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In Vivo Post–Cardiac Arrest Myocardial Dysfunction Is Supported by Ca2+/Calmodulin-Dependent Protein Kinase II–Mediated Calcium Long-Term Potentiation and Mitigated by Alda-1, an Agonist of Aldehyde Dehydrogenase Type 2

Originally published 2016;134:961–977



Survival after sudden cardiac arrest is limited by postarrest myocardial dysfunction, but understanding of this phenomenon is constrained by a lack of data from a physiological model of disease. In this study, we established an in vivo model of cardiac arrest and resuscitation, characterized the biology of the associated myocardial dysfunction, and tested novel therapeutic strategies.


We developed rodent models of in vivo postarrest myocardial dysfunction using extracorporeal membrane oxygenation resuscitation followed by invasive hemodynamics measurement. In postarrest isolated cardiomyocytes, we assessed mechanical load and Ca2+-induced Ca2+ release (CICR) simultaneously using the microcarbon fiber technique and observed reduced function and myofilament calcium sensitivity. We used a novel fiberoptic catheter imaging system and a genetically encoded calcium sensor, GCaMP6f, to image CICR in vivo.


We found potentiation of CICR in isolated cells from this extracorporeal membrane oxygenation model and in cells isolated from an ischemia/reperfusion Langendorff model perfused with oxygenated blood from an arrested animal but not when reperfused in saline. We established that CICR potentiation begins in vivo. The augmented CICR observed after arrest was mediated by the activation of Ca2+/calmodulin-dependent protein kinase II (CaMKII). Increased phosphorylation of CaMKII, phospholamban, and ryanodine receptor 2 was detected in the postarrest period. Exogenous adrenergic activation in vivo recapitulated Ca2+ potentiation but was associated with lesser CaMKII activation. Because oxidative stress and aldehydic adduct formation were high after arrest, we tested a small-molecule activator of aldehyde dehydrogenase type 2, Alda-1, which reduced oxidative stress, restored calcium and CaMKII homeostasis, and improved cardiac function and postarrest outcome in vivo.


Cardiac arrest and reperfusion lead to CaMKII activation and calcium long-term potentiation, which support cardiomyocyte contractility in the face of impaired postarrest myofilament calcium sensitivity. Alda-1 mitigates these effects, normalizes calcium cycling, and improves outcome.


Sudden cardiac arrest is a major cause of death in the Western world,1 and mortality after resuscitation remains >50%.2,3 This excess mortality is reflected in the post–cardiac arrest syndrome,2 a condition that comprises postarrest central nervous system dysfunction, postarrest ischemia/reperfusion, post–myocardial arrest dysfunction (PMAD), and continuation of the factors precipitating sudden cardiac arrest. PMAD is believed to be a form of myocardial stunning46 and has been observed for hypoxic arrest and after ventricular fibrillation.7 Recognizing that mortality remains high after resumption of circulation, the International Liaison Committee on Resuscitation has recommended more detailed resuscitation research, including the phenomenon of PMAD.1

PMAD may result from oxidative stress–induced cytosolic calcium overload. Calcium overload has traditionally been explained by 2 processes: reverse-mode operation of the sodium-calcium exchanger related to intracellular sodium overload during ischemia and sarcoplasmic reticulum ATPase dysfunction related to ATP depletion.8 These processes have been suggested to lead to reversible myofilament dysfunction and subsequently to irreversible cell death partly through Ca2+/calmodulin-dependent protein kinase II (CaMKII) signaling pathways.810 The relationship between acute calcium overload and long-term augmentation of calcium-induced calcium release (CICR) is critical but understudied. In particular, the biochemical pathways identified would be predicted to potentiate CICR. In addition, because normal CICR generates, on a beat-to-beat basis, systolic calcium levels in localized domains 10-fold higher than predicted to occur during postarrest calcium overload,11 any CICR augmentation might be positioned to play a dominant signaling role. Moreover, as it relates to excitation-contraction coupling, potentiated CICR transients would be expected to result in more forceful contractions, not the opposite. For example, in a post–cardiac arrest study, ryanodine receptor (RyR)–dependent calcium release was shown to support reversible injury.12 However, despite data in support of such a mechanism, no long-lasting changes in CICR have been demonstrated after reperfusion13 except for 1 early report.14 Alternatively, the increased CICR transients could be a consequence of a mismatch between cell contractility and external mechanical demand whereby reduced shortening, or diastolic stretch, of cells may raise intracellular Ca2+ levels15 or Ca2+ releasability from the sarcoplasmic reticulum.16 Two limitations of previous experimental models are that there is an absence of in vivo data and that much of the in vitro mechanistic data have come from the buffered saline–perfused Langendorff preparations that suffer from significant edema.17 In addition, CICR and CaMKII signaling have been linked to β-adrenergic signaling under pathological conditions,15 and cardiac arrest has been associated with dramatic elevations in catecholaminergic signaling, which is not present in the isolated heart model.18 We believed that a model preserving the neurohumoral axis might better reflect the in vivo environment and developed an in vivo model of cardiac arrest with resuscitation using extracorporeal membrane oxygenation (ECMO).19 Here, we report findings from this in vivo cardiac arrest–ECMO model relating to PMAD. Two novel approaches were introduced in this study. First, we examined cellular force in vitro using microcarbon fiber assessment simultaneously with CICR measurements. Second, we developed an approach to measuring calcium dynamics in vivo using a genetically engineered calcium indicator. Our data from this in vivo model support the contribution of calcium potentiation, CaMKII signaling, and oxidative stress to PMAD and suggest a potential new therapeutic candidate, Alda-1.


Studies were approved by the Stanford Administrative Panel on Laboratory Animal Care and conform to the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health (NIH).

Invasive Hemodynamics and Development of ECMO

Male Sprague-Dawley rats (weight, 250–350 g) were anesthetized and oxygenated/ventilated with 1% to 2% isoflurane, maintained at 37°C, and monitored throughout for depth of anesthesia. An ECMO blood circuit was built to support reanimation after cardiac arrest (Figure 1A).20 A conductance catheter (Millar Instruments, Houston, TX) was introduced into the left ventricle for pressure-volume loop measurements as described previously.21,22

Figure 1.

Figure 1. In vivo cardiac arrest models. A, Extracorporeal membrane oxidation (ECMO) circuit consisting of a venous reservoir (VR), roller pump (RP), miniature membrane oxygenator (MO) between the arterial (femoral artery [FA]) and venous (right atrium via external jugular [EJ]) systems perfused in a retrograde fashion. LV pressure was measured through the carotid artery with a Millar Catheter (not shown). B, Timeline of experiment followed by 15 minutes of downtime and then ECMO initiation. C, An example arrhythmic sudden cardiac arrest (CA) protocol. D, The ordinate for D through F is the same and shown in D only. Zoomed-in view of the beginning of rapid ventricular pacing (RVP) at 12-Hz pacing over the left ventricular apex, leading to hypotension. Blood pressure (black) is shown on top with the scale bar on the left, and heart rate (far-field ECG; gray) is shown underneath. E, Both ventricular fibrillation (VF) followed by pulseless electric activity (PEA) later in time in a zoomed-in view. F, Zoomed-in view of the heart off ECMO after full resuscitation. G and H, Representative pressure-volume loops from before (G) and after (H) arrest with ECMO resuscitation in the same animal. The end-systolic pressure-volume relationship (ESPVR; solid line) and end-diastolic pressure-volume relationship (dotted line) are superimposed. The ESPVR slope was 1.38 and 0.6 mm Hg/µL.

