Activated human platelet products induce proarrhythmic effects in ventricular myocytes

Journal of Molecular and Cellular Cardiology 51 (2011) 347–356

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Journal of Molecular and Cellular Cardiology j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / y j m c c

Original article

Activated human platelet products induce proarrhythmic effects in ventricular myocytes Jonas S.S.G. de Jong a, 1, Arie O. Verkerk b, 1, Marcel M.G.J. van Borren b, c, Olga M. Zakhrabova - Zwiauer b, h, Rienk Nieuwland d, Joost C.M. Meijers e, Jan-Willem N. Akkerman f, Arthur A.M. Wilde a, b, Hanno L. Tan a, b,⁎, Lukas R.C. Dekker a, g a

Department of Cardiology, Academic Medical Center, University of Amsterdam, Amsterdam, The Netherlands Heart Failure Research Center, Academic Medical Center, University of Amsterdam, Amsterdam, The Netherlands c Laboratory of Clinical Chemistry and Haematology, Jeroen Bosch Hospital, 's-Hertogenbosch, The Netherlands d Laboratory of Experimental Clinical Chemistry, Academic Medical Center, University of Amsterdam, Amsterdam, The Netherlands e Departments of Vascular Medicine and Experimental Vascular Medicine, Academic Medical Center, University of Amsterdam, Amsterdam, The Netherlands f Department of Clinical Chemistry and Haematology, University Medical Center Utrecht, The Netherlands g Department of Cardiology, Catharina Hospital, Eindhoven, The Netherlands h Interuniversity Cardiology Institute of the Netherlands, Utrecht, The Netherlands b

a r t i c l e

i n f o

Article history: Received 19 January 2011 Received in revised form 2 May 2011 Accepted 23 May 2011 Available online 30 May 2011 Keywords: Platelets Ventricular arrhythmias Action potential Membrane currents Ca2+ homeostasis

a b s t r a c t Sudden cardiac death remains one of the most prevalent modes of death and is mainly caused by ventricular fibrillation (VF) in the setting of acute ischemia resulting from coronary thrombi. Animal experiments have shown that platelet activation may increase susceptibility of ischemic myocardium to VF, but the mechanism is unknown. In the present study, we evaluated the effects of activated blood platelet products (ABPPs) on electrophysiological properties and intracellular Ca2+ (Ca2+i) homeostasis. Platelets were collected from healthy volunteers. After activation, their secreted ABPPs were added to superfusion solutions. Rabbit ventricular myocytes were freshly isolated, and membrane potentials and Ca2+i were recorded using patch-clamp methodology and indo-1 fluorescence measurements, respectively. ABPPs prolonged action potential duration and induced early and delayed afterdepolarizations. ABPPs increased L-type Ca2+ current (ICa,L) density, but left densities of sodium current, inward rectifier K+ current, transient outward K+ current, and rapid component of the delayed rectifier K+ current unchanged. ABPPs did not affect kinetics or (in)activation properties of membrane currents. ABPPs increased systolic Ca2+i, Ca2+i transient amplitude, and sarcoplasmic reticulum Ca2+ content. ABPPs did not affect the Na+−Ca2+ exchange current (INCX) in Ca2+-buffered conditions. Products secreted from activated human platelets induce changes in ICa,L and Ca2+i, which result in action potential prolongation and the occurrence of early and delayed afterdepolarizations in rabbit myocytes. These changes may trigger and support reentrant arrhythmias in ischemia models of coronary thrombosis. © 2011 Elsevier Ltd. All rights reserved.

1. Introduction Sudden cardiac death (SCD) remains one of the most prevalent modes of death in industrialized countries. It claims almost a million deaths annually in Western Europe and the United States [1,2]. Ventricular fibrillation (VF) in the setting of coronary artery disease is the most common underlying arrhythmia [2]. Acute coronary thrombosis is observed in 74–79% of SCD victims at autopsy [3,4]. Platelets play an important role in the occurrence of SCD. Animal experiments have shown that platelet activation increases susceptibility of ischemic myocardium to VF [5–8]. Coronary occlusion with a thrombus in pigs ⁎ Corresponding author at: Academic Medical Center, PO Box 22700, 1100 DE Amsterdam, The Netherlands. Tel.: + 31 20 5663264; fax: + 31 20 6975458. E-mail address: [email protected] (H.L. Tan). 1 Both authors contributed equally to the study. 0022-2828/$ – see front matter © 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.yjmcc.2011.05.016

induced VF significantly more often than a non-thrombotic occlusion, such as a balloon or ligature [8]. Similarly, in a dog model of regional myocardial ischemia, 51% of control animals developed VF after balloon inflation, but none after induction of severe thrombocytopenia [6]. Platelet activation results in the release of platelet products, the socalled secretome, which includes organic substances (e.g., ATP, serotonin, histamine) and more than 2000 proteins [9]. Many of these substances can alter electrophysiological properties of the heart in various animal species, supporting the notion that activated platelets exert proarrhythmic effects [10–12]. Conversely, platelet antagonists counteract ischemia-induced arrhythmias in animal studies [13,5]. We hypothesized that activated blood platelet products (ABPPs) affect cellular electrophysiological properties and, thereby, modulate vulnerability to VF. We studied this hypothesis by evaluating the effects of human ABPPs on action potentials (APs), ion channels, and intracellular Ca2+ (Ca2+i) homeostasis of isolated rabbit ventricular

