Reduces Palytoxin Toxicity in Primary Cultures of Skeletal Mus

Article pubs.acs.org/crt

The Stretch-Activated Channel Blocker Gd3+ Reduces Palytoxin Toxicity in Primary Cultures of Skeletal Muscle Cells Giorgia Del Favero,† Chiara Florio,† Barbara Codan,‡ Silvio Sosa,† Mark Poli,§ Orfeo Sbaizero,‡ Jordi Molgó,∥ Aurelia Tubaro,*,† and Paola Lorenzon† †

Department of Life Sciences and ‡Department of Industrial Engineering and Information Technology, University of Trieste, 34127 Trieste, Italy § United States Army Medical Research Institute of Infectious Diseases, Fort Detrick, Maryland 21701-5011, United States ∥ Institut Fédératif de Neurobiologie Alfred Fessard, Laboratoire de Neurobiologie et Développement, CNRS UPR 3294, 91198 Gif sur Yvette cedex, France S Supporting Information *

ABSTRACT: Palytoxin (PLTX) is one of the most toxic seafood contaminants ever isolated. Reports of human food-borne poisoning ascribed to PLTX suggest skeletal muscle as a primary target site. Primary cultures of mouse skeletal muscle cells were used to study the relationship between Ca2+ response triggered by PLTX and the development of myotoxic insult. Ca2+ imaging experiments revealed that PLTX causes a transitory intracellular Ca2+ response (transient phase) followed by a slower and more sustained Ca2+ increase (longlasting phase). The transient phase is due to Ca2+ release from intracellular stores and entry through voltage-dependent channels and the Na+/Ca2+ exchanger (reverse mode). The long-lasting phase is due to a massive and prolonged Ca2+ influx from the extracellular compartment. Sulforhodamine B assay revealed that the long-lasting phase is the one responsible for the toxicity in skeletal muscle cells. Our data analyzed, for the first time, pathways of PLTXinduced Ca2+ entry and their correlation with PLTX-induced toxicity in skeletal muscle cells. The cellular morphology changes induced by PLTX and the sensitivity to gadolinium suggest a role for stretch-activated channels.

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INTRODUCTION Palytoxin (PLTX) is the most well-known member of the PLTX family of marine toxins. It can be listed among the most toxic nonproteinaceous natural products known.1 Its discovery can be traced to the early 1970s when the group of Prof. Moore isolated it from Hawaiian soft corals of the genus Palythoa.2 PLTX and analogues are more and more frequently reported as seafood contaminants, not only in subtropical but also in temperate seas.3 The presence of PLTXs in fish and crustaceans has also been associated with human intoxications, some with lethal outcomes in tropical areas.4−7 Epidemiological data suggest skeletal muscle to be a common target in PLTX poisonings.1 In fact, the most frequently reported symptom after PLTX ingestion is myalgia, associated with rhabdomyolysis and myoglobinuria in the most severe cases. In addition, elevated serum levels of creatine phosphokinase and lactate dehydrogenase have often been reported.1,8 Similar alterations and increased plasma K+ levels have also been described during in vivo acute oral toxicity studies, supporting the muscular tissue as a sensitive target for PLTXs.9,10 At the cellular level, experimental evidence strongly suggests that PLTX binds to the Na+/K+ ATPase, impairing its activity.11−13 Current scientific consensus suggests that PLTX © 2012 American Chemical Society

binding triggers a conformational change, transforming the pump into an open ion channel with dramatic consequences to ionic homeostasis of cells.14,15 The activity of PLTX has been investigated in various different excitable cell models. In these models, PLTX causes a depolarization of the cellular membrane,16−18 which affects the mechanisms controlling intracellular calcium concentration ([Ca2+]i). The documented biological effects triggered by PLTX-induced depolarization include the disruption of the excitation−contraction coupling in cardiac cells,19 contraction of smooth muscles,20−25 and the release of neurotransmitters.26 The aim of this work was to further investigate the effects of PLTX on mouse skeletal muscle cells. The toxicity of PLTX was confirmed as a Ca2+-dependent mechanism. The novelty of our work is the evidence that the cytotoxicity is strictly related to a long-lasting and gadolinium-sensitive Ca2+ influx. With the support of 3D morphology analysis of cells incubated with the toxin, a possible involvement of stretch-activated Ca2+ channels in sustaining the long-lasting phase of the Ca2+ response is suggested. Received: May 4, 2012 Published: August 17, 2012 1912

