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Cell Calcium 55 (2014) 104– 118
Contents lists available at ScienceDirect
Cell Calcium
jo u rn al h om epage: www.elsev ier .com/ locate /ceca
ffects of endogenous cannabinoid anandamide
onxcitation–contraction coupling in rat ventricular myocytes
ina T. Al Kurya, Oleg I. Voitychukd, Ramiz M. Alia, Sehamuddin
Galadari c,eun-Hang Susan Yange, Frank Christopher Howarthb,
Yaroslav M. Shubad, Murat Oza,∗
Laboratory of Functional Lipidomics, Department of Pharmacology,
College of Medicine and Health Sciences, UAE University, Al Ain,
Abu Dhabi, Unitedrab EmiratesDepartment of Physiology, College of
Medicine and Health Sciences, UAE University, Al Ain, Abu Dhabi,
United Arab EmiratesDepartment of Biochemistry, College of Medicine
and Health Sciences, UAE University, Al Ain, Abu Dhabi, United Arab
EmiratesBogomoletz Institute of Physiology and International Center
of Molecular Physiology, National Academy of Sciences of Ukraine,
Kyiv-24, UkraineDepartment of Biological Sciences, Schmid College
of Science and Engineering, Chapman University, One University
Drive, Orange, CA 92866, USA
r t i c l e i n f o
rticle history:eceived 19 September 2013eceived in revised form5
November 2013ccepted 26 December 2013vailable online 5 January
2014
eywords:ndocannabinoidnandamideentricular myocytesontraction
a b s t r a c t
A role for anandamide (N-arachidonoyl ethanolamide; AEA), a
major endocannabinoid, in the cardio-vascular system in various
pathological conditions has been reported in earlier reports. In
the presentstudy, the effects of AEA on contractility, Ca2+
signaling, and action potential (AP) characteristics
wereinvestigated in rat ventricular myocytes. Video edge detection
was used to measure myocyte short-ening. Intracellular Ca2+ was
measured in cells loaded with the fluorescent indicator fura-2 AM.
AEA(1 �M) caused a significant decrease in the amplitudes of
electrically evoked myocyte shortening andCa2+ transients. However,
the amplitudes of caffeine-evoked Ca2+ transients and the rate of
recovery ofelectrically evoked Ca2+ transients following caffeine
application were not altered. Biochemical studiesin sarcoplasmic
reticulum (SR) vesicles from rat ventricles indicated that AEA
affected Ca2+-uptake andCa2+-ATPase activity in a biphasic manner.
[3H]-ryanodine binding and passive Ca2+ release from SR vesi-cles
were not altered by 10 �M AEA. Whole-cell patch-clamp technique was
employed to investigate the
ntracellular calciumentricular action potential
effect of AEA on the characteristics of APs. AEA (1 �M)
significantly decreased the duration of AP. Theeffect of AEA on
myocyte shortening and AP characteristics was not altered in the
presence of pertussistoxin (PTX, 2 �g/ml for 4 h), AM251 and
SR141716 (cannabinoid type 1 receptor antagonists; 0.3 �M)or AM630
and SR 144528 (cannabinoid type 2 receptor antagonists; 0.3 �M).
The results suggest thatAEA depresses ventricular myocyte
contractility by decreasing the action potential duration (APD) in
a
B1 an
manner independent of C
. Introduction
Endocannabinoids belong to a family of polyunsaturated
fattycid-based compounds that mimic most of the effects of
tetrahy-rocannabinol, the active ingredient of the marijuana
plant
Abbreviations: AP, action potential; APD, action potential
duration; APD60, actionotential duration at 60% level of
repolarization; AMP, amplitude; AA, arachidoniccid; APIII,
antipyrylazo III; BSA, bovine serum albumin; DMSO,
dimethylsulphox-de; DHP, dihydropyridine; NAEs,
N-acylethanolamines; AEA, N-arachidonoylthanolamide, anandamide;
NEM, N-ethylmaleimide; NO, nitric oxide; NT, normalyrode; PTX,
pertussis toxin; RCL, resting cell length; metAEA,
R-methanandamide;R, sarcoplasmic reticulum; THALF, time from peak
to half; TPK, time to peak; TRP,ransient-receptor potential.∗
Corresponding author at: Department of Pharmacology and
Therapeutics, Col-
ege of Medicine and Health Sciences, UAE University, P.O. Box
17666, Al Ain, Abuhabi, United Arab Emirates. Tel.: +971 3 7137523;
fax: +971 3 7672033.
E-mail address: murat [email protected] (M. Oz).
143-4160/$ – see front matter © 2014 Elsevier Ltd. All rights
reserved.ttp://dx.doi.org/10.1016/j.ceca.2013.12.005
d CB2 receptors.© 2014 Elsevier Ltd. All rights reserved.
Cannabis sativa. The most widely studied endogenous
cannabi-noids are N-arachidonoyl ethanolamide (AEA) or anandamide
and2-arachidonylglycerol [1,2]. In recent years extensive
researchfocusing on the biological actions of these molecules has
revealedthe existence of an endocannabinoid system that regulates
sev-eral physiological processes and pathological conditions [3].
It wassuggested that the endocannabinoid system consists of at
leastthe endocannabinoid receptors (such as CB1 and CB2
cannabi-noid receptors), the enzymes regulating the synthesis (such
asphospholipase-D, and monoacylglycerol), and the degradation(such
as fatty-acid amide hydrolase and lipases) processes, andthe
proteins involved in their transport across the biologicalmembranes
[1,3]. The CB1 receptors are located in the brainand several
peripheral tissues including the heart and the vas-
culature. The CB2 receptors, on the other hand, are
expressedprimarily in the immune system but recently their presence
inthe brain, myocardium, and smooth muscle cells have also
beendemonstrated [3].
dx.doi.org/10.1016/j.ceca.2013.12.005http://www.sciencedirect.com/science/journal/01434160http://www.elsevier.com/locate/cecahttp://crossmark.crossref.org/dialog/?doi=10.1016/j.ceca.2013.12.005&domain=pdfmailto:[email protected]/10.1016/j.ceca.2013.12.005
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Emerging evidence suggests a role for endocannabinoids in
theardiovascular system in various pathological conditions, such
asypertension, myocardial infarction and heart failure (for
recenteviews [4,5]). AEA has profound and rather complex actions
onlood pressure and cardiac function. For example, AEA has beenhown
to decrease arterial blood pressure in a triphasic manner bycting
on the contractility of smooth muscle, the release of
neu-otransmitters from peripheral nerve endings, and the
activationf autonomic reflex pathways [6]. Both cannabinoid
receptor-ependent and independent mechanisms have been shown to
playoles in AEA inhibition of smooth muscle contraction dependingn
the type of vascular structure, the presence of intact endothe-ium,
the metabolic products of endocannabinoids and the activityf
metabolizing enzymes [6–8].
Compared with a large body of information available on the
vas-ular effects of endocannabinoids, surprisingly few studies
haveocused on the role of endocannabinoids in the regulation of
con-ractility and Ca2+ signaling in cardiac muscle. Experiments
withEA performed in isolated Langendorff rat hearts and in
isolated,lectrically stimulated atrial appendages from human, rat,
and rab-it [9–12] have revealed a negative inotropic effect of
cannabinoidshat may underlie the ability of AEA to decrease cardiac
output asbserved in studies performed in vivo (for reviews
[4,7,8]). How-ver, mechanisms underlying these cardiac actions of
AEA remainargely unknown.
In the present study, we have hypothesized that the
negativenotropic actions of anandamide are due to altered Ca2+
homeo-tasis and AP characteristics in ventricular myocytes. Thus,
weave investigated the actions of AEA on contractile properties,
Ca2+
ignaling and AP waveforms in acutely dissociated rat
ventricularyocytes.
. Materials and methods
.1. Ventricular myocyte isolation
Ventricular myocytes were isolated from adult male Wistar
rats264 ± 19 g) according to previously described techniques [13].
Thistudy was carried out in accordance with the recommendationsn
the Guide for the Care and Use of Laboratory Animals of theational
Institutes of Health. The protocol was approved by theommittee on
the Ethics of Animal Experiments of the UAE Uni-ersity. Briefly,
the animals were euthanized using a guillotine andearts were
removed rapidly and mounted for retrograde perfu-ion according to
the Langendorff method. Hearts were perfused at
constant flow of 8 ml g heart−1 min−1 and at 36–37 ◦C with a
solu-ion containing (mM): 130 NaCl, 5.4 KCl, 1.4 MgCl2, 0.75 CaCl2,
0.4aH2PO4, 5 HEPES, 10 glucose, 20 taurine, and 10 creatine set to
pH.3 with NaOH. When the heart had stabilized perfusion was
contin-ed for 4 min with Ca2+-free isolation solution containing
0.1 mMGTA, and then for 6 min with cell isolation solution
containing.05 mM Ca2+, 0.75 mg/ml collagenase (type 1; Worthington
Bio-hemical Corp., USA) and 0.075 mg/ml protease (type X1V;
Sigma,ermany). Ventricles were excised from the heart, minced
andently shaken in collagenase-containing isolation solution
supple-ented with 1% BSA. Cells were filtered from this solution at
4-min
ntervals and resuspended in isolation solution containing 0.75
mMa2+.