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myocytes. We found that ABPPs cause changes in electrophysiological properties, which are conducive to arrhythmia occurrence. 2. Materials and methods 2.1. Platelets 2.1.1. Blood collection The study was approved by the institutional Medical Ethics Committee (reference number 06/002) and conforms to the principles outlined in the Declaration of Helsinki. Platelet donors were healthy volunteers (n = 5) who signed written informed consent. All blood collections were performed with a 19 G needle without vacuum at 10–11 h in the morning to rule out possible effects of circadian fluctuations in platelet function and activation. After the first few ml were discarded, 40 ml was drawn in custom prepared Falcon tubes containing 3.2% (0.109 M) sodium citrate (1:10). 2.1.2. Isolation of products of activated platelets Platelet-rich-plasma was prepared by centrifugation at 160 × g for 15 min at room temperature without brake, and was acidified with acid citrate dextrose to prevent platelet activation during isolation. Subsequently, platelets were pelleted at 800 × g during 15 min and the pellet washed with HEPES-buffered Tyrode solution (pH 6.5). After addition of the platelet inhibitor prostaglandin I2 (10 ng/ml, final concentration), the platelets were pelleted again at 800 × g during 15 min and suspended in HEPES-buffered Tyrode (pH 7.2) to a final concentration of 220–250 × 10 9 platelets/l. Platelets can be activated by different aggregating agents (agonists). Using the patch-clamp technique (see below), we tested which platelet activator left AP properties unaltered. Table 1 shows that thrombin receptor activating protein (TRAP) and collagen did not affect AP properties, making them suitable for our study. To induce the platelet secretion response, platelets were prewarmed to 37 °C in an aggregometer and stimulated with TRAP (15 μM final concentration) or collagen (2 μg/ml final concentration) for 5 min at a stirring speed of 900 rpm. Platelets were then pelleted by centrifugation at 13,000 rpm for 5 min and the supernatant that contained the secreted ABPPs was snap-frozen in liquid nitrogen and stored at −80 °C until use. In platelets used for control experiments, stirring and centrifugation were similar, but TRAP was added to the supernatant only after centrifugation and separation from the pellet. This control solution is named non-ABPPs in the remaining part of the manuscript. In a subset of samples, the release reaction was validated by analysis of the serotonin concentration in ABPPs and non-ABPPs. Measured serotonin concentrations in representative ABPPs and nonABPPs samples were 761 ng/ml and 157 ng/ml, respectively. These values were in accordance with previously published values [14,15]. The amount of Ca2+ in ABPPs and non-ABPPs was negligible compared to the amount of Ca2+ added to the secretome during further experiments

Table 1 Effects of platelet activators on action potentials at 1 Hz. 1 μM ADP (n = 5) RMP APA Vmax APD50 APD90 Pla ampl

50 μM ATP (n = 4)

99.6 ± 0.6 100.1 ± 1.9 100.9 ± 1.4 98.8 ± 2.9 93.3 ± 5.9 94.5 ± 4.0 124.8 ± 6.9* 87.4 ± 3.8* 115.9 ± 5.4* 91.3 ± 3.6 100.6 ± 1.3 93.3 ± 2.5*

1 μM TXA2 1 μM TRAP (n = 4) (n = 6)

1 μg/ml collagen (n = 8)

99.0 ± 0.9 98.0 ± 4.1 81.3 ± 8.4 83.2 ± 3.9* 84.0 ± 3.3* 96.1 ± 1.8

100.1 ± 0.3 99.4 ± 0.6 99.3 ± 1.0 101.5 ± 2.2 101.3 ± 1.5 99.4 ± 0.7

99.7 ± 0.5 98.9 ± 2.6 100.5 ± 2.2 96.1 ± 5.4 98.5 ± 3.5 99.8 ± 1.4

Mean ± SEM. Data are expressed as percentage of control values. RMP = resting membrane potential; APA = maximal action potential amplitude; Vmax = maximal upstroke velocity; APD50 and APD90 = action potential duration at 50% and 90% repolarization; Pla ampl = action potential plateau amplitude. *P b 0.05 paired t-test.

