Do rat cardiac myocytes release ATP on contraction?

Stefanie Gödecke,1 Thomas Stumpe,1 Hilmar Schiller,2 Hans-J. Schnittler,3 and Jürgen Schrader1

1Institut für Herz- und Kreislaufphysiologie, Heinrich-Heine-Universität, Düsseldorf; 2Cardion, Erkrath; and 3Institut für Physiologie, Medical Fakultät Carl-Gustav-Carus, Dresden, Germany

Submitted 15 February 2005 ; accepted in final form 21 April 2005


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
ATP is released by numerous cell types in response to mechanical strain. It then acts as a paracrine or autocrine signaling molecule, inducing a variety of biological responses. In this work, we addressed the question whether mechanical force acting on the membranes of contracting cardiomyocytes during periodic longitudinal shortening can stimulate the release of ATP. Electrically stimulated isolated adult rat cardiomyocytes as well as spontaneously contracting mouse cardiomyocytes derived from embryonic stem (ES) cells were assayed for ATP release with the use of luciferase and a sensitive charge-coupled device camera. Sensitivity of soluble luciferase in the supernatant of cardiomyocytes was 100 nM ATP, which is ~10-fold below the EC50 values for most purinergic receptors expressed in the heart (1.5–20 µM). Light intensities were not different between resting or contracting adult rat cardiomyocytes. Similar results were obtained with ES-cell-derived contracting mouse cardiomyocytes. ATP release was measurable only from obviously damaged or permeabilized cells. To increase selectivity and sensitivity of ATP detection we have targeted a recombinant luciferase to the sarcolemmal membrane using a wheat germ agglutinin-IgG linker. Contraction of labeled adult rat cardiomyocytes was not associated with measurable bioluminescence. However, when human umbilical vein endothelial cells were targeted with membrane-bound luciferase, shear stress-induced ATP release could be clearly detected, demonstrating the sensitivity of the detection method. In the present study, we did not detect ATP release from contracting cardiomyocytes on the single cell level, despite adequate sensitivity of the detection system. Thus deformation of the contracting cardiomyocyte is not a key stimulus for the release of cellular ATP.

cardiomyocytes; luciferase


EXTRACELLULAR ATP acts as a potent agonist on a variety of different cell types, including cardiomyocytes (20), inducing a broad range of physiological responses. The cellular effects mediated by ATP are determined by the subtypes of P2 purinergic receptors expressed in the particular cell type. In cardiomyocytes, the expression of ionotropic P2X1–P2X7 receptors and metabotropic P2Y1, P2Y2, P2Y4, P2Y6, and P2Y11 receptors have been described (37). The sensitivity of these receptors toward extracellular ATP is characterized by their EC50-ATP values, which are reported to be in the low micromolar range (37). The diversity of ATP receptors expressed in cardiomyocytes reflects the variety of ATP effects described for single cells as well as for the whole organ. In the single cardiomyocyte, micromolar levels of extracellular ATP increase plasma membrane permeabilities for cations (28) and for Cl (18), intracellular calcium transients (37, 43), the rate of phosphoinositide hydrolysis (29, 41), and the contraction amplitude (24, 27, 43). Moreover, ATP can stimulate phospholipase C (PLC) (27, 29), influence pH and inhibit glucose transport (11). On the organ level, ATP acts as a positive inotropic agent (24) and can induce various forms of arrhythmia (37). Furthermore, ATP can induce vasodilation by interacting with P2Y receptors of the vascular endothelium.

Low levels of ATP can be detected in the effluents of isolated, saline-perfused hearts (4). Because extracellular ATP is known to be rapidly degraded by ecto-ATPases present on endothelial cells and cardiomyocytes (44), much higher local concentrations of ATP can be assumed to exist locally at the site of release. Kuzmin et al. (21) estimated the baseline interstitial fluid ATP concentration in hearts in situ to be ~40 nM. However, a variety of conditions, such as electrical stimulation (1), hypoxia and ischemia (10, 21, 40), increased blood flow (39), {beta}-adrenergic stimulation (4), or mechanical stretch (36) have all been reported to be associated with a marked increase in the cardiac release of ATP.