dx.doi.org/10.1021/tx300203x | Chem. Res. Toxicol. 2012, 25, 1912−1920

Chemical Research in Toxicology

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Information). For all drugs, control experiments were performed to select a protocol (working concentrations and time exposures) that did not cause per se cell toxicity. The digital fluorescence-imaging microscopy system consisted of an inverted microscope (Zeiss Axiovert 135, Oberkochen, Germany) equipped with an intensified CCD camera (Hamamatsu Photonics, Hamamatsu, Japan). A temperature-controlled microincubator chamber (Medical Systems Corporation, Greenvale, NY) maintained the temperature at 37 °C during the experiments. Loaded cells were alternatively excited at 340 and 380 nm by a modified dual wavelength microfluorimeter (Jasco CAM-230, Tokyo, Japan). The ratio of corresponding images at 340 and 380 nm and the temporal plots of the fluorescence signal were calculated offline. Image acquisition was performed at two frames/s, and our time resolution was one ratio image/s. The temporal plots (i.e., fluorescence ratio vs time) were analyzed using Origin 6.0 software (Microcal Software Inc., Northampton, MA). [Ca2+]i variations were estimated by setting the baseline before the onset of each increase and by calculating the maximal value (peak) and the area under the temporal plot. Variation of the [Ca2+]i was expressed as [Ca2+]i increase with respect to basal value. Each set of experiments was carried out on at least three different cell preparations. Each experimental point was expressed as mean ± SE, with n being the number of cells tested. Atomic Force Microscopy. Morphology of single cells was evaluated using a Solver-Pro M (NT-MDT, Moscow, Russia) atomic force microscope (AFM). Living cells were scanned in a contact mode. The AFM cantilever was equipped with a PNP-DB silicon nitride tip (Nanoworld, Neuchâtel, Switzerland). The scan time was 10 min for 100 × 100 μm2, with a resolution of 256 × 256 points and pixel size 0.4 × 0.4 μm2. The AFM was equipped with liquid scanning setup, and measurements were performed at room temperature. Cells were maintained in NES solution (for details, see the section Ca2+ Imaging) for the entire experiment. With a Solver-Pro controller, it is possible to have multiple types of measurements during a single scan. For the purposes of this study, the variation of displacement along the z-axis has been recorded during the outward and return scan. Moreover, the variation of lateral force was also acquired. Statistical Analysis. Data are given as means ± SEs. The Student's t test and two-way ANOVA were used to examine statistical significance at the p < 0.05 level.

