Next Article in Journal
Plasma Copper Concentration Is Associated with Cardiovascular Mortality in Male Kidney Transplant Recipients
Next Article in Special Issue
Inhibition of Pyruvate Dehydrogenase in the Heart as an Initiating Event in the Development of Diabetic Cardiomyopathy
Previous Article in Journal
Structural Characterization of Polysaccharides from Coriandrum sativum Seeds: Hepatoprotective Effect against Cadmium Toxicity In Vivo
Previous Article in Special Issue
The Importance of SGLT-2 Inhibitors as Both the Prevention and the Treatment of Diabetic Cardiomyopathy
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Antioxidants
Volume 12
Issue 2
10.3390/antiox12020452
antioxidants-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Hyperkalemic or Low Potassium Cardioplegia Protects against Reduction of Energy Metabolism by Oxidative Stress

by
Hongting Diao
1,
Haiwei Gu
2,3,* and
Qin M. Chen
1,*
1
Department of Pharmacy Practice and Science, College of Pharmacy, The University of Arizona, Tucson, AZ 85721, USA
2
College of Health Solutions, Arizona State University Phoenix, Phoenix, AZ 85004, USA
3
Center for Translational Science, Florida International University, 11350 SW Village Parkway, Port St. Lucie, FL 34987, USA
*
Authors to whom correspondence should be addressed.
Antioxidants 2023, 12(2), 452; https://doi.org/10.3390/antiox12020452
Submission received: 12 December 2022 / Revised: 17 January 2023 / Accepted: 4 February 2023 / Published: 10 February 2023
(This article belongs to the Special Issue Oxidative Stress in Metabolic Cardiomyopathy)
Browse Figures
Review Reports Versions Notes

Abstract

:
Open-heart surgery is often an unavoidable option for the treatment of cardiovascular disease and prevention of cardiomyopathy. Cardiopulmonary bypass surgery requires manipulating cardiac contractile function via the perfusion of a cardioplegic solution. Procedure-associated ischemia and reperfusion (I/R) injury, a major source of oxidative stress, affects postoperative cardiac performance and long-term outcomes. Using large-scale liquid chromatography–tandem mass spectrometry (LC-MS/MS)-based metabolomics, we addressed whether cardioplegic solutions affect the baseline cellular metabolism and prevent metabolic reprogramming by oxidative stress. AC16 cardiomyocytes in culture were treated with commonly used cardioplegic solutions, High K+ (HK), Low K+ (LK), Del Nido (DN), histidine–tryptophan–ketoglutarate (HTK), or Celsior (CS). The overall metabolic profile shown by the principal component analysis (PCA) and heatmap revealed that HK or LK had a minimal impact on the baseline 78 metabolites, whereas HTK or CS significantly repressed the levels of multiple amino acids and sugars. H2O2-induced sublethal mild oxidative stress causes decreases in NAD, nicotinamide, or acetylcarnitine, but increases in glucose derivatives, including glucose 6-P, glucose 1-P, fructose, mannose, and mannose 6-P. Additional increases include metabolites of the pentose phosphate pathway, D-ribose-5-P, L-arabitol, adonitol, and xylitol. Pretreatment with HK or LK cardioplegic solution prevented most metabolic changes and increases of reactive oxygen species (ROS) elicited by H2O2. Our data indicate that HK and LK cardioplegic solutions preserve baseline metabolism and protect against metabolic reprogramming by oxidative stress.

1. Introduction

Cardiovascular disease is the principal cause of mortality worldwide. Most cardiac patients suffering from coronary artery disease have plaque buildup in the wall of the coronary arteries. With advancements in medical technology, an occluded coronary artery can be re-opened by the percutaneous coronary intervention technique. Open-heart surgery remains an option of treatment, especially for multivessel disease, diffuse or complex coronary disease, with or without valvular disease [1,2,3]. The development of cardiopulmonary bypass (CPB) using a heart–lung machine to sustain systematic circulation during operation has revolutionized cardiac surgery and reduced the mortality of patients with coronary artery disease. However, approximately 67% of CPB patients have post-operative complications, including atrial fibrillation, pulmonary dysfunction, or renal failure [4]. One contributing factor to these types of organ injury is procedure-associated ischemia and reperfusion (I/R), which produce oxidative stress.
In order to achieve surgical precision, cardiac contraction is minimized by the perfusion of a cardioplegic solution. This procedure causes controlled cessation of blood flow to the myocardium, i.e., myocardial ischemia. Upon completion of the surgical procedure, cardiac contraction and blood flow are resumed. Either ischemia or reperfusion results in an increased production of reactive oxygen species (ROS) [5,6]. Ischemia results in incomplete electron transport, impairment of mitochondrial respiration, and mitochondrial ROS production [7,8,9]. Reperfusion activates xanthine oxidase and NADPH oxidase in cardiac cells, which already inherit impaired mitochondria from ischemia, causing further increases of ROS [6,10]. Elevated levels of lipid peroxidation products, such as malondialdehyde, have been detected in the myocardium of patients following I/R from the coronary artery bypass graft surgery (CABG) [11]. Preventing oxidative stress is expected to reduce surgical-procedure-associated complications.
Cardioplegic solutions, along with induced hypothermia, during open-heart surgery are designed to prevent many detrimental effects of I/R [12,13,14]. The most commonly used cardioplegic solutions include modified St. Thomas, i.e., high potassium (high K+, HK) and low potassium (low K+, LK), Del Nido (DN), and histidine–tryptophane–ketoglutarate Custodiol® (HTK). Celsior (CS) was developed as a cardioplegia as well as a solution for donor organ preservation during cardiac transplant [15]. The HK, DN, and CS cardioplegic solutions, also known as extracellular cardioplegia, are hyperkalemic, delivering K+ via the coronary vasculature to reach the localized concentration of 15 to 25 mM [16]. Such high concentrations of K+ cause the resting membrane potential of the myocardium to change from −80 mV to −50 mV [17]. When the membrane potential reaches −50 mV, the voltage-dependent Na+ channels are inactivated, preventing the Na+-induced spike and rapid propagation of the action potential, thereby causing depolarized diastolic cardiac arrest. LK is used following the administration of HK to maintain the diastolic arrest. HTK, also called intracellular cardioplegia, is low in Na+. Depletion of Na+ in the extracellular spaces produces a hyperpolarization of the plasma membrane of cardiomyocytes, causing cardiac arrest in diastole [18]. Although these cardioplegic solutions have been widely used for open-heart surgery worldwide, little is known about the impact of these agents on the metabolism of cardiomyocytes.
Metabolomics provides an effective method for measuring overall chemical intermediates or metabolic state [19,20]. The application of metabolomics will not only help to identify new biomarkers for metabolic complications associated with cardioplegia but also will allow us to provide scientific evidence for choosing an ideal cardioplegic solution for cardiothoracic surgery, as such evidence is lacking. In this study, using large-scale liquid chromatography–tandem mass spectrometry (LC-MS/MS)-based metabolomics and AC16 human cardiomyocytes, we evaluated the metabolic impact of five commonly used cardioplegic solutions and addressed whether any of these solutions prevent metabolic changes by oxidative stress.

2. Material and Methods

2.1. Reagents

Cardioplegic solutions of HK, LK, DN, HTK (HTK Custodiol®, Bretschneider, Cardiolink Group, Barcelona, Spain), and CS (Sanofi, Bridgewater, NJ, USA) were collected as leftovers from cardiothoracic surgeries at the Banner-University Medical Center Tucson or Phoenix Children’s Hospital. HK, LK, and DN were formed at Banner-University of Arizona Hospital Pharmacy from Plasma-Lyte A (Baxter Healthcare Corporation, Deerfield, IL, USA).