Extracellular ATP in the cardiovascular system may originate from different cellular sources. ATP is well known to be a cotransmitter in perivascular sympathetic nerve endings (5). Activated platelets release substantial amounts of ATP during hemostasis (14). Furthermore, endothelial cells (3), vascular smooth muscle cells (26), erythrocytes (32), and inflammatory cells (8) were described as possible sources for extracellular ATP. Under ischemic conditions, myocytes release ATP (10, 11, 40), and it has also been speculated that exercising heart muscle cells are a source of extracellular ATP, similar to what has been reported for the working skeletal muscle cell (12, 25). Finally, cells disrupted, e.g., on hypoxia or vascular injury, release substantial amounts of their cytosolic nucleotides.

Mechanical strain leads to the release of ATP from a variety of cell types. Endothelial cells release ATP in response to fluid shear stress, whereas this is not the case for vascular smooth muscle cells (3). Mechanical perturbation of the cell membrane during hypoosmotic swelling mediates ATP release from prostate cancer cells (30), epithelial cells (15), and also neonatal cardiomyocytes (10). Furthermore, it has been reported that in cultured neonatal cardiomyocytes, ATP release (36) and a variety of stretch-induced biochemical events such as activation of protein kinases or an increase of protein synthesis (42) can be stimulated by application of mechanical force. During the cycle of systolic contraction and diastolic relaxation, the sarcolemmal membrane is subjected to continuous and pronounced mechanical deformations. In this study, we addressed the question whether this mechanical stimulus is sufficient to induce the release of ATP from the contracting heart muscle cell. This cardiomyocyte-derived ATP may then act in an autocrine way on the different P2 receptors expressed by this cell type, by modulating, for example, the contractile function of the cell.

We measured ATP release from contracting rat cardiomyocytes, derived as primary cultures of adult cells or genetically generated from mouse embryonic stem (ES) cells, respectively. ATP was assessed using soluble luciferin/luciferase reagent or a luciferase-protein A fusion protein targeted to the cell membrane in combination with a highly sensitive charge-coupled device (CCD) camera, allowing the observation of single cells. The sensitivity of the detection system was confirmed by using shear stress-stimulated human umbilical vein endothelial cells (HUVECs) as a positive control for local and transient release of ATP.


    METHODS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Preparation and culture of adult rat cardiomyocytes. Adult Ca2+-sensitive cardiomyocytes of Wistar rats (250–300 g) were prepared as described previously (34). After preparation cells were seeded on laminin (10 µg/ml)-coated coverslips (~28 mm) and incubated in serum-free medium 199 (M199), supplemented with nonessential amino acids, vitamins, gentamicin (50 µg/ml), penicillin (250 IU/ml), streptomycin (250 µg/ml), and insulin (1,000 IU/l). The medium was changed after 3 h to remove nonadherent cells. After at least 12 h, and a maximum of 2 days, cardiomyocytes were used for experiments.

ES cell culture and generation of genetically selected cardiomyocytes. ES cell-derived mouse cardiomyocytes were generated as described (19). Undifferentiated J1 ES cells were transfected via electroporation with a construct carrying {alpha}-myosin heavy chain (MHC)-aminoglycoside phosphotransferase (MHC-neor) and a phosphoglycerate kinase (pGK)-hygromycin-resistance transgene. Transfected cells were selected by incubation in growth medium containing hygromycin B (200 µg/ml), resulting in the isolation of clone CM7/1. For differentiation, 4 x 106 ES cells per 100-mm bacterial dish were plated and cultured in growth medium lacking leukemia inhibitory factor. After 4 days of being cultured, the resulting embryoid bodies were plated onto 35-mm cell culture dishes. When spontaneous contractile activity was noticed, growth medium supplemented with G418 (400 µg/ml) was added to eliminate noncardiomyocytes.

Preparation and culture of HUVECs. Human umbilical cords were obtained from a normal placenta. The umbilical vein was cannulated with blunt needles and perfused to wash the blood out. The vein was then filled with collagenase IA (Sigma) 0.025%, warmed at 37°C. Isolated endothelial cells were seeded in 25-cm2 flasks coated with 0.2% gelatin A in endothelial cell basal medium (Promocell) and were incubated at 37°C and 5% CO2 in a humidified incubator. Trypsinization was performed when endothelial cells reached 70–90% confluence.

Coupling of firefly luciferase to surface of adult rat cardiomyocytes or HUVECs. The principle underlying the coupling of firefly luciferase to the membranes of cardiomyocytes performed in this study is depicted in Fig. 3. N-Acetylglucosamine residues present on cell membranes in high copy numbers as parts of glycosylated membrane proteins are specifically recognized by the lectin portion of the WGA-IgG complex. The IgG component in turn interacts specifically with the Z-domains of the protein A-luciferase fusion protein.