.2. Measurement of ventricular myocyte shortening
Ventricular myocytes were allowed to settle on the glass bot-om
of a Perspex chamber mounted on the stage of an inverted
icroscope (Axiovert 35, Zeiss, Germany). Myocytes were
super-used (3–5 ml/min) with normal tyrode (NT) containing (mM):
140
m 55 (2014) 104– 118 105
NaCl, 5 KCl, 1 MgCl2, 10 glucose, 5 HEPES, and 1.8 CaCl2 (pH
7.4).Shortening of myocytes was recorded using a video edge
detectionsystem (VED-114, Crystal Biotech, USA). Resting cell
length (RCL)and amplitude of shortening (expressed as a % of
resting cell length)were measured in electrically stimulated (1 Hz)
myocytes main-tained at 35–36 ◦C. Data were acquired and analyzed
with SignalAverager software v 6.37 (Cambridge Electronic Design,
UK). Exper-imental solutions were prepared from stock immediately
prior toeach experiment.
2.3. Measurement of intracellular Ca2+ concentration
Myocytes were loaded with the fluorescent indicator fura-2
AM(F-1221, Molecular Probes, USA) as described previously [13].
Inbrief, 6.25 �l of a 1 mM stock solution of fura-2 AM (dissolved
indimethylsulphoxide) was added to 2.5 ml of cells to give a
finalfura-2 concentration of 2.5 �M. Myocytes were shaken gently
for10 min at 24 ◦C (room temperature). After loading, myocytes
werecentrifuged, washed with NT to remove extracellular fura-2
andthen left for 30 min to ensure complete hydrolysis of the
intracel-lular ester. To measure intracellular Ca2+concentration,
myocyteswere alternately illuminated by 340 and 380 nm light using
amonochromator (Cairn Research, UK) which changed the excita-tion
light every 2 ms. The resulting fluorescence emitted at 510 nmwas
recorded by a photomultiplier tube and the ratio of the emit-ted
fluorescence at the two excitation wavelengths (340/380 ratio)was
calculated to provide an index of intracellular Ca2+
concentra-tion. Resting fura-2 ratio, TPK Ca2+ transient, THALF
decay of the Ca2+
transient, and the amplitude of the Ca2+ transient were
measuredin electrically stimulated (1 Hz) myocytes.
2.4. Measurement of sarcoplasmic reticulum Ca2+ content
Sarcoplasmic reticulum (SR) Ca2+ release was assessed
usingpreviously described techniques [13,14]. After establishing
steadystate Ca2+ transients in electrically stimulated (1 Hz)
myocytesmaintained at 35–36 ◦C and loaded with fura-2, stimulation
waspaused for a period of 5 s. Caffeine (20 mM) was then applied
for10 s using a solution switching device customized for rapid
solutionexchange. Electrical stimulation was resumed and the Ca2+
trans-ients were allowed to recover to steady state. SR-releasable
Ca2+
was assessed by measuring the area under the curve of the
caffeine-evoked Ca2+ transient. Fractional release of SR Ca2+ was
assessedby comparing the amplitude of the electrically evoked
steady stateCa2+ transients with that of the caffeine-evoked Ca2+
transient andrefilling of SR was assessed by measuring the rate of
recovery ofelectrically evoked Ca2+ transients following
application of caffeine.
2.5. Assessment of myofilament sensitivity to Ca2+
In some cells shortening and fura-2 ratio were recorded
simul-taneously. Myofilament sensitivity to Ca2+ was assessed
fromphase-plane diagrams of fura-2 ratio versus cell length by
mea-suring the gradient of the fura-2-cell length trajectory during
laterelaxation of the twitch contraction. The position of the
trajectoryreflects the relative myofilament response to Ca2+ and
hence, canbe used as a measure of myofilament sensitivity to Ca2+
[15].
2.6. Preparation of cardiac sarcoplasmic reticulum vesicles
SR vesicles were obtained from rat ventricles by minor
mod-ifications of methods described earlier [16]. Adult male
Wistar
rats were anesthetized with intraperitoneal injection of
sodiumpentobarbital (100 mg/kg) and hearts were quickly removed
andrinsed in Ca2+-free Tyrode solution. The ventricles were blotted
onfilter paper to remove excess solution and homogenized in
cold
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aline solution of the following composition (mM): 154 NaCl; and0
Trizma-maleate; pH 6.8. The homogenate was centrifuged at000 × g
for 30 min and the pellet was discarded. The supernatantas
centrifuged at 10,000 rpm (8000 × g) for 30 min, and the
result-
ng supernatant was recentrifuged at 47,000 rpm (100,000 × g) for
h. The final pellet was resuspended in storage solution (mM):
154aCl; 10 Trizma-maleate; and 300 sucrose, pH: 6.8 then stored
at80 ◦C until used. The whole procedure was carried out in a
cold
oom, at 4 ◦C and in the presence of protease inhibitors (mM:
500enzamidine; 1 leupeptine; 1 pepstatin A and 200
phenylmethyl-ulphonyl fluoride). Membrane preparations for cardiac
SR vesiclesere generous gift from Dr. Susan Dunn (University of
Alberta,
dmonton, Canada).
.7. Measurements of Ca2+ uptake rate
SR Ca2+ uptake was measured with a spectrophotometer (Cory00,
Varian; Walnut Creek, CA, USA) using the Ca2+-sensitiveye
antipyrylazo III (APIII; Sigma–Aldrich, St. Louis, MO, USA),
asescribed previously [16]. SR membrane vesicles (100 �g) weredded
to 1 ml phosphate buffer medium containing (in mM): 100H2PO4, 3
MgCl2, 2 ATP, 0.02 ruthenium red, and 0.1 APIII, pH 7.1
37 ◦C). Ca2+ uptake was initiated by addition of 10 �M CaCl2 to
theedium and measured as changes in absorbance of APIII between
10 and 790 nm. Ruthenium red (10 �M) was used to block Ca2+
elease from the SR.
.8. Measurements of Ca2+-ATPase activity
ATPase activity was measured by minor modifications of cou-led
enzyme reaction method described earlier [17,18]. Briefly,he
reaction medium used for the assay of Ca2+-ATPase containedmM) 20
mM MOPS, pH 7.1, 80 mM KCI, 5 mM MgCl2, 0.1 mMaCl2, 0.1 mM EGTA,
100 �g of SR membrane protein/ml, and 2 �M23187 (Ca2+ ionophore)
was incubated for 5 min at 37 ◦C. A23187as added to the reaction to
render the vesicles permeable to Ca2+.t this point 2.5 mM [�-32P]
ATP was added, and the amount ofydrolysis was measured by a
colorimetric method [17]. Followinghe addition of ATP (3 mM) serial
samples were taken for determi-ation of Pi as described earlier
[18].
.9. Measurements of [3H]-ryanodine binding
[3H]-ryanodine binding was assayed by some modifications
ofreviously described methods [19]. Briefly, ventricles were
homog-nized in five volumes of 300 mM sucrose and 10 mM imidazole
(pH.1 at 4 ◦C) and aliquots of homogenate (about 100 �g of
protein)ere incubated at 37 ◦C in a buffer containing 25 mM
imidazole
pH 7.4 at 37 ◦C), 1 M KCl, 0.2–20 nM [3H]-ryanodine (7.4
Ci/mmol),. 9 mM EGTA, and 1 mM CaCl2. After 60 min, 0.4 ml aliquots
ofach sample were filtered under vacuum through Watman cellu-ose
nitrate filters with pores of 0.45 �M (presoaked in washinguffer
containing 25 mM imidazole and 1 M KCl), and washed twiceith 5 ml
of ice-cold washing buffer. The filters were dried and
xtracted in 5 ml of HydroflourTM (National Diagnostics, FL,
USA)cintillation fluid before counting for 3H. Triplicate 50-�l
samples ofhe incubation mixtures were also counted directly for
estimationsf total binding. Nonspecific binding was estimated from
paralleleasurements of binding in the presence of 10 �M unlabeled
ryan-
dine. The radioactivity was measured in Beckman LS 6000IC
liquidcintillation counter.
