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Research ArticleCardiovascular

Protamine Sulfate Induces Mitochondrial Hyperpolarization and a Subsequent Increase in Reactive Oxygen Species Production

Rabia Ramzan, Susanne Michels, Petra Weber, Annika Rhiel, Marc Irqsusi, Ardawan J. Rastan, Carsten Culmsee and Sebastian Vogt
Journal of Pharmacology and Experimental Therapeutics August 2019, 370 (2) 308-317; DOI: https://doi.org/10.1124/jpet.119.257725

Abstract

Protamine sulfate (PS) is widely used in heart surgery as an antidote for heparin, albeit its pharmacological effects are not fully understood and applications are often accompanied by unwanted side effects. Here we show the effect of PS on mitochondrial bioenergetics profile resulting in mitochondrial reactive oxygen species (ROS) production. Polarographic measurements were performed in parallel to membrane potential and ROS measurements by FACS analyzer using tetramethylrhodamine ethyl ester and MitoSOX fluorescent dyes, respectively. PS inhibited intact rat heart mitochondrial respiration (stimulated by ADP) to 76% (P < 0.001) from the baseline of 51.6 ± 6.9 to 12.4 ± 2.3 nmol O2⋅min−1⋅ml−1. The same effect was found when respiration was inhibited by antimycin A (101.0 ± 8.9 vs. 38.0 ± 9.9 nmol O2 ⋅min−1⋅ml−1, P < 0.001) and later stimulated by substrates of cytochrome oxidase (CytOx) i.e., ascorbate and tetramethyl phenylene diamine, suggesting that PS exerted its effect through inhibition of CytOx activity. Furthermore, the inhibition of mitochondrial respiration by PS was concentration dependent and accompanied by hyperpolarization of the mitochondrial membrane potential (Δψm), i.e., 18% increase at 50 µg/ml and an additional 3.3% increase at 250 µg/ml PS compared with control. This effect was associated with a strong consequent increase in the production of ROS, i.e., 85% and 88.6% compared with control respectively. We propose that this excessive increase in ROS concentrations results in mitochondrial dysfunction and thus might relate to the “protamine reaction,” contributing to the development of various cardiovascular adverse effects.

Introduction

During cardiovascular surgical procedures, a number of drugs are administered depending on the patients’ condition and the surgical procedure. Among these, protamine sulfate (PS) is commonly used as an antidote for heparin. Being a polycationic peptide, it reverses the anticoagulant effects of the highly negatively charged heparin by forming a complex with heparin (Viaro et al., 2002).

Although studies on healthy volunteers who received PS intravenously showed rapid decrease in protamine blood concentration, no significant changes were found in heart rate, mean arterial blood pressure, or cardiac output (Butterworth et al., 2002). However, increasing evidence from clinical and experimental data indicated a direct influence of PS on the myocardial tissue. Negative cardiovascular side effects of PS have been described in human patients (Conahan et al., 1981; Shapira et al., 1982) as well as in animals (Goldman et al.,1969). The drug administration resulted in decrease of cardiac oxygen consumption (Sethna et al., 1982; Wakefield et al., 1990).

Further clinical studies have provided information on adverse conditions associated with PS, e.g., systemic hypotension, bradycardia, pulmonary artery hypertension or hypotension, thrombocytopenia, and leucopenia (Wakefield et al., 1992). It has been established that PS mediates dose- and time-dependent detrimental effects on cardiac function independent of the presence of heparin (Hendry et al., 1987; Wakefield et al., 1989b, 1990).

However, the detailed molecular effects of PS on the cardiovascular system are not well understood. Fadali et al. (1976) found that hindering possible potential pathways leading to bradycardia and hypotension (e.g., ganglionic and adrenal medullary pathway blockade with hexamethonium chloride; postganglionic parasympathetic with atropine sulfate; alpha and beta adrenergic receptor by phenoxybenzamine and propranolol, respectively; or depletion of endogenous histamine) were ineffective in abolishing the unwanted cardiovascular effects of PS. In the same approach, the group isolated the vascular tree from the heart during extracorporeal circulation. The findings of the study showed that hypotension and bradycardia were produced by a direct effect of protamine on the myocardium and peripheral vascular system (Fadali et al., 1976). Earlier in vitro studies have also shown that PS application resulted in impaired mitochondrial respiration (Popinigis, 1974; Kossekova et al., 1975). Similar results were obtained with in vivo studies performed with dogs (Wakefield et al., 1989), as well as rabbit and rat hearts (Wakefield et al., 1990, 1992; Halpern et al., 1997). A rapid decline in ATP levels particularly in the presence of heparin occurred within minutes after application of PS to the cultured bovine pulmonary artery endothelial cells (Wakefield et al., 1989). Corresponding to these reports, a direct effect of PS on the contractile performance of isolated cardiomyocytes was also described previously (Hird et al., 1994).

The interaction of this drug to basic mitochondrial metabolic processes has an elementary relevance to the pathophysiological understanding. A previous study exposed that the site of inhibition of mitochondrial respiration by PS might be localized between cytochromes c and aa3, since submitochondrial inside out particles were only slightly affected by protamine (Konstantinov, 1975). Therefore, we suppose an elementary influence of PS on the regulatory triangle between mitochondrial respiration, mitochondrial membrane potential, and ROS production by its direct interaction with cytochrome c oxidase (CytOx) activity. Thus, this study addresses the effect of PS on the link between cytochrome c oxidase activity, mitochondrial membrane potential, and ROS production, representing a mitochondrial functional triangle that clarifies the known toxic effects of PS on the myocardium.

Material and Methods

All animal experiments had local approval (EX-8-2018) and were performed conforming to the guidelines from Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes.

Mitochondria were isolated by standard procedures of isolation as described previously (Ramzan et al., 2017). Briefly, whole heart tissue was excised rapidly from male Wistar rats (300–400 g) after decapitation and placed in ice-cold isolation medium (250 mM sucrose, 10 mM HEPES, 1 mM EDTA, and 0.2% fatty acid-free BSA, pH 7.4, at 4°C), washed, and homogenized by a Teflon potter in 30 ml of ice-cold isolation medium. Homogenate was centrifuged at 2500 rpm for 10 minutes. Supernatant was carefully collected and centrifuged at 9000 rpm for 15 minutes. The mitochondrial pellet was resuspended in 5-ml of ice-cold isolation medium and centrifuged again at 9000 rpm for 10 minutes. Finally, the pellet was resuspended in 200 µl of isolation medium and stored on ice until further use.