Animal Models of Disease

After institution of the ECMO circuit (Figure 1A), sudden cardiac arrest was initiated either by cessation of ventilation or by rapid ventricular pacing to induce ventricular fibrillation (Figure 1B–1E).19 Both processes led to rapid hypotension, followed by pulseless electric activity or asystole (Figure 1C). The arrest period was 15 minutes, after which ECMO was initiated and the heart resumed spontaneous beating (Figure 1F). We saw no differences between these 2 modes of cardiac arrest. Data are presented from the hypoxic cardiac arrest model because of the similarity to existing descriptions of isolated Langendorff heart ischemia/reperfusion. For selected experiments, rat hearts were explanted and retrogradely perfused on a Langendorff system. After stabilization (pre-arrest) for 10 minutes, hearts were subjected to 15 minutes of no-flow ischemia, and coronary perfusion was restored for 30 minutes.13 During explanted ischemia/reperfusion experiments, HEPES-buffered saline (See “Isolation of Left Ventricular Cardiac Myocytes”) was bubbled with 98% oxygen/2% isoflurane. In select experiments, heparinized blood was used to perfuse Langendorff hearts.

Isolation of Left Ventricular Cardiac Myocytes

Within 5 minutes of heart explantation from arrested animals weaned from ECMO (typically within 20 minutes after initiation of ECMO) or Langendorff hearts when specified, left ventricular cardiac myocytes were isolated according to our published protocols.23 Experiments were performed with HEPES-buffered solution containing (in mmol/L) 1 CaCl2, 137 NaCl, 5.4 KCl, 15 dextrose, 1.3 MgSO4, 1.2 NaH2PO4, and 20 HEPES (pH 7.4) maintained at 37.7°C with a feedback control system (IonOptix, Milton, MA).

Myocyte Unloaded Shortening and Relaxation

Myocyte contraction properties, including sarcomeric length, were evaluated with a sarcomeric imaging acquisition system (SarcLen; IonOptix) as previously described.23

Two-Carbon Fiber Technique

Carbon fibers (CFs; 12-µm diameter, kindly provided by Professor Jean-Yves LeGuennec)22 affixed to miniature hydraulic manipulators (SM-28; Narishige, Tokyo, Japan), computer controlled with a piezoelectric translator (P-621.1CL; Physik Instruments, Karlsruhe/Palmbach, Germany) and mounted on a custom-made railing system (IonOptix), were attached to single isolated ventricular myocytes. CFs were stretched axially by the piezoelectric translator controlled with custom software (Matlab; MathWorks, Natick, MA). CF bending and force were measured and analyzed with IonWizard.24 Individual systolic and diastolic cell results were plotted and fitted with a linear regression (Origin; OriginLab, Northampton, MA) to give both end-systolic and end-diastolic length-tension relationships. Data were normalized to the starting systolic force or as the Frank-Starling gain when specified.25

Calcium Transient Measurements

In vivo CICR measurement was performed with a GCaMP6f genetically encoded calcium indicator delivered by an AAV9 vector that was directly injected into the left ventricular apex of anesthetized rats (described in the “Delivery of AAV9-GCaMP6f” below). After 3 or 4 weeks, fluorescence transients were imaged with a custom-built fiberoptic excitation imaging system (AUST Development, LLC) attached to a camera system described previously26,27 and analyzed with ImageJ (NIH). For cell experiments, cardiomyocytes were loaded with 0.2 µmol/L Fluo-5f-AM (Molecular Probes, now part of Thermo Fisher, Waltham, MA) for 30 minutes and then allowed to incubate in dye-free HEPES-buffered saline for an additional 30 minutes to allow de-esterification of the calcium dye. Spatially averaged electrically evoked calcium transients were measured with a standard FITC cube (Chroma, Bella Falls, VT) using the HyperSwitch system (IonOptix). Fluorescence transients were normalized to ΔF/F units.26 Myocyte calcium transient properties, including ΔF/F, rise time, and decay time, were evaluated with a calcium imaging acquisition system (IonOptix) as previously described.23

Delivery of AAV9-GCaMP6f by Direct Intramyocardial Injection

Adult male Sprague-Dawley rats (weight, 275–300 g) were anesthetized with 2% isoflurane in oxygen. A left thoracotomy was performed after intubation and ventilation, and 150 μL normal saline containing AAV9-GCaMP6f (total titer 2e12, Penn Vector Core) was injected into 3 spots at equal volume at the apex of the left ventricular wall. After the successful injection, the chest was closed, and the animal was extubated and allowed to recover. GCaMP6f signal was detected at 3 to 4 weeks after delivery.

Pharmacological Reagents

Epinephrine infusion was performed with 23 µg·kg−1·min−1 for 30 minutes through venous access. In select experiments, control cardiomyocytes were exposed to 100 nmol/L epinephrine in the extracellular solution. For ryanodine experiments, 1 mg dissolved ryanodine (Sigma, St. Louis, MO) was injected into the femoral vein empirically to stop the heart. To inhibit CaMKII, 1 mg KN92 or KN93 (Sigma) or 100 µg myristoylated autocamtide-2–related inhibitory peptide (AIP; Calbiochem or Anaspec) was used. A myristoylated AC3-C peptide (KKALHAQERVDCL, synthesized by Anaspec), an inactive peptide inhibitor of CaMKII, was used as a control.28 To inhibit protein kinase A (PKA), 1 mg H89 (Sigma) was raised in 1 mL of 0.9% normal saline and infused over 10 minutes. Experimental protocols were initiated 10 minutes after infusion. Alda-1, 32.4 mg Alda-1 dissolved in 2 mL 50% polyethylene glycol/50% PBS, was injected intraperitoneally 30 to 60 minutes before the experimental protocol.

Sham Experiments

Sham experiments with animals on ECMO for 30 minutes yielded normal values in isolated cells (data not shown). Control animals refer to no ECMO unless otherwise stated.

Lucigenin-Enhanced Chemiluminescence

Total O2 was measured by lucigenin-enhanced chemiluminescence as described previously.29 Chemiluminescence was recorded by luminometer (Bio-Orbit, Turku, Finland) with 5 μmol/L lucigenin. In parallel experiments, ventricular tissue was incubated in NG-monomethyl-l-arginine (1 mmol/L), diphenylene iodonium (10 µmol/L), or Alda-1 (20 µmol/L). After measurement of baseline readings for 4 minutes, samples were equilibrated and dark adapted for 5 minutes, and chemiluminescence was recorded for 10 minutes. Recordings were performed by researchers blinded to sample identity. Results were expressed as counts per second per milligram of tissue dry weight.