.10. Measurement of Ca2+ release
SR vesicles (50 �g/ml) were passively loaded by incubation in �l
of 45CaCl2 (10 mCi/ml, 42.7 mCi/mg; NEN Life Science Products,
m 55 (2014) 104– 118
Inc.) in 1 ml of (mM): 100 KCl; 20 PIPES dipotassium salt; 2
CaCl2;at pH 7.1 at room temperature for 3 h. 45Ca2+ efflux was
initiatedby 50-fold dilution of 20 �l of 45Ca2+-loaded SR vesicles
intorelease buffer (mM): 100, KCl; 20 PIPES dipotassium salt; 1
EGTA;1 CaCl2; at 37 ◦C followed, after 10 s, by filtration through
HA filters(0.45 �m, Millipore Corp., Bedford, MA). Filters were
rinsed with5 ml ice-cold wash buffer (mM): 100 KCl; 20 PIPES
dipotassiumsalt; 1 EGTA; 1 CaCl2; 10 MgCl2; and 20 �M ruthenium
red. Filterswere allowed to air-dry overnight, and radioactivity
remainingwithin the membranes was counted in Beckman LS 6000IC
liquidscintillation counter. Stock solutions of A23187 (Calbiochem
Corp.,San Diego, CA, USA) were made in 100% DMSO; final
concentrationof DMSO was 0.12% (v/v). In control experiments, stock
solutionsof DMSO up to 0.3% (v/v) and ethanol up to 0.1% (highest
concen-tration used 0.07%) did not affect control conditions in
Ca2+ uptake,ryanodine binding, and 45Ca2+ release from SR
vesicles.
2.11. Measurement of action potentials
Action potentials (APs) were measured using whole-cell
patchclamp technique. In our recordings, only rod shaped
quiescentmyocytes with clear cross-striations were used. Recordings
weremade with an Axopatch 200B amplifier (Molecular Devices,
Down-ington, PA, USA) coupled to an A/D interface (Digidata
1322;Molecular Devices, Downington, PA, USA). Patch pipettes
werefabricated from filamented GC150TF borosilicate glass
(HarvardApparatus, Holliston, MA, USA) on a horizontal puller
(SutterInstruments Co., Novato, CA, USA). Electrode resistances
rangedfrom 2.0 to 3.0 M�, and seal resistances were 1–5 G�. After
gigaseal formation, the membrane was ruptured with gentle suc-tion
to obtain whole cell current-clamp configuration. APs wereelicited
in current-clamp mode by 4-ms, 0.9–1 nA injections ofsquare current
pulses at a frequency of 0.2 Hz. Basic extracellu-lar solutions
used for electrophysiological recordings contained(in mM): 144
NaCl, 5.4 KCl, 1.8 CaCl2, 1.2 MgCl2, 1 NaH2PO4, 10HEPES, 10
glucose, and pH 7.4 (adjusted with NaOH). Recordingpatch pipettes
were filled with intracellular solution containing(in mM): 150 KCl,
10 NaCl, 120 aspartate, 5 MgCl2, 0.1 CaCl2,1.1 EGTA, 10 HEPES, 4
Mg-ATP, 5 sucrose, and pH 7.2 (adjustedwith HCl). Experiments were
performed at room temperature(22–24 ◦C). Changes of external
solutions and application of drugswere performed using a multi-line
perfusion system driven bya micro-pump with a common outflow
connected to the cellchamber. In some experiments, applications of
drugs were per-formed using multi-barrel puffing micropipette with
a commonoutflow positioned in close proximity to the cell under
investiga-tion. In these experiments, the cell was continuously
superfusedwith the solution via a puffing pipette to reduce
possible arti-facts related to the switch from a static to a moving
solution andvice versa. Complete external solution exchange was
achieved in
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as routinely included in extracellular solution. In agreementith
earlier studies in rat cardiomyocytes [21,23], AP waveforms
emained unchanged in the presence of ethanol concentrationsp to
0.07% (v/v). AM251, AM630, methanandamide, and URB597ere purchased
from Tocris (Ellisville, MO, USA). Other reagents
nd chemicals used in our experiments were purchased
fromigma–Aldrich (St. Louis, MO, USA). Drugs were dissolved in
DMSOr ethanol. Stocks were kept at −20 ◦C until their use and final
con-entrations for ethanol and DMSO used in the experiments did
notxceed 0.07% (v/v). At low concentrations, DMSO had no
signifi-ant effect on the control conditions during shortening or
actionotential recordings.
Electrophysiological data were analyzed using pClamp
8.0Molecular Devices, Downington, PA, USA), Origin 8.0 (Origin
Laborp., Northampton, MA, USA) and Mat Lab R2011a (MathWorksorp.,
Natick, MA, USA) software. APD was measured at 60% of repo-
arization from AP amplitude. Cardiac myocytes were classified
toypes based on the shape of the AP in current-clamp mode.
.12. Data analysis
Each experiment was performed on several myocytes from dif-erent
batches. Since the results of identical experiments fromifferent
groups were qualitatively similar, the data were pooledor
statistical purposes with “n” denoting the total number of
cellsested for a particular data point. The results of the
experimentsere expressed as mean ± standard error of the mean
(S.E.M.). Sta-
istical analysis was performed using the paired t-test (within
theame cell analysis) and ANOVA tests (for analysis of data from
dif-erent cells). On all graphs (*) denote statistical significance
with
< 0.05, between specified values, or if not specified to the
respec-ive control.
. Results
.1. Effects of anandamide on ventricular myocyte shortening
In normal tyrode (NT) solutions, amplitudes of myocyte
short-ning in response to electrical stimulation (stimulated at 1
Hz,0 pulses delivered every 1 min) gradually decreased to 85–80%f
controls during experiments lasting up to 20 min. No furtherun down
of the shortening amplitudes was observed in NT solu-ion containing
0.007% ethanol (used as vehicle in 1 �M AEAolution; data not shown;
n = 7–10; compared to 0 time point;NOVA, F(1,15) = 0.018; P >
0.05) and, unless it was stated other-ise, ethanol (in the
concentration that was used in AEA containing
olution) was also included routinely in control (NT) solutions
dur-ng shortening and Ca2+ transient experiments.
In the first set of experiments, we tested the effect of AEAn
the contractility of acutely isolated rat ventricular myocytes.ig.
1A shows typical records of shortening in a myocyte super-used with
either NT (in the absence of AEA, NT contained 0.007%thanol in all
experiments) or NT + 1 �M AEA and during washoutith NT. The
amplitude of shortening was significantly (paired
-test; n = 12–14; P < 0.05) reduced up to 47.3 ± 2.6% of
controlsFig. 1B) when the concentration of AEA was increased in the
rangef 1 nM to 10 �M compared to NT (for 0.01 �M AEA; paired t-est,
n = 9; t(8) = 6.09; P < 0.05). Synthetic cannabinoid
WIN55,212-2ested at 1 �M concentration did not cause a significant
alter-tion on the amplitudes of myocyte shortening (paired t-test,
n = 7;
> 0.05).
The negative inotropic effect (decrease of shortening ampli-
udes) by AEA could be due to degradation products of AEA such
asrachidonic acid. For this reason, we have investigated the
effectsf methanandamide (metAEA), a non-hydrolyzable analog of
AEA
m 55 (2014) 104– 118 107
[24] and URB597, an inhibitor of fatty acid amide hydrolase
(FAAH,enzyme that metabolize AEA) [25]. In cardiomyocytes
pretreatedfor 10 min with 0.1–10 �M metAEA (Fig. 1C), the extent of
inhi-bition was not significantly different from that of AEA (n =
7–8,ANOVA, F(1,13) = 5.9, P > 0.05). Similarly, pretreatment
with 1 �MURB597 for 45 min at 37 ◦C did not alter the extent of AEA
inhibi-tion (Fig. 1D). In the presence and absence of URB597
treatment,AEA (1 �M) inhibited myocyte shortening to 68.2 ± 4.8% of
controlsand 62.7 ± 8.4% of controls, respectively. There were no
statisticallysignificant differences in the inhibitory effect of
AEA between con-trol (NT + 0.007% ethanol after 45 min
pretreatment) and URB597pretreated cells (n = 9–11, ANOVA, F(1,18)
= 7.8, P > 0.05). We havealso tested whether cyclooxygenase
products of AEA metaboliteswould mediate the observed actions of
this compound. The resultsindicated that the extent of shortening
amplitudes of cardiomy-ocytes was not significantly different after
incubating the cellswith 30 �M indomethacin, a cyclooxygenase
inhibitor, for 30 min(n = 8–11, ANOVA, F(1,17) = 6.1, P > 0.05).
Among other contractionparameters measured, resting cell length
(RCL) was not signifi-cantly altered (paired t-test; n = 12–14; P
> 0.05, data not shown)by 10 min superfusion with AEA (1 nM–1
�M). However, increasingthe concentration of AEA to 10 �M caused a
small but statisticallysignificant reduction of RCL in about 60% of
cells tested (pairedt-test; n = 12–14; P < 0.05, data not
shown).
In the next series of experiments we have tested whether
theeffect of AEA is mediated by the activation of cannabinoid
(CB)receptors. For this purpose, we have first tested the effects
of estab-lished antagonists of CB1 and CB2 receptors on the AEA
inhibitionof shortening amplitudes in cardiomyocytes. Two
structurally dif-ferent CB1 receptor antagonists (AM251 with a Ki
of 7.5 nM [26];Fig. 2A and SR141716 with a Ki of 1.8 nM [27]; Fig.