(data not shown). For subsequent patch-clamp studies, ABPPs were diluted 40× in most experiments to increase the superfusion volume and compensate for supraphysiological platelet activation; in experiments aimed at studying the effects of higher ABPPs concentrations, we used a 20× dilution. To study whether the observed effects of ABPPs could be mediated by a protein/peptide moiety, we determined concentrations of total protein in ABPPs from three platelet isolations using spectrophotometric determination with the Pierce bicinchoninic acid (BCA) protein assay. protein content was 0.32 ± 0.02 mg/ml, comparable to previously published values [9]. ABPPs diluted 40× contained approximately 8 μg/ml protein, which was expected to be fully eliminated by treatment with 0.4 mg/ml trypsin at 37 °C for 30 min; [33] thereafter, the trypsin effect was inhibited by 0.2 mg/ml trypsin inhibitor. We named this solution trypsin-treated ABPPs in the remaining part of the manuscript. 2.2. Cell preparation The investigation conformed to the Guide for the Care and Use of Laboratory Animals (NIH Publication 85-23, revised 1996) and was approved by the institutional animal experiments committee. Midmyocardial cells of New Zealand White rabbits were isolated by enzymatic dissociation from the most apical part of the left ventricular free wall as described previously [16]. Small aliquots of cell suspension were put in a recording chamber on the stage of an inverted microscope. Cells were allowed 5 min to attach to the bottom before superfusion was initiated. The temperature was 36–37 °C, except for sodium current (INa) recordings (20–21 °C). Quiescent rod-shaped cross-striated cells with a smooth surface were selected for measurements. 2.3. Electrophysiology 2.3.1. Data acquisition and analysis APs and sarcolemmal ion currents were recorded with the amphotericin-B-perforated patch-clamp and ruptured patch-clamp technique, respectively. Voltage control, data acquisition, and analysis were accomplished using custom software. Potentials were corrected for the estimated change in liquid junction potential, except for INa measurements, where it was 0.2 mV. Adequate voltage control was achieved with low-resistance pipettes (1.5–2.5 MΩ), and Rs and Cm compensation N80%. Membrane currents and potentials were filtered (low-pass, 1 kHz) and digitized (2 kHz), except for AP and INa measurements, where filtering and digitizing frequencies were 5 kHz and 20 kHz, respectively. Cell membrane capacitance (Cm) was determined as described previously [16]. 2.3.2. Current-clamp experiments APs were measured using a modified Tyrode solution containing (mM): NaCl 140, KCl 5.4, CaCl2 1.8, MgCl2 1.0, glucose 5.5, HEPES 5.0; pH 7.4 (NaOH). Pipette solution contained: K-gluconate 125, KCl 20, NaCl 10, amphotericin-B 0.22, HEPES 10; pH 7.2 (KOH). APs were elicited at 0.2–4 Hz by 3 ms, 1.5× threshold current pulses through the patch pipette. We analyzed resting membrane potential (RMP), maximal upstroke velocity (Vmax), AP amplitude (APA), AP plateau amplitude (defined as the potential difference between RMP and the potential at 50 ms after the AP upstroke), and AP duration at 20%, 50% and 90% repolarization (APD20, APD50, and APD90, respectively). Data from 10 consecutive APs were averaged. APs were measured in the absence and presence of ABPPs or nonABPPs in the same myocyte. In order to obtain steady-state conditions, AP recordings were started 4 min after application of ABPPs or nonABPPs. The occurrence of early afterdepolarizations (EADs) was tested at a pacing frequency of 0.2 Hz. Susceptibility to delayed afterdepolarizations (DADs) was tested by applying a 3-Hz pacing episode (10 s) followed by an 8-s pause.

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2.3.3. Voltage-clamp experiments INa, L-type Ca 2+ current (ICa,L), inward rectifier K + current (IK1), transient outward K + current (Ito1), rapid component of delayed rectifier K + current (IKr), and Na +-Ca 2+ exchange current (INCX) were measured with solutions described previously [17], and by voltageclamp protocols shown in the appropriate figures. ICa,L was measured in the presence of 0.25 mM DIDS to block the Ca 2+-activated Cl current. Ito1 was measured in the presence of 1 mM CdCl2, which blocks inward Ca 2+ currents [18], and thereby also prevents activation of the transient outward Ca 2+-activated Cl - current [19]. CdCl2 also strongly inhibits inward Na + currents [20]. INa, IKr, and IK1 were measured in the presence of 5 μM nifedipine to block ICa,L. INCX was measured as 10 mM Ni 2+-sensitive current. The presence of the slow component of the delayed rectifier K + current (IKs) in rabbit ventricular myocytes is debated (see ref. [21] and primary references cited therein). In our experiments, we did not observe residual tail current in the presence of the IKr blocker E-4031 (5 μM), neither did we observe currents sensitive to the IKs blocker chromanol-293B (90 μM) [21]. Therefore, tail currents after depolarizing voltage-clamp steps in our experiments were attributed to IKr. Membrane currents were defined as indicated in the typical examples shown in the appropriate figures. Ion currents were measured in paired experiments, i.e., in the absence and presence of ABPPs (40× dilution) in the same myocyte. Because non-ABPPs did not affect the AP significantly (see Section 3), the effects of non-ABPPs on ion currents were not measured with the exception of ICa,L. Because ion currents may show run-down and shifts in (in)activation shortly after patch rupture [22], control ion current measurements were started after a 10 min equilibration period and the effects of ABPPs or non-ABPPs were measured 4 min after application. Current densities were calculated by dividing current amplitudes by Cm.Voltage dependence of (in)activation was determined by fitting a Boltzmann function (y = [1 + exp{(V −V1/2)/ k}]) − 1 to the individual curves, yielding half-maximal voltage V1/2 and slope factor k. Current decay of INa and ICa,L were fitted with a double-exponential function to obtain the time constants of the fast and the slow components of current decay: y = [Af × exp(− t/τf)] + [As × exp(−t/τs)], where τf and τs are the time constants of fast and slow components and Af and As the fractions of the fast and slow component. 2.4. Ca 2+ transients Intracellular Ca 2+ (Ca 2+i) was measured in indo-1 loaded myocytes as described previously [23]. Dual wavelength emission of indo-1 was recorded ((405–440)/(505–540) nm, excitation at 340 nm) and free Ca 2+i was calculated [23]. Ca 2+i transients were elicited at 2 Hz using field stimulation [23] or at 0.2–3 Hz by 50-ms depolarizing voltage-clamp steps from −80 mV to 0 mV using the perforated patch-clamp technique. We analyzed diastolic and systolic Ca 2+i concentrations, Ca 2+i transient amplitudes, and systolic Ca 2+i rise. The rates of decay of the systolic and caffeine-induced Ca 2+ transients were obtained by fitting single exponential functions to the decay phase of the transient. The sarcoplasmic reticulum (SR)dependent rate of Ca 2+ uptake (SERCA activity; kSR) was calculated by subtracting the decay rate constant of caffeine evoked Ca 2+ transient (kcaff) from systolic Ca 2+ transient decay rate constant (ksys) [24] The SR Ca 2+ content was calculated by integrating the resulting INCX following a rapid application of 20 mM caffeine after a 20-s stimulus period (frequency: 0.2 or 3 Hz) while the membrane potential was kept at −80 mV [25]. The INCX, defined as current activated by caffeine application (for example, see Fig. 6E arrows), was not corrected for Ca 2+ removal by nonelectrogenic pathways, i.e., the sarcolemmal Ca 2+-ATPase, because in rabbit INCX contributes about 95% of Ca removal [25,26]. The SR Ca 2+ contents are expressed relative to total cell volume, which was calculated from Cm measure-