Oxidative Fluorescent Microtopography

O2 was detected in the ventricle with the fluorescent probe dihydroethidium (Molecular Probes) as described previously.29 Cryosections (30 μm) were incubated in physiological buffer for 30 minutes at 37°C with either Alda-1 (20 µmol/L) or mito-TEMPO (10 µmol/L) followed by 5 minutes of dark incubation with 2 μmol/L dihydroethidium. Images were obtained on a confocal microscope at ×60 (Bio-Rad MRC-1024 laser; filter settings: excitation filter, 488 nm; emission filter, 550 nm) and quantified (red intensity times area) with Image-Pro Plus software (Media Cybernetics, Rockville, MD). Analysis was performed by researchers blinded to sample identity. Mean fluorescence was calculated from 4 separate, high-power fields from each quadrant to produce n=1.

Protein Extracts and Western Blot

Heart tissues were homogenized in lysis buffer containing 50 mmol/L Tris-HCl (pH 7.5), 150 mmol/L NaCl, 1% NP-40, 0.5% sodium deoxycholate, 1% SDS, and Halt protease and phosphatase inhibitor cocktail (Thermo Fisher Scientific). Heart tissue homogenates were separated on 4% to 20% TGX gel (Bio-Rad) and transferred to Immobilon-FL polyvinylidene fluoride membrane (Millipore). Membranes were blocked in Odyssey TBS blocking buffer and incubated with respective primary antibodies: CaMKII (Santa Cruz Biotechnology), phospholamban (2D12), RyR (under nonreducing conditions; 34C; Thermo Fisher Scientific), phospho-CaMKII Thr286/287 (D21E4; Cell Signaling), phospholamban pThr17, phospholamban pSer16, RyR2 pSer2814 (Badrilla), and GAPDH (Sigma). IRDye 800CW goat anti-mouse IgG and IRDye 680LT goat anti-rabbit IgG were used as secondary antibodies (LI-COR). Quantitative Western images were acquired with an Odyssey Fc Imager, and images were analyzed with Image Studio Software (LI-COR).


For comparisons between independent samples, the t test using the Welch-Satterthwaite correction for unequal variances was used. One-way Kruskal-Wallis tests were performed for multiple-group comparisons. For significant differences, post hoc testing was performed. P values from post hoc testing were corrected for multiple comparisons with the Bonferroni correction. Exact P values are stated in most cases, and a value of P<0.05 was taken to indicate a statistically significant difference between means. All data are presented as mean±SEM.

For comparisons with nested designs (multiple cells from different numbers of rats), a mixed-effects model was used to account for the correlation between cells from the same rat. The treatment (PMAD versus control) was modeled as a fixed effect, and the animal was modeled as a random effect. We fitted the mixed models using maximum likelihood and tested the significance of the treatment fixed effect.


Myocardial Dysfunction Is Found in a Rodent Model of Cardiac Arrest and Resuscitation

Pressure-volume hemodynamic measurements were made before (Figure 1G) and after (Figure 1H) cardiac arrest. Analysis showed significant decrement in contraction and relaxation after cardiac arrest (end-systolic pressure-volume relationship, −44%; cardiac input, 48%; preload recruitable stroke work, −31%; τ +500%; see Table). These data establish PMAD in our model.

Table. In Vivo and Isolated Cell

ControlPMAD (P Value)
ESPVR, mm Hg/µL1.0±0.100.5±0.1 (0.02)
PRSW, mm Hg58±830±6 (0.02)
CI, (mm Hg/μL)·s93±664±6 (0.005)
τ, ms12±345±6 (7e−11)
ESLTR, au84±1134±6 (0.04)
EDLTR, au44±628±6 (0.02)
Peak PRSWcell, au8±23.0±1 (0.006)

au indicates arbitrary units; CI, contractility index; EDLTR, end-diastolic length-tension relationship; ESLTR, end-systolic length-tension relationship; ESPVR, end-systolic pressure-volume relationship; PMAD, post–myocardial arrest dysfunction; PRSW, preload recruitable stroke work; and τ, isovolumic relaxation constant.

We next examined unloaded sarcomere shortening (%SL) and peak contraction (vc) and relaxation (vr) velocity in isolated adult cells from control and PMAD hearts. For control versus PMAD cells, the %SL was 13±0.3 versus 9±0.3, respectively (P=1.61×106), a 23% decrease in the %SL in unloaded cells (data not shown; n=143 cells from 25 animals and 187 cells from 26 animals for control and PMAD, respectively, for data in the Table), whereas vc was 4.5±0.2 versus 2.6±0.1 µ/ms (P=6.78e−7) and vr was 2.7±0. versus 2.0±0.1 1 µ/ms (P=0.0078; data not shown; n=136 cells from 11 animals and 164 cells from 12 animals for control and PMAD, respectively, for above data).

We further characterized this contractile dysfunction in PMAD by measuring single-cell force and contractility using the 2-CF technique.30Figure 2A shows an adult left ventricular myocyte before and during contraction in response to electric stimulation. Figure 2B and 2C shows the stepwise stretch protocol used to assess myocyte function. Figure 2C shows CF bending for the same stretched cell.30 Corresponding to the derivation by regression of load-independent values in vivo, the end-systolic and end-diastolic length-tension relationships are defined in single cells as the slopes of the linear regressions of systolic and diastolic force (proportional to CF bending; see Methods) and plotted versus sarcomere length (Figure 2D and 2E). In the Table, pooled analysis of the slopes of the ESTLR and EDTLR for both control and PMAD cells is shown, demonstrating both to be depressed (60% and 36% of control, respectively) in PMAD cells (P<0.05). In addition, the Frank-Starling gain was used to compare the change in linear slopes and at each stretched length by dividing the active force by the passive force (Figure 2F).25 These cellular data recapitulated load-varying curves obtained with pressure-volume loops and demonstrated impaired force development. We also explored the force-frequency relationship for both control and PMAD cells (Figure 2G). This demonstrated that the force-frequency relationship was depressed in PMAD cells at all pacing frequencies investigated (0.5–4.0 Hz).

Figure 2.