Protein concentrations were estimated by the bicinchoninic acid assay determining the total concentration of protein. The assay relies on two reactions. First, the peptide bonds in protein reduce Cu2+ ions from the copper (II) sulfate to Cu+. The amount of Cu2+ reduced is proportional to the amount of protein present in the solution. Bicinchoninic acid chelate with each Cu+ ion forming purple-colored complexes that strongly absorb light at the wavelength of 562 nm. Bovine serum albumin was used as a standard. Stock concentrations of mitochondrial pellets were 29.4 ± 6.0 mg/ml (n = 5). Protamine sulfate from Herring source was purchased from Sigma-Aldrich.

Because tracking the changes in oxygen consumption is the easiest way to monitor drug alterations on mitochondrial function, we performed polarography oxygen consumption measurements of mitochondrial samples using a Clark-type oxygen electrode (Hansatech Oxygraph System, Norfolk, UK) at 25°C and standard atmospheric pressure (101.32 kPa) in respiration buffer (130 mM KCl, 3 mM HEPES, 0.5 mM EDTA, 2 mM KH2PO4, 0.5% BSA, pH 7.4 at room temperature).

In the measuring chamber, 10 µl of stock mitochondrial sample corresponding to 29.4 mg/ml protein was added to 490 µl of constantly stirred respiration buffer, giving a final concentration of 0.589 ± 0.12 mg/ml (n = 5 measurements). The oxygen consumption measurements were started, and substrates (Fig. 1) were added as described in the legends to the Fig. 2. Rates of oxygen consumption were calculated. The measured respiratory control ratios (RCR) were found to be 4.38 ± 0.49 (n = 5).

Fig. 1.
Fig. 1.

Simplified and modified scheme of the mitochondrial respiratory chain or electron transport chain with some inhibitors. ETC means a series of complexes (I–IV) that transfer electrons from electron donors to electron acceptors via redox reactions. Electron transfer is coupled with the transfer of protons (H+) across the mitochondrial inner membrane by complexes I, III, and IV only. This creates an electrochemical proton gradient that drives the synthesis of ATP through the ATP synthase (complex V). The ETC also includes peptides and enzymes, which are proteins or protein complexes essential for electron transport, i.e., Cyt b, c, and c1; coenzyme Q or ubiquinone, Fe–S proteins; FMN-flavin mononucleotide. Heme a is a coordination complex consisting of a macrocyclic ligand a porphyrin, chelating an iron atom. It is a part of the CytOx-holoenzyme complex. Heme a differs from heme a3 in a methyl side chain at ring position 8. It is oxidized to a formyl group and a hydroxyethylfarnesyl group. An isoprenoid chain has been attached to the vinyl side chain at ring position 2 of the iron tetrapyrrole heme. The complex IV contains two hemes, cytochrome a and cytochrome a3, and two copper centers, the CuA and CuB centers. The final acceptor of electrons in the ETC during aerobic respiration is molecular oxygen (dioxygen). Four electrons are transferred from the binding site at the CytOx to oxygen for water production. Antimycin A (in a frame) is a potent inhibitor of complex III, so that cytochrome c cannot shuttle electrons to the enzyme. External reductants like ascorbate and TMPD transfer electrons directly to complex IV and thus are used to test activity under “isolated” conditions. Protamine sulfate binds as a polycationic compound directly to the cytochrome c binding site and blocks electron transfer. In polarographic measurements, glutamate and malate are added to test mitochondrial respiratory function. The glutamate-aspartate antiporter imports glutamate from the cytosol into the matrix and exports aspartate from the matrix to the cytosol. Aspartate is converted by cytosolic aspartate aminotransferase to oxaloacetate. In the cytosol, malate dehydrogenase catalyzes the reaction of oxaloacetate and NADH to produce malate and NAD+. Herein, two electrons generated from NADH, and an accompanying H+, are attached to oxaloacetate to form malate. The malate-aspartate shuttle is a biochemical system for translocating electrons produced during glycolysis across the semipermeable inner membrane of the mitochondrion for oxidative phosphorylation in eukaryotes. These electrons enter the electron transport chain of the mitochondria via reduction equivalents to generate ATP. The shuttle system is required because the mitochondrial inner membrane is impermeable to NADH, the primary reducing equivalent of the electron transport chain. Therefore, malate transports reducing equivalents across the membrane. Finally, the malate-aspartate shuttle is sustaining the redox reactions, because NADH in the cytosol is oxidized to NAD+ and NAD+ in the matrix is reduced to NADH. The NAD+ in the cytosol can be reduced again by further glycolysis, and the NADH in the matrix can be used to pass electrons to the electron transport chain for ATP synthesis. The mitochondrial membrane potential (∆ψm) generated by proton pumps (Complexes I, III, and IV of the ETC) is an essential component in the process of energy storage during oxidative phosphorylation in the mitochondria. The sum of the membrane potential and the pH gradient are together known as the proton motive force (PMF). This indicates the total potential energy stored in the transmembrane gradients, which is available to drive protons back into the matrix space, and provide the power for biologically useful processes. PMF finally drives the ATP synthase to make ATP. Note, all ETC complexes (except complex II) participate in proton pumping to form ∆ψm. ATP, adenosine triphosphate; CN−, cyanide; CO, carbon monoxide; CuA, copper A; CuB, copper A; Cyt b, cytochrome b; Cyt c, cytochrome c; Cyt c1, cytochrome c1; CytOx, cytochrome c oxidase; ETC, electron transport chain; FAD, flavin adenine dinucleotide; FADH2, flavin adenine dinucleotide (reduced); Fe–S, iron–sulfur proteins; FMN, flavin mononucleotide; H2S, hydrogen sulfide; NAD, nicotinamide adenine dinucleotide; NADH, nicotinamide adenine dinucleotide (reduced); Q, Coenzyme Q or ubiquinone; TCA cycle, tricarboxylic acid cycle; TMPD, tetramethyl phenylene diamine.

Fig. 2.
Fig. 2.