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ments assuming a cell volume–capacitance ratio of 4.58 pF/pL [27]. Because non-ABPPs did not affect the Ca 2+i homeostasis significantly (data not shown) Ca 2+i transients were always measured in paired experiments, i.e., in the absence and presence of ABPPs in the same myocyte. 2.5. Statistics Data are mean ± SEM. Group comparisons were made using the t-test or two-way repeated measures ANOVA followed by pairwise comparison using the Student–Newman–Keuls test. Proportions of EAD and DAD occurrence were compared using Fisher's exact test. P b 0.05 defined statistical significance. 3. Results 3.1. ABPPs prolong action potential duration Fig. 1A shows representative APs at 1 Hz in control conditions (solid line) and in the presence of non-ABPPs (top, dashed line) and ABPPs (bottom, dashed line). Non-ABPPs did not change the AP configuration. However, ABPPs induced a more positive AP plateau potential and a longer AP duration. The effects of collagen-activated and TRAP-activated ABPPs on AP plateau amplitude and AP duration did not differ significantly (data not shown). Therefore, data obtained from collagen-activated and TRAP-activated ABPPs were pooled. Fig. 1B summarizes the effects of non-ABPPs and ABPPs on AP properties. On average, ABPPs caused a 5% more positive AP plateau potential and a 12% longer AP duration at 90% repolarization. No significant differences of ABPPs on RMP, Vmax or maximal AP amplitude were observed. AP prolongation and larger AP plateau amplitudes were present at all frequencies measured (data not shown). 3.2. ABPPs induce triggered activity 3.2.1. Early afterdepolarizations EADs typically occur at slow heart rates [28]. We studied susceptibility to EAD formation in control conditions and in the presence of ABPPs at 0.2 Hz. Fig. 1C shows representative APs in control conditions (solid line) and in the presence of ABPPs (dashed line). While EADs in control conditions were rare, EADs were observed in 12 out of 20 myocytes tested in the presence of ABPPs (P b 0.05; Fig. 1E). The EADs occurred as single rather than multiple afterdepolarizations and had a ‘take-off potential’ (the potential where repolarization turns into depolarization) between −40 and 0 mV. Application of 0.3 μM nifedipine in the presence of ABPPs caused significant AP shortening and reduction of AP plateau amplitude, and abolished the EADs in three out of three myocytes tested (data not shown). 3.2.2. Delayed afterdepolarizations DADs typically occur at fast heart rates [29]. We studied the susceptibility to DAD formation in control conditions and in the presence of ABPPs by applying a 3-Hz pacing episode (10 s) followed by an 8-s pause. Fig. 1D shows the last two stimulated APs in control conditions (solid line) and in the presence of ABPPs (dashed line). While DADs in control conditions were rare, DADs were found in 8 out of 17 myocytes tested in the presence of ABPPs (P b 0.05; Fig. 1E). The DADs did not result in triggered APs. Nifedipine (0.3 μM) abolished the DADs in three out of three myocytes tested (data not shown). 3.3. ABPPs increase the L-type Ca 2+ current After establishing that ABPPs prolong AP duration, we next sought to identify which sarcolemmal ion currents contribute to this effect by

