Role of endoplasmic reticulum oxidase 1α in H9C2 cardiomyocytes following hypoxia/reoxygenation injury
- Authors:
- Published online on: June 9, 2020 https://doi.org/10.3892/mmr.2020.11217
- Pages: 1420-1428
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Copyright: © Lai et al. This is an open access article distributed under the terms of Creative Commons Attribution License.
Abstract
Introduction
Prompt reperfusion is essential for recovery following acute myocardial infarction; however, reperfusion can also lead to ischemia/reperfusion (I/R) injury (1). Although myocardial I/R injury involves complex pathophysiological mechanisms that have not yet been fully elucidated, oxidative stress and intracellular Ca2+ overload are considered to be two of the main mechanisms contributing to myocardial I/R injury (2–4). Increases in reactive oxygen species (ROS) and intracellular Ca2+ levels are described as mutually causative, forming a vicious cycle (5). Intracellular Ca2+ homeostasis and ROS production are closely related to the endoplasmic reticulum (ER) and mitochondria. When faced with oxidative stress, ischemia, hypoxia, Ca2+ imbalance and other conditions, unfolded proteins accumulate in the (ER), and upon exceeding its capacity to deal with unfolded proteins, ER homeostasis is lost and the ER stress (ERS) response is activated (6). Accumulating studies have reported that ERS serves an important role in myocardial I/R injury (7,8).
The ER is associated with the mitochondria at multiple levels through mitochondrial-associated membranes (MAMs), which are specific protein-rich regions of the ER located in close proximity to the mitochondria. MAMs regulate several functions, including the synthesis and transport of phospholipids, Ca2+ transfer between organelles and cell signaling pathways (9–11). ER oxidase 1 (ERO1) is a glycosylated flavonase located at MAMs, of which there are two mammalian isoforms; the ERO1α isotype, which is found distributed throughout the body, and the ERO1β isotype, which is most abundant in pancreatic β cells and lymphocytes (12,13). Both isoforms respond to ERS, but only ERO1α is induced by hypoxia (14,15). ERO1α serves an important role in catalyzing the formation of protein disulfide bonds, ER redox and Ca2+ homeostasis (16), and the ERO1α-dependent ER-mitochondrial calcium flux has been observed to contribute to ERS (17). However, the exact role of ERO1α in myocardial I/R injury remains unclear.
In the present study, myocardial I/R injury was simulated using myocardial hypoxia/reoxygenation (H/R). The effects of ERO1α on myocardial H/R were observed by genetically knocking down the expression of ERO1α with short hairpin RNA (shRNA), and treatment with the ERS activator, TM or the ERS inhibitor, 4-Phenylbutyric acid (4-PBA).
Materials and methods
Lentiviral cell transfection
Using the ERO1α gene mRNA sequence (GenBank accession no. NM_138528.1), three shRNA candidate target sequences (1832, 1833 and 1834) were designed and synthesized by Shanghai GeneChem Co., Ltd. The above target sequences were cloned into the lentiviral vector pMAGic4.1 and scrambled shRNA was cloned into the pMAGic4.1 vector as the negative control. H9C2 cells (American Type Culture Collection) were plated in 6-well plates at a density of 1×106 cells/well, and upon reaching 70% confluence, the cells were transfected with either recombinant lentivirus ERO1α-shRNA (Table I) or scrambled shRNA with titers of 5×106 TU/ml. Following transfection for 48 h, the efficiency of ERO1α silencing was assessed using reverse transcription-quantitative PCR and western blotting. The shRNA with the best silencing effect was selected for subsequent experiments.
Exposure of H9C2 cardiomyocytes to H/R and treatments
H9C2 cardiomyocytes were purchased from the American Type Culture Collection and the H/R model was established using the AnaeroPack® method (18). Briefly, a hypoxic atmosphere was created by incubating an AnaeroPack® (Mitsubishi Gas Chemical Company, Inc.), which absorbs oxygen, and H9C2 cardiomyocytes in a sealed airtight container together at 37°C. Following incubation for 3 h, the AnaeroPack® container was opened to terminate the hypoxic conditions. The cells in the culture plates were removed and subsequently placed in a CO2 incubator at 37°C for 6 h. The morphology was observed under an inverted light microscope at ×200 magnification. In the 4-PBA + H/R group, H9C2 cardiomyocytes were treated with 0.5 mM 4-PBA (Sigma-Aldrich; Merck KGaA), a selective ERS inhibitor, 2 h prior to H/R induction. In the TM + ERO1α-shRNA + H/R, cardiomyocytes were pretreated with 2 µg/l TM (Cayman Chemical), an ERS activator, 24 h prior to H/R exposure to counteract the reduction in ERS following ERO1α knockdown.