(A–D) Measurements of mitochondrial oxygen consumption by polarography. Original protocols of oxygen measurements are shown on the left side and combined data of the same time course with statistical analysis (gray arrow at x-axis) are on the right. Freshly isolated rat heart mitochondria (0.589 ± 0.12 mg/ml) were added to the total volume of 500 µl respiratory buffer in the oxygraph chamber and measurements were started after closing the chamber. Substrates were added during each measurement as indicated. (A) Oxygen consumption measurements for calculating respiratory control ratio (n = 5). Mitochondrial preparation with a high respiratory control ratio (RCR) of at least 4.38 ± 0.49 indicates the higher intactness and good quality of mitochondria after isolation. (B) Inhibition of ADP stimulated respiration (state III) by protamine sulfate (n = 5). Addition of 250 µg/ml PS to the isolated mitochondria at state III, i.e., ADP-stimulated oxygen consumption state, strongly inhibits the rate from 51.6 ± 6.9 to 12.4 ± 2.3 nmol O2⋅min−1⋅ml−1, corresponding to approximately 76% reduction in oxygen consumption (P < 0.001 calculated through standard one-way ANOVA analysis). Furthermore, the PS inhibition of state III respiration was not released by the addition of 4 µM CCCP. (C) Dose-dependent effect of PS on ADP stimulated (state III) respiration (n = 5). The PS effect was found to be dose dependent. A direct correlation was found between the inhibition rate of mitochondrial oxygen consumption and the concentration of PS showing that mitochondrial respiration rate was reduced to half when doubling the concentration of the drug. (D) Effect of protamine sulfate on oxygen consumption activity of CytOx in isolated intact rat heart mitochondria (n = 5). The specific action of PS is related to the inhibition of cytochrome c oxidase (CytOx) activity. The antimycin A-inhibited mitochondrial respiration was stimulated by adding the substrates for CytOx, i.e., ascorbate and TMPD. Here, the electron transport to CytOx was stopped by adding 250 µg/ml PS, which clearly decreased the mitochondrial oxygen consumption rate in the presence of ascorbate and TMPD: 101.02 ± 8.9 to 38.0 ± 9.9 nmol O2⋅min−1⋅ml−1 after PS addition. AA, antimycin A; ADP, adenosine diphosphate; Asc, ascorbate; CCCP, carbonyl cyanide m-chlorophenyl hydrazone; Glu, glutamate; Mal, malate; PS, protamine sulfate; RHM, rat heart mitochondria; State II, substrate-induced mitochondrial respiration; State III, ADP-stimulated mitochondrial respiration; State IV, ADP completely phosphorylated to form ATP; TMPD, tetramethyl phenylene diamine (***P < 0.001; **P < 0.01; *P < 0.05 calculated through standard one-way ANOVA).

Mitochondrial membrane potential and ROS production were measured by flow cytometry. Freshly isolated rat heart mitochondria were resuspended in respiration buffer at a concentration of 0.589 ± 0.12 mg/ml. Substrates were added as mentioned above to a total volume of 500 µl respiratory buffer. Finally, samples were successively stained with the corresponding fluorescent dyes and measured immediately.

Mitochondrial reactive oxygen species were detected with 2.5 µM MitoSOX red indicator (Thermo Fisher Scientific, Darmstadt, Germany). Changes in mitochondrial membrane polarization were determined using 150 nM tetramethylrhodamine ethyl ester (TMRE; Thermo Fisher Scientific). Samples were measured with a total count of 50,000 events utilizing a Guava easyCyte 6-2L flow cytometer (Merck Millipore, Darmstadt, Germany) while data analysis and gating were performed with GuavaSoft 3.1.1 software. Data were plotted as dot plots using GraphPad Prism version 5.0 (GraphPad Software Inc., La Jolla, CA). Results were calculated as mean ± S.D. Finally, statistical analysis, including regression analysis, a standard one-way ANOVA, and post hoc Kruskal-Wallis tests were performed by using IBM SPSS Statistics 21.

Results

Determining the Quality of Isolated Mitochondrial Preparations by Cytochrome c Test.

Immediately after isolation, the quality of fresh mitochondrial preparations was assessed by performing cytochrome c test as previously described (Ramzan et al., 2017). The study of this basic parameter confirmed the intactness and quality of mitochondrial preparations. The freshly isolated mitochondrial samples always showed minimal stimulation at 10 µM cytochrome c, indicating a high integrity of the outer mitochondrial membrane. The increase in oxygen consumption rate after 10 μM cytochrome c addition in four different mitochondrial preparations was 9.0 ± 2.6 nmol O2⋅min−1⋅ml−1.

Determining the Quality of Isolated Mitochondrial Preparations by RCR Measurements.

Next, mitochondrial oxygen consumption was measured to assess the respiratory control ratio, which is considered as a standard measure of coupling between mitochondrial respiration and oxidative phosphorylation. This is another widely established parameter for the measurement of mitochondrial intactness and is obtained by dividing the oxidation rate calculated at state III (high steady-state increase in oxygen consumption rate by ADP in the presence of respiratory substrates, e.g., glutamate, malate) to the rate measured at state IV, which occurs when all the added ADP is converted to ATP and respiration spontaneously and markedly decreases to a slower rate, which is also nearly constant. Our mitochondrial preparations retained a respiratory control ratio (RCR) of at least 4.38 ± 0.49 (see example of the original data in Fig. 2A, left side on top), indicating the desired intactness and quality. The Kruskal-Wallis test showed statistically significant differences in oxygen consumption between different groups, χ2(4) = 21.313, P = 0.000, with a mean rank score of 3.00 for rat heart mitochondria (RHM), 8.80 for Glu+Mal, 20.60 for ADP, 12.20 for State IV, and 20.40 for carbonyl cyanide m-chlorophenyl hydrazine (CCCP).

Inhibition of Mitochondrial Respiration by PS.

To verify the effect of PS on mitochondrial respiration, 250 µg/ml PS was added, which strongly inhibited oxygen consumption rate at state III, i.e., ADP-stimulated oxygen consumption (Fig. 2B) from 51.6 ± 6.9 to 12.4 ± 2.3 nmol O2⋅min−1⋅ml−1, corresponding to approximately 76% reduction (P < 0.001). Interestingly, the inhibition of state III respiration by PS was not released by the addition of 4 µM CCCP (Fig. 2B). The Kruskal-Wallis test showed statistically significant differences in oxygen consumption between different groups, χ2(4) = 19.176, P = 0.001, with a mean rank score of 3.00 for RHM, 15.20 for Glu+Mal, 23 for ADP, 12.30 for PS, and 11.50 for CCCP.

Dose-Dependent Inhibition of Mitochondrial Respiration by PS.

Corresponding to clinically used concentrations of PS that induced adverse reactions on contractility in mammalian heart preparations or isolated myocardium (Hendry et al., 1987; Wakefield et al., 1990, 1992; Park et al., 1994), we performed titrations of PS at increasing concentrations beginning at 25 µg/ml followed by 50, 100, 200, and up to the higher concentration of 250 µg/ml (Fig. 2C) after inducing State III respiration.