2B) and twoCB2 receptor antagonists (AM630 with a Ki of 32.1 nM
[26] andSR144528 with Ki of 0.6 nM [27]) were tested. At 300 nM
concentra-tion, these antagonists were not able to reverse the
inhibitory effectof AEA on the shortenings amplitudes of
cardiomyocytes (n = 8–12,ANOVA, P > 0.05).
Since the activation of both CB1 and CB2 receptors are medi-ated
by Gi/o subtypes of G-proteins [26], we have also tested theeffect
of inhibitors of Gi/o proteins such as pertussis toxin (PTX)
andN-ethylmaleimide (NEM) on AEA induced inhibition of
cardiomy-ocyte shortening (Fig. 2E and F). Preincubation of
cardiomyocytesin either PTX (2 �g/ml, 3 h at 37 ◦C) or NEM (50 �M
for 30 min at37 ◦C; [28]) did not alter the extent of AEA effect on
cardiomy-ocyte shortening (n = 9–12, ANOVA, P > 0.05). However,
in positivecontrol experiments, preincubation of cardiomyocytes in
PTX orNEM effectively blocked the inhibitory effect of clenbuterol,
a �2adrenoreceptor agonist on cardiomyocyte shortening
(supplementFig. 1).
3.2. Effects of anandamide on intracellular Ca2+ levels
In the second set of experiments we have investigated the
effectsof 10 min bath application of 1 �M AEA on the resting
intracellularCa2+ levels and on the amplitudes and kinetics of Ca2+
transientselicited by electrical-field stimulation. Typical records
of Ca2+ trans-ients in a myocyte superfused with either NT or NT +
1 �M AEAand during washout with NT are shown in Fig. 3A. The
effects of1 �M AEA on resting fura-2 ratio, TPK Ca2+ transient,
THALF decayof the Ca2+ transient, and AMP of Ca2+ transient are
shown inFig. 3B–E, respectively. Although, AEA has been shown to
alter intra-cellular Ca2+ levels in various types of cells (for
reviews [29,30]),application of 1 �M AEA for 10 min did not cause a
significant alter-
ation in resting fura-2 ratio and TPK Ca2+ transient (paired
t-test;n = 21–24 cells, P > 0.05). However, THALF decay of the
Ca2+ tran-sient and AMP of the Ca2+ transient were significantly
reduced by1 �M AEA to 118.9 ± 5.5 ms and 0.243 ± 0.032 fura-2 ratio
units
-
108 L.T. Al Kury et al. / Cell Calcium 55 (2014) 104– 118
Fig. 1. Effects of AEA, metAEA and preincubation with URB597 or
indomethacin on AEA induced inhibition of ventricular myocyte
shortening. (A) Typical records of shorteningin an electrically
stimulated (1 Hz) ventricular myocyte superfused with either NT
(containing the vehicle, 0.007% ethanol) or NT + 1 �M AEA and
during washout with NT. (B)Bar graph showing the mean amplitudes
(AMP) of shortening expressed as a percentage of control values in
vehicle (NT + 0.007% ethanol), and in presence of AEA (1 nM–10
�M).Myocytes were maintained at 35–36 ◦C and superfused with AEA
for 10 min. Data are shown as means ± S.E.M., n = 12–14 cells.
*Indicates statistically significant differenceat the level of P
< 0.05. (C) Bar graph showing the mean amplitudes (AMP) of
shortening expressed as a percentage of control values in vehicle
(NT + 0.007% ethanol), and inpresence of metAEA (1 nM–10 �M). Data
are shown as means ± S.E.M., n = 7–8 cells. *Indicates
statistically significant difference at the level of P < 0.05.
(D) Bar graph showingthe effect of AEA on the mean amplitudes (AMP)
of shortening expressed as percentage of control values in NT
containing 0.007% ethanol or after 45 min incubation with1 �M
URB597 at 37 ◦C. The S.E.M. values are indicated on top of each
column. Data are mean ± S.E.M., n = 7–11 cells. (E) Bar graph
showing the effect of AEA on the meana ontainT –10 ce
cPc
3t
mimlu
mplitudes (AMP) of shortening expressed as percentage of control
values in NT che S.E.M. values are indicated on top of each column.
Data are mean ± S.E.M., n = 9
ompared to 129.3 ± 5.2 ms (paired t-test, n = 24 cells, t(23) =
3.73; < 0.05) and 0.326 ± 0.024 fura-2 ratio units (paired
t-test, n =24ells, t(23) = 5.79; P < 0.05) in controls,
respectively.
.3. Effect of anandamide on sarcoplasmic reticulum Ca2+
ransport
The effect of 1 �M AEA on Ca2+ transport was investigated
inyocytes exposed to 20 mM caffeine. Fig. 4A shows a typical
record
llustrating the protocol used in these experiments. Initially,
theyocyte was electrically stimulated (ES) at 1 Hz. Electrical
stimu-
ation was then turned off for 5 s. Caffeine was then applied for
10 ssing a rapid solution-exchanger device. Electrical stimulation
was
ing 0.007% ethanol or after 30 min incubation with 30 �M
indomethacin at 37 ◦C.lls.
then restarted, and the recovery of intracellular Ca2+ was
recordedduring a period of 60 s. SR Ca2+ content was assessed by
measuringcaffeine-evoked Ca2+ release (area under the
caffeine-evoked Ca2+
transient) and fractional release of Ca2+ by comparing the
ampli-tude of the electrically evoked steady-state Ca2+ transients
withthat of the caffeine-evoked Ca2+ transient in the presence of
eitherNT alone or NT with 1 �M AEA. Fractional release of SR Ca2+
was notsignificantly altered in 1 �M AEA compared to NT (0.749 ±
0.024in AEA versus 0.753 ± 0.028 in controls; paired t-test; n = 23
cells,
t(22) = 0.166; P > 0.05; Fig. 4B). The area of
caffeine-evoked Ca2+
transient (Fig. 4C) and recovery of the Ca2+ transient during
elec-trical stimulation following application of caffeine (Fig. 4D)
werealso not significantly altered in myocytes exposed to 1 �M
AEA
-
L.T. Al Kury et al. / Cell Calcium 55 (2014) 104– 118 109
0
20
40
60
80
100
N-ethylmaleimide trea ted
Pertuss is tox in trea ted
vehicletrea ted
SR14452 8
AEA - - + + AM630 - + + -
AEA AEA
AM63 0 DC
B
% A
MP
shor
teni
ng( n
orm
aliz
ed to
NT
)
A
***
AM25 1
0
20
40
60
80
100
*
% A
MP
shor
teni
ng( n
orm
aliz
ed to
NT
)
SR14171 6
0
20
40
60
80
100
% A
MP
sho
rten
ing
( nor
mal
ized
to N
T)
**
0
20
40
60
80
100
AEA - - + + SR144528 - + + -
% A
MP
shor
teni
ng( n
orm
aliz
ed to
NT
)
AEA - - + + AM251 - + + -
**
0
20
40
60
80
100 P>0.05
% A
MP
sho
rten
ing
(nor
mal
ized
to N
T)
0
20
40
60
80
100
vehicletrea ted
AEA - - + + SR141716 - + + -
FE
% A
MP
sho
rten
ing
( nor
mal
ized
to N
T)
P>0.05
Fig. 2. Effects of cannabinoid receptor antagonists, pertussis
toxin and N-ethylmaleimide on AEA-induced inhibition of
cardiomyocyte shortening. Bar graphs showing themean amplitudes
(AMP) of shortening expressed as a percentage of control values in
vehicle (NT + 0.007% ethanol), and (A) in presence of CB1 receptor
antagonist AM251(0.3 �M), (B) CB1 receptor antagonist SR141716 (0.3
�M), (C) CB2 receptor antagonist AM630 (0.3 �M), and (D) CB2
receptor antagonist SR144528 (0.3 �M). Data are shownas means ±
S.E.M., n = 9–11 cells. *Indicates statistically significant
difference at the level of P < 0.05. (E) Bar graph showing the
effect of AEA on the mean amplitudes (AMP) ofshortening expressed
as a percentage of control values in vehicle (NT + distilled
water), and pertussis toxin treated cells. (F) Bar graph showing
the effect of AEA on the meanamplitudes (AMP) of shortening
expressed as a percentage of control values in vehicle and
N-ethylmaleimide treated cells. Data are shown as means ± S.E.M., n
= 9–11 cells.*
mP
3
att[perbgo(s
Indicates statistically significant difference at the level of P
< 0.05.
yocytes compared to control cells (paired t-test; n = 21–23
cells, > 0.05).
