In vitro Assessment of Myocardial Protection following Hypothermia-Preconditioning in a Human Cardiac Myocytes Model

Summary

The distinct effects of different degrees of hypothermia on myocardial protection have not been thoroughly evaluated. The goal of the present study was to quantify the levels of cell death following different hypothermia treatments in a human cardiomyocyte-based model, laying the foundation for future in-depth molecular research.

Abstract

Ischemia/reperfusion-derived myocardial dysfunction is a common clinical scenario in patients after cardiac surgery. In particular, the sensitivity of cardiomyocytes to ischemic injury is higher than that of other cell populations. At present, hypothermia affords considerable protection against an expected ischemic insult. However, investigations into complex hypothermia-induced molecular changes remain limited. Therefore, it is essential to identify a culture condition similar to in vivo conditions that can induce damage similar to that observed in the clinical condition in a reproducible manner. To mimic ischemia-like conditions in vitro, the cells in these models were treated by oxygen/glucose deprivation (OGD). In addition, we applied a standard time-temperature protocol used during cardiac surgery. Furthermore, we propose an approach to use a simple but comprehensive method for the quantitative analysis of myocardial injury. Apoptosis and expression levels of apoptosis-associated proteins were assessed by flow cytometry and using an ELISA kit. In this model, we tested a hypothesis regarding the effects of different temperature conditions on cardiomyocyte apoptosis in vitro. The reliability of this model depends on strict temperature control, controllable experimental procedures, and stable experimental results. Additionally, this model can be used to study the molecular mechanism of hypothermic cardioprotection, which may have important implications for the development of complementary therapies for use with hypothermia.

Introduction

Ischemia/reperfusion-derived myocardial dysfunction is a common clinical scenario in patients after cardiac surgery^1,2. During nonpulsatile low flow perfusion and periods of total circulatory arrest, damage involving all types of heart cells still occurs. In particular, the sensitivity of cardiomyocytes to ischemic injury is higher than that of other cell populations. At present, therapeutic hypothermia (TH) affords substantial protection against an expected ischemic insult in patients undergoing cardiac surgery^3,4. TH is defined as a core body temperature of 14-34 °C, although no consensus exists regarding a definition of cooling during cardiac surgery^5,6,7. In 2013, an international panel of experts proposed a standardized reporting system to classify various temperature ranges of systemic hypothermic circulatory arrest⁸. Based on electroencephalography and metabolism studies of the brain, they divided hypothermia into four levels: profound hypothermia (≤ 14 °C), deep hypothermia (14.1-20 °C), moderate hypothermia (20.1-28 °C), and mild hypothermia (28.1-34 °C). The expert consensus provided a clear and uniform classification, allowing studies to be more comparable and provide more clinically relevant outcomes. This protection afforded by TH is based on its capacity to reduce the metabolic activity of cells, further limiting their rate of high-energy phosphates consumption^9,10. However, the role of TH in myocardial protection is controversial and may have multiple effects depending on the degree of hypothermia.

Myocardial I/R is well known to be accompanied by increased cell apoptisis¹¹. Recent reports have observed that programmed cardiomyocyte death increases during open-heart surgery, and may coincide with necrosis, thereby increasing the number of dead myocardial cells¹². Therefore, reducing cardiomyocyte apoptosis is a useful therapeutic approach in clinical practice. In the mouse atrial HL-1 cardiomyocyte model, therapeutic hypothermia was shown to reduce the mitochondrial release of cytochrome c and apoptosis-inducing factor (AIF) during reperfusion¹³. However, the effect of temperature in regulating apoptosis is controversial and appears to depend on the degree of hypothermia. Cooper and colleagues observed that compared to a normothermic cardiopulmonary bypass control group, the apoptosis rate of myocardial tissue from pigs with the deep hypothermic circulatory arrest was increased¹⁴. In addition, the results of some studies have suggested that deep hypothermia may activate the apoptosis pathway, while less aggressive hypothermia appears to inhibit the pathway^12,15,16. The reason for this result may be due to confounding effects associated with ischemic injury and a lack of understanding of the mechanisms by which temperature affects myocardial tissue. Therefore, the temperature limits at which apoptosis is enhanced or attenuated should be accurately defined.