The PS effect was found to be dose dependent. In fact, a direct correlation was found between the inhibition rate of mitochondrial oxygen consumption and the concentration of PS showing that mitochondrial respiration rate was reduced to half while doubling the concentration of the drug.

A simple linear regression analysis was performed to predict that an increase in PS decreases the consumption of oxygen. A significant regression equation was found, F(1,23) = 130.637, P < 0.000, with an R2 of 0.850. Furthermore, statistically significant differences were found by the Kruskal-Wallis test between different groups, χ2(4) = 36.610, P = 0.000, with a mean rank score of 5.90 for RHM, 24.70 for Glu+Mal, 38 for ADP, 33 for 25 µg/ml PS, 26.30 for 50 µg/ml PS, 17.60 for 100 µg/ml PS, 11.70 for 200 µg/ml PS, and 6.80 for 200 µg/ml PS.

CytOx Inhibition by PS.

It has already been described that PS acts at a target structure between cytochrome c and cytochrome aa3 (Konstantinov, 1975), thereby inhibiting cytochrome c oxidase (CytOx) activity. To assess this direct interaction of PS with CytOx alone and not to other members of the electron transport chain (ETC), we measured the effect of the drug on mitochondrial respiration in the presence of antimycin A, which binds to complex III, thus blocking the electron transfer between Complex III and IV (see Fig. 1). To specify PS action, we stimulated this antimycin A-inhibited respiration of mitochondria by adding the substrates for CytOx, i.e., ascorbate and TMPD, which directly transfers electrons to cytochrome c (Fig. 2D). Electron transport to CytOx was stopped, and mitochondrial oxygen consumption rate was clearly decreased when 250 µg/ml PS was added (respiration after AA: 6.9 ± 1.0 nmol O2⋅min−1⋅ml−1, ascorbate and TMPD: 101.02 ± 8.9 nmol O2⋅min−1⋅ml−1, after PS: 38.0 ± 9.9 nmol O2⋅min−1⋅ml−1, respectively). Statistically significant differences were found by the Kruskal-Wallis test between different groups, χ2(4) = 32.130, P = 0.000, with a mean rank score of 3.00 for RHM, 13 for Glu+Mal, 27 for ADP, 8 for antimycin (AA), 33 for ascorbate + TMPD, 21.60 for PS, and 20.40 for CCCP.

Mitochondrial Hyperpolarization and Stimulation of ROS Production by PS.

For establishing the link of PS-induced inhibition of mitochondrial respiration to another important mitochondrial basic process, i.e., membrane potential and consequent ROS generation, we used two different concentrations of the drug i.e., 50 (minimum) and 250 µg/ml (maximum) and correlated these with the control data (Fig. 3). Clearly, membrane potential (∆ψm) increased from 71.31% ± 2.14% (Control) to 86.97% ± 1.78% and 90.58% ± 1.89% of relative TMRE fluorescence (Fig. 3, A and B). Correspondingly, ROS production (measured as relative MitoSOX fluorescence) was also increased from 5.98% ± 1.80% (control) to 39.91% ± 15.6% in the presence of 50 µg/ml PS and 52.41% ± 14.5% in case of 250 µg/ml PS (Fig. 3, C and D). It is evident that PS addition resulted in marked increase in membrane potential as well as ROS production (P < 0.001 and P < 0.01 compared with controls). In addition, we measured membrane potential and ROS in correlation to state II (induced by the addition of substrates for complex I: glutamate + malate) and state III (generated by the addition of ADP in the presence of complex I substrates) conditions of mitochondria (Fig. 4). Initially, addition of ADP in the presence of substrates initiated oxidative phosphorylation and resulted in an increased mitochondrial respiration as described earlier (Fig. 2C), followed by partial depolarization of the membrane potential (Fig. 4, A and B) and low ROS (Fig. 4, C and D). When PS was added at state III of the mitochondria, again an increase in membrane potential (glutamate+malate = 69.66% ± 6.54% and glutamate+malate+ADP = 64.21% ± 4.72% vs. glutamate+malate+ADP+50 µg/ml PS = 83.03% ± 6.72% and glutamate+malate+ADP+250 µg/ml PS = 86.15% ± 4.31% relative TMRE fluorescence) as well as an increase in ROS production was measured (glutamate+malate = 3.93% ± 2.71% and glutamate+malate+ADP = 3.83% ± 1.93% vs. glutamate+malate+ADP+50 µg/ml PS = 19.73% ± 5.33% and glutamate+malate+ADP+250 µg/ml PS = 29.64% ± 6.08% relative MitoSOX fluorescence). This increased production of ROS after PS exposure appeared to be dose dependent (P < 0.001).

Fig. 3.
Fig. 3.

(A–D) Measurements of mitochondrial membrane potential (A and B) and ROS production (C and D) by flow cytometry at two different concentrations of PS in comparison with the control. (A and C) Respective histograms with gating of one representative measurement. (B and D) Percentage of mitochondria within the gate of n = 5 independent experiments. Freshly isolated rat heart mitochondria were resuspended in 500 µl of respiration buffer at a concentration of 0.589 ± 0.12 mg/ml. Substrates were added as indicated. Finally, fluorescence was measured immediately after adding 150 nM TMRE to the samples. For mitochondrial reactive oxygen species (ROS) measurements, 2.5 µM MitoSOX was added to the samples. Measurements were performed for n = 5 experiments. The data are shown as mean ± S.D. Distinctly, addition of PS even at 50 µg/ml concentration has increased mitochondrial membrane potential (∆ψm) from 71.65% ± 1.33% (Control) to 86.72% ± 2.31%, which stimulated further to 91.10% ± 2.1% of relative TMRE fluorescence at 250 µg/ml PS. Correspondingly, ROS production was also increased from 6.55% ± 0.34% (Control) to 48.31% ± 14.7% and 62.17% ± 8.19% relative MitoSOX fluorescence at the same two PS concentrations, respectively. Ctr, control; PS, protamine sulfate. (***P < 0.001; **P < 0.01; *P < 0.05 calculated through standard one-way ANOVA).

Fig. 4.
Fig. 4.