.4. Effect of anandamide on myofilament sensitivity to Ca2+
The effects of AEA on myofilament sensitivity to Ca2+ werelso
investigated. These experiments tested whether AEA decreaseshe
mechanical responses by altering the affinity of the contrac-ile
machinery of the ventricular myocytes to intracellular Ca2+
15]. A typical record of myocyte shortening and fura-2 ratio
andhase-plane diagrams of fura-2 ratio versus cell length in
myocytesxposed to NT are shown in Fig. 5A. The gradient of the
trajectoryeflects the relative myofilament response to Ca2+ and
hence, haseen used as a measure of myofilament sensitivity to Ca2+
[15]. The
radients of the fura-2-cell length trajectory during late
relaxationf the twitch contraction measured during the periods
500–600 msFig. 5B), 500–700 ms (Fig. 5C) and 500–800 ms (Fig. 5D)
were notignificantly altered in AEA compared to NT (containing
0.007%
ethanol) suggesting that myofilament sensitivity to Ca2+ is
notreduced by AEA (AEA-treatment was compared to NT
containing0.007% ethanol, paired t-test; n = 23–30 cells; P >
0.05).
3.5. Effect of anandamide on Ca2+-uptake and Ca2+-ATPaseactivity
in cardiac sarcoplasmic reticulum vesicles
Decreases in THALF decay and AMP of the Ca2+ transient byAEA
during twitch responses, can be due to previously reportedeffects
of endocannabinoids on Ca2+ release and uptake processesin myocytes
and neurons ([31,32]; for reviews [29,30]). For thispurpose, we
first measured Ca2+ uptake by SR vesicles isolatedfrom rat
ventricles. The net Ca2+ uptake is the sum of the SR Ca2+
influx (mainly by the activity of Ca2+-ATPase) and Ca2+ leak
from
SR (through ryanodine-sensitive channels). Our experiments
werecarried out in the presence of 10 �M ruthenium red, to
inhibitcompletely the activity of cardiac Ca2+ release channel.
Underthese conditions, the net Ca2+ uptake closely correlates with
the
-
110 L.T. Al Kury et al. / Cell Calcium 55 (2014) 104– 118
Fig. 3. Effects of AEA on amplitude and time-course of
intracellular Ca2+ in ventricular myocytes. (A) Typical records of
Ca2+ transients in an electrically stimulated (1 Hz)ventricular
myocyte superfused with either NT or NT + 1 �M AEA and during
washout with NT; scale bar indicates 0.2 fura-2 ratio unit (RU).
Also shown resting fura-2r ecay ow s meaP
Scusdu1(wcF(smawp1e
atio (340/380 nm; B), time to peak (TPK) Ca2+ transient (C),
time to half (THALF) dere maintained at 35–36 ◦C and superfused
with AEA for 10 min. Data are shown a
< 0.05.
R Ca2+-ATPase activity. Fig. 6A illustrates that AEA, in the
con-entration range 0.1–1 �M, causes a significant increase of
Ca2+
ptake rate, while elevating the concentrations of AEA to 10
�Mlowed down the uptake rate significantly (Fig. 6A).
Experimentalata were fitted by a single exponential function from
the rate ofptake and time constants � were presented in control
(treated0 min with vehicle, 0.07% ethanol) and in the presence of
AEAFig. 6A and B). In the presence of 0.1 and 1 �M AEA, � valuesere
significantly decreased (AEA-treatment was compared to NT
ontaining 0.007% ethanol; n = 8–9 samples from 3
experiments,(1,13) = 14.9; ANOVA, P < 0.05), whereas, 10 �M AEA
increasedn = 8–9, F(1,13) = 17.2; ANOVA, P < 0.05) � values
(Fig. 6B). Sub-equently, we have measured the activity of
Ca2+-ATPase in SRembranes (Fig. 6C and D). In the presence of 0.1
and 1 �M AEA,
ctivity of Ca2+-ATPase was significantly increased
(AEA-treatment
as compared to NT containing 0.007% ethanol; n = 8–9 sam-les
from 3 experiments, F(1,20) = 24.1; ANOVA, P < 0.05), whereas,0
�M AEA decreased (n = 8–9, F(1,20) = 7.3; ANOVA, P < 0.05)
thenzyme activity (Fig. 6C and D).
f the Ca2+ transient (D) and amplitude (AMP) of the Ca2+
transient (E). Myocytesns ± SEM, n = 21–24 cell. *Indicates
statistically significant difference at the level of
3.6. Effect of anandamide on [3H]-ryanodine binding
andCa2+-release in cardiac sarcoplasmic reticulum vesicles
Ryanodine binding is widely used method to investigate
theinteraction between drugs and SR release channels. In Fig.
7A,saturation binding curves for [3H]-ryanodine were presentedin
the presence and absence of the AEA. As the
[3H]-ryanodineconcentration was increased from 0 to 20 nM,
ryanodine bindingto SR vesicles was rapidly increased and saturated
at 10–20 nM.At a concentration of 10 �M, AEA caused a significant
inhibition ofthe specific binding of [3H]-ryanodine. In controls
and in presenceof 0.1, 1 and 10 �M AEA, maximum binding activities
(Bmax)were 352 ± 37, 365 ± 42, 339 ± 29 and 289 ± 24 pmol/mg
protein,respectively. In controls and in presence of 0.1, 1 and 10
�MAEA, KD values were 2.2 ± 0.6, 1.8 ± 0.7, 2.5 ± 0.7 and 2.3 ± 0.4
nM,
respectively. There was no statistically significant
differencebetween control and AEA treated groups with respect to
Bmax andKD values (ANOVA, n = 9–12, P > 0.05). Effect of AEA on
saturationbinding was further analyzed by Scatchard analysis (Fig.
7B), which
-
L.T. Al Kury et al. / Cell Calcium 55 (2014) 104– 118 111
F he effa of Ca2
e ± S.E.
y21o
Fat
ig. 4. Effect of AEA on sarcoplasmic reticulum Ca2+. (A) Typical
record illustrating t rat ventricular myocyte. Also shown are mean
amplitudes of SR fractional releasevoked intracellular Ca2+ after
application of caffeine (D). Data are shown as means
ielded Bmax values of 289 ± 31, 293 ± 28, 289 ± 33, 263 ± 36
and01 ± 24 pmol/mg protein in controls and in presence of 0.1, 1
and0 �M AEA, respectively. KD values for controls and in presencef
0.1, 1 and 10 �M AEA were 0.9 ± 0.2, 0.9 ± 0.1, 1.0 ± 0.2 and
ig. 5. Effect of AEA on myofilament sensitivity to Ca2+. (A)
Typical record of myocyte short myocytes exposed to NT. The arrow
indicates the region where the gradient was measurrajectory during
late relaxation of the twitch contraction during the periods
500–600 (B),
ects of electrical stimulation (ES) and rapid application of
caffeine on fura-2 ratio in+. (B) Area under the caffeine-evoked
Ca2+ transient (C) and recovery of electricallyM., n = 21–23
cells.
0.6 ± 0.1 nM, respectively. Although Bmax values tend to
decreaseslightly with increasing AEA concentrations, there were no
sta-tistically significant difference between controls and 0.1 �M
and1 �M AEA treated groups (ANOVA, n = 11–14, P < 0.05).
However,
ening and fura-2 ratio and phase-plane diagrams of fura-2 ratio
versus cell length ined. B-D shows the effect of 1 �M AEA on the
mean gradient of the fura-2-cell length500–700 (C) and 500–800 ms
(D). Data are shown as means ± S.E.M., n = 25–30 cells.
-
112 L.T. Al Kury et al. / Cell Calcium 55 (2014) 104– 118
0
2
4
6
8
10
AEA 10 μM
Ca2
+ (
μM)
AEA 0.1 μM
0
50
100
150
200
*
Ca2
+-A
TPa
se a
ctiv
ity(%
of
cont
rol)
*
0 2 4 6 8 100
5
10
15
20
25 Con trol AEA 0.1 μM AEA 1 μM AEA 10 μM
AEA0.1 μM
100 ms AEA0.1 μM
Time (min)
AEA10 μM
Con trol
DC
AEA10 μM
AEA1 μM
Con trol
AT
P hy
drol
yzed
(μM
Pi/m
g pr
otei
n)
0
50
100
150
200
250
*
*
AEA1 μM
BA
τ o
f C
a2+ u
ptak
e (s
)
Fig. 6. Effects of AEA on Ca2+ uptake and Ca2+-ATPase activity
in cardiac SR vesicles. (A) Effect of AEA on the uptake of Ca2+
from SR vesicles. Ca2+ uptake by SR membraneswas measured in
control conditions and in the presence of 0.1, 1 and 10 �M AEA.
Uptake of Ca2+ was initiated by addition of CaCl2 (10 �M). (B)
Effect of AEA on the rate of Ca2+
uptake by SR vesicles. Experimental data were fitted by a single
exponential function, from which the time constant (�) of Ca2+
uptake was calculated. Average time constantsof SR Ca2+ uptake in
control conditions, and in the presence of 0.1, 1 and 10 �M AEA
were presented in the bar graph. Data are the mean ± S.E.M., of 8–9
measurements.* ct of A 2+
C fit ofm value
tA
ov4
dvC(
3m
ate(cmgpnawftp
Indicates statistical difference from control values at the
level of P < 0.05. (C) Effea2+-ATPase activity. Slope of each
hydrolysis curve was calculated from the lineareans ± S.E.M. of 8–9
measurements. *Indicates statistically different from control
he difference in Bmax and KD values between controls and 10 �MEA
was statistically significant (ANOVA, n = 11–14, P > 0.05).