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Fig. 3. Schematic diagram of luciferase targeting to the sarcolemmal membrane. GlucNac, N-acetylglucosamine; WGA, wheat germ agglutinin. For details, see RESULTS. ?, ATP release is considered hypothetical.

 
Generation of wheat germ agglutinin-IgG complexes. Rabbit anti-mouse IgG (whole molecule) was conjugated to wheat germ agglutinin (WGA) with sulfosuccinnimidyl 4-[N-maleimidomethyl]cyclohexane-1-carboxylate (sulfo-SMCC) as a cross-linking molecule with the use of the Controlled Protein-Protein Cross-Linking Kit (Pierce). In brief, 7 mg rabbit IgG were activated with sulfo-SMCC according to the manufacturer's instructions for generating maleimide-IgG. Free sulfhydryl groups were introduced into 1 mg of WGA by reaction with N-succinimidyl-S-acetylthioacetate, followed by deprotection of the masked SH-groups with hydroxylamine. Immediately after deprotection, sulfhydryl-WGA was mixed with maleimide-IgG in approximately equal molar amounts. The cross-linked conjugate was dialyzed against cardiomyocyte stimulation buffer (see below) or PBS and concentrated using Centriprep YM-3 centrifugal filter devices (Amicon/Millipore). Final protein concentrations were adjusted to 100 µg/ml. Aliquots (100 µl) were rapidly frozen and stored at –80°C.

Generation of protein A-luciferase fusion protein. For construction of a fusion gene containing four synthetic Z-domains based on the IgG binding domain of protein A fused to the NH2 terminus of the firefly luciferase gene, the following cloning strategy was used: plasmid pGem-luc (Promega) was linearized with BamHI at the 5' end of the luciferase gene. A KpnI site was inserted by oligonucleotide ligation. The resulting plasmid pGem-K was linearized with NotI, treated with Klenov polymerase to generate blunt ends and cut with KpnI. A 388-bp FspI KpnI fragment isolated from plasmid pEZZ18 (Amersham Pharmacia) containing two synthetic Z-domains was integrated, thereby reconstituting the NotI site 5' to the Z-domains. To amplify the number of IgG binding motifs, the resulting plasmid pGem-Z was linearized with NotI and blunt ends were generated by Klenov treatment. A 373-bp FspI EcoRI fragment from pEZZ18 containing the sequence identical to the above-mentioned FspI KpnI fragment was inserted, resulting in plasmid pGem-ZZ. This construct contains an in- frame fusion of a short part of the 5' coding region of the lacZ {alpha}-peptide, followed by four synthetic Z-domains and the coding region of the luc gene under control of the inducible lac promoter.

Plasmid pGem-ZZ was transformed into competent E. coli HB101 cells. Bacterial lysates of isopropyl-{beta}-D-thiogalactopyranoside-induced cultures were prepared as described by others for an almost identical fusion protein (2). However, due to the high degree of luciferase activity loss during IgG-agarose affinity purification, we omitted the final affinity purification step. Instead, after the filtration step, the bacterial lysate was dialyzed against cardiomyocyte stimulation buffer (see below) or PBS and concentrated with the use of centrifugal filter devices (Centriprep YM-3, Amicon/Millipore). Final protein concentrations were adjusted to 7.5 µg/µl. Aliquots (100 µl) were snap-frozen and stored at –80°C. Luciferase activity of bacterial extracts was determined in a total volume of 50 µl by mixing 40-µl reaction buffer, which contained (in mmol/l) 20 tricine, 0.033 coenzyme A, 1.07 (MgCO3)4Mg(OH)2, 0.1 EDTA, 33.3 DTT, 2.67 MgSO4, 0.53 ATP, and 0.47 luciferin, pH 7.3, with 10 µl of various dilutions of bacterial extract. Luminescence was monitored in 10-s intervals at 25°C with a Berthold luminometer (model Biolumat LB 9500 T). Luciferase activity was measured as 9 x 1010 relative light units·mg protein–1·s–1.

The specific activity of the luciferase was not altered by addition of the NH2-terminal Z-domains, as could be shown by comparative activity assays and Western blot experiments using anti-luciferase antibodies (Promega) on bacterial pGEM-luc and pGem-ZZ extracts (data not shown).