(A–D) Measurements of mitochondrial membrane potential (A and B) and ROS production (C and D) by flow cytometry. (A and C) Respective histograms with gating of one representative measurement. (B and D) Percentage of mitochondria within the gate of n = 5 independent experiments. Mitochondrial membrane potential and ROS measurements at state II, state III, and at two different concentrations of PS added at state III. Addition of ADP in the presence of glutamate and malate initiated oxidative phosphorylation and resulted in an increased mitochondrial respiration followed by partial depolarization of the membrane potential and consequently low ROS. Addition of PS at mitochondrial state III again resulted in an increased membrane potential and increase in ROS (∆ψm: 65.6% ± 4.0% and 62.0% ± 3.8% vs. 82.5% ± 9.1% and 86.5% ± 6.0% of relative TMRE fluorescence; ROS: 2.3% ± 2.2% and 3.2% ± 2.4% vs. 16.7% ± 4.4% and 26.5% ± 6.0% relative MitoSOX fluorescence). ADP, adenosine diphosphate; Glu, glutamate; Mal, malate; PS, protamine sulfate (***P < 0.001; **P < 0.01; *P < 0.05 calculated through standard one-way ANOVA).

A simple linear regression analysis was performed to predict that addition of PS increases the mitochondrial membrane potential as well as ROS production. Significant regression equations were found in both cases: F(1,13) = 72.378, P < 0.000, with an R2 of 0.848 for mitochondrial hyperpolarization, and F(1,13) = 31.746, P < 0.000, with an R2 of 0.709 for ROS production.

Additionally simple linear regression analysis was performed to predict that an increase in PS increases the mitochondrial membrane potential as well as ROS production. Significant regression equations were found in these cases as well: F(1,13) = 28.435, P < 0.000, with an R2 of 0.686 for membrane potential, and F(1,13) = 70.586, P < 0.000, with an R2 of 0.844 for ROS production. The Kruskal-Wallis test showed statistically significant differences among different groups at the P < 0.000 level.

Mitochondrial Hyperpolarization and Stimulation of ROS Production by PS after CytOx Inhibition.

To evaluate the direct effect of PS on CytOx under substrate stimulation, mitochondrial respiratory chain was blocked at complex III (bc1-complex) by 2.5 µM antimycin A, which distinctly decreased the membrane potential (glutamate+malate+ADP = 64.21% ± 4.73% vs. glutamate+malate+ADP+AA = 39.62% ± 9.89% relative TMRE fluorescence). Under these conditions of inhibited respiratory chain, CytOx (Complex IV) was stimulated by the addition of its substrates, i.e., ascorbate and TMPD. Corresponding to previous results, when PS was added to the mitochondria in the presence of glutamate+malate+ADP+AA+ascorbate+TMPD, a significant increase in the membrane potential (from 49.96% ± 11.22% to 77.32% ± 3.47% of relative TMRE fluorescence) as well as in the ROS production (from 6.19% ± 4.22% to 23.60% ± 8.79% relative MitoSOX fluorescence) was found (Fig. 5). Statistically significant differences were found among different groups at P < 0.000 level by the Kruskal-Wallis test.

Fig. 5.
Fig. 5.

(A–D) Measurements of mitochondrial membrane potential (A and B) and ROS production (C and D) by flow cytometry. (A and C) Respective histograms with gating of one representative measurement. (B and D) Percentage of mitochondria within the gate of n = 5 independent experiments. Comparison of mitochondrial membrane potential and ROS measurements after inhibiting the respiratory chain by antimycin A, followed by the addition of substrates (ascorbate +TMPD) for CytOx in the presence and absence of 250 µg/ml PS. Once again, the addition of PS under these conditions as well, significantly increased both the membrane potential and ROS production. AA, antimycin A; ADP, adenosine diphosphate; Asc, ascorbate; Glu, glutamate; Mal, malate; PS, protamine sulfate; ROS, reactive oxygen species; TMPD, tetramethyl phenylene diamine (***P < 0.001, calculated through standard one-way ANOVA).

Discussion

Protamine sulfate is a small polycationic protein originally isolated from the sperm of salmon (Bolan and Klein, 2013). Usually the recommended dose is 1 mg of protamine to antagonize 100 Units of heparin. Other than some difficulties in calculations, protamine dosage is usually based on body weight (3 to 4 mg/kg) or in a ratio to the heparin dosage (Lindblad, 1989; DiNardo et al., 2011) and depends on the time elapsed since the heparin infusion was discontinued. We adjusted our PS concentrations according to the previous studies (Hendry et al., 1987; Wakefield et al., 1990, 1992; Park et al., 1994), so our experimental conditions were relevant to the clinical scenario. Although it is routinely used, the optimal rate of drug administration is also a matter of consideration (Kumar and Cucchiara, 2013).

Concerning the potential cytotoxic effects of PS, the heart and certain blood cells (white blood cells and platelets) appear to be a relevant target for this widely used pharmacological compound. Notably, the heart appears to be the primary target for protamine since it contains a high number of mitochondria. Heart energy supply relies mainly on fat metabolism (Lemieux and Hoppel, 2009), and mitochondrial dysfunction is one of the major causes for heart failure (Rosca and Hoppel, 2013). Excessive production of reactive oxygen species (ROS) plays an important role in cellular injury (Knowlton et al., 2014), e.g., in cardiac arrhythmias (Jeong et al., 2012). Mitochondria are the main source of ROS generation in the cell (Suski et al., 2012), which participate in the process of ischemia and reperfusion, hence contribute to the development of myocardial infarction and stroke (Kalogeris et al., 2014). Among various forms of ROS, the superoxide anion is considered to be the most frequent of mitochondrial ROS (Murphy, 2009).

The central parameter that controls the generation of reactive oxygen species in mitochondria remains to be the membrane potential (Turrens, 2003; Nicholls, 2004). An exponential increase in the production of ROS has been measured in mitochondria with a membrane potential of >140 mV (Korshunov et al., 1997). An increase in membrane potential could be a result of ATP synthase inhibition or closure of the mitochondrial permeability transition pore (Wojtczak et al., 1999). It has been reported that mitochondria show significant ROS production when they are not synthesizing ATP and consequently have a high membrane potential and an increased pool of reduced coenzyme Q (Murphy, 2009).

Data from previous studies indicate that protamine affects mitochondria at the membrane level (Popinigis et al., 1971). Although a biphasic effect of protamine on intact mitochondria has been reported previously, stating stimulation of respiration at lower concentrations while inhibition of respiration at higher concentrations, this biphasic effect was found to be mainly dependent on the media used to study mitochondrial respiration (Popinigis, 1974; Kossekova et al., 1975). Since we used nearly physiologic KCl buffer, which is routinely used for oxidative phosphorylation studies, in our measurements we found a dose-dependent inhibitory effect of the drug on mitochondrial respiration (Fig. 2C) conforming to previous studies (Person and Fine, 1961) performed with fresh rat heart homogenate. Here, we found that the respiration was inhibited even at low concentrations of the drug, and the extent of this inhibition was increased at increasing concentrations of PS. This could indeed explain the known side effects of the drug on energy metabolism and cardiovascular functions (Horrow, 1985). Figure 2D shows that protamine sulfate inhibits mitochondrial respiration by inhibiting the activity of cytochrome c oxidase as reported previously (Person and Fine, 1961).