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Fig. 1. ABPPs prolong action potential (AP) duration. (A) Top, representativeAPs at 1 Hz in control conditions and in the presence of non-ABPPs. Bottom, representativeAPs at 1 Hz in control conditions and in the presence of ABPPs. (B) Average effects of non-ABPPs and ABPPs on AP parameters measured at 1 Hz. RMP = resting membrane potential, Vmax = maximal upstroke velocity, APA = maximal AP amplitude, Pla = AP plateau amplitude, APD20, APD50 and, APD90 = AP duration at 20%, 50%, and 90% repolarization. *P b 0.05 in paired t-test. (C) Representative example of an early afterdepolarization (EAD) induced by ABPPs. (D) Representative example of a delayed afterdepolarization (DAD) induced by ABPPs. (E) Incidence of EADs and DADs in control conditions and presence of ABPPs. *P b 0.05 in Fisher's exact test.

studying the major inward and outward ion currents in the absence and presence of ABPPs. 3.3.1. Na + current INa was measured using a two-step protocol (Fig. 2A). During the first depolarizing pulse (P1), INa activates; the second pulse (P2) is used for measuring voltage dependency of inactivation. Fig. 2B shows representative INa recordings upon depolarizing pulses to − 30 mV. Neither mean INa densities (Fig. 2C) nor current decay (Fig. 2B) were affected by ABPPs. Also, voltage dependency of INa activation and inactivation were similar in the absence and presence of ABPPs (Fig. 2D). 3.3.2. L-type Ca 2+ current ICa,L was measured using a two-step protocol (Fig. 3A). During the first depolarizing pulses (P1), ICa,L activates; the second pulse (P2) is used for measuring voltage dependency of inactivation. Fig. 3B shows representative ICa,L recordings upon depolarizing pulses to 0 mV. Mean ICa,L densities (Fig. 3D) increased in response to ABPPs. For example, at 0 mV peak ICa,L averaged −14.2 ± 1.2 and − 16.5 ± 1.9 pA/pF in the absence and presence of ABPPs (P b 0.05, n = 6, paired t-test), respectively. Neither current decay (Fig. 3C) nor (in)activation

properties (Fig. 3E) were affected by ABPPs. The increase in ICa,L density was not larger at higher ABPPs concentrations (20× dilution; Fig. 3F), while non-ABPPs did not affect ICa,L density (Fig. 3E) or gating properties (data not shown). 3.3.3. K + currents Ito1 was measured using a two-step protocol (inset). During the first depolarizing pulse (P1), Ito1 activates; the second pulse (P2) is used for measuring voltage dependency of inactivation. Fig. 4A shows representative Ito1 recordings upon depolarizing voltage steps from −80 to 60 mV. Neither mean Ito1 densities (Fig. 4B) nor (in)activation properties (data not shown) were affected by ABPPs. IK1 was measured as steady-state current at the end of hyperpolarizing voltage-clamp steps from − 40 mV. Fig. 4C shows representative IK1 recordings upon hyperpolarizing pulses to −110 mV.Both inward and outward IK1 components were not significantly different in the absence and presence of ABPPs (Fig. 4D). IKr was activated by depolarizing voltage-clamp steps from −50 mV and defined as the tail current upon stepping back to the holding potential. Fig. 4E shows representative IKr recordings upon depolarizing pulses to − 10 mV. Neither mean IKr densities (Fig. 4F) nor activation properties (data not shown) were affected by ABPPs.

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Fig. 2. No effects of ABPPs on the Na+ current (INa). (A) Protocol used. (B) Representative INa in control conditions and in presence of ABPPs. Inset, average time constants of current decay (n = 7) in the absence (black bars) and presence (white bars) of ABPPs. (C) Average current–voltage (I–V) relationships of INa (n = 7) in control conditions (closed symbols) and in the presence of ABPPs (open symbols). (D) Voltage dependence of INa (in)activation (n = 7) in control conditions (closed symbols) and in the presence of ABPPs (open symbols). Solid lines: Boltzmann fits of the average data.

Fig. 3. ABPPs increase L-type Ca2+ current (ICa,L) density. (A) Protocol used. (B) RepresentativeICa,L in control conditions and in presence of ABPPs. Insets: protocols used. (C) Average time constants of current decay (n = 6) in the absence (black bars) and presence (white bars) of ABPPs. (D) Average I–V relationships of ICa,L (n = 6) in control conditions (closed symbols) and in the presence of ABPPs (open symbols). *P b 0.05 in paired t-test. (E) Voltage dependence of ICa,L (in)activation in control conditions (closed symbols) and in the presence of ABPPs (open symbols). Solid lines: Boltzmann fits of the average data. (F) Average effects of non-ABPPs (n = 5), 80× diluted (n = 5), 40× diluted (n = 6), and 20× diluted (n = 5) ABPPs on ICa,L measured at 0 mV. Data are normalized to its control conditions, i.e., in absence of non-ABPPs or ABPPs.

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Fig. 4. No effects of ABPPs on K+ currents. (A) Representative transient outward K+ current (Ito1) in control conditions and in presence of ABPPs. (B) Average I–V relationships of Ito1 (n = 7) in control conditions (closed symbols) and in the presence of ABPPs (open symbols). (C) Representative inward rectifier K+ current (IK1) in control conditions and in the presence of ABPPs. (D) Average I–V relationships of IK1 (n = 5) in control conditions (closed symbols) and in the presence of ABPPs (open symbols). (E) Representative rapid delayed rectifier K+ current (IKr) in control conditions and in the presence of ABPPs. (F) Average I–V relationships of IKr (n = 5) in control conditions (closed symbols) and in the presence of ABPPs (open symbols). Insets: protocols used.