To gain a better understanding of the mechanisms associated with the efficacy of hypothermia and provide a rational basis for its implementation in humans, it is essential to identify a culture condition similar to in vivo conditions that can produce damage similar to that observed for the clinical condition in a reproducible manner. An essential step towards achieving this goal is to establish the optimal conditions for inducing cardiomyocyte apoptosis. Accordingly, in the present study, we explored the methodological details regarding oxygen-glucose deprivation experiments with cultured cells, a facile in vitro model of ischemia-reperfusion. Furthermore, we evaluated the effect of different hypoxic-ischemic times on cardiomyocyte apoptosis, and verified our hypothesis regarding the effect of different temperature conditions on cell apoptosis in vitro.

Protocol

Information regarding commercial reagents and instruments are listed in the Table of Materials.

The AC16 human cardiomyocyte cell line was derived from the fusion of primary cells from adult ventricular heart tissue with SV40-transformed human fibroblasts¹⁷, which were purchased from BLUEFBIO (Shanghai, China). The cell line develops many biochemical and morphological features characteristic of cardiomyocytes. In addition, the cell line is widely used to evaluate myocardial damage and myocardial function in vitro^18,19.

1. Cell culture

NOTE: The basal culture medium consists of serum-free Dulbecco’s modified Eagle’s medium (DMEM), 10% fetal bovine serum (FBS), 1% cardiac myocyte growth supplement, and 1% penicillin/streptomycin solution. Store the medium at 4 °C and prewarm to 37 °C before use.

1. Remove the cryopreserved human cardiomyocyte (HCM) cells from liquid nitrogen and thaw them in a water bath at 37 °C.
2. Gently shake the vial (<1 minute) in a 37 °C water bath until only a small piece of ice remains in the vial.
3. Transfer the vial into a sterile laminar flow hood. Wipe the outside of the vial with a cotton ball dipped in 70% alcohol.
4. Transfer 4 mL of prewarmed complete growth medium dropwise into the centrifuge tube containing the thawed cells.
5. Centrifuge the cell suspension at 200 × g for 10 minutes. After centrifugation, discard the supernatant and resuspend the pellet in 5 mL of complete medium.
6. Maintain the cells at 37 °C in a humidified incubator under an atmosphere with 95% air and 5% CO₂.
   NOTE: Before harvesting the cells for experiments, the cells are allowed to grow until reaching approximately 60-70% confluency.

2. Establishment of an oxygen-glucose deprivation (OGD) model

NOTE: Two hours before the study period, replace the growth medium with serum-free medium, and the cells were reincubated in a humidified incubator for 2 h at 37 °C under an atmosphere with 5% CO₂.

1. Aspirate the medium from a 6-well plate and gently wash the cells three times with phosphate buffered saline (PBS).
2. Add 2 mL of fresh sugar-free DMEM per well.
3. Culture the cells at a constant temperature in a three-gas incubator under an atmosphere with a mixture of 95% N₂, 5% CO₂, and 0.1% O₂ at 37 °C for 1, 2, 4, 8, 12, or 16 h.
4. After the hypoxia treatment, aspirate the medium from the 6-well plate and wash the cells with PBS (pH 7.4) three times.
5. Add 2 mL of complete DMEM per well.
6. Maintain the cells at 37 °C in a humidified incubator under an atmosphere with 95% air and 5% CO₂.

3. Time-temperature protocol

NOTE: A standard time-temperature protocol is used during cardiac surgery, as described previously by others^20,21. Treat HCMs according to the following protocol (Figure 1): timepoint 1 (T1) indicates the end of induction, timepoint 2 (T2) indicates the end of maintenance and timepoint 3 (T3) indicates the end of rewarming. Analyze control cells maintained under continuous normothermic conditions (37 °C). The temperature conditions are created using a tri-gas incubator, which allows precise temperature regulation.

1. Two hours before the experiments, aspirate the culture medium from a 6-well plate and add 2 mL of fresh serum-free DMEM per well.
2. Re-incubate the cells in a humidified incubator for 2 h at 37 °C under an atmosphere with 5% CO₂.
3. After 2 h, aspirate the medium from the 6-well plate and wash the cells with PBS (pH 7.4) three times.
4. Add 2 mL of fresh serum-free DMEM per well.
5. Reincubate the cells in the tri-gas incubator.
   NOTE: Replace the medium to remove unattached cells and debris.
6. Immediately change the temperature by placing the culture dishes a tri-gas incubator.
7. After 1 h of cooling, quickly aspirate the medium from the 6-well plate and add 2 mL of fresh sugar-free DMEM per well.
8. Culture the cells in a tri-gas incubator under an atmosphere comprising 95% N₂, 5% CO₂, and 0.1% O₂ at 37 °C for 12 h to establish hypoxia.
9. Set the temperature as described below.
   NOTE: The protocol starts with 10 h of a low-temperature treatment, followed by a rewarming phase for 2 h up to 37 °C, and 24 h of normothermia (37 °C). At every time point, three Petri dishes are removed for analysis.
10. After low-temperature treatment, aspirate the medium from the 6-well plate and wash with PBS (pH 7.4) three times.
11. Add 2 mL of complete DMEM per well.
12. Maintain the cells at 37 °C in a humidified incubator under an atmosphere with 95% air and 5% CO₂.