Attachment of luciferase to cell surfaces. For experiments with cell surface-bound luciferase, rat cardiomyocytes were cultured as described above on laminin-coated glass coverslips. Culture medium was carefully removed and the coverslips were rinsed twice with cardiomyocyte stimulation buffer that contained (in mmol/l) 137 NaCl, 5.4 KCl, 1.0 NaH2PO4, 0.8 Mg2SO4, 2.0 CaCl2, 5.5 glucose, and 5.0 Tris, pH 7.4. Cells were incubated in stimulation buffer containing WGA-IgG complex (diluted 1:10) for 30 min at room temperature, allowing the WGA residues to bind to carbohydrates on the cell surfaces. Subsequently, cells were carefully washed with stimulation buffer and overlaid with a 1:10 dilution of bacterial extract containing the luciferase fusion protein. During a 30-min incubation period at room temperature, the Z-domains of the fusion protein could bind to the surface-coupled IgG residues. After being washed, the coverslips were brought into the perfusion/stimulation chamber containing stimulation buffer, and cells were allowed to equilibrate for 30 min before being monitored with a CCD camera.

For experiments with HUVECs, cells were cultured as described above on gelatin-coated glass coverslips. Loading of cells was performed with WGA-IgG complex and luciferase extracts in PBS, diluted 1:10 in endothelial cell basal medium.

Bioluminescence measurement on adult rat cardiomyocytes. Coverslips with luciferase-labeled adult cardiomyocytes were introduced into a perfusion chamber, allowing both microscopical observation and electrical stimulation of the cells. The exact geometry of the chamber has been described previously (35). After an equilibration period of 30 min, the chamber was perfused at constant flow (0.5 ml/min) with cardiomyocyte stimulation buffer (see above) containing 150 µM D-luciferin (Sigma), 100 µg/ml coenzyme A (Sigma), and in some experiments, various ATP concentrations. In some experiments, cells were electrically stimulated to contract using the lowest possible stimulation strength plus 10% to minimize cell damage (stimulator model G270, Strotmann, Aachen, Germany; 1 Hz, biphasic impulses between 10 and 20 µA) using platinum electrodes. Light emission was monitored using a Zeiss Axiovert 35 microscope in combination with a CCD camera (Micromax 512 BFT System, Visitron Systems, Puchheim, Germany). Luminescence values were calculated using MetaMorph Software (Universal Imaging).

For experiments with soluble luciferase, coverslips with adult cardiomyocytes were introduced into the perfusion chamber and perfused with contraction buffer containing 2 mg/ml luciferin/luciferase reagent (Sigma). It was especially important to use freshly diluted luciferin/luciferase reagent for each measurement because luciferase activity is generally known to be very delicate and dependent on buffer composition. In particular, the high-chloride concentration of the cardiomyocyte buffer (140 mM NaCl) necessary for cell viability constitutes suboptimal reaction conditions for the enzyme (7).

Stimulation was performed as described above. For measurements with ES cell-derived mouse cardiomyocytes grown on 35-mm cell culture dishes, the culture medium was exchanged with medium containing luciferin/luciferase reagent (2 mg/ml). Measurements were started after the cells began to contract spontaneously. All measurements were performed with prewarmed buffers at 37°C.

Bioluminescence measurement on HUVECs under fluid shear stress. Shear stress-induced ATP release from HUVECs was detected either luminometrically or with a CCD camera. For luminometric detection of ATP, HUVECs were grown to confluency on a gelatin- coated cover slides (0.75 cm2) and loaded with recombinant luciferase. The slides were then inserted into a self-designed, parallel-plate flow chamber that was integrated into a luminescence device (Berthold Biolumat LB 9500). A peristaltic pump was used to repeatedly apply 5-s shear-stress pulses of 15 dyn/cm2 under constant removal of the flow-through and online luminometric measurements at 37°C. The cross-section of the flow chamber was 0.4 x 0.05 cm2. Wall shear stress was calculated using the equation 3yQ/2a2b (22), where y is the viscosity (poise) of the medium at 37°C, Q is the volumetric flow rate (in ml/s), a is the half-channel height (in cm), and b is the channel width (in cm). Luminescence rates were recorded automatically in 2-s intervals.

For experiments performed using image analysis (CCD camera), a cone-and-plate rheological device as described by Schnittler et al. (31) was used to create defined levels of laminar shear stress (6) at 37°C in a closed system. HUVECs were seeded onto cross-linked gelatin-coated perfectly plan-parallel and round glass coverslips (diameter 28 mm, Assistent), grown to confluency, and loaded with recombinant luciferase. The rheological device is equipped with a microscope coupled to a Micromax 512 BFT System allowing online observation of shear stress-exposed cells by time lapse recording. Basal luminescence was measured for 30 s before the beginning of the experiment. Repeated 5-s shear stress impulses of 15 dyn/cm2 were applied to the cells, and luminescence was then recorded and normalized for the basal luminescence.