Additionally, structural changes in mitochondria may be associated with the effects of PS, since contraction of intermembrane space together with swollen cristae and extreme vacuolization were observed after incubation of mitochondria with the drug (Popinigis et al., 1971; Wakefield et al., 1990). These structural changes might result in desorption of cytochrome c from the cristae of the inner mitochondria membrane in the presence of protamine, thus blocking oxidative phosphorylation (Kossekova et al., 1975). Moreover, the measured membrane hyperpolarization in the presence of PS independent of different experimental conditions (Fig. 3B; Fig. 4B; Fig. 5B) and concomitant inhibition of mitochondrial respiration was not released by CCCP (Fig. 2, B and D), which may be due to the fact that protamine blocks proton leakage and inhibits other exchange diffusion barriers, in turn building up a high pH gradient between intramembrane space and matrix with an apparent pronounced increase in pH within the mitochondrial matrix (Popinigis et al., 1970). Consequently, all the complexes of the ETC except CytOx are reduced. Furthermore, cytochrome c, which is not involved in electron transport because of PS action could serve as an amplifier of ROS production (Akopova et al., 2012).

Deterioration of metabolism results in the reduction of blood supply (Nicholls, 2004), which causes slower heart beat (bradycardia) and low blood pressure (systemic or pulmonary artery hypotension). Contrarily an increased blood pressure within the arteries of the lungs results in pulmonary artery hypertension where mitochondrial dysfunction plays an essential role in the pathogenesis of the disease (Freund-Michel V et al., 2014). Overall, all these factors are the major hallmarks of PS unwanted side effects on the cardiovascular system (Fig. 6). The combination of reduced ATP synthesis together with excessive formation of ROS eventually results in heart failure (Sabbah, 2016). Correspondingly, protamine-induced thrombocytopenia (decreased number of blood platelets) and leucopenia (decreased number of white blood cells) have already been reported (Al-Mondhiry et al., 1985). In the case of blood platelets, it has been described that an increase in ROS production leads to thrombus formation (Zharikov and Shiva, 2013), which in turn may lead to thrombocytopenia.

Fig. 6.
Fig. 6.

Proposed mechanism of protamine sulfate- induced cardiovascular adverse reactions. After administration into the blood circulation, PS forms a complex with heparin depending on the dosage ratio. Since the mechanism of elimination or metabolic pathway of PS is not known yet, it is suggested that the rest of the drug that is not bound to heparin easily enters into mitochondria because of its chemical properties, i.e., being highly basic small molecular size protein (approximately 4500 Da). Once accumulating inside the mitochondrial intermembrane space, PS binds between the binding site of cytochrome c and cytochrome a3 at the outer surface of the inner mitochondrial membrane, thereby inhibiting mitochondrial respiration. At the same time, ETC remains highly reduced. Consequently, an increased membrane potential is maintained through reduced complexes I and III, while simultaneously electrons may leak from the ETC and react with dioxygen contributing to the generation of ROS (Murphy,Spacebar2009). Excessive ROS production at high membrane potential (Turrens, 2003) is followed by slowed or inhibited oxidative phosphorylation in mitochondria. These excessive ROS act as signaling molecules to start various pathways in the heart as well as in white blood cells and platelets (the only blood cells containing/harboring mitochondria). Concomitantly, although the membrane potential is maintained high, it cannot be used by ATP synthase to produce ATP because of its limited capacity since the respiration is inhibited. Finally, this results in proton and electron leakage, membrane disruption, and energy deficiency. In conclusion, inhibition of mitochondrial respiration by protamine sulfate at the site of cytochrome oxidase results in the inhibition of oxidative phosphorylation on one side and an increase in the reduced components of prior electron transport chain by electron transmission on the other side, followed by an increase in membrane potential and consequent increase in ROS production. The decrease in ATP levels and chronic increase in ROS production contribute to the development of drug related adverse conditions, which ultimately may lead to the reduction of myocardial contractility. ATP, adenosine triphosphate; ETC, electron transport chain; OxPhos, oxidative phosphorylation; ROS, reactive oxygen species.

Finally, the inhibitory effect of PS on mitochondrial respiration not only affects the mitochondrial membrane potential but also the redox state of mitochondria to such an extent that both are strongly increased while the mitochondrial respiration remains inhibited between the cytochrome c and cytochrome a binding site. Extensive increase in ROS levels results in oxidative stress. Correspondingly, monitoring the redox status of patients before and after cardiovascular surgical procedures by determining redox parameters, i.e., total antioxidant status, total glutathione concentrations, glutathione peroxidase activity, and both nitrite and nitrate concentrations in the peripheral blood of patients, can be of great impact (Schuh et al., 2018) for optimal treatment in the postoperative time period. In future clinical studies, when PS is used, a combination of mitochondria-specific antioxidants or ROS scavengers to reverse the side effect of PS should be evaluated. Nevertheless, in clinical routine, measures should be taken to anticipate and attenuate the potential risks of PS-induced oxidative stress before its use in relation to mitochondrial bioenergetics profile.

Acknowledgments

The authors thank Muhammad Mudassir Iqbal and Muhammad Shahbaz Aslam for their help and support in statistical analysis.

Authorship Contributions

Participated in research design: Ramzan, Vogt.

Conducted experiments: Ramzan, Michels, Weber, Rhiel.

Performed data analysis: Ramzan, Rastan, Vogt.

Wrote or contributed to the writing of manuscript: Ramzan, Michels, Irqsusi, Rastan, Culmsee, Vogt.