Morphological analysis following Hoechst 33258 staining
A total of 1×105 H9C2 cardiomyocytes/well were incubated in 24-well plates. Cells were fixed by 4% paraformaldehyde at 4°C for 1 h, washed twice with PBS and stained with Hoechst 33258 (Beyotime Institute of Biotechnology) at room temperature for 5 min. Hoechst 33258 staining solution was subsequently aspirated and washed twice with PBS for 5 min each. Stained cell nuclei were visualized using an IX70 fluorescence microscope (Olympus Corporation) (magnification, ×200). A total of five fields were randomly selected from each well, and the proportion of apoptotic cells was calculated as the ratio of nuclear pyknosis cells to the total cells.
Flow cytometric analysis of apoptosis
The Annexin V-FITC Apoptosis Assay kit (Beyotime Institute of Biotechnology) was used to detect cell apoptosis. Briefly, following treatment, cells were transferred to individual tubes and a solution containing 195 µl binding buffer, 5 µl Annexin V-FITC and 10 µl propidium iodide was added to each test tube. The tubes were subsequently mixed and incubated at room temperature in the dark for 15 min. Apoptotic cells were detected using a FACSScan flow cytometer (FACSVerse; BD Biosciences). The data analysis was conducted using CellQuest software version 3.3 (BD Biosciences).
Detection of caspase-3 activity
Caspase-3 activity was measured using a caspase-3 activity kit (Beyotime Institute of Biotechnology). Briefly, cells were suspended in lysis buffer on ice for 15 min and subsequently centrifuged at 16,000 × g for 10 min. The supernatants were then incubated with 20 ng Ac-DEVD-pNA in a 96-well plate for 2 h at 37°C (19). The absorbance of pNA was measured at 405 nm using an Infinite® 200 PRO microplate reader (Tecan Group, Ltd.).
Measurement of intracellular Ca2+ levels
Cells were washed twice with PBS buffer and then incubated with Fluo-3/AM working fluid for 30 min in the dark at 37°C. The fluorescence intensity was determined using a confocal laser scanning microscope (TCS SP5; Leica Microsystems GmbH), with an excitation wavelength of 488 nm and an emission wavelength of 525 nm (magnification, ×400). A total of five fields were randomly selected from each dish. Semi-quantitative analysis was conducted using ImageJ 1.49 software (National Institutes of Health).
Measurement of intracellular ROS levels
Intracellular ROS levels were detected by ROS fluorescent probe dichlorodihydrofluorescein diacetate (DCFH-DA). Briefly, 1×106 H9C2 cardiomyocytes/ml were incubated with 10 µM DCFH-DA at 37°C for 30 min and then washed three times with PBS buffer. The fluorescent intensity was detected with an excitation wavelength of 488 nm and an emission wavelength of 525 nm using an Infinite® 200 PRO microplate reader (Tecan Group, Ltd.). The results represent the percentage variation relative to the untreated control.
Extraction of the cytoplasmic and mitochondrial components
The cytoplasmic and mitochondrial proteins were extracted using a cell mitochondria isolation kit (Beyotime Institute of Biotechnology). Briefly, cells were incubated in lysis buffer at 4°C and centrifuged at 3,000 × g for 10 min. Subsequently, the supernatants were centrifuged for a second time at 4°C for 10 min at 13,000 × g. The cytoplasmic components were present in the supernatant, whereas the cell pellet contained the mitochondrial components.
Measurement of mitochondrial membrane potential (Δψm)
A total of 100 µl (1 mg/ml) purified mitochondria was added to 900 µl JC-1 staining solution and subsequently incubated at 37°C for 30 min. To detect the JC-1 monomer, the excitation and emission wavelength was set to 490 and 530 nm, respectively. To detect the JC-1 polymer, the excitation and emission wavelength was set to 525 and 590 nm, respectively (20). The fluorescent intensity was determined using an Infinite® 200 PRO microplate reader (Tecan Group, Ltd.).
Reverse transcription-quantitative PCR (RT-qPCR)
Total RNA was extracted using TRIzol® reagent (Takara Bio, Inc.). Total RNA was reverse transcribed into cDNA using the PrimeScript RT reagent kit (Takara Bio, Inc.). qPCR was subsequently performed using the SYBR® Premix Ex Taq™ kit (Takara Bio, Inc.), according to the manufacturer's protocol, and a qTower2.2 quantitative PCR instrument (21). Primer sequences of the genes used for the qPCR are presented in Table II. Relative expression of genes was calculated using the comparative cycle threshold (Ct) (2−ΔΔCt) method with 18s RNA as the internal control (22).
