after Q treatment, contributing to explain the ability of this flavonoid to protect cardiomyocytes from oxidative stress.

GSH is the major nonenzymatic regulator of intracellular redox homeostasis, ubiquitously present in all cell types at a millimolar concentration (27). It is synthesized enzymatically by glutamate-cysteine ligase and GSH synthetase, with the former being the rate-limiting enzyme (24). GSH exists in cell, in both reduced (GSH) and oxidized (GSSG) forms. Under normal cellular redox conditions, the major portion is in reduced form. Our data showed both a significant upregulation of glutamate-cysteine ligase expression level and a marked increase of GSH content after Q treatment, indicating a clear role of Q in increasing thiol-related compound levels. Thus our results strongly suggest that the elevation of GSH by Q is probably due to an induction of glutamate-cysteine ligase and not of GSH reductase, the enzyme responsible for the production of GSH from GSSG. In fact, we have not evidenced any significant upregulation of GSH reductase after Q treatment. Accordingly, Myhrstad et al. (33) have shown that Q is able to increase the intracellular concentration of GSH by ∼50% in COS-1 cells. They also demonstrated that Q increased the expression of both the regulatory and the catalytic subunit of glutamate-cysteine ligase. In a recent study, Q and other flavonoids were found to efficiently protect neuronal cells from oxidative glutamate toxicity and other forms of oxidative injury (20). Interestingly, the increased production of GSH was one of three mechanisms suggested by the authors for the protective effects of Q (20).

GSH is also a cofactor for GST, an abundant cellular enzyme in mammalian tissues. GST is generally viewed as a phase 2 enzyme, primarily involved in the detoxification of electrophilic xenobiotics through the formation of GSH-electrophile conjugates (17, 48). Recently, several studies have also reported that GST plays an important role in protecting cells against ROS-mediated injury, catalyzing the decomposition of lipid hydroperoxides generated from oxidative damage of cellular lipids (61, 63).

In this study, we have demonstrated the induction of GST by Q in cardiomyocytes, which may contribute to the increased cell resistance to H2O2-elicited toxicity.

The observed strong induction of NQO1 by Q may also be involved in the cytoprotective effects on H2O2-induced cytotoxicity. In this context, NQO1 may act as an antioxidant enzyme via its ability to maintain the cellular levels of ubiquinol and vitamin E, two important nonprotein antioxidants (42).

HO catalyzes the rate-limiting step in the heme degradation, yielding to biliverdin, carbon monoxide, and free iron. Biliverdin is subsequently reduced to bilirubin by biliverdin reductase. Three isoforms of HO have been characterized: the inducible HO-1, the constitutive HO-2, and the recently cloned HO-3, which is only marginally active in heme degradation (43). In this study, we demonstrated the ability of Q to induce HO-1 gene expression and to increase HO activity. HO-1 is also one of the heat shock proteins (HSP32) and is highly upregulated by various factors causing oxidative stress, such as heat shock, cytokines, hypoxia, and ischemia-reperfusion (25, 49, 56). The induction of HO-1 is considered beneficial, as it may protect against apoptotic cell death in fibroblasts and endothelial cells (8, 12, 38). Moreover, HO-1-deficient mice developed larger myocardial infarcts in response to hypoxia, whereas cardiac-specific HO-1 overexpression has been shown to protect against ischemia-reperfusion injury (64, 65). Clark et al. (9) demonstrated that HO-1 upregulation before ischemia ameliorates myocardial function and reduces infarct size on reperfusion of isolated rat hearts.

Previous studies have demonstrated that Q is able to protect H9c2 cardiomyoblasts against oxidative stress, not only by a direct antioxidant mechanism, but also through the modulation of prosurvival protein kinases, such as Akt/PKB and ERK1/2 (5). The data here presented complete these findings, showing also the enhancement of antioxidant/detoxifying enzyme activities by Q.

Considering the ability of Q to increase GST, NQO1, TR, and HO activities and GSH content, important cellular defenses against oxidative stress, we investigated whether their induction after Q treatment leads to protection against ROS-mediate cytotoxicity. The results clearly showed that treatment of cardiomyocytes with Q resulted in a complete protection against H2O2-mediated cytotoxicity and intracellular accumulation of ROS already at the shortest treatment time.

The protective role of Q against H2O2 toxicity at short exposure times could be ascribed to its well-known antioxidant activity. The ability to decrease ROS levels and maintain cell viability also at longer times could be due to both modulation of intracellular signaling pathways, as previously demonstrated (5), and regulation of some important phase 2 enzymes, evidencing its indirect antioxidant activity. These results suggest that cardiac cells exposed to Q through diet may be protected against oxidative stress, having higher levels of phase 2 detoxifying enzymes. In conclusion, cardiomyocytes could be better prepared to subsequent toxic insults after Q nutritional intake. In particular, Q intake may have a role in the counteraction and prevention of cardiac stress related to diseases.

GRANTS

This study was supported by Ministry of Education, University and Research (Italy), and Fondazione del Monte di Bologna e Ravenna (Italy).

Footnotes

  • 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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