2.2. Cell Lines, Culture Conditions, and Treatments

AC16 human cardiomyocytes were purchased from Millipore Sigma-Aldrich and cultured in Dulbecco’s modified Eagle’s medium mixed with Ham’s F12 (DMEM/F12, GIBCO, Life Technologies, Carlsbad, CA, USA) supplemented with 12.5% fetal bovine serum (FBS, R&D Systems, Minneapolis, MN, USA) and antibiotics (1% of 100 U penicillin/50 g/mL streptomycin, GIBCO) for culture in a 5% CO2 incubator at 37 °C with weekly subculture. The cells were seeded at a density of 0.3 × 106 cells per well in 6-well plates and grown to 90% confluence before experiments.
Cells were treated with cardioplegic solutions according to the ratios used clinically. HK or LK (0.4 mL) was mixed with 1.6 mL DMEM, whereas DN (1.6 mL) was mixed with 0.4 mL DMEM. For HTK or CS, 2 mL pure crystalloid solution was added to AC16 cells. Cells in 6-well plates were serum-starved overnight in 0.5%FBS/DMEM/F12 and treated with a cardioplegic solution for 3 h followed by exposure to 200 μM H2O2 for 1 h.

2.3. Metabolomics

The samples were collected for LC-MS/MS metabolomics as described [21,22,23,24]. Following a quick rinse with phosphate-buffered saline (PBS), cells were placed in 1 mL/well of 8:2 (v:v) methanol: H2O for 30 min incubation on dry ice. This serves to quench metabolism and extract the metabolites. The cells were scraped from the culture plates and transferred to centrifuge tubes. Another 0.7 mL/well of 8:2 (v:v) methanol: H2O was added to the plates on the dry ice to scrape and combined with the corresponding centrifuge tubes. Cell-free extracts were collected after 10 min centrifugation at 13,000 rpm under 4 °C. The soluble fractions of the extracts were dried at 4 °C using a speed vacuum. The samples were processed for LC-MS/MS as described [25].
LC-MS/MS analyses were performed using an Agilent 1290 UPLC-6490 QQQ-MS system (Santa Clara, CA, USA). Each sample was injected twice by an auto-sampler set at 4 °C, 10 μL for analysis using a negative ionization mode or 4 μL for analysis using a positive ionization mode. Both chromatographic separations were performed in the hydrophilic interaction chromatography mode on a Waters XBridge BEH Amide column (150 × 2.1 mm, 2.5 μm particle size, Waters Corporation, Milford, MA, USA), with a flow rate of 0.3 mL/min and the column compartment temperature of 40 °C. The mobile phase contained Solvent A (10 mM ammonium acetate and 10 mM ammonium hydroxide in 95% H2O/5% ACN) and Solvent B (10 mM ammonium acetate and 10 mM ammonium hydroxide in 5% H2O/95% ACN). After an initial 1 min isocratic elution of 90% Solvent B, the percentage of Solvent B decreased to 40% by t = 11 min. Solvent B was maintained at 40% for 4 min (t = 15 min) before gradually returning to 90% in preparation for the next injection. The mass spectrometer was equipped with an electrospray ionization source, and targeted data acquisition was performed in the multiple-reaction monitoring (MRM) mode. Agilent Masshunter Workstation software (Santa Clara, CA, USA) was used to control the operation of the LC-MS/MS system, whereas the extracted MRM peaks were integrated using the Agilent MassHunter Quantitative Data Analysis (Santa Clara, CA, USA).

2.4. Cell Morphology

AC16 cells were seeded in 6-well plates for treatments as described in Section 2.2. The cells were fixed in 4% paraformaldehyde for 5 min, washed twice with PBS, and stained with 0.1% Coomassie blue (dissolved in 10% acetic acid and 50% methanol) for 10 min. After washing off the dye, cell morphology was recorded under an inverted microscope with 10× lens (Rebel, Echo, San Diego, CA, USA).

2.5. ROS Detection

AC16 cells were seeded in a clear bottom 96-well dark plate for treatment with cardioplegic solutions or various concentrations of lidocaine for 3 h. After rinsing two times with warm (37 °C) Hanks’ Balanced Salt Solution (HBSS, 1.26 mM CaCl2, 0.49 mM MgCl2, 0.406 mM MgSO4, 5.33 mM KCl, 0.44 mM KH2PO4, 4.17 mM NaHCO3, 137.93 mM NaCl, and 0.338 mM Na2HPO4), the cells were incubated in dark with 10 µM 2′,7′–dichlorofluorescein diacetate (DCFDA) for 30 min at 37 °C in a 5% CO2 incubator. After the removal of DCFDA, 100 µL HBSS was added to the cells along with 200 µM H2O2 for measurements of ROS according to the manufacturer’s instructions (ROS detection kit, Abcam, Cambridge, UK). DCF fluorescence was recorded at Ex485/Em535 nm by a plate reader (CLARIOstar Plus, BMG Labtech, Ortenberg, Germany).

2.6. Statistical Analysis

Changes in metabolites or pathways between control versus treatment were analyzed using the web-based software MetaboAnalyst 5.0 (https://www.metaboanalyst.ca/MetaboAnalyst/ModuleView.xhtml, accessed 3 June 2022). PCA, K-means, heatmap clustering, and pathway analysis overview of altered metabolic profiles were analyzed using MetaboAnalyst 5.0. Bar graphs were presented as means ± standard deviations (SD) of the relative abundance of metabolites and analyzed via one-way ANOVA and corrected by Dunnett’s multiple comparisons test with GraphPad Prism 9 software. The data with two grouping variables were analyzed with two-way ANOVA and corrected by Turkey’s multiple comparisons test. p value or adjusted p value < 0.05 was set as the threshold for significant difference.

3. Results

3.1. Effects of Cardioplegic Solutions on the Metabolome

Table 1 lists the components of five cardioplegic solutions commonly used in the US and internationally for CPB surgeries (HK, LK, DN, and HTK) or cardiac transplants (CS). To evaluate the impact of these solutions on cellular metabolism, we treated AC16 cardiomyocytes according to their clinical applications, with the final concentration of each component listed in Table 1. The length of treatment was 4 h, mimicking the average CPB surgical time. A total of 94 metabolites were detected among the control (Ctrl) and cells treated with a cardioplegic solution, as shown in the heatmap (Figure 1A). When compared with the Ctrl, a similar pattern of metabolites was found in HK- and LK-treated groups, but HTK- and CS-treated cells exhibited a clear difference, showing decreases in 12 amino acids and 4 sugars (Figure 1A). The metabolic profile of DN-treated cells is closer to that of Ctrl, HK, or LK (Figure 1A,B). The overall metabolic profiles as summed by the principal component analysis (PCA) revealed that HK- or LK-cardioplegia-treated cells showed no significant difference from the Ctrl (Figure 1B).
PCA indicates significant deviations in the profile of HTK- or CS-treated cells (Figure 1B). The differences in the metabolites are displayed in a volcano plot (Figure 2A,B). The pathway analyses showed that HTK and CS caused significant perturbations of the pathways of: (1) alanine, aspartate, and glutamate metabolism; (2) histidine metabolism; (3) arginine biosynthesis; (4) tricarboxylic acid (TCA) cycle; (5) aminoacyl-tRNA biosynthesis; and (6) valine, leucine, and isoleucine biosynthesis (Figure 2C,D). CS also affected the glyoxylate and dicarboxylate metabolism pathway (Figure 2D). These results indicate that unlike HTK or CS, HK or LK cardioplegic solution preserves the basal metabolic profile.