Figure 2. In vivo and isolated cell load–dependent parameters in control and cardiac arrest. A, Isolated twitching myocyte before (top) and during twitch in response to electric activation demonstrating carbon fiber (CF) bending (CFB). B The sarcomere length (SL; µm) in response to 1-Hz pacing vs time as CF stretch was applied to 3 different end-point lengths demonstrates the cellular Frank-Starling effect. Between each step, the CFs were returned to baseline. Each stretch produced increased sarcomere length change. C, CFB (proportional to force) vs time, demonstrating the increase in both diastolic (passive) and systolic (active) single cell force. D, Stretch-induced peak systolic (□) and baseline diastolic (◼) tension from a control cell. The linear fits to the data represented the end-diastolic length-tension relationships (EDLTR; black line) and end-systolic length-tension relationships (ESLTR; gray line). In this case, the ESLTR was 22 µN/µm (R2=0.96), and the EDLTR was 6.6 µN/µm (R2=0.95). E, Stretch-induced peak systolic (□) and baseline diastolic (◼) tension from a post–myocardial arrest dysfunction (PMAD) cell. In this case, the ESLTR was 11 µN/µm (R2=0.92), and the EDLTR was 6 µN/µm (R2=0.98). F, Preload recruitablestroke work (PRSWcell) vs afterloadcell force for control (□) and PMAD (◼), n= 8 cells from 3 hearts and 8 cells from 2 hearts for control and PMAD, respectively. Exponential models were fit to the data: Frank-Starling gain (FSG)=y0+A1e(−x/t1). For control and PMAD, y0, A1, and t1 were 0.94 and 1.47, 9.5 and 2.7, and 0.78 and 0.54 second, respectively. R2 values were 0.96 and 0.92. N=8 cells from 3 hearts and 8 cells from 2 hearts for control and PMAD, respectively. G, Force-frequency relationship for pooled baseline force (µm) vs pacing frequency for control (◼) and PMAD (□) plotted with SE; n= 8 cells from 3 hearts and 8 cells from 2 hearts for control and PMAD, respectively. H, Example traces for simultaneous calcium-induced calcium release (bottom, gray) and cellular force (top, black) measurements, shown for an isolated cell electrically paced at 1 Hz. The SL was ≈1.87, ≈1.94, and ≈2.0 µm (at slack length and first and second stretch, respectively).

Cardiac Myocytes From Postarrest Animals Exhibit Markedly Enhanced Peak Calcium Transients

Force development is critically regulated by CICR. Previous reports examining postarrest CICR have found little change but did so using high-affinity calcium indicators,13,31 whereas rapid calcium transients were subject to dye kinetic filtering that could both mask important phenotypic differences and alter cellular contractility.26 Our findings were consistent with this effect for Fluo-4 but not for the lower affinity dye Fluo-5f (online-only Data Supplement Figure I). Therefore, we combined measurements of mechanical load and CICR simultaneously in dynamically contracting cardiomyocytes (Figure 2H). It has been reported that CICR does not change with acute stretch in either amphibian cells29 or mammalian heart cells. We observed this to be the case (Figure 2H).

We next explored CICR after cardiac arrest using the approach outlined earlier in Figure 2H. In Figure 3A, a single sweep of a steady-state pacing train is shown for a PMAD and a control cell. Consistent with one report,14 we found postarrest cells to have markedly enhanced peak calcium transients. In pooled analysis (Figure 3B), a significant 290% increase in CICR amplitude was present (ΔF/Fpeak from control [n=97 cells from 11 animals] and PMAD [n=72 cells from 12 animals] were 99±8 and 272±13, respectively; P=1.53×106). We found no difference in the time to peak (tp) of the calcium transient (52±4 and 46±10 milliseconds; P=0.11; same n as above). The time to 90% decay (td) of the CICR in PMAD was prolonged (340±9 versus 456±9 milliseconds; P=4e−15), as was the time constant of decay (τ; 152±5 versus 216±7 milliseconds). We cannot exclude that this effect is from dye kinetic filtering as the dye nears saturation in the PMAD group, rather than a true finding. The augmented CICR was present in isolated cells long after in vivo cardiac arrest (at least 8 hours, which was the longest time point investigated); hence, we labeled this phenomenon calcium long-term potentiation. In contrast, consistent with predicted effects of high-affinity indicators,25 we found that Fluo-4 minimized this difference in CICR (online-only Data Supplement Figure I). At times, the calcium potentiation led to spontaneous delayed calcium increases, a cellular surrogate candidate for delayed afterdepolarizations. In Figure 3C, the top panel shows sarcomere shortening under load, and the bottom panel displays CICR for the same fiber. After stretch (arrow), a delayed calcium increase occurred and induced chaotic contraction without coordinated force generation. In this case, with rapid pacing, the fibrillatory behavior converted to organized contraction. All cells investigated from all PMAD hearts exposed to load displayed this phenomenon (n=12 from 4 rat hearts). Rarely, load-independent spontaneous delayed calcium increases were seen, as shown in Figure 3D. Such behavior was never seen in control cells.

Figure 3.

Figure 3. Cardiac calcium memory after cardiac arrest. A, Example calcium-induced calcium release (CICR) transients measured by Fluo-5f in control vs post–myocardial arrest dysfunction (PMAD) and PMAD in which arrest was generated in the presence of KN93 (PMAD-KN93) or ryanodine (PMAD-Rya), as well as a control animal exposed to intravenous infusion of epinephrine (Epi) for 30 minutes. Cells were paced steadily at 1 Hz. B, Pooled analysis of ΔF/Fpeak for all conditions tested. The ΔF/Fpeak was significantly higher in animals exposed to epinephrine infusion and cardiac arrest. Ryanodine and KN93 reduced ΔF/Fpeak to control levels in PMAD ventricular myocytes. Wild-type was no different than sham. N=114, 135, 89, 16, 17, 9, 11, 15, 11, 31, 20, 51, 32, 119, 19, and 11 cells for control, PMAD, PMAD– autocamtide-2–related inhibitory peptide (AIP), PMAD-AIP-INactive (AC3-C), PMAD-KN93, PMAD-KN92 PMAD-Rya, Epi infusion, control-KN93 (Cntr-KN93), control ryanodine (Cntr-Rya), epi infusion with ryanodine (Epi-Rya), PMAD-Langendorff-Saline (PMAD-Lan-Sal), PMAD-Langendorff-PMAD blood (PMAD-Lan-Blood), brainstem herniation (BSH), BSH-ryanodine (BSH-Rya), and PMAD-H89. Cntr-Sham experiments demonstrated no difference compared with control. P values compared with control for PMAD, Epi infusion, BSH, PMAD-Lan-Blood were 6.4e−24, 2.2e−18, 2e−25, and 2e−14 (corrected for multiple comparisons), respectively. C, Example CICR transients (red) and sarcomere shortening (black) before and after carbon fiber stretch. Top set showed stretch followed by a delayed calcium release (black arrow), initiating fibrillatory behavior in both traces. Bottom showed termination of “arrhythmia” 30 seconds later through rapid pacing. D, Example CICR transients from 2 different PMAD cells. Top shows CICR in response to electric stimulation followed by delayed calcium transients. Bottom shows spontaneous calcium transients competing for paced rhythm. E, Pooled analysis of velocity of contraction (Vc) vs sarcomere length for control and PMAD. Baseline (filled) and stretched (open) for pairs of similar stretches from ≈1.8 to ≈ 2.0 µm. PMAD (red) vs control (black); n= 8 and 9 cells from 3 and 4 hearts for PMAD and control hearts, respectively. F, Pooled data comparing unloaded sarcomere shortening (%SL) and ΔF/Fpeak of CICR under control, PMAD, epinephrine infusion in animal but not cells (EPI inf), and epinephrine in isolated cells (EPI cell) but not in whole animal, as well as PMAD in the presence of Alda-1 (PMAD-Alda, n=25 cells from 4 animals). N for control, PMAD, and Epi infusion is the same as above. G, Baseline force (BSF) in example cells paced steadily at 1 Hz in control, PMAD, PMAD-KN93, and PMAD-Rya cells. H, Pooled data for BSF measurements at 1-Hz steady-state pacing in control and PMAD in the presence and absence of KN93 or ryanodine. N= 8, 8, 5, and 4 cells for control, PMAD, PMAD-KN93, and PMAD-Rya.