MATERIALS AND METHODS

Materials. Bovine fetal calf serum (FCS) was from PAA Laboratories (Mascia Brunelli, Milan, Italy). PLTX with a declared purity of more than 90% was from Wako Chemicals GmbH (Neuss, Germany). The ion channel blockers cyclopiazonic acid, gadolinium, lanthanum, and cadmium were purchased from Sigma (Milan, Italy). Verapamil (Ver), KB-R7943, and SKF-96365 were from Tocris Bioscience (Bristol, United Kingdom). PLTX was diluted in water/ethanol (1:1); cyclopiazonic acid, and KB-R7943 in Me2SO; all the other chemicals were diluted in twicedistilled water. Stock solutions were stored at −20 °C and prepared to use volumes not exceeding 5 μL to avoid any significant variation of the composition of the extracellular milieu. Primary Culture of Skeletal Muscle Cells. Cultures were established from mouse satellite cells (for details, see 27). Briefly, myoblasts were isolated from the hind leg muscles of 7 day old male Balb/c mice killed by cervical dislocation as approved by the Animal Care Committee of the University of Trieste and consistent with European legislation. Muscle tissue was minced and then treated with collagenase and trypsin. Expansion and enrichment of desmin-positive cells were achieved by repeated plating and cultivation in Dulbecco's modified Eagle's medium (DMEM) with D-valine containing 20% FCS. Myoblasts were maintained as exponentially growing cells in the presence of medium consisting of HAM's F-10 containing 20% FCS, Lglutamine (2 mM), penicillin (100 units/mL), and streptomycin (100 mg/mL). To induce cell differentiation and fusion of myoblasts into myotubes, the medium was replaced with DMEM supplemented with 2% horse serum and L-glutamine, penicillin, and streptomycin as above. Myotubes were maintained at 37 °C in CO2 (5%)-enriched air. Experiments were performed on myotubes between the fifth and the seventh day of differentiation. In these experimental conditions, spontaneous contractile activity was detectable as cell twitches that were clearly visible using a bright field microscope (20× objective). Sulforhodamine B (SRB) Assay. Cells were seeded in 96-well microplates 5−7 days before the experiments. Once differentiated, cells were treated with PLTX according to the experimental protocol. At the end of cell treatments, toxin-containing medium was removed. Cells were rinsed twice with PBS (100 μL; pH 7.4) and fixed with 50 μL of 50% trichloroacetic acid (1 h, 4 °C). After fixation, cells were rinsed twice with twice-distilled water (100 μL), and an equal volume of SRB (0.4% in 1% acetic acid) was added. After 30 min, cells were rinsed three times with acetic acid (1%) and solubilized with 200 μL of Tris (10 mM).28 Sample absorbance was read on an Automated Microplate Reader EL 311s (Bio-Tek Instruments, Winooski, VT) with a single wavelength of 570 nm. For all drugs, control experiments were performed to select a protocol (working concentrations and time exposures) that did not cause per se cell toxicity. Each experimental point was expressed as mean of at least four independent experiments performed in triplicate ± SE. Ca2+ Imaging. Videoimaging experiments were carried out according to Lorenzon and co-workers.29 Cells were plated on glass coverslips coated with Matrigel. Loading of the Ca2+ indicator fura-2 pentacetoxymethylester (fura-2 AM) was performed by incubating cells (30 min, 37 °C) in a normal external solution (NES) in mM: KCl, 2.8; NaCl, 140; CaCl2, 2; Hepes, 10; MgCl2, 2; and glucose, 10; pH 7.35) supplemented with 1 mg/mL of bovine serum albumin and 5 μM fura-2 AM. After they were loaded, cells were rinsed and maintained in NES for an additional 15 min at 37 °C to allow deesterification of the dye. Experiments were performed in NES or in a Ca2+-free solution containing in mM: NaCl, 140; KCl, 2.8; MgCl2, 4; glucose, 10; and Hepes−NaOH, 10; pH 7.35. PLTX and calcium blockers were gently applied to the bathing solution by loading appropriate volumes (≤5 μL) of concentrated solution into a 2 mL syringe connected to the microincubator chamber via a small tube. Aspiration to the syringe of 0.5 mL of incubation medium (out of 1 mL) followed by reintroduction of this mixture into the chamber yielded accurate and rapid (≤1 s) delivery and mixing of the agents. Preliminary control experiments were carried out to exclude any artifact due to solvent vehicles (Figure S1 in the Supporting

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RESULTS PLTX Toxicity in Skeletal Myotubes. Measurement of cell viability by SRB assay was performed after either 24 h of continuous exposure to PLTX or after short incubation (1, 2, or 4 h), followed by a recovery in toxin-free medium up to 24 h. After 24 h of incubation with increasing concentrations of PLTX from 0.004 to 9 nM, different effects on skeletal myotubes were detected at different concentrations. Starting from 0.01 nM PLTX, the spontaneous contractile activity of the cultured myotubes was completely abolished (data not shown), but no significant effects on cell viability were detected up to 0.1 nM PLTX. Concentrations higher than 0.1 nM PLTX caused a progressive decrease in cell viability with an EC50 = 0.54 ± 0.07 nM. Nearly 100% cell death was observed at 3 nM (Figure 1A). Exposure to PLTX triggered a toxic effect, which developed even after shorter incubation times (1, 2, or 4 h; Figure S2 in the Supporting Information). Moreover, 1, 2, or 4 h of exposure to 1 nM PLTX triggered a persistent toxic effect, which persisted even after removal of the compound. At the end of the recovery period, no statistically significant difference in cell viability was detected among cells exposed for short times (1, 2, and 4 h) and those exposed continuously for 24 h (Figure 1B). PLTX Effect on [Ca2+]i in Skeletal Myotubes. In skeletal myotubes, [Ca2+]i was measured using videoimaging. PLTX caused a significant biphasic increase in [Ca2+]i in all observed 1913



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Figure 1. Toxicity of PLTX on mouse skeletal muscle cells assessed using the SRB assay. (A) Cell survival after 24 h of exposure to increasing concentrations of PLTX. (B) Cell survival after 1, 2, or 4 h of exposure to 1 nM PLTX and recovery up to 24 h in toxin-free medium as compared to 24 h of continuous exposure.