    RESULTS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In initial experiments, freshly isolated adult rat cardiomyocytes were attached to a laminin-coated coverslip, allowing the cells to contract on electrical stimulation. Cells were then brought into cuvettes containing soluble luciferase/luciferin reagent dissolved in cardiomyocyte stimulation buffer, which then was inserted into a luminometer and stimulated electrically with two platinum electrodes. To avoid cell damage, stimulation parameters were chosen to be just above the stimulation threshold (biphasic pulse, 200 µs, 10 µA). ATP release was clearly detectable on stimulation (data not shown). However, microscopical observation of the cells revealed that several cardiomyocytes were apparently damaged during electrical stimulation, visible by the loss of their typical rod-shaped phenotype. Hence, it was not possible to distinguish in these experiments whether the ATP was selectively released from beating cardiomyocytes or originated from cells damaged during the experimental procedure.

Because overall measurements may have yielded erroneous results, we decided to measure ATP release from single beating cardiomyocytes using a sensitive CCD camera system. A specially designed cardiomyocyte perfusion chamber allowing both electrical stimulation and microscopical observation was used (35). As shown in Fig. 1, AC, for a typical experiment, we defined regions that matched individual cells (regions 13). Only rod-shaped cells, which contracted on electrical stimulation (biphasic pulse, 200 µs, 10 µA), were chosen for further experiments (regions 1 and 2). Cells that evidently had lost their rod-shaped phenotype at the beginning of the experiment and did not contract on stimulation (region 3) were excluded from the analysis. Bioluminescence determined from resting, not electrically stimulated, cells (Fig. 1B) was compared with the bioluminescence of identical cells stimulated electrically to contract (Fig. 1C). Altogether, 21 resting and 16 contracting cells derived from 5 independent experiments were analyzed. As depicted in Fig. 1D, light intensities were 1,117.9 ± 83.7 arbitrary units per pixel for the resting cells and 1,065.3 ± 78.3 arbitrary units per pixel for the contracting cells. We conclude that no detectable ATP release is observed in contracting cardiomyocytes compared with resting cells. The weak ATP signal observed in rounded cells (region 3) was independent of electrical stimulation and was most probably a consequence of lost membrane integrity and slow leakage of intracellular ATP due to cell damage.



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Fig. 1. Bioluminescence measurement over resting and contracting adult rat cardiomyocytes using soluble luciferase for ATP detection. A: light-microscopical image of two intact (regions 1 and 2) and one hypercontracted (region 3) adult rat cardiomyocyte perfused with contraction buffer containing luciferin/luciferase reagent. B: charge-coupled device (CCD) camera image of the same cells without electrical stimulation. C: CCD camera image of the same cells under electrical stimulation leading to contraction of cells 1 and 2 but not cell 3. D: average light intensities measured over resting (n = 21) and contracting (n = 16) cells from 5 independent experiments. AU, arbitrary units. The scale bar in A is 25 µm.

 
To arrive at an estimate of sensitivity of our measuring system, we mixed different concentrations of ATP to the luciferin/luciferase solution in our perfusion chamber and recorded luminescence. ATP concentrations in the midnanomolar range (100 nM, data not shown) could clearly be detected.

To exclude the possibility that membrane properties of isolated cardiomyocytes might have been altered due to enzymatic digestion, a further series of experiments were carried out using genetically selected cardiomyocytes derived from mouse ES cells. A stable ES cell line containing a vector with both MHC-neor and a pGK-hygromycin resistance transgene was selected to generate spontaneously contracting cardiomyocytes, as described in detail by Klug et al. (19). For ATP measurements, embryoid bodies were plated on 35-mm-thick cell culture dishes and used for CCD camera monitoring 4–7 days after the appearance of spontaneously contracting cell regions. At that time, cells had not yet formed a confluent monolayer. Discrete clusters of cells could be distinguished, of which ~50% showed spontaneous contraction. The representative experiments demonstrated in Fig. 2, AC, show an ATP signal over both cell clusters (Fig. 2B), whereas only cluster 1 showed contractile activity. Calculation of average intensities obtained from three independent experiments revealed that there was no difference between the marked regions, indicating that the ATP detected was not released as a consequence of contraction (Fig. 2D). Inhibition of cell contraction with 10 mM BDM (Fig. 2C) reduced light intensities. This effect, however, can be fully explained by a direct inhibitory effect of BDM on the luciferase observed in separate luminometric measurements (data not shown). Release of intracellular ATP by disruption of the cell integrity with digitonin was clearly visible. Digitonin dose-dependently increased light intensities 3- and 25-fold over resting values (Fig. 2D).