Simultaneous measurements of CICR and force generation allowed us to test whether length-dependent activation, a putative mechanism for the Frank-Starling effect,30 was impaired in PMAD by comparing contraction velocity immediately before and after stretch (Figure 3E). Care was taken to equalize baseline and stretched sarcomere lengths. The poststretch change in velocity was similar between control and PMAD, but the absolute velocity was also lower in PMAD (ratio of poststretch to prestretch vc, 1.7±0.2 versus 1.8±0.2 for control versus PMAD; P=0.5). This is important because, although PMAD is associated with reduced baseline myofilament calcium sensitivity, the response to stretch (length-dependent activation) is preserved.

To examine the multidimensional relationship between force, frequency, and CICR, we plotted these together in 3-dimensional space. This demonstrated the reduced myofilament sensitivity seen at all frequencies and the extent to which the potentiation of the calcium transient abrogates the reduced contractile force (Figure 4A and 4B).

Figure 4.

Figure 4. Calcium force-frequency relationships after cardiac arrest. A, Plot of force (z axis), pacing frequency (x axis), and calcium expressed as ΔF/Fpeak (y axis) for control cells at slack length loaded with Fluo-5f and simultaneously stretched with carbon fiber. N=4 cells from 2 control hearts. The curve showed a steep curve shifted to lower ΔF/Fpeak. The color represents force and is coded from low to high with blue to red, B, Similar plot in postarrest cells. N=4 cells from 2 PMAD. Compared with A, the curve is shifted to a flatter curve despite also being shifted to higher ΔF/Fpeaks (rightward) at all pacing frequencies. Force was color coded with the same palette as in A, and all scale bars were identical.

Because previous studies in vitro have not found an augmented CICR after arrest, we compared our data with data from cells isolated from saline-perfused bubble-oxygenated Langendorff resuscitated hearts. Cells from these hearts showed a more minor depression of %SL and no difference in CICR compared with controls (Figure 3B), in agreement with most reported results.6,13 However, when the control Langendorff heart was perfused with oxygenated and pH-controlled blood from a cardiac arrest animal, augmented CICR was demonstrated (PMAD-Lan-Blood, Figure 3B), suggesting a dependency of effect not on neural innervation of the heart but on an intact circulation. Because cardiac arrest is known to be associated with marked increases in both catecholamines18,32 and reactive oxygen species, we hypothesized these to be key mechanisms.

Potentiation of CICR Begins In Vivo

Although responses in isolated cells are often used as a surrogate for in vivo effects and the control animals gave us confidence that this effect was related to the cardiac arrest, the possibility remained that this was an artifact of that isolation process for those cells. To assess whether that CICR was augmented in vivo, we transduced rat hearts with AAV9-GCaMP6f vectors and measured in vivo CICR using a custom-built fiberoptic imaging system (AUST Development, LLC) tunneled under the skin to the heart (Figure 5A and 5B). Using this system, we were able to make sequential measurements in the same rat (eg, a week apart in Figure 5C). An example fluorescent image (acquired through the fiberoptic placed on the heart) from baseline and peak calcium transient is shown in Figure 5B. We then performed a cardiac arrest experiment as described above with the additional measurement of CICR by way of genetically engineered calcium indicator fluorescence. CICR after arrest was potentiated in this in vivo measurement, similar to our findings at the cellular level from this same model and the blood-perfused preparation described above, but in contrast to the traditional saline-perfused Langendorff preparation. Furthermore, the proarrhythmic potential of delayed calcium afterdepolarizations in this postarrest setting was seen, in agreement with our in vitro data (Figure 5E). We found the peak ΔF/F to be ≈300% increased after cardiac arrest compared with the internal baseline prearrest control (n=2 hearts). This increase, when corrected for heart rate by atrial pacing to similar heart rates, is less dramatic and approximates ≈200%.

Figure 5.

Figure 5. In vivo calcium-induced calcium release (CICR) by AAV9-GCaMP6f fluorescence. A, Example fluorescence measurements of CICR (y axis, ΔF/F, vs x axis, time). In vivo fluorescence imaging was performed 3 or 4 weeks after GCaMP6f delivery in the same live rat (3w P.D. or 4w P.D.). B, Example image of raw fluorescence seen through fiberoptic on the heart by the camera at baseline and peak fluorescence during a calcium transient. C, Example cardiac arrest experiment from a different rat while measuring CICR fluorescence continuously through the experiment (y axis, ΔF/F, vs x axis, time). Shown are measurements at baseline, after cardiac arrest, on extracorporeal membrane oxidation (ECMO) demonstrating delayed calcium release initiating fibrillatory behavior similar to what was seen in isolated cells, and finally off ECMO.

Catecholamine Excess Reproduces Calcium Potentiation

It has been demonstrated that catecholamines are dramatically elevated with in vivo cardiac arrest.18 To test the hypothesis directly in vivo that catecholamine signaling was responsible for the augmented CICR, we blunted catecholamine signaling with β- and α-blockade (labetalol). However, elevated catecholamines are also required for survival, and the presence of signaling antagonists in vivo prevented adequate resuscitation, as might be expected. Therefore, to examine the role of catecholaminergic activation in the promotion of cardiac myocyte calcium potentiation in vivo, a stable, ventilated animal was infused with epinephrine for 30 minutes demonstrating its profound elevation after arrest. Infusion induced a typical adrenergic response, characterized by an increase in heart rate and mean arterial pressure. On termination of epinephrine infusion before cell isolation, blood pressure and heart rate normalized (blood pH was also normal) within 10 minutes. In isolated cardiomyocytes from this model, a pronounced increase in CICR was seen similar to after cardiac arrest (Epi, Figure 3A and B). We further explored whether cardiac myocyte calcium potentiation was sustained after brainstem herniation, a condition known to induce a physiological endogenous catecholamine surge.33 This led to a similar dramatic increase in heart rate and blood pressure. In cells isolated from these hearts, a similar increase in ΔF/Fpeak was seen (271±21; n=110 cells from n=9 animals; P=1e−14 versus control; Figure 3B, brainstem herniation, and online-only Data Supplement Figure 2II).

We also added epinephrine directly to the cell bath of control cells, which, as expected, acutely manifested a markedly increased contractile performance (%SL, 20±0.5) in association with a more modest increase CICR (ΔF/Fpeak, 141±0.53%). This was in marked contrast to the prolonged administration in vivo as described earlier.