Figure 2. Effect of ACh and PLTX on the [Ca2+]i of mouse skeletal muscle cells. (A) Each representative temporal plot was from a cell exposed first to 20 μM ACh and, after washout (trace interruptions), to 6 or 1 nM PLTX. At both concentrations, PLTX induced a biphasic [Ca2+]i increase. (B) Amplitude of the transient and the long-lasting phases of the [Ca2+]i increase at 6 and 1 nM PLTX.

cells. Analysis of the [Ca2+]i increase was carried out at 6 nM PLTX, the concentration that evoked the maximal Ca2+ response in terms of responsive cells,30 and at 1 nM, a submaximal concentration reducing cell survival. At both concentrations, changes in [Ca2+]i levels began with a transitory increase (transient phase) that usually peaked within 20 s and resembled the Ca2+ response elicited by 20 μM acetylcholine (ACh), the functional agonist of skeletal muscle cells, applied to the same cells before toxin incubation. The transient phase was followed by a slower increase that reached a plateau within 300 s and remained unchanged even 30 min after the toxin application (long-lasting phase; Figure 2A). After the onset of the second phase of the [Ca2+]i increase, the removal of the toxin from the external bathing solution did not reduce significantly the Ca2+ signal (data not shown; n = 22 cells). During the long-lasting phase, cells maintained the responsiveness to ACh. Indeed, 30 min after PLTX exposure, the percentage of cells responsive to 20 μM ACh was 100% as in untreated counterparts, but the amplitude of the AChinduced Ca2+ responses was significantly lower, indicating that PLTX-treated cells were still viable but functionally impaired (data not shown). The fluorescence increase reached during transient and longlasting phases were concentration dependent. The corresponding peaks were 79.2 ± 5.1 (n = 65 cells) and 84.9 ± 3.8% higher than the basal level measured before the toxin addition at 6 nM (n = 65 cells) and 57.2 ± 3.9 (n = 30 cells) and 58.3 ± 3.5% at 1 nM (n = 30 cells). At both concentrations, amplitudes of transient and long-lasting phase were similar, but the amount of [Ca2+]i mobilized in the cytoplasm during the second phase exceeded that elicited during the transient phase. Indeed, because of the longer duration, the area subtended by the longlasting phase was 60.5 ± 7.2-fold the area of the transient phase

at 6 nM PLTX (n = 65 cells) and 40.2 ± 3.6-fold at 1 nM PLTX (n = 30 cells; for further details, see Table S1 in the Supporting Information). Because the amplitude of the transient phase induced by 6 and 1 nM PLTX resembled that elicited by 20 μM ACh (see above), it results that the amount of Ca2+ mobilized by the toxin is significantly higher than that induced by the physiological neurotransmitter due the longlasting phase (Figure 2A and Table S1 in the Supporting Information). Contribution of Extracellular and Intracellular Ca2+ to the [Ca2+]i Increase. The role of extracellular Ca2+ influx was assessed by exposing cells to 6 nM PLTX, while bathed in a nominally Ca2+-free solution, where Ca2+ was replaced with Mg2+. This experimental condition significantly modified the pattern of Ca2+ response elicited by PLTX. In the Ca2+-free solution, while the transient phase of the [Ca2+]i increase was not altered (52.3 ± 4.3% higher than basal [Ca2+]i; n = 28 cells), the absence of extracellular Ca2+ almost abolished the long-lasting phase, whose amplitude was reduced to 11.9 ± 1.0% (n = 28 cells; Figure 3). The contribution of Ca2+ release from intracellular stores was investigated by applying 6 nM PLTX to cells preincubated with 1 μM cyclopiazonic acid (30 min at 37 °C), a well-known blocker of the sarcoplasmic Ca2+-ATPase pump.31 By blocking the pump, cyclopiazonic acid causes emptying of the intracellular Ca2+ reservoir and abolishes any possible contribution of Ca2+ release from intracellular stores. Preincubation with cyclopiazonic acid significantly decreased (more than 80%) the amplitude of the transient phase of the [Ca2+]i increase triggered by PLTX but did not decrease the amplitude of the 1914



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Figure 4. Contribution of extracellular and intracellular Ca2+ to PLTX toxicity. Cell survival measured after 30 min and 1 h of incubation with 1 nM PLTX in Ca2+-free solution or in the presence of 1 μM Cyc, as compared to control (*p < 0.05 in comparison to 1 nM PLTX).