3.2. Metabolic Profiles of Oxidative Stress and Impact of Cardioplegic Solutions

H2O2 at the dose of 200 μM was chosen here for metabolomics experiments since it does not cause detectable cytotoxicity, as measured by cell morphology, cell number, or general metabolic activity via CCK-8 assay [24,26]. This dose mimics the range of oxidants in the myocardium during I/R, 10–30 μM, based on the common belief that intracellular oxidants are 10–100-fold lower than extracellular H2O2 concentration [27,28].
A significant separation of metabolic profiles was discovered between the Ctrl and H2O2-treated group by PCA (Figure 3A). The heatmap shows 35 metabolites that were significantly up- (16) or down- (19) regulated, as determined by the Student’s t-test with FDR correction (Figure 3B). Among the significantly decreased metabolites are NAD and acetylcarnitine (Figure 3B). Increased metabolites include glucose derivatives, i.e., glucose 6-P, glucose 1-P, fructose, mannose, and mannose 6-P. Several sugar alcohols in the pentose phosphate pathway also increased, including L-arabitol, adonitol, Ribose 5-P, and xylitol (Figure 3B). Pathway analyses revealed significant alteration of nine metabolic pathways by H2O2 treatment (p < 0.05): (1) nicotinate and nicotinamide metabolism; (2) pentose phosphate pathway; (3) TCA cycle; (4) amino sugar and nucleotide sugar metabolism; (5) pentose and glucuronate interconversions; (6) alanine, aspartate, and glutamate metabolism; (7) purine metabolism; (8) glycolysis; and (9) butanate metabolism (Figure 3C).
Compared with the Ctrl or H2O2-treated cells, pretreatment with HK or LK was able to shift the metabolic profile of H2O2-treated cells towards that of Ctrl (Figure 4A). Among the 17 metabolites of 9 significantly changed pathways due to H2O2 treatment, HK or LK pretreatment was able to block H2O2 from inducing increases or decreases in the majority of these metabolites (Figure 4B). Specifically, the decrease in NAD or acetylcarnitine was attenuated by HK or LK treatment (Figure 5A,B). H2O2-induced increases in glucose 6-P, glucose 1-P, mannose 6-P, and oxaloacetic acid were inhibited by HK or LK (Figure 5B,C). The increases in the metabolites in the pentose phosphate pathway, i.e., xylitol, L-arabitol, adonitol, and D-ribose 5-P, due to H2O2 treatment were prevented by HK or LK (Figure 5D). These data indicate the capacity of HK or LK for protection against metabolic shifts by oxidative stress.

3.3. Protection against ROS Generation

To explore the mechanism of HK or LK for protection against metabolic shift by oxidative stress, we tested whether the cardioplegic solutions affect ROS production by H2O2. AC16 cells were able to tolerate five cardioplegic solutions without morphological evidence of toxicity (Figure 6A). When the baseline •OH generation was measured by treatment of a cardioplegic solution using DCFH-DA assay, the level of DCF fluorescence in HK- or LK-treated cells was similar to that of the control without any treatment (Figure 6B). The positive control of 200 μM of H2O2 showed a time-dependent elevation of DCF fluorescence over 240 min (Figure 6B). DN, HTK, or CS induced elevation of DCF fluorescence, with CS producing nearly half the amount of ROS as H2O2 (Figure 6B). We then investigated whether any of these cardioplegic solutions are capable of preventing H2O2 from generating •OH. When pretreating cells with a cardioplegic solution, we found an inhibition of H2O2 induced •OH generation by HK or LK, but not DN, HTK, or CS (Figure 6C).

4. Discussion

Using LC-MS/MS-based metabolomics, we evaluated the impact of five cardioplegic solutions on cellular metabolism and metabolic alterations by oxidative stress. Our data indicate that HK and LK cardioplegia solutions did not cause significant changes in the baseline metabolic profile, whereas HTK and CS induced significant metabolic shifts. Induction of oxidative stress by H2O2 treatment resulted in decreases in NAD and acetylcarnitine but increases in glucose 6-P, glucose 1-P, mannose 6-P, oxaloacetic acid, and four sugar alcohols from the pentose phosphate pathway. Our data suggest that HK and LK cardioplegic solutions can provide protection against alterations in energy metabolism due to oxidative stress at the cellular level. Such protection correlates with an inhibition of ROS generation.
The cardioplegic solutions are capable of supporting an extended period of myocardial ischemia with organ preservation during cardiac surgery. The choice of which cardioplegic solution is based on the preference and experience of each surgeon. HK, LK, and DN, the so-called blood cardioplegia, require isogenic mixing of the patient’s blood for their clinical application, whereas HTK or CS is perfused into the myocardium as a pure crystalloid. Experimental or clinical evidence suggests that blood cardioplegia is superior to the crystalloids for cardiac protection [29,30,31,32,33,34,35]. Among the many advantages of blood cardioplegia are oxygen-carrying capacity, continuous supply of metabolic substrates, physiologic buffering, and consistent osmotic pressure [36,37]. We mimicked the clinical application by mixing HK, LK, and DN with the culture medium. This approach has maintained the supply of nutrients, including glucose, amino acids, vitamins, and minerals, which were absent in the crystalloid solution HTK or CS. Therefore, the large deviation in the basal metabolic profile by HTK or CS could result from nutrient deprivation in the cell culture experiments. Nevertheless, our finding of HK or LK having a minimal impact on the baseline metabolic profile yet protecting against metabolic switch by oxidants supports the clinical observation of the superiority of the blood cardioplegia.
The metabolic shift detected here by oxidative stress resembles that of surgery-induced ischemia. A decrease in NAD level has been reported in myocardial tissue due to CABG surgery [38,39]. Metabolic reprogramming indeed occurs with CPB since a nuclear magnetic resonance study of the left atrial tissue showed increases in glucose, pyruvate, citrate, and lactate, indicating a reduction of glycolytic energy metabolism [40]. On the other hand, increases in sugar alcohols have been linked to apoptosis of cardiomyocytes during ischemia [41,42,43]. Although xylitol, adonitol, arabitol, and ribose-5-P may not be inducers of apoptosis, the correlation of their elevations with cell death suggests the functionality as a biomarker of cell injury. Inhibition of H2O2-induced metabolic changes by HK and LK suggest the cytoprotective capacity of HK and LK against ischemic injury or oxidative stress.
Ischemia causes a shift from aerobic to anaerobic metabolism, where ATP production is decreased due to O2 deprivation for the respiratory chain. Glycolysis, the main ATP source in the ischemic myocardium, is progressively limited by the restraint of glyceraldehyde 3-phosphate dehydrogenase flux due to NADH accumulation [44]. High levels of NADH and FADH2 inhibit β-oxidation, leading to an accumulation of long-chain acyl CoA, acylcarnitine esters, and free fatty acids [45,46]. These lipophilic compounds can act as “detergents”, altering membrane functions, such as Na+/K+ ATPase, and sarcoplasmic reticular Ca2+-stimulated ATPase [47]. Upon reperfusion, these changes contribute to the elevation of cytosolic Ca2+ [48]. Oxidative stress can cause Ca2+ influx and elevation of cytosolic Ca2+. Elevation of cytosolic Ca2+ either by ischemic reperfusion or oxidative stress causes mitochondrial membrane permeability transition and a further increase in oxidant production [5,6].
HK and LK deliver 20 mg/L or 0.085 mM lidocaine, whereas DN introduces 104 mg/L or 0.444 mM lidocaine to the cells in the culture. In contrast, lidocaine is absent in HTK and CS. Lidocaine is a sodium channel blocker that inhibits the entry of Na+ into cells through a fast voltage-gated Na+ channel (NaV). A number of publications have reported the cytotoxicity of lidocaine in a variety of cell types in the mM concentration range [49,50]. Cytotoxicity may result from high doses that allow lidocaine to act as a surfactant and disrupt the plasma membrane [50]. This may explain the lack of full protection of DN against the metabolic change of oxidative stress.
In cardiomyocytes, lidocaine appears to serve as a cytoprotective agent. Lidocaine protects H9C2 cardiomyocytes against loss of viability by hypoxia [51]. Protection against myocardial infarction has been reported in mice with lidocaine injection [52]. A similar discovery was reported with regional ischemia in rats [53]. CPB in a canine model showed that lidocaine protected against mortality due to global myocardial ischemia and reperfusion [54]. An early study using a canine model of regional ischemic reperfusion showed a reduction of infarct size by lidocaine in association with decreased lipid peroxidation [55]. The protective effect of lidocaine against myocardial infarction was also observed in a porcine experimental model [56]. These protective effects may result from the inhibition of Nav and the prevention of intracellular Na+ increase. Na+ efflux is coupled to Ca2+ influx via the sodium–calcium exchanger (NCX). Inhibition of intracellular Ca2+ elevation by lidocaine has been demonstrated by pacing isolated rat hearts [57]. In cardiomyocytes, late openings of the sodium channel by H2O2 treatment may contribute to an increase in cytosolic Ca2+ [58]. Lidocaine may block H2O2 from inducing elevation of cytosolic Ca2+ and, therefore, production of ROS from the mitochondria. A preliminary experiment indeed showed protection against ROS generation by lidocaine (Figure 7). Early evidence indicates that lidocaine preserved mitochondrial oxidative phosphorylation during ischemic reperfusion [59,60]. Consistent with the protective role of lidocaine, an inhibitor of NCX was capable of preventing intracellular Ca2+ overload or mitochondrial Ca2+ increase and enhancing cardiac recovery following ischemia [61,62]. Therefore, the observed protective effect of HK or LK may be related to the presence of lidocaine and the inhibition of ROS generation.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/antiox12020452/s1.