Western blotting
Total cell proteins were extracted using RIPA Lysis Buffer (Beyotime Institute of Biotechnology). Mitochondria proteins were extracted using the Cell Mitochondria Isolation kit (Beyotime Institute of Biotechnology) according to the manufacturer's protocol. Protein concentration was determined via BCA protein assay kit (Beyotime Institute of Biotechnology). A total of 50 µg of extracted protein were electroblotted onto a PVDF membrane following separation on a 10% SDS-PAGE. The membranes were blocked with 5% non-fat milk for 1 h at room temperature prior to being incubated with the following primary antibodies overnight at 4°C: Anti-ERO1α (1:1,000; cat. no. sc-365526) purchased from Santa Cruz Technology, and anti-78 kDa glucose-regulated protein (GRP78; 1:2,000; cat. no. ab108615), anti-C/EBP homologous protein (CHOP; 1:2,000; cat. no. ab179823), anti-caspase-12 (1:2,000; cat. no. ab62484), anti-cytochrome c (1:2,000; cat. no. ab133504) anti-cytochrome c oxidase subunit IV (COX IV; 1:2,000; cat. no. ab153709) and anti-GAPDH (1:5,000; cat. no. ab181602) purchased from Abcam. Following the primary antibody incubation, membranes were washed three times with wash buffer and incubated with horseradish peroxidase-conjugated IgG secondary antibodies (1:2,000; cat. nos. A0208 and A0216, respectively; Beyotime Institute of Biotechnology) for 2 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence system (Beyotime Institute of Biotechnology), with GAPDH as the loading control (COX IV was used as a mitochondrial loading control). Semi-quantitative analysis was conducted using ImageJ 1.49 software (National Institutes of Health).
Statistical analysis
Statistical analysis was performed using SPSS version 19.0 software (IBM Corp.) and data are presented as the mean ± SD. To determine statistical differences amongst >2 groups, one-way ANOVA was used, followed by Tukey's post hoc test for multiple group comparisons. P<0.05 was considered to indicate a statistically significant difference.
Results
Expression levels of ERO1α protein increase following H/R induction
H9C2 cardiomyocytes were exposed to hypoxia for 3 h and reoxygenation for 1, 3, 6 and 12 h, and ERO1α protein expression levels were subsequently assessed using western blotting. In the control (CON) group, H9C2 cardiomyocytes did not undergo hypoxia and reoxygenation. The cells in the CON group grew well and most of them were fusiform. Compared with the CON group, hypoxia-reoxygenation caused some cells to appear irregular or round in shape, and floating cells and cell debris increased significantly (Fig. 1A). ERO1α protein expression levels significantly increased following H/R induction compared with the CON group (Fig. 1B); and ERO1α expression levels reached their highest in H9C2 cardiomyocytes following 3 h of hypoxia and 6 h of reoxygenation. In addition, ERS marker GRP78 protein expression levels peaked following 3 h of reoxygenation. Therefore, subsequent experiments in the present study were performed in cells following 3 h of hypoxia and 6 h of reoxygenation.
Consistent with ERO1α expression, H/R-stimulated H9C2 cardiomyocytes demonstrated increased the proportion of apoptotic cells. The nuclei of H9C2 cardiomyocytes in the CON group were round and homogeneously stained, whereas those in the H/R group were observed to have significant apoptotic characteristics, such as cell shrinkage and chromatin condensation (Fig. 1C). Counting the cells with apoptotic characteristics found that the percentage of apoptotic cells in the H/R group was significantly increased compared with the CON group.
Transfection with specific ERO1α-shRNAs decreases ERO1α expression levels in H9C2 cardiomyocytes
The role of ERO1α following H/R was chosen for further investigation because previous studies had reported that ERS served an important role in myocardial I/R injuries (23,24). ERO1α-shRNA carrier vectors were observed to contain the expected sequence (Table SI), indicating their successful construction. Following transfection of H9C2 cardiomyocytes with three different ERO1α-shRNAs (1832, 1833 and 1834), the mRNA expression levels of ERO1α were significantly decreased in the H9C2 cardiomyocytes (Fig. 2A). Although all three lentiviral shRNAs significantly inhibited ERO1α protein expression (Fig. 2B), the knockdown efficiency of shRNA 1832 was more effective compared with shRNA 1833 and shRNA 1834. Therefore, the lentivirus shRNA 1832 was selected for use in subsequent experiments.