In our experimental model (at least 5 days of cell differentiation), the prevalent class of voltage-dependent Ca2+ channels expressed by myotubes is the L type.32 For this reason, we studied the response of PLTX in the presence of Ver (20 μM).33 Ver decreased by nearly 60% the amplitude of the transient phase of the Ca2+ response elicited by 6 nM PLTX in comparison to controls. However, it did not affect the longlasting phase (n = 34 cells; compare Figure 5A,B and Figure 5F). The nonselective blockers of the voltage-dependent Ca2+ channels cadmium (500 μM) and lanthanum (100 μM) had similar effects, inhibiting the transient (n = 12 cells) but not the long-lasting phase (n = 12 cells; compare Figure 5A,C and Figure 5F). These results ruled out the possibility that voltageoperated Ca2+ channels other than L type are involved in the long-lasting phase of the [Ca2+]i increase. To test the role of the Na+/Ca2+ exchanger, cells were exposed to 6 nM PLTX in the presence of 5 μM KB-R7943, which prevents the reverse mode of the exchanger.34 As observed with Ver, KB-R7943 inhibited the transient phase by ∼30% (n = 42 cells) and did not alter the sustained phase of PLTX-induced Ca2+ response (Figure 5A vs D and Figure 5F). The long-lasting phase of the [Ca2+]i increase remained unchanged even when 6 nM PLTX was added to the cells in presence of both blockers (n = 46 cells; Figure 5A vs E and Figure 5F). Extracellular Ca2+ Influx through Voltage-Independent Ca 2+ Channels and Its Contribution to the Cytotoxicity. In skeletal muscle cells, Ca2+ influx from the extracellular compartment occurs also through store-operated and stretch-activated channels. Store-operated channels could be potentially activated by PLTX as a consequence of the store emptying induced by the toxin during the transient phase. Stretch-activated Ca2+ channels could be activated by the osmotic stress and swelling induced by PLTX due to action on the Na+/K+ pump.13,35 During Ca2+ imaging and cytotoxicity experiments, we observed that exposure to PLTX also caused a significant alteration of cell morphology under our experimental conditions. For a more precise morphological evaluation of the alteration of cellular shape induced by 6 nM PLTX, atomic force microscopy was used. The variation of displacement along the z-axis was recorded during the outward and return scan. Cells were scanned before toxin addition, as control (Figure 6A), and after toxin application to the external bathing solution (Figure

Figure 3. Contribution of extracellular Ca2+ influx and Ca2+ release from intracellular stores to the [Ca2+]i increase induced by PLTX. (A) Representative time−course plots of [Ca2+]i measured in mouse skeletal muscle cells after the addition of 6 nM PLTX in control, Ca2+free conditions and in the presence of 1 μM cyclopiazonic acid (Cyc). Trace interruption stands for 30 min. (B) Effect of the absence of extracellular Ca2+ and depletion of the intracellular Ca2+ stores on amplitude of the transient and the long-lasting phases of the [Ca2+]i increase (*p < 0.05 in comparison to 6 nM PLTX; n = 28 and 12 cells).

long-lasting phase (n = 12 cells; Figure 3). Taken together, these results indicate that the transient phase of the Ca2+ response induced by PLTX is mainly sustained by Ca2+ release from the intracellular compartments, while the long-lasting phase is due to the extracellular Ca2+ influx. Contribution of Extracellular and Intracellular Ca2+ to PLTX-Induced Cytotoxicity. The role of extracellular and intracellular Ca2+ in PLTX-induced cytotoxicity was measured using the SRB assay. This assay was performed after either 30 min or 1 h of incubation with 1 nM PLTX. Preincubation with cyclopiazonic acid did not counteract the cytotoxicity. Control experiments performed in the presence of cyclopiazonic acid alone excluded any effect per se. Thus, cytotoxicity seemed independent from the transient phase of [Ca2+]i increase sustained by Ca2+ release from intracellular stores. On the contrary, the cytotoxicity of PLTX was prevented in the Ca2+free solution, when the long-lasting phase of Ca2+ influx was abolished (Figure 4). Extracellular Ca2+ Influx through Voltage-Dependent Channels and Na+/Ca2+ Exchanger. We studied whether voltage-dependent channels and the Na+/Ca2+ exchanger (reverse mode) were involved in PLTX-induced long-lasting phase of the [Ca2+]i increase and cytotoxicity. 1915