Author Contributions

H.D. designed and performed the experiments, analyzed the data, including metabolomic data, and wrote the manuscript; H.G. carried out LC-MS/MS-based metabolomic analysis; Q.M.C. supervised the project and wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

The research programs under the direction of Qin M. Chen have been supported by National Institutes of Health (NIH) grants (R01 GM125212, R01 GM126165), Holsclaw Endowment and the University of Arizona, College of Pharmacy start-up fund.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The authors confirm that the data supporting the findings of this study are available within the article and its Supplementary Material. Raw data that support the findings of this study are available from the corresponding author, upon reasonable request.

Acknowledgments

We thank Yan Jin for her technical assistance in metabolomics analyses, Lenee Shrestha, Wujing Dai, and Daniel Wurm for discussion, and Banner-University of Arizona Medical Center Tucson and Phoenix Children’s Hospital for providing the leftover cardioplegic solutions. Benjamin Yuchen is acknowledged for English editing. The research programs under the direction of Qin M. Chen have been supported by NIH R01 GM125212, R01 GM126165, Holsclaw Endowment, and the University of Arizona College of Pharmacy start-up fund.

Conflicts of Interest

The authors declare that there are no conflict of interest.

Abbreviations

CABG: coronary artery bypass graft surgery; CPB, cardiopulmonary bypass surgery; CCK-8, cell counting kit-8; FDR, false discovery rate; glucose-1-P, glucose-1-phosphate; glucose-6-P, beta-D-glucose-6-phosphate; LC-MS/MS, liquid chromatography with tandem mass spectrometry; mannose-6-P, mannose-6-phosphate; NAD, nicotinamide adenine dinucleotide; NaV, voltage gated Na+ channel; NCX, sodium–calcium exchanger; D-ribose-5-P, D-ribose-5-phosphate; ROS, reactive oxygen species; TCA, tricarboxylic acid cycle.