Cardiac Myocyte Calcium Potentiation Is Dependent on CaMKII and Ryanodine but Not on PKA

Because a link between subacute and chronic β-adrenergic signaling– and CaMKII-, rather than the acute PKA-, dependent pathways has been observed,3436 we tested whether CaMKII inhibition could block the induction of CICR potentiation in our in vivo cardiac arrest model. As shown in Figure 3B, AIP, a CaMKII inhibitor, given before and during cardiac arrest, abolished the CICR increase in both the PMAD and epinephrine-infusion models. The lower-affinity inhibitor KN93 blunted cardiac myocyte calcium potentiation in PMAD (Figure 3A and 3B) but not in epinephrine infusion. In the presence of AIP or KN93, no delayed afterdepolarizations or stretch-induced cellular fibrillatory behavior was seen. Because CaMKII activation is partly mediated by calcium-dependent pathways and CaMKII is known to be localized to the dyad, we tested whether inhibiting CICR during cardiac arrest would blunt cardiac myocyte calcium potentiation. We found that both for epinephrine infusion and PMAD, simultaneous ryanodine application blocked the augmentation in CICR (Figure 3B), suggesting that calcium release by RyR was a critical contributor (In the presence of ryanodine, animals were supported with ECMO). In contrast, the PKA inhibitor H89 did not prevent the increase in CICR either before or during cardiac arrest (Figure 3B), suggesting that CaMKII activation was a critical component. The sham inhibitor KN92 had no effect on CICR in PMAD (Figure 3B).

Cardiac Myocyte Calcium Potentiation Supports Impaired Cardiac Function

We measured both %SL (Figure 3F) and force (Figure 3G and 3H) in cells isolated from PMAD hearts that were exposed to CaMKII inhibitors, KN93, AIP, or ryanodine, during the initial cardiac arrest to inhibit CICR augmentation. We found that both %SL and baseline force were lower under these conditions, suggesting that the augmented CICR is necessary to support cardiac contractility after cardiac arrest.

CaMKII-Dependent Phosphorylation in PMAD

In the brain, CaMKII is a mediator of long-term potentiation, a form of synaptic plasticity, and is associated with enhanced calcium signaling.37,38 CaMKII activity is important for calcium-regulated/mediated activities such as excitation-contraction coupling and excitation-transcription coupling.39 CaMKII activity may also become Ca/CaM autonomous, and such activity is increased in myocardial disease, contributing to apoptosis, arrhythmia, and disrupted excitation-contraction coupling and excitation-transcription coupling, leading to pathological hypertrophy.39 CaMKII and its targets, RyR2 and phospholamban, have also been shown to be critical in cardiac ischemia/reperfusion injury.40,41 To assess CaMKII activation/autophosphorylation as a potential mediator of cardiomyocyte calcium potentiation in our in vivo cardiac arrest model, we examined the phosphorylation status of CaMKII Thr287 (PT287) and total CaMKII protein in ventricular tissue lysates after cardiac arrest (Figure 6A). After 30 minutes of cardiac arrest and 30 minutes of ECMO reperfusion, the ratio of pCaMKII PT287 to total CaMKII nearly doubled in the PMAD group compared with the control group (Figure 6C), whereas the total CaMKII protein remained unchanged (Figure 6D), indicating an activation of CaMKII. This activation was reduced by 30% in the presence of the CaMKII inhibitor AIP given 30 to 40 minutes before the onset of cardiac arrest (Figure 6C). We also examined the phosphorylation state of RyR2 and phospholamban, the targets of both CaMKII and PKA, at the sarcoplasmic reticulum. CaMKII-dependent phosphorylation of phospholamban at Thr17 (PT17) showed similar patterns (Figure 6E and 6F). The ratio of PT17 to total phospholamban increased nearly 2-fold in the cardiac arrest group compared with the control group, and PT17 phosphorylation was abolished by AIP treatment before cardiac arrest. In contrast, PKA-dependent phosphorylation of phospholamban at Ser16 is not different between the cardiac arrest group and the control group (Figure 6J).

Figure 6.

Figure 6. Ca2+/calmodulin-dependent protein kinase II (CaMKII)-dependent phosphorylation is increased in post–myocardial arrest dysfunction (PMAD) and can be inhibited by autocamtide-2related inhibitory peptide (AIP) and Alda-1. A and B, Example Western blots showing CaMKII-dependent phosphorylation and protein level of CaMKII, phospholamban (PLN), and ryanodine receptor 2 (RyR2) in the control group, PMAD model (30 minutes of cardiac arrest and 30 minutes of extracorporeal membrane oxidation reperfusion), and PMAD in the presence of AIP (A) and Alda-1 (B), both given 30 minutes before the onset of cardiac arrest. C through I,Quantitative summary of Western blot analysis. CaMKII-dependent phosphorylation of CaMKII at T287 (C) and phosphorylation of PLN at T17 (E) increased significantly in PMAD and was inhibited by AIP, whereas the protein level of CaMKII (D) and PLN (F) did not change significantly. CaMKII autophosphorylation (PT287) in PMAD was also inhibited by Alda-1 (C). CaMKII-dependent RyR2 phosphorylation at S2814 did not vary considerably in PMAD or with AIP or Alda-1 treatment, illustrated as PS2814/RyR2 in G. However, the RyR2 protein level surged 7-fold in PMAD compared with the control group (RyR2/GAPDH in H); overall pRyR2 PS2814 was also shown to increase dramatically (normalized to GAPDH in I). Neither AIP nor Alda-1 treatment reduced the protein level or total RyR2 phosphorylation at S2814 (H and I). J,Western blot and quantitative assessment showed that protein kinase A phosphorylation of PLN at S16 and PLN protein level were not different between the PMAD and control groups. K. Representative Western blot and quantitative summary showed that CaMKII was activated after PMAD but not epinephrine (Epi) infusion for 30 minutes. Graph data represent the mean±SEM of each group. P values were reported by ordinary 1-way ANOVA with multiple comparisons (CI and K) or t test (J). *P<0.05. **P<0.005.

The most striking change during PMAD was found for RyR2, on the detection of both total protein and phosphorylation at Ser2814 (PS2814; Figure 6A). However, the ratio of PS2814 to total RyR2 did not vary significantly (Figure 6G). Signals for RyR2 protein and PS2814 surged ≈7-fold in PMAD (Figure 6H and 6I) and were not significantly altered by AIP or Alda-1 treatment. In addition, a single band of RyR2 protein was detected in hearts from the control group, whereas a doublet of bands became prominent after cardiac arrest, suggesting some form of oligomerization or protein modification (see Discussion). Because similar calcium potentiation was observed in cardiomyocytes isolated from rats treated with epinephrine and in PMAD (Figure 3B), the phosphorylation status of CaMKII was examined in heart tissues collected after 30 minutes of epinephrine infusion. CaMKII phosphorylation was also increased in this model, but to a lesser extent than seen during cardiac arrest (Figure 6K), leading us to consider the contributory role of oxidative stress (reactive oxygen species) in the effects seen in vivo.

Cardiac Arrest Increased Myocardial O2•− Generation Predominantly From Mitochondria

We measured myocardial superoxide (O2) production using lucigenin-enhanced chemiluminescence. We found it to be elevated after epinephrine infusion but significantly more elevated after PMAD (70% increase in total chemiluminescence compared with control; P<0.001; Figure 7A). When ventricular tissue was incubated with the nitric oxide synthase inhibitor NG-monomethyl-l-arginine or the NADPH oxidase inhibitor diphenylene iodonium, total O2 levels were not significantly different, suggesting that the major contribution of O2 was mitochondrial. To confirm the chemiluminescence findings and to investigate further the cellular source of O2, we measured its production using dihydroethidium oxidative fluorescence microtopography (Figure 7B), again finding significant O2 production in PMAD. Finally, to confirm the source of O2, we used the mitochondria-targeted antioxidant with O2 and alkyl scavenging properties Mito-TEMPO. Mito-TEMPO led to a significant reduction in O2 generation (P<0.01; Figure 7B), confirming the dominant mitochondrial source.