3.3.4. NCX current INCX was measured in Ca 2+-buffered conditions as the Ni 2+sensitive current during a descending voltage ramp protocol. The effects of Ni 2+ on INCX are reversible [30]; therefore,INCXmeasurements

in the absence and presence of ABPPs were possible in the same cell. Fig. 5A shows representative traces of INCX in the absence and presence of ABPPs. ABPPs neither changed INCX in the reverse (outward) mode nor in the forward (inward) mode (Fig. 5B). 3.4. ABPPs affect Ca 2+ homeostasis

Fig. 5. No effects ofABPPs on Na+-Ca2+ exchange current (INCX). (A) Representative INCX in control conditions and in presence of ABPPs. (B) Average I–V relationships of INCX (n = 6) in control conditions (closed symbols) and in the presence of ABPPs (open symbols).

We found that ABPPs induced both EADs and DADs. There are multiple mechanisms underlying EAD formation: [29,31,32] (1) reactivation of INa, (2) reactivation of ICa,L, and (3) INCX following spontaneous SR Ca 2+ release. The mechanism underlying DADs is INCX activated by spontaneous SR Ca 2+ release [29,33]. Thus, both types of afterdepolarizations might be due to spontaneous SR Ca 2+ release. This typically occurs in Ca 2+i overload conditions [34,35]. Accordingly, we studied the effect of ABPPs on Ca 2+i transients. To exclude possible effects of alterations in AP shape on Ca 2+i transients, the transients were studied in voltage-clamp mode with pulses of similar duration [36,24]. Fig. 6A shows typical Ca 2+i measurements of a myocyte stimulated at 0.2 Hz in control conditions (black line) and in the presence of ABPPs (red line); Fig. 6B shows the normalized Ca 2+i signals. Fig. 6 (A and B) suggests that the amplitude of systolic Ca 2+i is increased and that the rate of decay of Ca 2+i is faster in presence of ABPPs. The extent of the alterations in Ca 2+i measured at a 0.2-Hz frequency are summarized in Fig. 6C. In presence of ABPPs, the average amplitude of the Ca 2+i transient (TA) is increased to 204% of control conditions, while the diastolic Ca 2+i concentrations did not differ significantly (Fig. 6C). Consequently, the average systolic Ca 2+i concentration was significantly increased (Fig. 6C). The increase in systolic Ca 2+i occurred within 3 min of superfusion with ABPPs, and dissipated after 4 min washout (Fig. 8). The systolic Ca 2+i transient is highly dependent on SR Ca 2+ content [24]. The larger systolic Ca 2+i thus suggests that the SR Ca 2+ content is increased in presence of ABPPs. Indeed, the calculated SR Ca 2+ content using the integrated INCX in response to caffeine induced SR Ca 2+ discharge (Fig. 6E, left and middle panels) was increased in presence of ABPPs (Fig. 6D). In

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Fig. 6. ABPPs augment SERCA function. (A) Typical Ca2+i transients at 0.2 Hz in control conditions and in presence of ABPPs. (B) Normalized and superimposed Ca2+ transients. (C) Average Ca2+i transient parameters (n = 6) at 0.2 Hz. TA = transient amplitude, Ksys = systolic Ca2+ transient decay rate constant. *P b 0.05 in paired t-test. (D) Average SR Ca2+ content (n = 5) calculated from INCX activated by caffeine-induced rise of Ca2+i (see E), and the average SR-dependent rate of Ca2+ removal (kSR, n = 5) in control conditions and in the presence of ABPPs at 0.2 Hz. *P b 0.05 in paired t-test. (E) Left, typical caffeine-induced rise of Ca2+i (upper panels) and associated INCX (lower panels). Right, INCX–Ca2+i relationships of the measurements in the left panel. (F) Average slopes of the INCX–Ca2+i relationships in control conditions and in presence of ABPPs (n = 5).

presence of ABPPs, the average SR Ca 2+ content was increased to 125% of control conditions. Furthermore, the rate constant of decay of the Ca 2+i transient (ksys) was significantly increased in presence of ABPPs (Fig. 6C). On average, Ca 2+i transient decay rate constant was 2.5 times higher than in control conditions. Given that SERCA is the primary route for Ca 2+ removal [24,26], the accelerated rate of the Ca 2+i transient decay is indicative of enhanced SERCA function. Indeed kSR was increased in presence of ABPPs (Fig. 6D). Moreover, the systolic Ca 2+i rise was much faster in presence of ABPPs (Fig. 6C). On average, the systolic Ca 2+i rise increased to 164% of control conditions. These effects were present at all frequencies measured (data not shown), except for diastolic Ca 2+i concentration which was significantly increased at 3 Hz, but not at the other frequencies (data not shown). Finally, we determined the relationship between Ca 2+i and INCX obtained during the decay phase of caffeine evoked Ca 2+ transients [24] in control and ABPPs conditions. The data were fitted in each cell with linear regressions over the same Ca 2+i range (Fig. 6E, right panel, dashed lines). The average slopes of the Ca 2+i and INCX relationships did not change in presence of ABPPs (Fig. 6F). This indicates that Na +–Ca 2+ exchanger function does not change under these conditions, which agrees without patch-clamp findings where