4. CCK-8 viability assay

1. Warm the 0.25% trypsin-ethylenediaminetetraacetic acid (EDTA) solution and PBS to 37 °C prior to use.
2. Aspirate the medium, rinse the cells with 1 mL of PBS and then add 0.5 mL of 0.25% trypsin along the wall of the well. Incubate at 37 °C until almost all HCMs are detached (approximately 1 min).
3. Add 1 mL of DMEM complete medium to the wells to neutralize the trypsin.
4. Transfer the cell suspension to a 15 mL centrifuge tube and pellet the HCMs by centrifugation at 500 × g for 3 min. Aspirate the supernatant without disturbing the pellet.
5. Dispense 100 µL aliquots of the cell suspension (5000 cells/well) into a 96-well plate. Preincubate the plate for 24 h in a humidified incubator (at 37 °C.)
6. Use the cultured cells in the 96-well plates to generate different treatment groups.
7. Incubate the plate for an appropriate length of time (16 h) in an incubator.
8. Add 10 µL of CCK-8 solution to each well of the plate.
9. Incubate the plate for 1 hour in the incubator.
10. Measure the absorbance at 450 nm using a microplate reader.
    CAUTION: Be careful not to introduce bubbles to the wells, as they interfere with OD reading measurements.

5. Flow cytometry for apoptosis analysis

1. Perform trypsinization and centrifugation steps by following steps 4.2-4.4.
   NOTE: Cells are harvested with trypsin without EDTA. To assess apoptosis, both floating and adherent cells are collected.
2. Centrifuge the cells for 5 min at 1000 × g. Discard the supernatant and resuspend the pellet in 1 mL of PBS.
3. Count the cells using a hemocytometer. Using a pipette, transfer 100 µL of Trypan Blue-treated cell suspension to a hemocytometer. Using a hand tally counter, count the live, unstained cells in one set of 16 squares and then use PBS to generate a cell suspension at 1×10⁷ cells/ml.
4. Obtain 200 µL of the cell suspension (5 × 10⁵ -1 × 10⁶ cells).
5. Centrifuge for 5 min at 1000 × g and resuspend the pellet in Annexin V-FITC binding solution.
6. Add 5 µL of Annexin V-FITC to dye the cells.
7. Add 10 µL of propidium iodide into the cell suspension.
8. Gently mix the cells and incubate them for 20 minutes at room temperature in the dark.
   NOTE: The cells are resuspended 2-3 times during the incubation to improve staining.
9. Start the flow cytometer and make sure the software is working appropriately.
10. Open two dot plot windows in the flow cytometry software.
11. Select forward scattered light (FSC) on the X axis and side scattered light (SSC) on the Y axis to exclude cell debris and/or clumps in terms of their size and granularity, respectively.
12. Select the PE (590 mm) detection channel and FITC (530 mm), which is used to measure the fluorescence intensity of the cells.
13. Place the blank (HCMs that have not been dyed) sample tube on the flow cytometer.
14. Click Record to collect particles from the suspension in the blank sample tube and then gate the cell population for further analysis in the first dot plot.
15. Place the single-stained samples on the tube support arm. Click Record to collect particles from the suspension and then gate the cell population for further analysis in the first dot plot.
16. Collect the HCMs in other sample tubes and optimize the measurement by adjusting the voltages of the fluorescence channels.
    NOTE: Unstained and single-stained samples are used as compensation controls during the experiment.
17. Display the statistics of each sample tube and calculate the rate of apoptosis of each sample.