Figure 7.

Figure 7. Cardiac arrest hearts demonstrated increased myocardial O2generation, which can be inhibited by Alda-1. A, Myocardial O2 production, measured by lucigenin, was mildly increased in hearts treated with epinephrine but markedly elevated after cardiac arrest (P<0.01). The NADPH oxidase inhibitor diphenylene iodonium (DPI) and the nitric oxide synthase inhibitor NG-monomethyl-l-arginine (L-NMMA) did not change O2 generation compared with uninhibited controls; however, the aldehyde dehydrogenase type 2 activator Alda-1 did inhibit O2 (*P<0.01). B, Dihydroethidium (DHE) fluorescence microscopy allowed topographical assessment of O2 production in the ventricular wall and showed a mild increase in O2 production with epinephrine infusion compared with a marked increase after cardiac arrest (†P<0.01). Alda-1 inhibited O2 generation. Mito-TEMPO, used as the mitochondria-targeted antioxidant with superoxide and alkyl scavenging properties, confirmed that the majority of O2 being generated was from mitochondria (*P<0.01 vs uninhibited). Myocytes exhibited green autofluorescence. White arrows denoted endothelial O2-producing cells. Scale bar: 5 μm. PMAD indicates post–myocardial arrest dysfunction.

Alda-1 Reduces Calcium Potentiation and Improves Cardiac Performance in PMAD

Because the augmented CICR triggered by catecholamines and maintained by CaMKII was supportive of cardiac function and survival in the early stages, we looked toward downstream effects of oxygen radicals as a potential therapeutic intervention. Recently, activation of aldehyde dehydrogenase type 2 by Alda-1 has been shown to play a pivotal role in cardioprotection in the setting of myocardial infarction through detoxification of O2-induced reactive aldehydes (such as 4-hydroxynonenal acetaldehyde) to less reactive acids (such as 4-hydroxynon-2-enoic acid and acetic acid).42 We hypothesized the reactive aldehydes were playing an important role in PMAD. In keeping with this hypothesis, we found 4-hydroxynonenal adducts to be significantly increased after cardiac arrest, and administration of Alda-1 in vivo during cardiac arrest significantly decreased both 4-hydroxynonenal adducts (online-only Data Supplement Figure III) and O2 to levels seen after Mito-TEMPO inhibition (P<0.01 versus control; Figure 7A and 7B). Alda-1 also had a dramatic beneficial impact on PMAD both in vitro and in vivo. At the cellular level, Alda-1 treatment blunted the increase in CICR seen in PMAD by 30%, similar to CaMKII inhibition (Figure 3B). However, although myofilament calcium sensitivity was unchanged with CaMKII inhibition, it was enhanced with Alda-1 (Figure 3F). In addition, immunoblotting of CaMKII and autophosphorylated CaMKII revealed that in hearts exposed to Alda-1 during cardiac arrest, T287 autophosphorylation was reduced by 30%, similar to the effect of AIP (Figure 6B and 6C). As expected, CaMKII inhibition had no impact on 4-hydroxynonenal adduct formation, whereas Alda-1 reduced 4-hydroxynonenal adduct formation after cardiac arrest (online-only Data Supplement Figure III). We also found that Alda-1 reduced malondialdehyde fluorescence after myocardial infarction in a separate pig model (data not shown).

Alda-1 Improved Outcome After Cardiac Arrest

We found weaning from resuscitation (defined as reaching 20% of the prearrest mean arterial pressure) was successfully achieved in 5 of 8 animals (62.5%) treated with Alda-1 compared with only 4 of 30 animals (13%) in the absence of Alda-1 (Figure 8D). Moreover, with Alda-1 administration, we found that whole-animal cardiac contractility, as indexed by the preload recruitable stroke work, contractility index, and τ, returned to near control levels (Figure 8A-8C and theTable). Alda-1 had no effect on these same parameters before arrest (data not shown).

Figure 8.

Figure 8. Alda-1 improved in vivo contractility and recovery rate to spontaneous circulation after cardiac arrest. A, Summary data for preload recruitable stroke work (PRSW) for control, PMAD, and PMAD-Alda (P=0.04). B, Summary data for cardiac input (CI) for control, PMAD, and PMAD-Alda. C, Summary data for tau for control, PMAD, and PMAD-Alda. D, Alda-1 improved chances of weaning from ECMO resuscitation. P values in graphs are with respect to control using post hoc analysis. Comparisons between groups were performed with Kruskall-Wallis nonparametric tests. E, Proposed mechanism for PMAD, mediated by calcium overload, Ca2+/calmodulin-dependent protein kinase II (CaMKII), and oxidative stress (reactive oxygen species [ROS]). Catecholamines released during cardiac arrest and the ischemic/reperfusion process triggered cytosolic Ca2+ influx and sarcoplasmic reticulum Ca2+ release/uptake. The activation of ryanodine receptor 2 (RyR2; by oxidative stress or other unknown mechanism**) also increased calcium-induced calcium release (CICR). Elevated cytosolic Ca2+ -activated CaMKII further enhanced CICR and led to mitochondrial Ca2+ overload. This Ca2+ overload contributed to excess ROS (O2) and reactive aldehyde (4- hydroxynonenal [4-HNE]) generation, more CaMKII activation, and other cellular damages. AIP inhibited CaMKII autophosphorylation but did not improve cardiac function after arrest. Alda-1 reduced oxidative stress, inhibited CaMKII autophosphorylation, and improved cardiac function. ALDH2 indicates aldehyde dehydrogenase type 2; βAR, β-adrenergic receptor; LTCC, L-type calcium channel; MCU, mitochondria calcium uniporter; NCX, Na+/Ca2+ exchanger; NHE, Na+/H+ exchanger; PKA, protein kinase A; and PLN, phospholamban.


For >3 decades, PMAD has been recognized as the critical factor in recovery and survival after cardiac arrest.5,6 To date, most research focused on ischemia/reperfusion has used variants of a Langendorff isolated heart model. Here, we present an in vivo model of PMAD, which, in leaving the neurohumoral axis intact, uncovered a significant increase in CICR in vivo that continued in isolated cardiac myocytes. A similar increase in CICR was reproduced when blood from an arrested animal was used as perfusate in the Langendorff system. We did not see this potentiation of CICR in the saline-perfused Langendorff model. This cardiac myocyte calcium “memory” seems to be triggered by a combination of catecholamines and reactive oxygen species and maintained by autophosphorylation of CaMKII with subsequent phosphorylation of RyR2 and phospholamban.