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Figure 5. Contribution of L type calcium channel and Na+/Ca2+ exchanger (reverse mode) to the [Ca2+]i increase induced by PLTX. Representative time−course plots of [Ca2+]i after the addition of 6 nM PLTX in control conditions (A), in the presence of 20 μM Ver (Ver, B; n = 34 cells), 100 μM La3+ and 500 μM Cd2+ (C; n = 12), 5 μM KB-R7943 (D; n = 42 cells), or both (E; n = 46 cells). Trace interruption stands for 30 min. (F) Effect of the blockers, used alone or combined, on the amplitude of transient and long-lasting phases of the [Ca2+]i increase (*p < 0.05 in comparison to 6 nM PLTX).

the presence of Gd3+ alone excluded any effect of the ions per se on cell viability.

6B−F) up to 50 min postexposure. In cells that remained attached up to the end of the recording, experiments revealed a mean increase of cellular volume of 9.0 ± 3.4% (n = 5 cells), which we attribute to cell swelling. Moreover, in all of the cells analyzed, a concomitant loss of cell adhesion was detected approaching the end of acquisition as revealed by comparison between outward and return scan (data not shown). The contribution of store-operated and stretch-activated channels to the Ca2+ influx triggered by 6 nM PLTX was investigated using SKF-96365 or gadolinium ions (Gd3+), widely used as blockers of both voltage-independent channel types.36 Ca2+ imaging experiments showed that in cells preincubated with 20 μM SKF-96365 (30 min at 37 °C), the first transient of the Ca2+ response elicited by PLTX was significantly decreased (by ∼75%) as compared to controls. The long-lasting phase was reduced, but the effect of SKF96365 was not statistically significant (n = 18; Figure 7). Gd3+ did not alter the amplitude of the transient phase of the PLTXinduced response (n = 45 cells). However, Gd3+ markedly decreased the long-lasting phase of the [Ca2+]i increase (n = 45 cells; Figure 8). Removal of Gd3+ resulted in the restoration of the long-lasting phase induced by PLTX (n = 12 cells; Figure 9). To assess the effect of Gd3+ on PLTX toxicity, SRB assay was performed in the presence of the ion. In these experimental conditions, the cytotoxicity of PLTX was significantly reduced, with a 30% increase in cell survival after up to 1 h of exposure to 1 nM PLTX (Figure 10). Control experiments performed in

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DISCUSSION In this study, we investigated the relationship between PLTXinduced alteration of Ca2+ homeostasis and cytotoxicity on primary cultures of mouse skeletal muscle cells, a predictive in vitro model for the study of the actions of this marine toxin on one of its primary toxicological targets. PLTX reduced the viability of differentiated skeletal muscle cells with an EC50 = 0.54 nM after 24 h of continuous incubation. This value is in agreement with literature reports for the potency of this family of compounds on excitable cells.37 In addition, PLTX significantly reduced cell survival 24 h after toxin removal, even after exposures as short as 1−4 h, indicating that limited exposures to PLTX are sufficient to trigger irreversible toxic effects in skeletal muscle cells. To support this hypothesis, it is striking that [Ca2+]i increase induced by PLTX persisted even after the removal of the toxin from the medium. All together, our data suggest that once PLTX comes into contact with skeletal muscle cells, it causes irreversible toxicity, possibly through massive [Ca2+]i increase. These results at the cellular level further support the view that skeletal muscle tissue is likely to be an important target for PLTX, as already hypothesized on the basis of in vivo studies and epidemiological data.1,9,10 Exposure of skeletal muscle cells to PLTX causes a profound alteration of Ca2+ homeostasis. Ca2+ imaging experiments, carried out at fast-time resolution (1 ratio image/s), for up to 1916



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Figure 7. Contribution of voltage-activated Ca2+-channels to the [Ca2+]i increase. (A) Representative temporal plot of [Ca2+]i after addition of 6 nM PLTX in control conditions or in the presence of 20 μM SKF-96365. Trace interruption stands for 30 min. (B) Effect of SKF-96365 on the two phases of the PLTX-induced [Ca2+]i increase (*p < 0.05 in comparison to 6 nM PLTX; n = 18 cells).