Footnotes

Abbreviations

AA
antimycin A
CCCP
carbonyl cyanide m-chlorophenyl hydrazine
CytOx
cytochrome c oxidase
ETC
electron transport chain
PS
protamine sulfate
RCR
respiratory control ratios
RHM
rat heart mitochondria
ROS
reactive oxygen species
TMPD
tetramethyl phenylene diamine
TMRE
tetramethylrhodamine ethyl ester

References

  1. ↵
    1. Akopova OV,
    2. Kolchinskaya LI,
    3. Nosar VI,
    4. Bouryi VA,
    5. Mankovska IN, and
    6. Sagach VF
    (2012) Cytochrome C as an amplifier of ROS release in mitochondria. Fiziol Zh 58:3–12.
    OpenUrl
  2. ↵
    1. Al-Mondhiry H,
    2. Pierce WS, and
    3. Basarab RM
    (1985) Protamine-induced thrombocytopenia and leukopenia. Thromb Haemost 53:60–64.
    OpenUrlPubMed
  3. ↵
    1. Kitchens C,
    2. Konkle B, and
    3. Kessler C
    1. Bolan CD and
    2. Klein HG
    (2013) Blood component and pharmacologic therapy for hemostatic disorders, in Consultative Hemostasis and Thrombosis Konkle, 3rd ed (Kitchens C, Konkle B, and Kessler C eds) pp 496–525, W.B. Saunders, Philadelphia.
  4. ↵
    1. Butterworth J,
    2. Lin YA,
    3. Prielipp R,
    4. Bennett J, and
    5. James R
    (2002) The pharmacokinetics and cardiovascular effects of a single intravenous dose of protamine in normal volunteers. Anesth Analg 94:514–522.
    OpenUrlCrossRefPubMed
  5. ↵
    1. Conahan TJ III.,
    2. Andrews RW, and
    3. MacVaugh H III.
    (1981) Cardiovascular effects of protamine sulfate in man. Anesth Analg 60:33–36.
    OpenUrlPubMed
  6. ↵
    1. Davis P,
    2. Cladis F, and
    3. Motoyama E
    1. DiNardo JA,
    2. Shukla AC, and
    3. McGowan FX Jr.
    (2011) Anesthesia for congenital heart surgery, in Smith’s Anesthesia for Infants and Children, 8th ed (Davis P, Cladis F, and Motoyama E eds) pp 605–673, Mosby, Philadelphia.
  7. ↵
    1. Fadali MA,
    2. Papacostas CA,
    3. Duke JJ,
    4. Ledbetter M, and
    5. Osbakken M
    (1976) Cardiovascular depressant effect of protamine sulphate: experimental study and clinical implications. Thorax 31:320–323.
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Freund-Michel V,
    2. Khoyrattee N,
    3. Savineau JP,
    4. Muller B, and
    5. Guibert C
    (2014) Mitochondria: roles in pulmonary hypertension. Int J Biochem Cell Biol 55:93–97.
    OpenUrlCrossRefPubMed
  9. ↵
    1. Goldman BS,
    2. Joison J, and
    3. Austen WG
    (1969) Cardiovascular effects of protamine sulfate. Ann Thorac Surg 7:459–471.
    OpenUrlCrossRefPubMed
  10. ↵
    1. Halpern P,
    2. Hochhauser E,
    3. Puzhevsky A,
    4. Rudick V,
    5. Sorkine P, and
    6. Vidine B
    (1997) The isolated post ischaemic rat heart is more vulnerable to protamine sulphate than the non-ischaemic heart. Cardiovasc Surg 5:235–240.
    OpenUrlCrossRefPubMed
  11. ↵
    1. Hendry PJ,
    2. Taichman GC, and
    3. Keon WJ
    (1987) The myocardial contractile responses to protamine sulfate and heparin. Ann Thorac Surg 44:263–268.
    OpenUrlCrossRefPubMed
  12. ↵
    1. Hird RB,
    2. Crawford FA Jr.,
    3. Mukherjee R,
    4. Zile MR, and
    5. Spinale FG
    (1994) Effects of protamine on myocyte contractile function and beta-adrenergic responsiveness. Ann Thorac Surg 57:1066–1074; discussion 1074–1075.
    OpenUrlCrossRefPubMed
  13. ↵
    1. Horrow JC
    (1985) Protamine: a review of its toxicity. Anesth Analg 64:348–361.
    OpenUrlPubMed
  14. ↵
    1. Jeong EM,
    2. Liu M,
    3. Sturdy M,
    4. Gao G,
    5. Varghese ST,
    6. Sovari AA, and
    7. Dudley SC Jr.
    (2012) Metabolic stress, reactive oxygen species, and arrhythmia. J Mol Cell Cardiol 52:454–463.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Kalogeris T,
    2. Bao Y, and
    3. Korthuis RJ
    (2014) Mitochondrial reactive oxygen species: a double edged sword in ischemia/reperfusion vs preconditioning. Redox Biol 2:702–714.
    OpenUrl
  16. ↵
    1. Knowlton AA,
    2. Chen L, and
    3. Malik ZA
    (2014) Heart failure and mitochondrial dysfunction: the role of mitochondrial fission/fusion abnormalities and new therapeutic strategies. J Cardiovasc Pharmacol 63:196–206.
    OpenUrl
  17. ↵
    1. Konstantinov AL
    (1975) [Protamine as a specific inhibitor of electron transport between cytochromes c and a on the outer surface of the mitochondrial membrane]. Biokhimiia 40:401–407.
    OpenUrlPubMed
  18. ↵
    1. Korshunov SS,
    2. Skulachev VP, and
    3. Starkov AA
    (1997) High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria. FEBS Lett 416:15–18.
    OpenUrlCrossRefPubMed
  19. ↵
    1. Kossekova GP,
    2. Mitovska MI, and
    3. Dancheva KI
    (1975) Protamine inhibition of the oxidative phosphorylation in intact, cytochrome c-depleted and restored mitochondria. Acta Biol Med Ger 34:539–547.
    OpenUrlPubMed
  20. ↵
    1. Kitchens C,
    2. Konkle B, and
    3. Kessler C
    1. Kumar M and
    2. Cucchiara B
    (2013) Hematologic interventions for acute central nervous system disease, in Consultative Hemostasis and Thrombosis Konkle, 3rd ed (Kitchens C, Konkle B, and Kessler C eds) pp 737–753, W.B. Saunders, Philadelphia.
  21. ↵
    1. Lemieux H and
    2. Hoppel CL
    (2009) Mitochondria in the human heart. J Bioenerg Biomembr 41:99–106.
    OpenUrlCrossRefPubMed
  22. ↵
    1. Lindblad B
    (1989) Protamine sulphate: a review of its effects: hypersensitivity and toxicity. Eur J Vasc Surg 3:195–201.
    OpenUrlPubMed
  23. ↵
    1. Murphy MP