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Fig. 2. Bioluminescence measurement over resting and contracting embryonic stem (ES) cell-derived mouse cardiomyocytes using soluble luciferase for ATP detection. A: light-microscopical image of one spontaneously contracting (region 1) and one noncontracting cluster (region 2) of mouse cardiomyocytes genetically selected from ES cells, perfused with contraction buffer containing luciferin/luciferase reagent. B: CCD camera image of the same cell regions. C: CCD camera image of the same cell regions perfused with contraction buffer containing luciferin/luciferase reagent and 10 mM butanedione monoxime (BDM) to completely block contraction. D: average light intensities measured over resting (n = 5) and contracting (n = 11) cell regions from 3 independent experiments. In 2 experiments, digitonin was added to release intracellular ATP.

 
Recent studies have shown that detection of ATP released from astrocytoma cells in the bulk extracellular medium was possible only after pharmacological inhibition of ecto-ATPase activity. However, detection of submicromolar ATP concentrations in the cell surface environment by surface-targeted luciferase was possible without pharmacological intervention, indicating an efficient and rapid degradation of released ATP by surface ectonucleotidases (17). Because ectonucleotidase activity has been reported on the cardiomyocyte cell surface (23, 44), we decided to bring luciferase molecules closer to the region of potential ATP release, namely the sarcolemmal membrane. For this purpose, we have targeted a recombinant luciferase fusion protein containing four IgG binding motifs of Staphylococcus aureus protein A to the cell surface by using a chemically coupled WGA-IgG linker molecule. WGA is a lectin that binds mainly to N-acetylglucosamine (GlcNAc) residues from glycosylated molecules in the glycocalyx, and it has been shown to bind to the surface of isolated rat cardiomyocytes (33). Figure 3 schematically depicts the principle of the WGA-IgG-mediated luciferase binding. For measurement of potential ATP release, isolated rat cardiomyocytes were loaded with WGA-IgG and ZZ-luciferase, as described in METHODS.

For CCD camera monitoring, regions were defined matching the positions of rod-shaped, excitable cells (Fig. 4A). Light intensities were sampled and corrected for luminescence values from neighboring cell-free control regions. Bioluminescence measurements were performed in the absence (Fig. 4B) and presence of electrical stimulation (Fig. 4C). Altogether, 31 contracting and 26 resting cells taken from 5 independent experiments were analyzed. The corrected light intensities were 0.655 ± 6.08 arbitrary units/pixel for resting cells and –0.052 ± 8.922 arbitrary units/pixel for contracting cells. No increase in light intensity could be detected in the immediate vicinity of contracting cells.



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Fig. 4. Bioluminescence measurement over resting and contracting adult rat cardiomyocytes using cell surface-attached luciferase for ATP detection. A: light-microscopical image of adult rat cardiomyocytes loaded with membrane attached protein A-luciferase and perfused with contraction buffer containing luciferin. Scale bar is 25 µm. B: CCD camera image of the same cells without electrical stimulation. C: CCD camera image of the same cells with electrical stimulation. D: CCD camera image showing the membrane-bound luciferase visualized by perfusion with buffer containing luciferin and 100 µM ATP.

 
To confirm membrane binding of ZZ-luciferase, ATP was added at the end of each experiment (Fig. 4D). To determine the sensitivity of our system, the cardiomyocytes were loaded with ZZ-luciferase and light emission was recorded over resting cells after the addition of various concentrations of ATP. From the data shown in Fig. 5, it is evident that threshold concentration for ATP producing measurable bioluminescence was 3.3 x 10–5 M. However, the sensitivity was quite variable in individual experiments, and in some experiments, even 330 nM could be clearly detected.



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Fig. 5. Bioluminescence of cell surface attached luciferase bound to adult rat cardiomyocytes. Bioluminescence of 26 quiescent and 31 contracting cells from 5 independent experiments in luciferin-containing buffer was determined by CCD camera monitoring. The sensitivity of the system was determined by monitoring luciferase-loaded cells after addition of a range of ATP concentrations. Numbers in parentheses refer to the cells monitored for each ATP concentration, taken from 2–5 independent preparations. ANOVA, post hoc test Bonferroni (P < 0.0001 for 10–4 and 10–3 M ATP); unpaired Student's t-test (P < 0.0001 for 3.3 x 10–5, 10–4, and 10–3 M ATP) vs. quiescent cardiomyocytes.