In the next set of experiments, we have tested the effect of
AEAn the passive release of Ca2+ from SR. For this purpose, cardiac
SResicles were loaded with 45Ca2+ and effect of AEA was tested on
the5Ca2+ content (Fig. 7C). AEA, in the concentration range 0.1–10
�M,id not cause any alteration in the 45Ca2+ content of cardiac
SResicles loaded with 45Ca2+. However, incubation in 2 �M
A23187,a2+ ionophore, effectively reduced 45Ca2+ content of SR
vesiclesANOVA, n = 8–9, F(1,15) = 25.6; P < 0.05).
.7. Effects of anandamide on the action potentials of
ventricularyocytes
Generation of the cardiac AP requires a specific
temporalctivation pattern of several ion channels. AEA has been
showno influence the functional properties of several of these
channelsither directly or indirectly thereby affecting cardiac
excitabilityfor a review [29]). In this set of experiments,
patch-clampedardiomyocytes were exposed to the AEA while
continuouslyonitoring their Vrest and APs in the current clamp
mode. The
eneration of APs was evoked by 0.9–1 nA depolarizing
currentulses of 4 ms duration. Since the intracellular pipette
solution didot contain Ca2+-chelating agents, the generation of
each AP wasccompanied by myocyte contraction. Therefore, current
pulses
ere applied at a frequency of 0.2 Hz. During a typical
experiment
ollowing protocols were employed: first, whole-cell
configura-ion was established and 4–5-min dialysis of the myocytes
withipette solution was allowed to ensure the equilibrium
conditions
EA on the activity of Ca -ATPase in SR vesicles. (D) Effect of
AEA on the slope of the data, and the slopes were presented as
percent of controls. Data are shown ass at the level of P <
0.05.
between the intracellular pipette solution and intracellular
milieu.Subsequent to achieving stable recordings of baseline
electricalactivity (Vrest and AP parameters), myocytes were exposed
to AEAfor 10–15 min and subsequently it was washed out.
In our initial experiments, effects of AEA were tested on
thepassive membrane properties of cardiomyocytes. The passive
prop-erties of the ventricular cells from controls were not
significantlydifferent from those of the AEA treated cells. Resting
membranepotentials (mean ± SEM) were −76.3 ± 1.7 and −74.2 ± 1.6 mV
incontrol (n = 11) and AEA treated (n = 14) myocytes,
respectively.The mean cell capacitance in the control group was
109.6 ± 12.8 pF,whereas in the AEA treated cells was 104.7 ± 11.6
pF. Theinput resistance was 82.3 ± 13.4 M� in the control cells
and85.6 ± 11.8 M� in AEA-treated cells. In control cells, these
passivemembrane properties were not altered significantly in
experi-ments lasting up to 25–30 min. In 9 control cells measured,
restingmembrane potentials, cell capacitance, and input resistance
after25 min of experiment were −73.9 ± 3.8 mV, 114.3 ± 12.7 pF,
and83.4 ± 11.7 M�, respectively. These values were not
significantlydifferent than control values obtained within the
first 5 min ofpatch-clamp experiment (n = 9; paired t-test, P >
0.05).
In agreement with earlier studies in rat ventricular
myocytes(for a review [33]) and according to their duration of APs,
twomain types of waveforms were observed; endocardial (with longAP
durations, Fig. 8A; 44.6 ± 2.9 ms; n = 23) and epicardial (with
short AP durations; Fig. 8B; 14.9 ± 1.6; n = 26) myocytes.
Restingmembrane potentials were −77.6 ± 1.3 mV in endocardial
cells, and−78.5 ± 1.4 mV in epicardial cells. Similar to earlier
findings [33],the amplitudes of APs (122.6 ± 8.9 mV versus 110.7 ±
7.1 mV) and
-
L.T. Al Kury et al. / Cell Calciu
0 5 10 15 200
100
200
300
Con trol AEA 0.1 μM AEA 1 μM AEA 10 μM
[3H]-Ryanodine (nM)
[3H
]-R
yano
dine
bou
nd (
fmol
/mg)
0
10
20
30
40
A2318 7 2 μM
AEA10 μM
AEA1 μM
AEA0.1 μM
[3H]- Ryanod ine boun d (fmol/ mg)
C
Ca2
+ c
onte
nt in
SR
ves
icle
s(n
mo l
/mg
prot
ein)
Con tro l
*
0 10 0 20 0 30 0
100
200
300
B
AB
ound
/Fre
e(f
mol
/mg/
nM)
Fig. 7. Effects of AEA on [3H]-ryanodine binding and Ca2+
release from SR vesicles.(A) Effect of AEA (0.1–10 �M) on the
saturation binding of [3H]-ryanodine to SRvesicle membranes. (B)
Scatchard plots derived from the saturation binding exper-iments.
Ratio of specific [3H]-ryanodine binding to the free [3H]-ryanodine
wasplotted against the [3H]-ryanodine concentration used in
saturation experiments.Data were from 3 separate experiments (3–4
trials in each experiment). Each datapoint represents the mean of
9–14 trials. (C) Effect of AEA on Ca2+ release from SRmembranes.
The loaded vesicles were preincubated for 30 min in AEA (0.1–10
�M)o8
dd(
erocRept
tudes without causing significant alteration to the time
course
r A23187 (2 �M) prior to the release assay. Data are shown as
means ± S.E.M., from to 9 measurements.
V/dtmax values (177.3 ± 8.4 V/s versus 141.6 ± 7.2 V/s) in
endocar-ial cells were significantly higher than those of
epicardial cellsANOVA, n = 23–26, P < 0.05).
In the concentration range 0.1–1 �M, AEA consistently short-ned
the duration of AP in both cell types (measured at 60%
ofepolarization, APD60; Fig. 8A and B) with a hyperpolarizing
shiftn Vrest. Changes in AP shortening in response to AEA (1 �M)
appli-ation were noticeable within 10–15 s (insets to Fig. 8A and
B).ecoveries were usually partial and required longer time.
Similar
ffect of AEA was observed in 5 cells recorded at physiological
tem-eratures (35–36 ◦C). APD60 decreased from 33 ± 4 ms in
controlso 21 ± 3 ms in the presence of AEA (ANOVA, n = 5, P <
0.05).
m 55 (2014) 104– 118 113
Effects of AEA on Vrest (Fig. 8C), and APD60 (Fig. 8F) reacheda
statistically significant level at 1 �M AEA (ANOVA, n = 8–11,P <
0.05). However, AP amplitude (Fig. 8D) and maximal rate of
rise(dV/dtmax) of AP (Fig. 7E) were not altered significantly in
both typesof cells at 0.1 �M and 1 �M AEA (ANOVA, n = 10–12, P >
0.05).
At higher concentrations (10 �M), AEA caused a 5–10
mVdepolarization (Fig. 9A–C; ANOVA, n = 18, F(1,16) = 8.3; P <
0.05 inendocardial cells, and ANOVA, n = 24, F(1,22) = 0.6; P <
0.05 in epi-cardial cells) and decreased significantly the AP
amplitudes anddV/dtmax in the majority of both types of myocytes
(Fig. 9D and E;ANOVA, n = 18–24, P < 0.05). AEA (10 �M) also
caused a significantdecrease in APD60 (Fig. 9F). However, in a
subgroup of endocardial(5 out of 18) and epicardial (4 out of 24)
cells, AEA, although causeda significant depolarization and caused
a significant decrease in APamplitude and maximum rate of rise of
AP (supplement Figs. 3 and4), it did not alter APD60 values
significantly (supplement Fig. 4).
In the next series of experiments, we have investigated
theeffects of CB1 receptor antagonist AM251 (0.3 �M) and CB2
recep-tor antagonist AM630 (0.3 �M) on AEA-induced changes in the
APduration of endocardial and epicardial myocytes. The effect of
AEAon APD60 remained unaltered in the presence of 0.3 �M
AM251(ANOVA, n = 5–7, P > 0.05; Fig. 10A). Similarly,
pretreatment withCB2 receptor antagonist AM630 (0.3 �M) did not
cause a significanteffect on AEA-induced changes in AP duration in
both endocar-dial and epicardial myocytes (ANOVA, n = 5–7, P >
0.05; Fig. 10B).Actions of cannabinoid receptors (CB1 and CB2) are
mediated by theactivation of Gi/o proteins sensitive to PTX
treatments [3]. There-fore, we have tested whether PTX pretreatment
(2 �g/ml, 3 h at37 ◦C) alters AEA-induced changes in AP duration in
endocardialand epicardial myocytes. The results of experiments were
pre-sented in Fig. 10C. PTX did not cause a significant alterations
inthe effects of AEA on both endocardial and epicardial
myocytes(ANOVA, n = 5–7, P > 0.05; Fig. 10C).