References

  1. Spadaccio, C.; Benedetto, U. Coronary artery bypass grafting (CABG) vs. percutaneous coronary intervention (PCI) in the treatment of multivessel coronary disease: Quo vadis?—A review of the evidences on coronary artery disease. Ann. Cardiothorac. Surg. 2018, 7, 506–515. [Google Scholar] [CrossRef] [PubMed]
  2. Doenst, T.; Haverich, A.; Serruys, P.; Bonow, R.O.; Kappetein, P.; Falk, V.; Velazquez, E.; Diegeler, A.; Sigusch, H. PCI and CABG for Treating Stable Coronary Artery Disease: JACC Review Topic of the Week. J. Am. Coll. Cardiol. 2019, 73, 964–976. [Google Scholar] [CrossRef]
  3. Welt, F.G.P. CABG versus PCI—End of the Debate? N. Engl. J. Med. 2022, 386, 185–187. [Google Scholar] [CrossRef] [PubMed]
  4. Pahwa, S.; Bernabei, A.; Schaff, H.; Stulak, J.; Greason, K.; Pochettino, A.; Daly, R.; Dearani, J.; Bagameri, G.; King, K.; et al. Impact of postoperative complications after cardiac surgery on long-term survival. J. Card. Surg. 2021, 36, 2045–2052. [Google Scholar] [CrossRef]
  5. Raedschelders, K.; Ansley, D.M.; Chen, D.D. The cellular and molecular origin of reactive oxygen species generation during myocardial ischemia and reperfusion. Pharmacol. Ther. 2012, 133, 230–255. [Google Scholar]
  6. Chen, Q.M. Nrf2 for protection against oxidant generation and mitochondrial damage in cardiac injury. Free Radic. Biol. Med. 2022, 179, 133–143. [Google Scholar] [CrossRef]
  7. Chen, Y.R.; Zweier, J.L. Cardiac mitochondria and reactive oxygen species generation. Circ. Res. 2014, 114, 524–537. [Google Scholar] [CrossRef] [PubMed]
  8. Kalogeris, T.; Bao, Y.; Korthuis, R.J. Mitochondrial reactive oxygen species: A double edged sword in ischemia/reperfusion vs preconditioning. Redox Biol. 2014, 2, 702–714. [Google Scholar] [CrossRef]
  9. Bugger, H.; Pfeil, K. Mitochondrial ROS in myocardial ischemia reperfusion and remodeling. Biochim. Biophys. Acta Mol. Basis Dis. 2020, 1866, 165768. [Google Scholar] [CrossRef]
  10. Granger, D.N.; Kvietys, P.R. Reperfusion injury and reactive oxygen species: The evolution of a concept. Redox Biol. 2015, 6, 524–551. [Google Scholar]
  11. Lazzarino, G.; Raatikainen, P.; Nuutinen, M.; Nissinen, J.; Tavazzi, B.; Di Pierro, D.; Giardina, B.; Peuhkurinen, K. Myocardial release of malondialdehyde and purine compounds during coronary bypass surgery. Circulation 1994, 90, 291–297. [Google Scholar] [CrossRef]
  12. Habertheuer, A.; Kocher, A.; Laufer, G.; Andreas, M.; Szeto, W.Y.; Petzelbauer, P.; Ehrlich, M.; Wiedemann, D. Cardioprotection: A review of current practice in global ischemia and future translational perspective. Biomed. Res. Int. 2014, 2014, 325725. [Google Scholar] [CrossRef] [PubMed]
  13. Ji, M.J.; Hong, J.H. A Cardioplegic Solution with an Understanding of a Cardiochannelopathy. Antioxidants 2021, 10, 1878. [Google Scholar] [CrossRef]
  14. Francica, A.; Tonelli, F.; Rossetti, C.; Tropea, I.; Luciani, G.B.; Faggian, G.; Dobson, G.P.; Onorati, F. Cardioplegia between Evolution and Revolution: From Depolarized to Polarized Cardiac Arrest in Adult Cardiac Surgery. J. Clin. Med. 2021, 10, 4485. [Google Scholar] [CrossRef]
  15. Llosa, J.C.; Rodriguez Lambert, J.L.; Naya, J.L.; Gosalbez, F.; Valle, J.M. Celsior, a novel cardioplegic solution for arrest and storage in heart transplantation. Transplant. Proc. 2000, 32, 2589–2590. [Google Scholar] [CrossRef] [PubMed]
  16. Dobson, G.P.; Faggian, G.; Onorati, F.; Vinten-Johansen, J. Hyperkalemic cardioplegia for adult and pediatric surgery: End of an era? Front. Physiol. 2013, 4, 228. [Google Scholar] [CrossRef] [PubMed]
  17. Chambers, D.J.; Fallouh, H.B. Cardioplegia and cardiac surgery: Pharmacological arrest and cardioprotection during global ischemia and reperfusion. Pharmacol. Ther. 2010, 127, 41–52. [Google Scholar] [CrossRef]
  18. Takahashi, A.; Chambers, D.J.; Braimbridge, M.V.; Hearse, D.J. Optimal myocardial protection during crystalloid cardioplegia. Interrelationship between volume and duration of infusion. J. Thorac. Cardiovasc. Surg. 1988, 96, 730–740. [Google Scholar] [CrossRef]
  19. Griffin, J.L.; Atherton, H.; Shockcor, J.; Atzori, L. Metabolomics as a tool for cardiac research. Nat. Rev. Cardiol. 2011, 8, 630–643. [Google Scholar] [CrossRef]
  20. Turer, A.T. Using metabolomics to assess myocardial metabolism and energetics in heart failure. J. Mol. Cell Cardiol. 2013, 55, 12–18. [Google Scholar] [CrossRef]
  21. Gu, H.; Zhang, P.; Zhu, J.; Raftery, D. Globally Optimized Targeted Mass Spectrometry: Reliable Metabolomics Analysis with Broad Coverage. Anal. Chem. 2015, 87, 12355–12362. [Google Scholar] [CrossRef] [PubMed]
  22. Shi, X.; Wang, S.; Jasbi, P.; Turner, C.; Hrovat, J.; Wei, Y.; Liu, J.; Gu, H. Database-Assisted Globally Optimized Targeted Mass Spectrometry (dGOT-MS): Broad and Reliable Metabolomics Analysis with Enhanced Identification. Anal. Chem. 2019, 91, 13737–13745. [Google Scholar] [CrossRef] [PubMed]
  23. Jasbi, P.; Shi, X.; Chu, P.; Elliott, N.; Hudson, H.; Jones, D.; Serrano, G.; Chow, B.; Beach, T.G.; Liu, L.; et al. Metabolic Profiling of Neocortical Tissue Discriminates Alzheimer’s Disease from Mild Cognitive Impairment, High Pathology Controls, and Normal Controls. J. Proteome Res. 2021, 20, 4303–4317. [Google Scholar] [CrossRef] [PubMed]
  24. Zhu, C.; Gu, H.; Jin, Y.; Wurm, D.; Freidhof, B.; Lu, Y.; Chen, Q.M. Metabolomics of oxidative stress: Nrf2 independent depletion of NAD or increases of sugar alcohols. Toxicol. Appl. Pharmacol. 2022, 442, 115949. [Google Scholar] [CrossRef]
  25. He, H.; Shi, X.; Lawrence, A.; Hrovat, J.; Turner, C.; Cui, J.Y.; Gu, H. 2,2′,4,4′-tetrabromodiphenyl ether (BDE-47) induces wide metabolic changes including attenuated mitochondrial function and enhanced glycolysis in PC12 cells. Ecotoxicol. Environ. Saf. 2020, 201, 110849. [Google Scholar] [CrossRef]
  26. Dai, W.; Qu, H.; Zhang, J.; Thongkum, A.; Dinh, T.N.; Kappeler, K.V.; Chen, Q.M. Far Upstream Binding Protein 1 (FUBP1) participates in translational regulation of Nrf2 protein under oxidative stress. Redox Biol. 2021, 41, 101906. [Google Scholar] [CrossRef]
  27. Grill, H.P.; Zweier, J.L.; Kuppusamy, P.; Weisfeldt, M.L.; Flaherty, J.T. Direct measurement of myocardial free radical generation in an in vivo model: Effects of postischemic reperfusion and treatment with human recombinant superoxide dismutase. J. Am. Coll. Cardiol. 1992, 20, 1604–1611. [Google Scholar] [CrossRef]
  28. Sies, H. Hydrogen peroxide as a central redox signaling molecule in physiological oxidative stress: Oxidative eustress. Redox Biol. 2017, 11, 613–619. [Google Scholar] [CrossRef]
  29. Matte, G.S.; del Nido, P.J. History and use of del Nido cardioplegia solution at Boston Children’s Hospital. J. Extra Corpor. Technol. 2012, 44, 98–103. [Google Scholar]
  30. Jacob, S.; Kallikourdis, A.; Sellke, F.; Dunning, J. Is blood cardioplegia superior to crystalloid cardioplegia? Interact. Cardiovasc. Thorac. Surg. 2008, 7, 491–498. [Google Scholar] [CrossRef]
  31. Li, Y.; Lin, H.; Zhao, Y.; Li, Z.; Liu, D.; Wu, X.; Ji, B.; Gao, B. Del Nido Cardioplegia for Myocardial Protection in Adult Cardiac Surgery: A Systematic Review and Meta-Analysis. ASAIO J. 2018, 64, 360–367. [Google Scholar] [CrossRef] [PubMed]