 
To demonstrate that membrane-targeted luciferase can sensibly detect cellular ATP release, we measured shear stress-induced ATP secretion reported for cultured HUVECs (3). Figure 6A shows a typical luminometrical detection of shear stress-induced ATP release from HUVECs loaded with membrane-bound luciferase. Sharp luminescence peaks can be observed immediately on stimulation. According to the literature (3), repeated shear stress impulses caused reduced ATP signals. Perfusion with medium containing 1 µM ATP at the end of the experiment showed that the observed ATP release falls into the micromolar range. As shown in Fig. 6B, shear stress-induced ATP release from HUVECs can also be detected by the CCD camera.



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Fig. 6. Bioluminescence of shear stress-induced ATP release from human umbilical vein endothelial cells (HUVECs) measured with cell surface-attached luciferase. A: luminometric detection of ATP released from HUVECs loaded with membrane-attached protein A-luciferase and stimulated with 5-s pulses of shear stress (15 dyn/cm2) in a flow-through system. At the end of the experiment, the medium in the chamber was exchanged from basal medium to basal medium containing 1 µM ATP. Luminescence was measured continuously in 2-s intervals. B: shear stress-induced ATP release from luciferase-tagged HUVECs. Luminescence was measured with a CCD camera at selected time points over 30 s.

 

    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Release of ATP from the contracting heart has been reported in several studies (3, 9, 13). Besides nonmyocyte cells (reviewed in Ref. 37), cardiomyocytes have also been speculated to be a source of extracellular ATP. Recently, Dutta and coworkers (10) have described an ATP conductive pathway in neonatal rat cardiomyocytes that responds to hypoxia and ischemia but also mechanical stretch through osmotic swelling with pronounced ATP release. The maxi-anion channel involved in this pathway is also present, albeit less frequently in adult rat cardiomyocytes. This study provides clear evidence that mechanical stretch of the sarcolemmal membrane during regular contraction is not associated with a detectable release of ATP from isolated cardiomyocytes. This was measured at the single cell level with soluble luciferase as well as luciferase tagged to the cell surface of cardiomyocytes in combination with a sensitive CCD camera allowing high-resolution detection.

Measurement of ATP release from a whole organ such as the heart comprises a variety of different cell types, including, aside from cardiomyocytes, nerve terminals, coronary endothelial cells, smooth muscle cells, and red blood cells, all of which have been described to be potential sources of ATP (37). It is therefore very difficult to clearly assign a measured extracellular ATP signal to a defined cell type in experiments using, e.g., isolated perfused hearts. Furthermore, experimental manipulations like electrical stimulation almost inevitably lead to the damage of an undefined amount of cells, which as a consequence release their intracellular nucleotides to the surrounding medium. ATP within cardiomyocytes is normally present in millimolar concentrations, so that few damaged cells can give rise to a substantial extracellular ATP signal. Considerations similar to those for the intact organ apply to isolated cells in culture. Also, in this reduced system, it is technically very difficult to distinguish between a nonselective release mechanism and a selective release from intact cells. In fact, in initial experiments, we have observed extracellular ATP on electrical stimulation when using luminometric detection of signals derived from cardiomyocytes attached to coverslips. However, we also observed that on electrical stimulation, some cardiomyocytes hypercontracted and lost their rod-shaped structure, indicative of cell damage and ATP release.

To circumvent these obvious problems, we have decided to analyze ATP release by microscopical observation of single cells. Only single cardiomyocytes showing regular contraction on electrical stimulation were chosen to investigate the release of ATP, thereby clearly excluding nonmyocytes, as well as damaged cells as potential contaminating sources. The amount of ATP released by a single cell experiment is naturally lower than the quantities released by overall measurements in cell culture. To compensate for this drop in signal intensity we utilized a liquid nitrogen-cooled CCD camera, presently the most sensitive device to detect low light intensities, together with spatial allocation of the signal to a defined region: one contracting cardiomyocyte.