4. Discussion
The results of this study indicate for the first time that
impairedCa2+ signaling underlies the negative inotropic actions of
AEA inrat ventricular myocytes and that direct interaction of AEA
withion channel(s) shaping APs, rather than the activation of
knowncannabinoid receptors, mediates, at least in part, the effects
of AEAon myocyte contractility.
Administration of AEA causes complex hemodynamic
changesinvolving phases of both increased and decreased blood
pressureas well as changes in heart rate and contractility (for
reviews[4,7]). The results of earlier studies suggested that the
vascularactions of endocannabinoids involve multiple systems and
com-plex set of cellular and molecular mechanisms [6,7]. In
additionto receptor-mediated and direct actions of endocannabinoids
onmuscular structures, neuronal and endothelial cells have also
beenshown to be influenced by AEA and its metabolic products.
Further-more, the presence of off-target sites has been reported to
causeexperimental variations observed in these studies [34].
Use of video edge detection in individual myocytes has sev-eral
advantages over in vivo experiments and traditional in vitrosystems
such as Langendorff-perfused heart preparation, sinceit allows
measurement of contractility at single-cell level in arelatively
isolated environment and excludes the influence ofautonomic nerve
endings, gap-junctions, neurotransmitter uptakesystem, and coronary
perfusion status. In our experiments, AEAcaused a significant
reduction in the maximal shortening ampli-
of myocyte contraction. These findings provide evidence that
thenegative-inotropic effect of AEA results from a direct
interactionof AEA with ventricular myocytes, rather than actions of
AEA on
-
114 L.T. Al Kury et al. / Cell Calcium 55 (2014) 104– 118
Fig. 8. Effect of AEA on the excitability of ventricular
myocytes. Representative recordings show the APs in controls (dark
gray area), in the presence of 1 �M AEA (light grayarea) and after
washout (striped area) in the ventricular endocardial (A) and
epicardial (B) myocytes; the insets on panels A and B show the time
course of the action potentialduration (APD60) and resting
potential (Vrest) changes in response to AEA application (indicated
by horizontal bars). (C)–(F) show summary of AEA effects on
amplitude andshape of the AP in cardiomyocytes; quantification of
the changes in Vrest (C), AP amplitude (D), AP maximal rate of rise
(E) and AP duration (F), characterized byAPD60 incontrols (dark
gray bars) and in response to 0.1 �M (cross hatched bars) or 1 �M
AEA (light gray bars). Data are shown as means ± S.E.M., from 8 to
12 myocytes for eachg
nr
iotaHbCksti
bccmdiwpamd
roup.
erve endings and neurotransmitter uptake systems that have
beeneported in various neuronal structures [35,36].
Negative-inotropic actions of AEA might be attributed to
thempaired release of Ca2+ from the SR. In fact, AEA and other
vari-us cannabinoid receptor agonists have been reported to
modulatehe ryanodine sensitive intracellular Ca2+ stores and
Ca2+-ATPasectivity in various cell types [31,37–39], for recent
reviews [30,40].owever, binding of ryanodine to SR membranes was
not alteredy AEA. Similarly, in the presence of AEA (0.1 and 1 �M),
passivea2+ release remained unchanged. Furthermore, the amplitude
andinetics of caffeine-induced Ca2+ release from intracellular
Ca2+
tores were not changed by AEA. Collectively, these results
indicatehat ryanodine-sensitive intracellular Ca2+ stores are not
involvedn negative-inotropic effects of AEA observed in this
study.
However decreases in THALF decay and AMP of the Ca2+ transienty
AEA during twitch responses can be due to increased uptake
ofytosolic Ca2+ to SR. In fact, AEA, in 0.1 and 1 �M
concentrations,aused a significant increase in Ca2+-ATPase activity
in cardiac SRembranes suggesting that increased Ca2+ uptake
contributes to
ecrease in Ca2+ transients. At higher concentrations, AEA (10
�M)nhibited the activity of this enzyme (Fig. 6B). Interestingly,
NAEs
ith varying carbon chain lengths [37] and fatty acid-based
com-
ounds such as arachidonic acid (metabolic product of AEA) haslso
been shown to modulate the activity of Ca2+-ATPase in a
similaranner in cardiac and skeletal SR membranes [41]. It is
likely that
ecreased resting cell length found at high concentrations (10
�M)
of AEA is due to increased resting Ca2+ levels in some of the
car-diomyocytes.
However, methanandamide (metAEA), a non hydrolyz-able analog of
AEA [24], also showed similar biphasicactions, suggesting that AEA,
but not its metabolic prod-ucts produces observed actions on
Ca2+-ATPase (supplementFig. 2). Potentiation of the Ca2+-ATPase
activity without alteringCa2+ release and ryanodine-binding to the
Ca2+ release channelcan account for the decrease in THALF decay and
amplitude of theCa2+ transient caused by AEA (1 �M). Decreases in
THALF decayand amplitudes of the Ca2+ transient by low
concentration of AEA(1 �M) during twitches, but not
caffeine-induced Ca2+ transients,may suggest that compared to
caffeine-induced responses, fastCa2+ transients during electrical
stimulations are relatively moresensitive to alterations in
Ca2+-ATPase activity.
Activation of cannabinoid receptors alters the levels of sec-ond
messengers such as cAMP, cGMP and protein kinase C [30,42]which are
known to be involved in tuning the Ca2+ sensitivity ofthe
contractile proteins. However, sensitivity of contractile pro-teins
to intracellular Ca2+ remained unchanged in the presenceof AEA
suggesting that phosphorylation and de-phosphorylationof the
contractile proteins do not play a significant role in
negative-inotropic actions of AEA. An earlier study in isolated
ratatria demonstrated that AEA caused negative inotropic effects
bydecreasing cAMP and increasing nitric oxide (NO) levels [11].
How-ever, AEA still decreased contractile performance in the
presence
-
L.T. Al Kury et al. / Cell Calcium 55 (2014) 104– 118 115
Fig. 9. Effects of high AEA concentration (10 �M) on the
excitability of ventricular myocytes. Representative recordings of
APs in controls (dark gray area) and in the presenceof 10 �M AEA
(light gray area) and after washout (striped area) in the
ventricular endocardial (A) and epicardial (B) myocytes; the insets
on panels A and B show the timecourse of the APD60 and resting
potential (Vrest) changes in response to AEA application (indicated
by horizontal bars). (C)–(F) show summary of AEA effects on
amplitudea P ampc as m
oaascrAlICsu
acete(dAutncnit
r
nd shape of the AP in cardiomyocytes; quantification of the
changes in Vrest (C), Aontrols (dark gray bars) and in response to
10 �M (light gray bars). Data are shown
f L-NAME, a NO synthase inhibitor, excluding a NO-mediated
neg-tive inotropic effect on human atrial muscle [10]. Similarly,
innother study in rat isolated heart, the negative inotropic
actions ofynthetic cannabinoid HU-210 were not correlated with the
intra-ellular concentrations of cAMP and cGMP [43]. Collectively,
theseesults, in agreement with our findings, suggest that the
effects ofEA on myocyte contractility are not related to changes in
intracel-
ular Ca2+ release machinery or sensitivity of myofilaments to
Ca2+.n addition, in the presence of AEA, resting levels of
intracellulara2+ and cell length of ventricular myocytes remained
unaltereduggesting that AEA does not significantly affect Ca2+
homeostasisnder resting conditions.
During excitation–contraction coupling, alterations in
themplitudes and kinetics of cardiac AP are closely associated
withorresponding changes in the contractility of myocytes.
Earlierlectrophysiological studies on rat ventricular myocytes
indicatedhat there are two distinctly different groups of cells;
displayingither epicardial (short duration) or endocardial (long
duration) APsfor a review [33]). In our study, low AEA
concentrations (0.1 �M),id not cause alterations in amplitudes and
kinetics of APs (Fig. 8).t low AEA concentrations, there was a
slight decrease in Vrest val-es, which reached a statistically
significant level at 1 �M AEA. Athis concentration, AEA decreased
durations of APs without sig-ificantly affecting the amplitudes and
dV/dtmax of APs. At higheroncentrations (10 �M), AEA-induced
changes in APD accompa-ied with depolarization of the Vrest and
decreases of dV/dtmax
n the endocardial and epicardial ventricular myocytes,
suggestinghat AEA acts on multiple ion channels with different
potencies.
In cardiac muscle, extracellular Ca2+ required to trigger
Ca2+
elease from SR enters through L-type voltage-dependent Ca2+
litude (D) AP maximal rate of rise (E) and AP shape (F),
characterized by APD60 ineans ± S.E.M., from 13 to 20 myocytes.
channels opened during the AP. Our unpublished results
usingvoltage-clamp mode of whole-cell patch clamp technique
indi-cate that, in agreement with the changes in the amplitudesand
dV/dtmax of APs, AEA (1–10 �M) caused significant inhi-bition of
voltage-dependent Na+ and L-type Ca2+ channels incardiomyocytes.