  32. Ribeiro, R.V.P.; Friedrich, J.O.; Ouzounian, M.; Yau, T.; Lee, J.; Yanagawa, B.; Canadian Cardiovascular Surgery Meta-Analysis Working Group. Supplemental Cardioplegia During Donor Heart Implantation: A Systematic Review and Meta-Analysis. Ann. Thorac. Surg. 2020, 110, 545–552. [Google Scholar] [CrossRef] [PubMed]
  33. Karduz, G.; Yaman, M.O.; Altan, M.; Sahin, G.; Toraman, F.; Aksu, U. Thomas and del Nido cardioplegia are superior to Custodiol cardioplegia in a rat model of donor heart. Scand. Cardiovasc. J. 2021, 55, 122–128. [Google Scholar] [CrossRef] [PubMed]
  34. Sharma, A.; Dixit, S.; Mittal, S.; Sharma, M.; Sharma, D.; Mawar, K.K. DelNido cardioplegia versus St Thomas cardioplegia solution in double valve replacement: A single centre experience. Perfusion 2021, 36, 476–481. [Google Scholar] [CrossRef] [PubMed]
  35. Lenoir, M.; Bouhout, I.; Jelassi, A.; Cartier, R.; Poirier, N.; El-Hamamsy, I.; Demers, P. Del Nido cardioplegia versus blood cardioplegia in adult aortic root surgery. J. Thorac. Cardiovasc. Surg. 2021, 162, 514–522.e2. [Google Scholar] [CrossRef]
  36. Velez, D.A.; Morris, C.D.; Budde, J.M.; Muraki, S.; Otto, R.N.; Guyton, R.A.; Vinten-Johansen, J. All-blood (miniplegia) versus dilute cardioplegia in experimental surgical revascularization of evolving infarction. Circulation 2001, 104, I296–I302. [Google Scholar] [CrossRef]
  37. Fang, Y.; Long, C.; Lou, S.; Guan, Y.; Fu, Z. Blood versus crystalloid cardioplegia for pediatric cardiac surgery: A meta-analysis. Perfusion 2015, 30, 529–536. [Google Scholar] [CrossRef]
  38. Carlucci, F.; Tabucchi, A.; Biagioli, B.; Simeone, F.; Scolletta, S.; Rosi, F.; Marinello, E. Cardiac surgery: Myocardial energy balance, antioxidant status and endothelial function after ischemia–reperfusion. Biomed. Pharmacother. 2002, 56, 483–491. [Google Scholar] [CrossRef]
  39. Nguyen, B.A.; Suleiman, M.S.; Anderson, J.R.; Evans, P.C.; Fiorentino, F.; Reeves, B.C.; Angelini, G.D. Metabolic derangement and cardiac injury early after reperfusion following intermittent cross-clamp fibrillation in patients undergoing coronary artery bypass graft surgery using conventional or miniaturized cardiopulmonary bypass. Mol. Cell. Biochem. 2014, 395, 167–175. [Google Scholar] [CrossRef]
  40. Maltesen, R.G.; Rasmussen, B.S.; Pedersen, S.; Hanifa, M.A.; Kucheryavskiy, S.; Kristensen, S.R.; Wimmer, R. Metabotyping Patients’ Journeys Reveals Early Predisposition to Lung Injury after Cardiac Surgery. Sci. Rep. 2017, 7, 40275. [Google Scholar] [CrossRef]
  41. Galvez, A.S.; Ulloa, J.A.; Chiong, M.; Criollo, A.; Eisner, V.; Barros, L.F.; Lavandero, S. Aldose reductase induced by hyperosmotic stress mediates cardiomyocyte apoptosis: Differential effects of sorbitol and mannitol. J. Biol. Chem. 2003, 278, 38484–38494. [Google Scholar] [CrossRef]
  42. Chiong, M.; Parra, V.; Eisner, V.; Ibarra, C.; Maldonado, C.; Criollo, A.; Bravo, R.; Quiroga, C.; Contreras, A.; Vicencio, J.M.; et al. Parallel activation of Ca(2+)-induced survival and death pathways in cardiomyocytes by sorbitol-induced hyperosmotic stress. Apoptosis 2010, 15, 887–903. [Google Scholar] [CrossRef]
  43. Son, N.H.; Ananthakrishnan, R.; Yu, S.; Khan, R.S.; Jiang, H.; Ji, R.; Akashi, H.; Li, Q.; O’Shea, K.; Homma, S.; et al. Cardiomyocyte aldose reductase causes heart failure and impairs recovery from ischemia. PLoS ONE 2012, 7, e46549. [Google Scholar] [CrossRef] [Green Version]
  44. Wu, J.; Jin, Z.; Zheng, H.; Yan, L.J. Sources and implications of NADH/NAD(+) redox imbalance in diabetes and its complications. Diabetes Metab. Syndr. Obes. 2016, 9, 145–153. [Google Scholar]
  45. Jaswal, J.S.; Keung, W.; Wang, W.; Ussher, J.R.; Lopaschuk, G.D. Targeting fatty acid and carbohydrate oxidation—A novel therapeutic intervention in the ischemic and failing heart. Biochim. Biophys. Acta 2011, 1813, 1333–1350. [Google Scholar] [CrossRef]
  46. Ojuka, E.; Andrew, B.; Bezuidenhout, N.; George, S.; Maarman, G.; Madlala, H.P.; Mendham, A.; Osiki, P.O. Measurement of beta-oxidation capacity of biological samples by respirometry: A review of principles and substrates. Am. J. Physiol. Endocrinol. Metab 2016, 310, E715–E723. [Google Scholar] [CrossRef] [PubMed]
  47. Aitken-Buck, H.M.; Krause, J.; Zeller, T.; Jones, P.P.; Lamberts, R.R. Long-Chain Acylcarnitines and Cardiac Excitation-Contraction Coupling: Links to Arrhythmias. Front. Physiol. 2020, 11, 577856. [Google Scholar] [CrossRef] [PubMed]
  48. Marsh, J.D.; Smith, T.S. Calcium overload and ischemic myocardial injury. Circulation 1991, 83, 709–711. [Google Scholar] [CrossRef]
  49. Kubrova, E.; Su, M.; Galeano-Garces, C.; Galvan, M.L.; Jerez, S.; Dietz, A.B.; Smith, J.; Qu, W.; van Wijnen, A.J. Differences in Cytotoxicity of Lidocaine, Ropivacaine, and Bupivacaine on the Viability and Metabolic Activity of Human Adipose-Derived Mesenchymal Stem Cells. Am. J. Phys. Med. Rehabil. 2021, 100, 82–91. [Google Scholar] [CrossRef] [PubMed]
  50. Nishimura, Y.; Kanada, A.; Yamaguchi, J.Y.; Horimoto, K.; Kobayashi, M.; Tatsuishi, T.; Kanemaru, K.; Ueno, S.Y.; Oyama, Y. Cytometric analysis of lidocaine-induced cytotoxicity: A model experiment using rat thymocytes. Toxicology 2006, 218, 48–57. [Google Scholar] [CrossRef]
  51. Jin, H.; Yu, J. Lidocaine protects H9c2 cells from hypoxia-induced injury through regulation of the MAPK/ERK/NF-kappaB signaling pathway. Exp. Ther Med. 2019, 18, 4125–4131. [Google Scholar] [PubMed]
  52. Kaczmarek, D.J.; Herzog, C.; Larmann, J.; Gillmann, H.J.; Hildebrand, R.; Schmitz, M.; Westermann, A.; Harendza, T.; Werdehausen, R.; Osthaus, A.W.; et al. Lidocaine protects from myocardial damage due to ischemia and reperfusion in mice by its antiapoptotic effects. Anesthesiology 2009, 110, 1041–1049. [Google Scholar] [CrossRef]
  53. Canyon, S.J.; Dobson, G.P. Protection against ventricular arrhythmias and cardiac death using adenosine and lidocaine during regional ischemia in the in vivo rat. Am. J. Physiol Heart Circ. Physiol. 2004, 287, H1286–H1295. [Google Scholar] [CrossRef] [PubMed]
  54. Dias, R.R.; Stolf, N.A.; Dalva, M.; Dias, A.R.; Moreira, L.F.; Oliveira, S.A. Inclusion of lidocaine in cardioplegic solutions provides additional myocardial protection. J. Cardiovasc. Surg. (Torino) 2004, 45, 551–555. [Google Scholar]
  55. Lesnefsky, E.J.; VanBenthuysen, K.M.; McMurtry, I.F.; Shikes, R.H.; Johnston, R.B., Jr.; Horwitz, L.D. Lidocaine reduces canine infarct size and decreases release of a lipid peroxidation product. J. Cardiovasc. Pharmacol. 1989, 13, 895–901. [Google Scholar] [CrossRef]
  56. Lee, R.; Nitta, T.; Schmid, R.A.; Schuessler, R.B.; Harris, K.M.; Gay, W.A., Jr. Retrograde infusion of lidocaine or L-arginine before reperfusion reduces myocardial infarct size. Ann. Thorac. Surg. 1998, 65, 1353–1359. [Google Scholar] [CrossRef] [PubMed]
  57. Stefenelli, T.; Wikman-Coffelt, J.; Wu, S.T.; Parmley, W.W. Intracellular calcium during pacing-induced ventricular fibrillation. Effects of lidocaine. J. Electrocardiol. 1992, 25, 221–228. [Google Scholar] [CrossRef]
  58. Ma, J.; Song, Y.; Shryock, J.C.; Hu, L.; Wang, W.; Yan, X.; Zhang, P.; Belardinelli, L. Ranolazine attenuates hypoxia- and hydrogen peroxide-induced increases in sodium channel late openings in ventricular myocytes. J. Cardiovasc. Pharmacol. 2014, 64, 60–68. [Google Scholar] [CrossRef]