ERO1α knockdown decreases H/R-induced ERS
The mRNA and protein expression levels of ERS-related molecules, including ERO1α, GRP78 and CHOP were detected using RT-qPCR and western blotting. The expression levels of ERO1α, GRP78 and CHOP mRNA and protein significantly increased following H/R compared with the CON group, whereas 4-PBA inhibited the expression levels of ERO1α, GRP78 and CHOP protein (Fig. 3A). Furthermore, ERO1α knockdown reversed the H/R-induced increase in mRNA and protein expression levels of GRP78 and CHOP (Fig. 3B and C). Together, these data strongly suggested that ERS may serve an important role in myocardial H/R injury, and that ERO1α may have a crucial role in H/R-induced ERS in H9C2 cardiomyocytes.
ERO1α knockdown inhibits H/R-induced cell apoptosis
The percentage of apoptotic cells, caspase-3 activity and the expression levels of caspase-12 and cleaved-caspase-12 (c-caspase-12) were investigated to determine the role of ERO1α in H/R injury. H9C2 cardiomyocytes underwent hypoxia for 3 h, then reoxygenation for 6 h, and the percentage of apoptotic cells was evaluated using flow cytometry. The percentage of apoptotic cells in the H/R group was significantly increased compared with the CON group (Fig. 4A); however, the percentage of apoptotic cells of the ERO1α-shRNA + H/R group was decreased compared with the H/R group.
Apoptosis requires the activation of cysteine protease, which both promotes and executes cell death (25). Excessive ERS was reported to induce apoptotic signaling in I/R injury (26). Thus, the expression levels of caspase-12 and c-caspase-12 proteins, and caspase-3 activity were subsequently investigated (Fig. 4B and C). The expression levels of c-caspase-12 in the H/R group were significantly increased compared with the CON group; however, ERO1α knockdown reduced c-caspase-12 expression compared with the H/R group. These data suggested that ERO1α knockdown may inhibit H/R-induced apoptosis in H9C2 cardiomyocytes.
ERO1α knockdown decreases H/R-induced increases in intracellular ROS and Ca2+ levels
Under stress conditions, increases in intracellular Ca2+ levels promote apoptosis (27), thus the intracellular Ca2+ influx in response to H/R was measured. Intracellular Ca2+ levels in H9C2 cardiomyocytes were markedly increased in the H/R group, whereas ERO1α knockdown decreased the intracellular Ca2+ levels (Fig. 5A and B). Furthermore, oxidative stress is related to apoptosis-induced H/R injury (28), thus intracellular ROS levels were analyzed to investigate the association between ERO1α and oxidative stress. ERO1α knockdown significantly decreased intracellular ROS levels in the H/R group (Fig. 5C).
ERO1α knockdown alleviates H/R-induced mitochondrial dysfunction through reducing ERS
To investigate whether ERO1α knockdown protected mitochondrial function in H/R injury, cardiomyocytes were pretreated with 2 µg/l TM, an ERS activator, 24 h prior to H/R exposure to counteract the reduction in ERS following ERO1α knockdown. Intracellular ROS levels, Δψm, and expression levels of cytochrome c in mitochondrial and cytosolic fractions were evaluated. TM pretreatment of ERO1α-shRNA-transfected H9C2 cardiomyocytes undergoing H/R injury did not reverse the reduction in intracellular ROS levels observed following ERO1α knockdown (Fig. 5C). Compared with the H/R group, decreased expression levels of cytochrome c were observed in the cytoplasm in the ERO1α-shRNA + H/R group, whereas increased levels were found in the mitochondria. However, no significant differences were observed between the TM + ERO1α-shRNA + H/R and the ERO1α-shRNA + H/R groups (Fig. 6A). After H/R, the Δψm significantly decreased, while ERO1α knockdown had a trend towards an increase in Δψm in the ERO1α-shRNA + H/R group; however, there was no significant difference when comparing the H/R and ERO1α-shRNA + H/R groups (Fig. 6B). Taken together, these findings suggested that ERO1α may be responsible for the H/R-induced mitochondrial damage of H9C2 cardiomyocytes, which is associated with ERS.
Discussion
In mammalian cells, the ER has an important role in the proper folding and assembly of polypeptide chains, Ca2+ storage and post-translational modifications (29). ERS occurs following I/R-induced increases in ROS, which results in the accumulation of misfolded or unfolded proteins in the ER lumen (30). Thus, there is increasing evidence to suggest that ERS is associated with I/R injury (31–33).