Collectively, these results suggest that
duringexcitation–contraction coupling, shortening of AP due to the
inhi-bition of L-type Ca2+ channels and decreases in Ca2+-induced
Ca2+
release from sarcoplasmic reticulum causes
negative-inotropiceffect of AEA reported in earlier studies. In
line with this hypoth-esis, although caffeine induced contractures
and myofilamentsensitivity to Ca2+ remained unchanged, electrically
induced Ca2+
transients were significantly depressed by AEA; further
suggest-ing that Ca2+-induced Ca2+ release was impaired in the
presence ofAEA.
Although, this is the first patch clamp study investigating
AEAaction on the cardiac APs, an earlier report using
intracellularrecording methods in rat papillary muscle fibers
reported that AEA,in the concentration range of as low as 1–100 nM
potently inhibitedAP durations in an AM251 sensitive-manner,
suggesting that theactivation of CB1 receptors mediates the
negative inotropic actionsof AEA [44]. However, under our
experimental conditions, changeson neither amplitudes nor kinetics
of epicardial and endocardialAPs were detectable until 1 �M
concentration of AEA. Importantly,the effects of AEA on the
duration of both types of APs werenot reversed in the presence of
CB receptor antagonists tested;
AM251 and AM630 (Fig. 10). In addition, AEA continued to
affectAPD after PTX-pretreatment (Fig. 10 and supplement Fig. 4).
It islikely that differences in methods (patch clamp versus
intracellu-lar sharp electrode recording resulting in lower resting
membrane
-
116 L.T. Al Kury et al. / Cell Calciu
Fig. 10. Effects of cannabinoid receptor antagonists and
pertussis toxin treatmentson AEA-induced changes in myocyte
excitability. (A) Effect of CB1 receptor antag-onist AM251 (1 �M)
on AEA-induced changes in AP duration in endocardial andepicardial
myocytes. APD60 was presented in control (dark gray bars) and in
thepresence of 1 �M AEA (light gray bars), 1 �M AM251 (dark gray
cross hatched bars),and 1 �M AEA + 1 �M AM251 (light gray hatched
bars). (B) Effect of CB2 receptorantagonist AM630 (1 �M) on
AEA-induced changes in AP duration in endocardialand epicardial
myocytes. APD60 was presented in control (dark gray bars) and inthe
presence of 1 �M AEA (light gray bars), 1 �M AM630 (dark gray cross
hatchedbars), and 1 �M AEA + 1 �M AM630 (light gray hatched bars).
(C) Effect of pertus-sis toxin (PTX) treatments on AEA-induced
changes in AP duration in endocardialand epicardial myocytes.
Changes are presented in control (dark gray bars), in pres-ence of
1 �M AEA (light gray bars), in control + PTX (dark gray cross
hatched bars),and in presence of 1 �M AEA + PTX (light gray hatched
bars). Data are shown asmeans ± S.E.M., from 5 to 7 myocytes of
each type; *indicate statistically significantdifference at the
level of P < 0.05.
m 55 (2014) 104– 118
potentials) and preparations (ventricular myocytes versus
intactmuscle fibers with nerve endings and gap junction
connections)used in these studies may account for some of the
discrepancies.In earlier studies AEA has been reported to inhibit
noradrenalinerelease from atrium subjected to electrical field
stimulation [30]and enhance vagal activity (for a review [6]) in in
vivo and in in vitromuscle preparations. However, this uptake
mechanism is not likelyto be involved in our studies on isolated
ventricular myocytes.
Involvement of cannabinoid-receptors in the negative
inotropicactions of cannabinoids has been reported in several
earlier studies[9–12]. However, the results of these investigations
have not beenconclusive (for reviews [6–8]). Both cannabinoid
receptor depend-ent and independent mechanisms have been suggested
[6]. WhileFord et al. [9] showed in rat cardiac muscle, that the
inhibitoryeffect of AEA on contractility was not reversed in the
presenceof CB1 (SR141716A) and CB2 (SR144528) receptor
antagonists,Bonz et al. [10] reported that AEA, metAEA, and HU-210
decreasedcontractile performance in human atrial muscle via
activation ofCB1 receptors. In another study in rat atria, AEA was
suggestedto have negative and positive inotropic effects mediated
by theactivation of CB1 and CB2 receptors, respectively [11]. Under
ourexperimental conditions, two structurally different CB1
antago-nists AM251 (0.3 �M) and SR141716 (0.3 �M) were not able
toreverse the inhibitory effect of AEA on cardiomyocyte
shorten-ing. Similarly, two different CB2 antagonists AM630 (0.3
�M) andSR144528 (0.3 �M) failed to antagonize AEA-induced decrease
ofcardiomyocyte shortening. However, at higher concentrations
suchas 1 �M, these antagonists themselves showed inhibitory
actionson cardiomyocyte shortening (n = 9–12; data are not shown).
To ourknowledge, negative inotropic actions of relatively high
concen-trations of AM251 and AM630 have not been reported
previously.However, negative inotropic actions of SR141716 and
SR144528on the contractile functions of isolated rat heart have
also beenattributed to direct actions of SR141716 and SR144528 on
thecontractility of cardiomyocytes [45]. Earlier studies on cardiac
mus-cle and other preparations also indicate that cannabinoid
receptorantagonists with different chemical structures can have
off-targetbinding sites ([9,29,46,47], for a review [29]). In
summary, basedon the insensitivity of the effect of AEA on myocyte
shorteningto CB1 and CB2 antagonists, as well as to the
pretreatments withPTX and NEM, it is likely that AEA decreases
myocyte shorteningand shortens AP duration in a manner that is
independent of CB1and CB2 cannabinoid receptors. In agreement with
these results,AEA, at similar or higher concentrations, has been
shown to inhibitdirectly the functions of voltage-gated Na+
channels in neuronalstructures [48–50], L-type Ca2+ channels
[51,52] and various typesof K+ channels (for a review [34]).
AEA belongs to a group of fatty acid-based molecules
calledlong-chain N-acylethanolamines (NAEs) which are produced
abun-dantly in response to tissue necrosis and cellular stress
[53,54].In fact, accumulation of NAEs was first observed in
experimentalmyocardial infarction induced by ligation of coronary
arteries incanine heart ([37,55], for a review [56]). It was
demonstrated thatNAE content increases up to 500 nmol/g
(approximately 500 �M) ininfarcted areas of canine heart during
ischemia [55]. Although AEAconstitutes minor (1–3%) portion of
total NAE levels [56], the resultsof this study may have important
implications regarding the con-tractile responses of ventricular
myocytes to ischemia and cellularstress [53,54,56]. We have
previously reported that major NAEsspecies such as
N-stearylethanolamine and N-oleoylethanolamineproduced during
ischemia have significant effects on the ampli-tudes and kinetics
of APs and accompanying ionic currents that
could account for the negative inotropic actions of these
com-pounds on ventricular myocytes [57]. Similar to AEA, other
NAEs,metabolic degradation products of NAEs, and structurally
relatedcompounds have also been shown to modulate functions of
-
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L.T. Al Kury et al. / Cell
oltage-gated Ca2+ [57,51,58,59], Na+[48,51], and ATP modulated+
channels [60]. In this context, it is important to note that
metAEA,etabolically stable analog of AEA, also decreased the
shortening ofyocytes. Furthermore, AEA continued to inhibit myocyte
shorten-
ng after pretreatment of these cells with URB597, a potent
inhibitorf fatty acid amide hydrolase, suggesting that degradation
productsf AEA are not involved in observed effects of this
compound.
Shortening of AP duration by AEA can be beneficial or harm-ul,
depending on the underlying pathology. Thus, during acuteschemia,
in which the duration of the cardiac APD is alreadyhortened, a
further decrease should be proarrhythmic [61]. How-ver, the APD
shortening should be beneficial in preventing thoserrhythmias
caused by triggered activities observed in conditionsuch as heart
failure [61,62]. In conclusion, the results indicateor the first
time that AEA inhibits myocyte contractility by act-ng on multiple
mechanisms. In the present study, AEA decreasedhe duration of APs
and modulated the activity of Ca2+-ATPase in
CB1 and CB2 receptor-independent manner. Considering mas-ive
release of various NAEs including AEA during ischemia andypoxic
conditions, further understanding of their target proteinsnd action
mechanisms would help in the development of betterreatment
modalities for these pathological conditions.
onflict of interest
Authors have no conflict of interest in this study.
cknowledgments
This study was supported by the United Arab Emirates Univer-ity
Research Funds. Research in our laboratory is also supported byABCO
partner of Sigma–Aldrich. We cordially thank Mr. Muham-ad A.
Qureshi for his technical help in myocyte isolation andyocyte
shortening experiments. This article has been submitted
o fulfill in part the thesis requirements for Ms. Lina Al
Kury.
ppendix A. Supplementary data
Supplementary data associated with this article can be found,
inhe online version, at
http://dx.doi.org/10.1016/j.ceca.2013.12.005.
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