For the detection of ATP we used the luciferin-luciferase reaction known to be a highly sensitive method for the detection of ATP in aqueous solution. It should be noted, however, that the enzyme activity is impaired by suboptimal reaction conditions, such as a temperature of 37°C or physiological salt solutions necessary to provide a normal physiological environment for contracting cardiomyocytes. Luciferase activity has been reported to have an optimal reaction temperature of 25°C and to be very sensitive to salt conditions, such as high Cl concentrations (6). In our experiments, we have added high concentrations of soluble luciferase to the supernatant (2 mg/ml) to compensate for lowered enzyme activity. The limit of detection in these experiments was ~100 nM ATP. In the experiments with genetically selected, ES-cell-derived mouse cardiomyocytes, there was always a detectable but contraction-independent release of ATP over the cells. Whether this basal ATP release is a general feature of ES-cell derived cardiomyocytes or whether it relates to the genetic selection procedure used is presently not known. Nevertheless, our detection system is sensitive enough to measure ATP release with high local resolution.

Several possible mechanisms might have hampered the detection of released ATP by soluble luciferase. The released ATP is rapidly diluted by diffusion and in addition is rapidly degraded by ectonucleotidases present on the cardiomyocyte surface (44). Furthermore, the release of ATP might be spatially confined to specific regions of the cell surface. Therefore, measurement of ATP in the bulk supernatant might underestimate the actual ATP concentrations at the location of release, namely the cell surface. We have thus attempted to measure ATP by targeting the luciferase to the cell surface. To this end, we have modified a method developed by Beigi et al. (2) using antibodies against surface molecules of a target cell to tag a luciferase-protein A fusion protein to the cell surface. This elegant method, however, has several limitations, most prominently the availability of the appropriate antibodies against surface properties of target cells. We therefore have used a more general feature of cell surfaces to couple the luciferase to the membrane, namely the appearance of glycosylated residues, which can be recognized by lectins. Because glycosylated residues are a general characteristic feature of cell surfaces of every cell type, lectins chemically cross-linked to unspecific rabbit IgG instead of a specific antibody can be used to target protein A-fusion proteins to any cell type. In our particular case, we used WGA, a lectin specifically recognizing N-acetylglucosamine residues. The WGA-IgG linker molecule was then employed to target protein A-tagged luciferase to the cell membrane of cardiomyocytes. With the use of this method, we routinely were able to measure ATP when added in the midmicromolar range; in two of five experiments, we clearly detected 1 µM ATP. Beigi et al. (2) report in their work a detection level in the midnanomolar range. It should be mentioned, however, that the cell numbers in these experiments were between 1 x 106 (for promyelocytes) and 7 x 107 (for platelets) for each measurement, whereas for reasons discussed above, we were measuring single cells. The lower sensitivity of the tagged luciferase for exogenously added ATP is most likely due to the lower absolute quantity of enzyme molecules bound to the cell surface. Occasionally, we observed light signals over rounded and therefore damaged cells (see Fig. 1, region 3), indicating leakage of intracellular ATP. The tagged luciferase, however, is sufficiently sensitive to measure cellular ATP secretion. Nonlytic release of ATP from HUVECs after stimulation with fluid shear stress served as a positive control and clearly demonstrated that our detection system can measure the release of "physiological" amounts of ATP at the cellular level.

Baseline concentration of ATP in the interstitial space of the rat heart has been determined to be ~40 nM (21). Given the sensitivity of our measurements, it can be concluded that the ATP concentration at the cell surface does not fall within the range of EC50-ATP values reported for cardiomyocyte P2 receptors. In case of P2X receptors EC50-ATP values between 1 µM for P2X1 and P2X2 and 20 µM for P2X4 were reported (37), whereas for P2Y2 receptors the EC50-ATP values were found to be 3 µM for P2Y1 (16) and in the range of 1.5–5.8 µM for P2Y2 (38).

In summary, the reported release of ATP from intact beating hearts under normoxic conditions is unlikely to be derived from cardiomyocytes. ATP is rather released from nonmuscle cells, such as coronary endothelium or nerve terminals. Cardiomyocytes, however, may contribute to the general release of nucleotides under conditions of ischemia, anoxia, or other conditions of cell damage, but not as a consequence of mechanical deformation of the sarcolemmal membrane during contraction.


    FOOTNOTES
 

Address for reprint requests and other correspondence: S. Gödecke, Institut für Herz- und Kreislaufphysiologie, Heinrich-Heine-Universität, Düsseldorf, Universitätsstrasse 1, D-40225 Düsseldorf, Germany (e-mail: stefanie.goedecke{at}uni-duesseldorf.de)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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