ERO1α is a conserved glycoprotein, which has been demonstrated to accept electrons from reduced protein disulfide isomerase and transfer them to oxygen molecules, catalyze the formation of oxygen-mediated protein disulfide bonds and contribute to ERS (34,35). ERO1α activity may be an important factor contributing to the large production of ROS in cells, as it has been previously found that ERO1α dysfunction may result in a rapid decrease in ER-derived oxidative stress (16). However, in homocysteine-induced ERS, ERO1α demonstrated a negative regulatory effect (36). In the present study, ERO1α expression levels in H9C2 cardiomyocytes were confirmed using western blotting, and H/R induction was subsequently used to simulate I/R in these cells to further verify whether ERO1α expression was altered as a result of oxidative stress. ERO1α expression levels in H9C2 cardiomyocytes were markedly increased following H/R, reaching their highest levels following 6 h of reoxygenation, whereas 4-PBA decreased the expression levels. In addition, the number of apoptotic cells was significantly increased following H/R induction. Therefore, it was hypothesized that ERO1α may serve an important role in H/R development.
To further understand ERO1α function in H9C2 cardiomyocytes following H/R, lentiviral shRNA was used to reduce ERO1α expression levels. In previous studies, it has been reported that following myocardial I/R, ERS promoted apoptosis in cardiomyocytes through the CHOP and caspase-12 signaling pathways (37); that c-caspase-12 expression levels, and caspase-3 activity were increased following H/R (38,39); and that caspase-12, which indirectly activates cytoplasmic caspase-3, was considered to be a crucial mediator of ERS-induced apoptosis (40). Consistent with these studies, the data from the present study revealed that the expression levels of GRP78, CHOP and c-caspase-12, as well as caspase-3 activity, were significantly increased. However, following the transfection of H9C2 cardiomyocytes with ERO1α-shRNA, the expression levels and activity significantly decreased. These results indicated that ERO1α may have an important role in ERS, and that the downregulation of ERO1α may decrease ERS and apoptosis in myocardial cells following H/R injury.
The ER is the main storage location of Ca2+ in cells and participates in dynamic Ca2+ exchange with the mitochondria (41); Ca2+ is transferred to the mitochondrial matrix to stimulate mitochondrial ATP synthesis by activating the tricarboxylic acid cycle. During I/R, the increase in intracellular ROS levels and oxidative stress from multiple different sources leads to a large amount of Ca2+ dissociating from the ER to the mitochondria (42). This subsequently promotes mitochondrial Ca2+ overload, which triggers cell apoptosis by opening the mitochondrial permeability transition pore (43). Oxidative stress promotes ERS, and persistent ERS has been observed to promote mitochondrial dysfunction, which in turn induces oxidative stress (44). As previously mentioned, ERO1α is strongly associated with the generation of ROS, and it has been demonstrated that due to excessive oxidation, ERO1α enhances inositol triphosphate receptor (IP3R) activity, and promotes IP3R-mediated Ca2+ release and transfer to the mitochondria, facilitating apoptosis (45,46). Thus, further investigation into the relationship between ERO1α and mitochondrial function following H/R is required.
The Δψm reflects the functional status of the mitochondria. In the present study, it was observed that oxidative stress led to a decrease in Δψm, accompanied by significantly increased ERO1α expression levels. Compared with the H/R group, following ERO1α-shRNA transfection, the concentration of Ca2+ decreased, while ERO1α knockdown had a trend towards an increase in Δψm. ERO1α knockdown reduced the release of cytochrome c from the mitochondria to the cytoplasm. However, the pretreatment of ERO1α-transfected H9C2 cardiomyocytes with TM following H/R injury did not reverse the reduced intracellular ROS levels, the ratio of cytoplasmic/mitochondrial cytochrome c achieved by ERO1α knockdown. These results suggested that the downregulation of ERO1α may attenuate oxidative stress, and decrease the intracellular Ca2+ concentration and the percentage of apoptotic cells in H9C2 cardiomyocytes following H/R.
In conclusion, the findings in the present study indicated that ERO1α may serve as a positive mediator of the apoptotic pathway during H/R; however, the exact mechanism by which ERO1α achieves this function remains to be investigated. In addition, the effects of ERO1α on the H9C2 cardiomyocytes in vitro may not reflect the in vivo scenario; thus, future studies should encompass animal models for further validation of these results.
Supplementary Material
Supporting Data
Acknowledgements
Not applicable.
Funding
This study was supported by the Science and Technology Innovation Team Project of Changzhi Medical College (grant no. CX201409).
Availability of data and materials
The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.
Authors' contributions
LNL and XJZ conceived and designed the experiments; YL, YYL, SLC and NZ performed the experiments; and DW and LNL were involved in analyzing the data and drafting the manuscript. All authors read and approved the final manuscript.
Ethics approval and consent to participate
Not applicable.
Patient consent for publication
Not applicable.
Competing interests
The authors declare that they have no competing interests.
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