Figure 6. Atomic force microscopy of a skeletal muscle cell before and after PLTX action. Representative three-dimensional images of a single skeletal muscle cell under control conditions (A) and after addition of 6 nM PLTX (complete image acquisition every 10 min, B−F). Axes: x, 0−100 μm; y, 0−100 μm; and z, 0−6 μm.

According to the currently accepted molecular mechanism of PLTX action in excitable cells, the interaction of the toxin with the Na+/K+ pump and the consequent massive influx of Na+ and membrane depolarisation causes opening of the voltagedependent channels and activation of the reverse mode of the Na+/Ca2+ exchanger.13,41 In view of this, the contribution of voltage-dependent Ca2+ channels and the Na+/Ca2+ exchanger in PLTX-induced intracellular calcium rise was evaluated. Blockers of voltage-dependent Ca2+ channels (Ver, cadmium, and lanthanum) and the Na+/Ca2+ exchanger in the reverse mode (KB-R7943), even combined, inhibited only the transient phase of the PLTX-induced Ca2+ response. These results preclude any role for these two mechanisms of Ca2+ influx in the long-lasting increase of [Ca2+]i and in PLTX cytotoxicity and thus are in agreement with the lack of protective effect of Ver on excitable cells.23,37,42 PLTX is known to cause alteration of the cytoskeleton in several cell models43 and to trigger morphological modifications as a consequence of the ionic imbalance following its interaction with the Na+/K+ pump.13,44 Taking this as a starting point, we described, for the first time on skeletal muscle cells, the cellular morphology changes induced by PLTX using AFM microscopy. Progressive cell swelling, recorded up to 50 min following PLTX exposure, led to a 9% increase of cell volume when compared to pre-exposure measurement. This increase was functionally significant as demonstrated by the concomitant loss of cell adhesion detectable comparing outward and return scans. We also investigated the contribution of storeoperated and stretch-activated channels on [Ca2+]i increase,

30 min after toxin exposure, revealed that PLTX triggers a peculiar biphasic [Ca2+]i increase in skeletal muscle cells, as we partially described previously.30 A first short-lasting increase of the [Ca2+]i (“transient phase”) was followed by a second persistent [Ca2+]i increase, which remained stable up to 30 min after toxin exposure (“long-lasting phase”). Our results demonstrate that during the long-lasting phase, the amount of Ca2+ mobilized is significantly higher than that mobilized by the physiologic agonist ACh. This suggests that the functional relevance of the Ca2+ response induced by PLTX is in the nanomolar range, and furthermore, it explains the cytotoxic effect of PLTX on this cell type. Indeed, it has been demonstrated that Ca2+ plays a crucial role in the mechanism of action of PLTX in muscle tissues.20,25,38,39 Accordingly, in our cell model, exposure to the toxin in an extracellular Ca2+free solution prevented the long-lasting [Ca2+]i. increase, as well as cytotoxicity. It is important to note that the absence of cytotoxicity under Ca2+-free condition cannot be attributed to the altered binding of PLTX because: (i) replacing Ca2+ with Mg2+ maintains the ionic strength of the milieu40 and (ii) the role of Ca2+ as cofactor for the PLTX binding to its molecular target is guaranteed by Mg2+.16,18 On the other hand, depletion of intracellular calcium stores with cyclopiazonic acid did not reduce the long-lasting phase and did not prevent PLTXinduced cytotoxicity, indicating that only extracellular Ca2+ and not Ca2+ released from intracellular stores is involved in the cytotoxic mechanism of PLTX. 1917



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Figure 9. Contribution of stretch-activated Ca2+ channels to the [Ca2+]i increase induced by PLTX. (A) Representative temporal plot of [Ca2+]i after addition of 6 nM PLTX in control conditions, in the presence of 100 μM Gd3+ and after washout. Trace interruption represents washout. (B) PLTX-induced [Ca2+]i increase in the presence of Gd3+ and after its washout (*p < 0.05 in comparison to 6 nM PLTX; n = 12 cells).