    (2009) How mitochondria produce reactive oxygen species. Biochem J 417:1–13.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Nicholls DG
    (2004) Mitochondrial membrane potential and aging. Aging Cell 3:35–40.
    OpenUrlCrossRefPubMed
  25. ↵
    1. Park WK,
    2. Pancrazio JJ, and
    3. Lynch C III.
    (1994) Mechanical and electrophysiological effects of protamine on isolated ventricular myocardium: evidence for calcium overload. Cardiovasc Res 28:505–514.
    OpenUrlCrossRefPubMed
  26. ↵
    1. Person P and
    2. Fine AS
    (1961) Reversible inhibitions of cytochrome system components by macromolecular polyions. Arch Biochem Biophys 94:392–404.
    OpenUrlPubMed
  27. ↵
    1. Popinigis J
    (1974) The problem of permeability barrier in mitochondrial respiration: dual effect of protamine on succinate (plus rotenone) oxidation. FEBS Lett 41:46–49.
    OpenUrlPubMed
  28. ↵
    1. Popinigis J,
    2. Rzeczycki W, and
    3. Swierczyński J
    (1970) The effect of some proton conducting systems on protamine inhibited respiration. FEBS Lett 8:149–152.
    OpenUrlPubMed
  29. ↵
    1. Popinigis J,
    2. Takahashi Y,
    3. Wakabayashi T,
    4. Hull RM, and
    5. Williams CH
    (1971) Some aspects of mitochondrial structure. FEBS Lett 19:221–224.
    OpenUrlPubMed
  30. ↵
    1. Ramzan R,
    2. Schaper AK,
    3. Weber P,
    4. Rhiel A,
    5. Siddiq MS, and
    6. Vogt S
    (2017) Mitochondrial cytochrome c oxidase is inhibited by ATP only at very high ATP/ADP ratios. Biol Chem 398:737–750.
    OpenUrl
  31. ↵
    1. Rosca MG and
    2. Hoppel CL
    (2013) Mitochondrial dysfunction in heart failure. Heart Fail Rev 18:607–622.
    OpenUrlCrossRefPubMed
  32. ↵
    1. Sabbah HN
    (2016) Targeting mitochondrial dysfunction in the treatment of heart failure. Expert Rev Cardiovasc Ther 14:1305–1313.
    OpenUrl
  33. ↵
    1. Schuh AK,
    2. Sheybani B,
    3. Jortzik E,
    4. Niemann B,
    5. Wilhelm J,
    6. Boening A, and
    7. Becker K
    (2018) Redox status of patients before cardiac surgery. Redox Rep 23:83–93.
    OpenUrl
  34. ↵
    1. Sethna DH,
    2. Moffitt E,
    3. Gray RJ,
    4. Bussell J,
    5. Raymond M,
    6. Conklin C, and
    7. Matloff JM
    (1982) Effects of protamine sulfate on myocardial oxygen supply and demand in patients following cardiopulmonary bypass. Anesth Analg 61:247–251.
    OpenUrlPubMed
  35. ↵
    1. Shapira N,
    2. Schaff HV,
    3. Piehler JM,
    4. White RD,
    5. Still JC, and
    6. Pluth JR
    (1982) Cardiovascular effects of protamine sulfate in man. J Thorac Cardiovasc Surg 84:505–514.
    OpenUrlPubMed
  36. ↵
    1. Suski JM,
    2. Lebiedzinska M,
    3. Bonora M,
    4. Pinton P,
    5. Duszynski J, and
    6. Wieckowski MR
    (2012) Relation between mitochondrial membrane potential and ROS formation. Methods Mol Biol 810:183–205.
    OpenUrlCrossRefPubMed
  37. ↵
    1. Turrens JF
    (2003) Mitochondrial formation of reactive oxygen species. J Physiol 552:335–344.
    OpenUrlCrossRefPubMed
  38. ↵
    1. Viaro F,
    2. Dalio MB, and
    3. Evora PR
    (2002) Catastrophic cardiovascular adverse reactions to protamine are nitric oxide/cyclic guanosine monophosphate dependent and endothelium mediated: should methylene blue be the treatment of choice? Chest 122:1061–1066.
    OpenUrlCrossRefPubMed
  39. ↵
    1. Wakefield TW,
    2. Bies LE,
    3. Wrobleski SK,
    4. Bolling SF,
    5. Stanley JC, and
    6. Kirsh MM
    (1990) Impaired myocardial function and oxygen utilization due to protamine sulfate in an isolated rabbit heart preparation. Ann Surg 212:387–393; discussion 394.
    OpenUrlPubMed
  40. ↵
    1. Wakefield TW,
    2. Hinshaw DB,
    3. Burger JM,
    4. Burkel WE, and
    5. Stanley JC
    (1989a) Protamine-induced reductions of endothelial cell ATP. Surgery 106:378–385.
    OpenUrlPubMed
  41. ↵
    1. Wakefield TW,
    2. Ucros I,
    3. Kresowik TF,
    4. Hinshaw DB, and
    5. Stanley JC
    (1989b) Decreased oxygen consumption as a toxic manifestation of protamine sulfate reversal of heparin anticoagulation. J Vasc Surg 9:772–777.
    OpenUrlCrossRefPubMed
  42. ↵
    1. Wakefield TW,
    2. Wrobleski SK,
    3. Nichol BJ,
    4. Kadell AM, and
    5. Stanley JC
    (1992) Heparin-mediated reductions of the toxic effects of protamine sulfate on rabbit myocardium. J Vasc Surg 16:47–53.
    OpenUrlCrossRefPubMed
  43. ↵
    1. Wojtczak L,
    2. Teplova VV,
    3. Bogucka K,
    4. Czyz A,
    5. Makowska A,
    6. Wieckowski MR,
    7. Duszyński J, and
    8. Evtodienko YV
    (1999) Effect of glucose and deoxyglucose on the redistribution of calcium in ehrlich ascites tumour and Zajdela hepatoma cells and its consequences for mitochondrial energetics. Further arguments for the role of Ca(2+) in the mechanism of the crabtree effect. Eur J Biochem 263:495–501.
    OpenUrlCrossRefPubMed
  44. ↵
    1. Zharikov S and
    2. Shiva S
    (2013) Platelet mitochondrial function: from regulation of thrombosis to biomarker of disease. Biochem Soc Trans 41:118–123.
    OpenUrlAbstract/FREE Full Text
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Research ArticleCardiovascular

Protamine Sulfate Action on Mitochondrial Bioenergetics

Rabia Ramzan, Susanne Michels, Petra Weber, Annika Rhiel, Marc Irqsusi, Ardawan J. Rastan, Carsten Culmsee and Sebastian Vogt
Journal of Pharmacology and Experimental Therapeutics August 1, 2019, 370 (2) 308-317; DOI: https://doi.org/10.1124/jpet.119.257725
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