The role of CAMKII in cardiomyopathy has been described in seminal investigations.34,4345 Its activation is important in the generation of electrical storm46,47 and in reperfusion-related arrhythmias,48 whereas its inhibition can prevent the occurrence of ventricular arrhythmias.49 In our study, we found that inhibition of CaMKII also reduced cellular stretch–induced arrhythmias. We propose that oxidative stress and aldehydic adduct formation contribute to acute myofilament dysfunction and impaired myofilament calcium sensitivity, which is partially compensated for by this increase in CICR. Consistent with this hypothesis, the small-molecule activator of aldehyde dehydrogenase type 2, Alda-1, dramatically improved cardiac myofilament calcium sensitivity at the whole-heart and cellular levels by acting upstream of calcium signaling to reduce both mitochondrial O2 production and toxic aldehydic adduct formation. In so doing, myofilament calcium sensitivity after arrest was improved, and autophosphorylation of CaMKII and CaMKII augmentation of CICR were simultaneously reduced. This proposed mechanism (Figure 8E) has similarities to neuronal plasticity in which CaMKII senses phasic changes and then augments the same postsynaptic signals.37 We speculate that, just as in the brain, CaMKII induces long-term potentiation38 in the heart. The unique features of CaMKII autophosphorylation and its ability to follow calcium changes phasically37 allow integration of calcium signals in the “postsynaptic” dyadic cleft.34,35,41,43,5052

In an attempt to track the timing of signaling events that accompanied the calcium potentiation during cardiac arrest and ECMO reperfusion, we traced the dynamics of CaMKII signaling in our in vivo PMAD model. We varied the time of cardiac arrest and reperfusion/resuscitation and indeed observed variations in the phosphorylation status of CaMKII (PT287) and phospholamban (PT17; online-only Data Supplement Figure IV). It is worth noting that the surge of total RyR2 protein and PS2814 detection were prominent in almost all the heart samples collected after cardiac arrest/reperfusion despite variation in CaMKII activation. It is likely that in addition to CaMKII, RyR2 is responsive to other signaling pathways involved in oxygen homeostasis and return of spontaneous contraction in this cardiac arrest rat model. The overall elevation of RyR2 in its active form (PS2814) would independently increase sarcoplasmic reticulum calcium release and contribute to augmented CICR. The elevation of RyR2 signal seen in the Western blots could indicate a rapid increase in RyR2 protein after cardiac arrest and reperfusion, although it is unclear which signals may lead to such induction. Alternatively, RyR2 proteins may be posttranslationally modified in response to cardiac arrest/reperfusion and become stabilized against degradation. It is also possible that cardiac arrest and oxidative stress resulted in an RyR2 conformational change, which leads to increased antigenicity. The Western blots of RyR2 and PS2814 were performed under nonreducing conditions in which the RyR2 protein could retain some folded structure. It has been reported that oxidative stress can induce conformational changes in RyR2.53 Such conformational changes could make the epitope more accessible to the antibodies and result in higher detectable signals.

The sarcoplasmic reticulum Ca2+-ATPase regulator phospholamban is a common target of both CaMKII (PT17) and PKA (phosphorylation at Ser16). Examination of these phosphorylation sites after in vivo cardiac arrest/reperfusion demonstrated clearly that the CaMKII but not PKA pathway was activated after cardiac arrest because the ratio of PT17 to phospholamban surged while the ratio of phosphorylation at Ser16 to phospholamban remained unchanged. Inhibition of CaMKII with AIP before cardiac arrest reduced CaMKII activation and abolished phospholamban PT17 phosphorylation and calcium potentiation. However, in vivo and in vitro contractility was not improved. It is possible that CaMKII-mediated phospholamban PT17 phosphorylation is not contributing to PMAD. A previous study on the phospholamban S16A/T17A (nonphosphorylatable phospholamban) mutant mice showed that a CaMKII-dependent increase of phospholamban PT17 opposes rather than contributes to ischemia/reperfusion damage.40 Ideally, genetic models such as phospholamban S16A/S17A mutant mice, CaMKII mutant mice (overexpression and knockouts), and RyR2 S2814D, S2814A mutant mice would be studied in this in vivo cardiac arrest model to define the molecular pathway and to clarify the roles of individual contributors. However, the technical challenge of reproducing cardiac arrest and ECMO resuscitation in the mouse has not yet been met.

Our work has limitations. First, these studies have not yet been performed in humans. Second, we did not explore the role of the action potential because of technical limitations related to photostability of fluorescence voltage dyes. Finally, quantifying CaMKII oxidation was technically challenging and is not reported here.


Using new approaches for simultaneous force and CICR measurement in vitro and GECI enabled calcium fluorescence measurements in vivo, we present a novel mechanism of PMAD (Figure 8E). We propose that hypoxia triggers a surge in both catecholamines and reactive oxygen species that triggers an increase in CICR that is further maintained by autophosphorylation of CaMKII. In addition, stretch of cardiac cells may increase calcium uptake and sarcoplasmic reticulum releasability, aiding CICR.16,54 This increased CICR supports contraction in the face of reduced myofilament sensitivity. We show that inhibiting aldehydic adduct formation during arrest/reperfusion increases myofilament sensitivity, reduces CaMKII activation, and improves survival. These mechanisms should be explored further as potential targets for human therapeutic trials.


The authors thank Dr Peter Lee for his construction of the MATLAB control device for CF stretch. The authors also thank Dr Kathia Zaleta for assisting in experiments.


*Drs Woods, Shang, and Taghavi contributed equally.

Sources of Funding, see page 975

The online-only Data Supplement is available with this article at

Circulation is available at

Correspondence to: Christopher E. Woods, MD, PhD, Division of Cardiology, Arrhythmia Section, Palo Alto Medical Foundation, 1501 Trousdale Dr, Second Floor, Burlingame, CA 94010 or Euan A. Ashley, MB ChB, DPhil, FRCP, Division of Cardiovascular Medicine, Stanford University, 300 Pasteur Dr, Falk CVRB, Stanford, CA 94305. E-mail or


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Clinical Perspective

What Is New?

  • We developed a rodent model of cardiac arrest using extracorporeal membrane oxygenation resuscitation. We observed calcium-induced calcium release in vivo before and after resuscitation using a genetically encoded calcium sensor and a novel fiberoptic catheter imaging system. In cardiomyocytes isolated from this model, we assessed mechanical load and calcium-induced calcium release simultaneously using the microcarbon fiber technique and observed reduced myocardial performance but potentiation of calcium-induced calcium release.

  • This “memory” phenomenon was Ca2+/calmodulin-dependent protein kinase II dependent and mechanistically similar to long-term potentiation, a form of hippocampal learning. Because aldehydic adduct formation and oxidative stress were high after arrest, we used a small-molecule activator of aldehyde dehydrogenase type 2 that indirectly restored calcium signaling and improved cardiac performance.

What Are the Clinical Implications?

  • Despite return of circulation, survival after sudden cardiac arrest is low, in part as a result of post–myocardial arrest dysfunction. Our data demonstrate that this myopathy is associated with an increase in calcium-induced calcium release, which seems to support the function of the heart short term but carries the potential for long-term negative consequences. In our model, a small-molecular activator of aldehyde dehydrogenase restored heart function and improved survival.

  • This may represent a novel therapeutic approach to a condition with a high mortality.


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