INCX was measured in Ca 2+-buffered conditions as the Ni 2+-sensitive current during a descending voltage ramp protocol (Fig. 5). In addition, the unaltered relationship between INCX and Ca 2+i (Fig. 6F) combined with the finding that decay rate constants of caffeine induced Ca 2+ transients were not significantly different (0.67 ± 0.05 (control) versus 0.90 ± 0.23 (ABPPs) 1/s) suggests that sarcolemmalmediated Ca 2+ removal by Ca 2+ ATPases and other Ca 2+ transport pathways are virtually unaltered by ABPPs [24]. 3.5. Trypsin reduced ABPPs-induced effects on action potentials and Ca 2+i homeostasis Previously, it was observed that trypsin partially reduced the effects of rabbit ABPPs on Ca 2+i homeostasis in spontaneously beating chick embryonic heart cell aggregates [37], suggesting that peptides are partially responsible for the effects of ABPPs. To study whether our observed effects of ABPPs could be mediated by proteins/peptides, we studied the effects of trypsin-treated ABPPs. While Tyrode solution, to which 0.4 mg/ml trypsin and 0.2 mg/ml trypsin inhibitor were added, did not affect AP characteristics or Ca2+i homeostasis (data not shown), trypsin-treated ABPPs still increased APD90 significantly. However,

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180

Tyrode solution trypsin-treated ABPPs

*

Percentage of control

160 140 120

ABPPs trypsin-treated Tyrode solution

* *

*

100 80 60 40 20 0

systolic Ca2+i

Ca2+i transient amplitude

Fig. 7. ABPPs (incubated or not with trypsin) increased systolic Ca2+i and Ca2+i transient amplitude significantly. *P b 0.05 in paired t-test versus control (Tyrode solution). Tyrode solution treated with 0.4 mg/ml trypsin and 0.2 mg/ml trypsin inhibitor alone did not affect Ca2+i homeostasis (n = 9). ABPPs not treated with trypsin increase systolic Ca2+i and Ca2+i transient amplitude by 33% and 43%, respectively (n = 6). Trypsin-treated ABPPs increase systolic Ca2+i and Ca2+i transient amplitude by 14% and 18%, respectively (n = 12).

while untreated ABPP induced AP prolongation of 12% (Fig. 1B), trypsintreated ABPPs prolonged APs only by 4% (n = 7), suggesting that AP prolongation is partially mediated by peptides. Likewise, trypsin treatment significantly blunted the effect of ABPPs on Ca 2+i transients (Fig. 7). While untreated ABPPs increased systolic Ca2+i and Ca2+i transient amplitudes by 33% and 43%, respectively (n = 6), trypsintreated ABPPs increased systolic Ca 2+i and Ca2+i transient amplitudes only by 14% and 18%, respectively (n = 12). These data suggest that these effects of ABPPs are in large part mediated by peptides. 4. Discussion We found that human ABPPs have acute effects on electrophysiological properties and Ca 2+i homeostasis of rabbit ventricular myocytes. In myocytes exposed to ABPPs AP duration prolonged and EADs and DADs occurred (Fig. 1). Analysis of membrane currents revealed that ICa,L density was increased, while INa,Ito1, IK1, IKr, and INCX densities were unchanged (Figs. 2–5). ABPPs did not affect kinetics and (in)activation properties of membrane currents. ABPPs increased systolic Ca 2+i, Ca 2+i transient amplitude, enhanced SERCA mediated Ca 2+ uptake rate, and increased SR Ca 2+ content (Fig. 6). These effects

Fig. 8. ABPPs increase systolic Ca2+i within 3 min after superfusion. This effect is completely reversed after 4 min washout.*P b 0.05 in paired t-test versus control (Tyrode solution).

on Ca 2+i and the finding that the relationship between Ca 2+i and INCX is unaltered, are consistent with the notion that increased SERCA function is responsible for larger systolic Ca 2+i transient in presence of ABPPs. 4.1. ABPPs prolong action potential duration and induce EADs and DADs ABPPs prolong AP duration (Fig. 1B) and increase ICa,L(Fig. 3). The increase of the inward ICa,L seems a plausible explanation for AP prolongation. It also explains the increased incidence of EADs (Fig. 1C and E), as EADs occur at low heart rates under conditions of long APs and may be due to reactivation of ICa,L[28,32]. This is supported by our finding that the ICa,L blocker nifedipine abolished ABPPs-induced EADs, consistent with the finding that nifedipine abolished EADs induced by delayed rectifier K + current blockers [38] and the ICa,L modulator FPL-64716 [39]. On the other hand, the ABPPs-induced increase in SR Ca 2+ release (Fig. 6) could partially counteract AP prolongation by increasing the rate of inactivation of ICa,L[34]. However, such an increase in the amplitude of the Ca 2+i transient would also increase the efflux of Ca 2+ from the cell as Na +–Ca 2+ exchange is more strongly activated [34]. The latter would importantly result in an increase in the inward current carried by NCX, which in turn is also expected to prolong AP duration. Thus, although INCX density in Ca 2+-buffered conditions is not affected by ABPPs (Fig. 5), we think that the functional INCX is increased due to increased Ca 2+i transient amplitudes. Moreover, ABPPs induce DADs (Fig. 1D and E). This agrees with our finding of enhanced SR Ca 2+ content in response to ABPP application. If the SR Ca 2+ content becomes high, a condition known as Ca 2+ overload, spontaneous SR Ca 2+ release may occur [34,35]. Such spontaneous Ca 2+ release can occur during diastole and activates INCX, thereby producing DADs [40]. The ICa,L blocker nifedipine abolished the ABPPs-induced DADs, which agrees with the finding that nifedipine abolished DADs induced by the ICa,L modulator FPL-64716 [39]. This effect may be due to reduced Ca 2+ influx that reduces Ca 2+ overload. Thus, increases in ICa,L, functional INCX, and SR Ca 2+ content all provide plausible explanations for AP prolongation, and the occurrence of EADs and DADs in the presence of ABPPs. ABPPs did not affect INa, Ito1, IK1, and IKr, but taking into account that multiple ion channels and exchangers exist in cardiac myocytes, we cannot exclude that other current/exchanger mechanisms, not included in the present study, may contribute to the observed ABPPs effects. 4.2. Clinical implications Arrest of blood flow to myocytes results in a complex ischemic reaction that includes an increase in outward potassium current [41], a triphasic increase of extracellular potassium [42], a decrease of AP duration [43], depolarization of RMP and Ca2+ loading of the cell [44]. Furthermore, cell-to-cell connections are uncoupled [45] and reactive oxygen species are released, resulting in changes in ion channel function [46] and mitochondrial dysfunction [44]. These events create a hostile environment that renders the myocardium prone to arrhythmias [47]. VF during ischemia results from reentrant excitation. Reentry may be facilitated by increased spatial dispersion in refractory periods, which, in turn, are largely determined by AP duration. Onset of reentry can be evoked by triggered beats originating from EADs and DADs [35]. The ischemic tissue distal to a coronary thrombus is exposed to a plethora of platelet products [15]. Most of the products released by platelets are packaged in preformed storage granules. Platelets contain three types of granules: dense granules, α-granules and lysosomal granules [48]. Many of the substances released by platelets can alter electrophysiological properties of the heart in various animal species [10–12]. The proarrhythmic effects of ABPPs could well result from the summed result of smaller effects of individual substances. However, one study points to small trypsin-sensitive peptides

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produced by activated platelets that could increase Ca 2+i transients in chick cardiac cells [37]. In line with this study, we found that trypsin attenuated the effects of human ABPPs on AP prolongation and Ca 2+i increase, indicating that trypsin-sensitive peptides indeed contributed to the observed effects of human ABPPs. However, trypsin-treated ABPPs still produced significant effects on AP and Ca 2+i characteristics, indicating that trypsin resistant substances also contributed to ABPPs-induced AP prolongation and Ca 2+i transient increase. With regards to the effects of platelet products, the fact that arrhythmias during ischemia often present in the first seconds and minutes after the onset of ischemia fits best with the kinetics of dense granule release as these granules are released rapidly after platelet activation. Clearly, future studies must resolve which compound contributes most to the proarrhythmic properties of activated platelets.

[3] [4]

[5]

[6]

[7]

[8]

4.3. Limitations of the study [9]

Practical reasons dictated that the human blood products were snap-frozen in liquid nitrogen and stored at − 80 °C until use. We cannot exclude that this procedure has affected some unstable products arising from platelet activation or easily degraded products. Moreover, we limited ourselves to study the effects of ABPPs in nonischemic myocytes from the midmyocardial layer of rabbit hearts. In mammalian hearts, however, regional differences in electrophysiology and Ca 2+ handling exist [49,50]. In addition, (pseudo)ischemic conditions will result in time-dependent changes without steadystate conditions [46]. Including these factors in the present study would have complicated the interpretation of the ABPPs effects.

[10] [11]

[12] [13]

[14] [15]

4.4. Conclusion [16]

Exposure of isolated rabbit myocytes to products released from activated human platelets resulted in AP prolongation, early and late afterdepolarizations, and changes in Ca 2+ homeostasis caused by increased ICa,L. These changes may contribute to the initiation and maintenance of reentrant arrhythmias after coronary thrombotic occlusion.

[17]

[18]

[19]

Funding

[20]

This work was supported by the Netherlands Heart Foundation (grant number NHS2007B020) and the Netherlands Organization for Scientific Research (NWO, ZonMW-Vici; grant number 918.86.616 to H.L.T).

[21]

Conflict of interest

[23]

None. Disclosures

[22]

[24]

[25]

None. [26]

Acknowledgments [27]

The authors wish to thank J. Zegers and A.C.G. van Ginneken for kindly providing the data acquisition and analysis program, respectively, and J. Bourier and Z. Kingma for biotechnical assistance. References

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