  59. Baron, D.W.; Sunamori, M.; Harrison, C.E. Preservation of oxidative phosphorylation by lidocaine in ischemic and reperfused myocardium. Adv. Myocardiol. 1983, 4, 567–573. [Google Scholar]
  60. Kyo, S.; Laraia, P.J.; Magrassi, P.; Levine, F.H.; Tsai, C.H.; Austen, W.G.; Buckley, M.J. Myocardial protection by lidocaine during cardioplegia. J. Surg. Res. 1983, 34, 533–542. [Google Scholar] [CrossRef]
  61. Egar, J.; Ali, A.; Howlett, S.E.; Friesen, C.H.; O’Blenes, S. The Na+/Ca2+ exchange inhibitor SEA0400 limits intracellular Ca2+ accumulation and improves recovery of ventricular function when added to cardioplegia. J. Cardiothorac. Surg. 2014, 9, 11. [Google Scholar] [CrossRef] [PubMed]
  62. Namekata, I.; Shimada, H.; Kawanishi, T.; Tanaka, H.; Shigenobu, K. Reduction by SEA0400 of myocardial ischemia-induced cytoplasmic and mitochondrial Ca2+ overload. Eur. J. Pharmacol. 2006, 543, 108–115. [Google Scholar] [CrossRef] [PubMed]
Antioxidants 12 00452 g001 550
Figure 1. Effects of 5 Cardioplegic Solutions on Cellular Metabolism. AC16 human cardiac myocytes were seeded in 6-well plates and cultured in DMEM containing 12.5% FBS. When reaching 90% confluency, cells were placed in 0.5% FBS/DMEM/F12 for overnight serum starvation. The cells were treated with HK, LK, DN, HTK, and CS for 4 h before extraction of aqueous metabolites from cell lysates for LC-MS/MS-based metabolomics. The overall metabolic profile differences between control (Ctrl) and cells treated with a cardioplegia solution (n = 4) are shown in the heatmap (A) and by a principal component analysis (PCA) score plot (B).
Figure 1. Effects of 5 Cardioplegic Solutions on Cellular Metabolism. AC16 human cardiac myocytes were seeded in 6-well plates and cultured in DMEM containing 12.5% FBS. When reaching 90% confluency, cells were placed in 0.5% FBS/DMEM/F12 for overnight serum starvation. The cells were treated with HK, LK, DN, HTK, and CS for 4 h before extraction of aqueous metabolites from cell lysates for LC-MS/MS-based metabolomics. The overall metabolic profile differences between control (Ctrl) and cells treated with a cardioplegia solution (n = 4) are shown in the heatmap (A) and by a principal component analysis (PCA) score plot (B).
Antioxidants 12 00452 g001
Antioxidants 12 00452 g002 550
Figure 2. Metabolites and Metabolic Pathways Altered by HTK and CS. AC16 cells seeded in 6-well plates were used for the collection of aqueous metabolites as described in Figure 1 for LC-MS/MS-based metabolomics. A total of 67 or 65 metabolites showed statistically significant differences as determined by Student’s t-test (n = 4, p < 0.05) between control versus HTK or CS and are displayed in the volcano plots (A,B) generated by the MetaboAnalyst 5.0 software. The altered metabolic pathways are displayed with the x-axis representing the pathway impact score computed from pathway topological analysis and the y-axis being the -log10 of the p-value obtained from pathway enrichment analysis (C,D). The metabolic pathways perturbed by the HTK and CS treatment are displayed in the gradient from yellow to red, reflecting the p-value from statistical analysis, as shown in the y-axis, from non-significant (≤1 for −log10, p ≥ 0.1, light yellow) to significant (≥1 but ≤2 for −log10, p ≤ 0.01 to p ≥ 0.001, orange) to highly significant (≥2 for −log10, p ≤ 0.0001, red). The size of the circles reflects the number of metabolites altered in the metabolic pathway (C,D).
Figure 2. Metabolites and Metabolic Pathways Altered by HTK and CS. AC16 cells seeded in 6-well plates were used for the collection of aqueous metabolites as described in Figure 1 for LC-MS/MS-based metabolomics. A total of 67 or 65 metabolites showed statistically significant differences as determined by Student’s t-test (n = 4, p < 0.05) between control versus HTK or CS and are displayed in the volcano plots (A,B) generated by the MetaboAnalyst 5.0 software. The altered metabolic pathways are displayed with the x-axis representing the pathway impact score computed from pathway topological analysis and the y-axis being the -log10 of the p-value obtained from pathway enrichment analysis (C,D). The metabolic pathways perturbed by the HTK and CS treatment are displayed in the gradient from yellow to red, reflecting the p-value from statistical analysis, as shown in the y-axis, from non-significant (≤1 for −log10, p ≥ 0.1, light yellow) to significant (≥1 but ≤2 for −log10, p ≤ 0.01 to p ≥ 0.001, orange) to highly significant (≥2 for −log10, p ≤ 0.0001, red). The size of the circles reflects the number of metabolites altered in the metabolic pathway (C,D).
Antioxidants 12 00452 g002
Antioxidants 12 00452 g003 550
Figure 3. Metabolites and Metabolic Pathways Altered by H2O2 treatment. AC16 human cardiac myocytes were seeded in 6-well plates and cultured in DMEM containing 12.5% FBS. When reaching 90% confluency, cells were serum-starved in 0.5% FBS DMEM/F12 overnight. The cells were treated with 200 μM of H2O2 for 1 h before extraction of aqueous metabolites by 80% methanol from cell lysates for LC-MS/MS-based metabolomics. The overall metabolic profile differences between ctrl and H2O2-treated cells are shown by the PCA score plot (A). The heatmap shows significant differences (n = 4, p < 0.05) in 35 metabolites between Ctrl and H2O2-treated cells (B). The altered metabolic pathways by H2O2 treatment are displayed with the x-axis representing the pathway impact score computed from pathway topological analysis and the y-axis being -log10 of the p-value obtained from the pathway enrichment analysis (C). Metabolic pathway analyses by MetaboAnalyst 5.0 software depict the impact of H2O2 treatment, with the x-axis, y-axis, color, or size of the circle reflecting the information as described in Figure 2C.
Figure 3. Metabolites and Metabolic Pathways Altered by H2O2 treatment. AC16 human cardiac myocytes were seeded in 6-well plates and cultured in DMEM containing 12.5% FBS. When reaching 90% confluency, cells were serum-starved in 0.5% FBS DMEM/F12 overnight. The cells were treated with 200 μM of H2O2 for 1 h before extraction of aqueous metabolites by 80% methanol from cell lysates for LC-MS/MS-based metabolomics. The overall metabolic profile differences between ctrl and H2O2-treated cells are shown by the PCA score plot (A). The heatmap shows significant differences (n = 4, p < 0.05) in 35 metabolites between Ctrl and H2O2-treated cells (B). The altered metabolic pathways by H2O2 treatment are displayed with the x-axis representing the pathway impact score computed from pathway topological analysis and the y-axis being -log10 of the p-value obtained from the pathway enrichment analysis (C). Metabolic pathway analyses by MetaboAnalyst 5.0 software depict the impact of H2O2 treatment, with the x-axis, y-axis, color, or size of the circle reflecting the information as described in Figure 2C.
Antioxidants 12 00452 g003
Antioxidants 12 00452 g004 550