Figure 8. Contribution of stretch-activated Ca2+ channels to the [Ca2+]i increase. (A) Representative time course of [Ca2+]i after addition of 6 nM PLTX under control conditions or in the presence of 100 μM Gd3+. Trace interruption stands for 30 min. (B) Effect of Gd3+ on the two phases of the PLTX-induced [Ca2+]i increase (*p < 0.05 in comparison to 6 nM PLTX; n = 45 cells).

using SKF-96365 and Gd3+. SKF-96365 and Gd3+ are extensively used to block store-operated and stretch-activated Ca2+ channels, respectively.45−48 We found that SKF-96365 did not affect the long-lasting phase elicited by PLTX, excluding the activation of the store-operated channels during this phase. In contrast, Gd3+ abolished the long-lasting phase of the [Ca2+]i increase, suggesting a possible role for stretch-activated channels in the mode of action of PLTX. It is worth noting that Gd3+, as a trivalent cation of the lanthanide family, is also known to inhibit voltage-dependent Ca2+ conductance in neural cells.49 In our cell system, voltage-dependent Ca2+ blockade seems unlikely. In fact, even at the concentration of 100 μM, commonly used to investigate stretch-activated channels activity, Gd3+ did not affect the transient phase as Ver and La3+/Cd2+ did. The selective inhibitory action of Gd3+ on the long-lasting phase was observed up to 3-fold higher concentrations (data not shown). In addition to the inhibition of the long-lasting phase, Gd3+ also significantly reduced, although not completely, the toxic effects of PLTX on skeletal muscle cells, suggesting a role for the stretch-activated channels in the chain of events culminating in cell death. The long-lasting activation of stretch-activated Ca2+ channels triggered by PLTX could be the basis of the long-lasting (up to 40 min) contractions observed in smooth muscle,23,25 as well as the development of symptoms associated with PLTX intoxication, such as muscle spasms and myalgia in humans1 and hind limb paralysis in mice.9,10 In conclusion, we confirmed that the increase of [Ca2+]i is crucial in determining PLTX-induced cytotoxicity and demon-

Figure 10. Contribution of stretch-activated Ca2+ channels to the cell toxicity. Cell survival measured after 30 min and 1 h of incubation with 1 nM PLTX in the presence of 100 μM Gd3+ as compared to controls (*p < 0.05 in comparison to 1 nM PLTX).

strated that, at least in skeletal muscle cells, cytotoxicity is strictly related to a long-lasting and irreversible Ca2+ influx possibly related to the cellular morphologic alteration induced by the toxin. We suggest that the osmotic swelling and cytoskeletal disorganization following PLTX-induced ionic imbalance could ultimately trigger stretch-activated Ca2+ channels, which we propose to be involved in PLTX-induced toxicity. This novel aspect of the mechanism of action of PLTX may account for the widespread toxicity of the molecule on both excitable and nonexcitable tissues and represents an intriguing starting point for further investigations of PLTX effects at both the cellular level and in vivo. 1918



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ASSOCIATED CONTENT

S Supporting Information *

Representative temporal plots of the effect of vehicles on the [Ca2+]i of mouse skeletal muscle cells, time dependency of PLTX toxicity measured by SRB assay, and mean values of areas subtended by the temporal plots of the [Ca2+]i transients induced by PLTX and ACh. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Tel: +39-040-5588835. Fax: +39-040-5583215. E-mail: [email protected]. Funding

This work was supported by the Italian Ministry of Education, University and Research (PRIN). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Dr. Emanuela Testai and Prof. Takeshi Yasumoto for precious suggestions and discussion. ABBREVIATIONS ACh, acetylcholine; AFM, atomic force microscope; Cyc, cyclopiazonic acid; DMEM, Dulbecco's modified Eagle's medium; EC50, effective concentration on the 50% of the treated cells; FCS, bovine fetal calf serum; fura-2 AM, fura-2 pentacetoxymethylester; [Ca2+]i, intracellular calcium concentration; NES, normal external solution; PLTX, palytoxin; SRB, sulforhodamine B; Ver, verapamil

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REFERENCES

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Reduces Palytoxin Toxicity in Primary Cultures of Skeletal Mus

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