6. Mitochondrial depolarization assessment

1. Perform trypsinization and centrifugation by following steps 4.2-4.4.
2. Resuspend the cells in 500 µL of complete medium, and then adjust the cell density adjusted to 1 × 10⁵ - 6 × 10⁶ cells
3. Add 0.5 mL of JC-1 working solution to each tube.
4. Incubate the cells in a cell incubator at 37 °C for 20 minutes.
   NOTE: During the incubation period, add 1 mL of 5× JC-1 staining buffer to 4 mL of distilled water to prepare the JC-1 staining buffer.
5. After incubation, centrifuge the cells at 600 × g for 3 min at 4 °C.
6. Discard the supernatant and resuspend the pellet in 1 mL of JC-1 staining buffer.
7. Repeat steps 6.5 to 6.6 three times.
8. Resuspend the HCMs in 1 mL of ice-cold staining buffer in a 1.5 mL centrifuge tube and use the cells for analysis within 30 min.
9. Select the PE (590 nm) detection channel and FITC (530 nm) to measure the fluorescence intensity of JC-1 dye in the cells.
   NOTE: Set up the flow cytometer by following steps 5.10-5.17.

7. Reactive oxygen species assay

1. Perform trypsinization and centrifugation by following steps 4.2-4.4.
2. Stain the cells in culture medium with 10 µM DCFDA and adjust the cell density to 1 × 10⁶ - 1 × 10⁷ cells
3. Incubate for 30 minutes at 37 °C.
4. Gently pipette the cells up/down every 3-5 min.
5. After the incubation, wash the cells three times with serum-free cell culture medium.
6. Analyze the cells on a flow cytometer by following steps 5.10-5.17.
   NOTE: DCF is excited at 488 nm and the emission intensity measured at 530 nm.

8. Measurement of Caspase 3/ Caspase 8 Activity

1. Perform trypsinization by following steps 4.2-4.4.
2. Collect the cells by centrifugation at 600 × g at 4 °C for 5 minutes.
3. Add 100 µL of lysate buffer per 2 × 10⁶ cells.
4. Lyse the cells for 15 minutes on ice.
5. After incubating for 15 min in an ice bath, centrifuge the sample at 1.6 × 10⁴ x g at 4 °C for 15 minutes.
6. Transfer the supernatant to an ice-cold centrifuge tube for use.
   NOTE: The protein concentration in the sample should be at least 1-3 mg/mL.
7. Remove Ac-DEVD-pNA (2 mM) and place it on an ice bath for use.
8. Accurately add 40 µL of buffer solution to the enzyme-labeled plate, add 80 µL of the sample, and finally add 10 µL of Ac-DEVD-pNA (2 mM).
9. Incubate the sample at 37 °C for 120 minutes.
10. Measured the A405 value on a microplate reader according to the manufacturer's instructions.
    NOTE: The absorbance produced by the pNA generated by caspase-3/caspase-8 is calculated by subtracting the A405 value of the blank control from that of the sample.

Results

The effect of OGD exposure on the viability of HCMs was determined by CCK-8 assay. Compared with that observed in the control group, cell viability was significantly decreased in a time-dependent manner (Figure 2A). The apoptosis rates of HCMs at different times after reperfusion showed a specific trend, where from 0 to 16 h, the apoptosis rates gradually increased and reached the maximum rate at the 16 h time point (Figure 2B). As OGD for 12 h reduced cell acti...

Discussion

The complexities of intact animals, including the interactions between different types of cells, often prevent detailed studies of specific components of I/R injury. Therefore, it is necessary to establish an in vitro cell model that can accurately reflect the molecular changes after ischemia in vivo. Research on OGD models has been previously reported^13,22, and many sophisticated methods have been established^23,

Materials

Name	Company	Catalog Number	Comments
Annexin V-FITC cell apoptosis detection kit	Bio-Technology,China	C1062M
Cardiac myocyte growth supplement	Sciencell,USA	6252
Caspase 3 activity assay kit	Bio-Technology,China	C1115
Caspase 8 activity assay kit	Bio-Technology,China	C1151
DMEM, no glucose	Gibco,USA	11966025
Dulbecco's modified eagle medium	Gibco,USA	11960044
Fetal bovine serum	Gibco,USA	16140071
Flow cytometry	CytoFLEX,USA	B49007AF
Human myocardial cells	BLUEFBIO,China	BFN60808678
Mitochondrial membrane potential assay kit with JC-1	Bio-Technology,China	C2006
Penicillin/Streptomycin solution	Gibco,USA	10378016
Reactive oxygen species assay kit	Bio-Technology,China	S0033S
Three-gas incubator	Memmert,Germany	ICO50
Trypsin-EDTA (0.25%)	Gibco,USA	25200056