Figure 4. Effect of Cardioplegic Solutions on Metabolites Changed by H2O2 Treatment. AC 16 cells at 90% confluency were serum-starved in 0.5% FBS DMEM/F12 overnight. The cells were treated with HK, LK, DN, HTK, or CS for 3 h as described in the Section 2, followed by exposure to 200 μM of H2O2 for 1 h before extraction of aqueous metabolites by 80% methanol for LC-MS/MS-based metabolomics. K-means clustering shows the overall metabolic profile altered by H2O2 and pretreatment with each cardioplegia solution (A, n = 4). The heatmap displays the metabolites in the 9 metabolic pathways significantly altered by H2O2 treatment for all groups of comparisons (B, n = 4).
Figure 4. Effect of Cardioplegic Solutions on Metabolites Changed by H2O2 Treatment. AC 16 cells at 90% confluency were serum-starved in 0.5% FBS DMEM/F12 overnight. The cells were treated with HK, LK, DN, HTK, or CS for 3 h as described in the Section 2, followed by exposure to 200 μM of H2O2 for 1 h before extraction of aqueous metabolites by 80% methanol for LC-MS/MS-based metabolomics. K-means clustering shows the overall metabolic profile altered by H2O2 and pretreatment with each cardioplegia solution (A, n = 4). The heatmap displays the metabolites in the 9 metabolic pathways significantly altered by H2O2 treatment for all groups of comparisons (B, n = 4).
Antioxidants 12 00452 g004
Antioxidants 12 00452 g005 550
Figure 5. HK or LK Protects against H2O2 from Inducing Changes in the Metabolites. The metabolites altered by H2O2 treatment belonging to nicotinate and nicotinamide metabolism, TCA cycle, glycolysis/gluconeogenesis, and pentose phosphate pathway were analyzed for the protective effect of HK or LK cardioplegia solution. * indicates significant difference (n = 4, p < 0.05) between control and H2O2-treated cells, whereas # indicates significant difference (n = 4, p < 0.05) between HK or LK with H2O2 treatment in comparison with H2O2 treatment alone. These data were analyzed for the significant differences by one-way ANOVA using GraphPad Prism 9 software.
Figure 5. HK or LK Protects against H2O2 from Inducing Changes in the Metabolites. The metabolites altered by H2O2 treatment belonging to nicotinate and nicotinamide metabolism, TCA cycle, glycolysis/gluconeogenesis, and pentose phosphate pathway were analyzed for the protective effect of HK or LK cardioplegia solution. * indicates significant difference (n = 4, p < 0.05) between control and H2O2-treated cells, whereas # indicates significant difference (n = 4, p < 0.05) between HK or LK with H2O2 treatment in comparison with H2O2 treatment alone. These data were analyzed for the significant differences by one-way ANOVA using GraphPad Prism 9 software.
Antioxidants 12 00452 g005
Antioxidants 12 00452 g006 550
Figure 6. Effects of Cardioplegic Solutions on Cell Morphology and ROS Generation. AC16 cells were seeded in 6-well (A) or 96-well plates (B,C) for pretreatment with cardioplegic solutions for 3 h. After 1 h exposure to 200 μM H2O2, cell morphology was recorded under an inverted light microscope after fixing and staining in 0.1% Coomassie blue (A). Following pretreatment with a cardioplegic solution, the cells were incubated with 100 μM DCFH-DA for 30 min before recording ROS generation with 200 μM H2O2 treatment by a fluorescence microplate reader (B,C). The bar graphs represent the means ± SD from 4 replicates of one representative experiment (C). * indicates a significant difference (p < 0.05) of H2O2-treated cells from Ctrl or a cardioplegic-solution-treated cells, whereas # indicates a significant difference (p < 0.05) of Ctrl from DN-, HTK-, or CS-treated cells (B). Antioxidants 12 00452 i001 indicates significant difference from H2O2-treated cells (C). These data were analyzed for the significant differences by one-way ANOVA using GraphPad Prism 9 software.
Figure 6. Effects of Cardioplegic Solutions on Cell Morphology and ROS Generation. AC16 cells were seeded in 6-well (A) or 96-well plates (B,C) for pretreatment with cardioplegic solutions for 3 h. After 1 h exposure to 200 μM H2O2, cell morphology was recorded under an inverted light microscope after fixing and staining in 0.1% Coomassie blue (A). Following pretreatment with a cardioplegic solution, the cells were incubated with 100 μM DCFH-DA for 30 min before recording ROS generation with 200 μM H2O2 treatment by a fluorescence microplate reader (B,C). The bar graphs represent the means ± SD from 4 replicates of one representative experiment (C). * indicates a significant difference (p < 0.05) of H2O2-treated cells from Ctrl or a cardioplegic-solution-treated cells, whereas # indicates a significant difference (p < 0.05) of Ctrl from DN-, HTK-, or CS-treated cells (B). Antioxidants 12 00452 i001 indicates significant difference from H2O2-treated cells (C). These data were analyzed for the significant differences by one-way ANOVA using GraphPad Prism 9 software.
Antioxidants 12 00452 g006
Antioxidants 12 00452 g007 550
Figure 7. Lidocaine prevents ROS generation by H2O2. AC16 human cardiac myocytes were seeded in 96-well plates and were pretreated with indicated concentrations of lidocaine for 3 h before 30 min incubation with 10 μM DCFH-DA. H2O2 was added to the final 200 μM for recording the fluorescence of hydroxy radical formation. The bar graphs represent the means ± SD from 3 independent experiments. * indicates statistical significance (p < 0.05) from H2O2 -treated alone group as determined by two-way ANOVA.
Figure 7. Lidocaine prevents ROS generation by H2O2. AC16 human cardiac myocytes were seeded in 96-well plates and were pretreated with indicated concentrations of lidocaine for 3 h before 30 min incubation with 10 μM DCFH-DA. H2O2 was added to the final 200 μM for recording the fluorescence of hydroxy radical formation. The bar graphs represent the means ± SD from 3 independent experiments. * indicates statistical significance (p < 0.05) from H2O2 -treated alone group as determined by two-way ANOVA.
Antioxidants 12 00452 g007
Table
Table 1. Key Components of Common Cardioplegic Solutions.
Table 1. Key Components of Common Cardioplegic Solutions.
Ratio (Blood: Crystalloid) Con (mM)K+Ca2+Na+Mg2+MannitolLidocaineHistidine
Type
Blood
cardioplegia		4:1HK19.31.31477.0–0.085–
4:1LK9.30.81531.2–0.085–
1:4DN25.60.2152.615.83.60.444–
Crystalloid cardioplegia–* HTK100.02154––198
–** CS150.251001360–30
* HTK contains 2 mM tryptophan and 1 mM ketoglutarate; ** CS contains 20 mM glutamate and 3 mM glutathione. The concentrations were calculated on the basis of the dilution of HK, LK, and DN in DMEM/F12.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Diao, H.; Gu, H.; Chen, Q.M. Hyperkalemic or Low Potassium Cardioplegia Protects against Reduction of Energy Metabolism by Oxidative Stress. Antioxidants 2023, 12, 452. https://doi.org/10.3390/antiox12020452

AMA Style

Diao H, Gu H, Chen QM. Hyperkalemic or Low Potassium Cardioplegia Protects against Reduction of Energy Metabolism by Oxidative Stress. Antioxidants. 2023; 12(2):452. https://doi.org/10.3390/antiox12020452

Chicago/Turabian Style

Diao, Hongting, Haiwei Gu, and Qin M. Chen. 2023. "Hyperkalemic or Low Potassium Cardioplegia Protects against Reduction of Energy Metabolism by Oxidative Stress" Antioxidants 12, no. 2: 452. https://doi.org/10.3390/antiox12020452

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop