The cytolytic and cytotoxic activities of palytoxin

Toxicon 57 (2011) 449–459

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The cytolytic and cytotoxic activities of palytoxin Mirella Bellocci 1, *, Gian Luca Sala 1, Simone Prandi Dipartimento di Scienze Biomediche, Università di Modena e Reggio Emilia, Via G. Campi 287, I-41125 Modena, Italy

a r t i c l e i n f o

a b s t r a c t

Article history: Received 24 July 2010 Received in revised form 15 December 2010 Accepted 20 December 2010 Available online 29 December 2010

Palytoxin (PlTX) is one of the largest compound present in nature and, with its strong ability to modify the normal function of different biological systems, is also classified as one of the most potent biotoxins. Many alterations are triggered by PlTX, directly or indirectly related to its interaction with Naþ,Kþ-ATPase and the consequent conversion of this ion pump into a non-specific cation channel. The resulting perturbation of Naþ, Kþ, Ca2þ and Hþ ion fluxes is the driving force of PlTX-induced cytotoxic events, culminating with system disruption and, finally, cell death. The modifications in the distribution of these ions across the plasma membrane play key roles in the promotion of the PlTXinduced cytolytic and cytotoxic responses. In this scenario, PlTX-specific cytolysis can be part, but might not necessarily represent a unique aspect of the cytotoxic effects of the toxin. Owing to the complex array of responses, some of them being cell-type-specific and/ or affected by experimental conditions, the distinction between cytolytic and cytotoxic events becomes ill-defined, but the two responses show distinct features, whose further characterization could contribute to a better understanding of the molecular mechanism of cellular effects induced by PlTX. Ó 2010 Elsevier Ltd. All rights reserved.

Keywords: Palytoxin Cytolysis Cytotoxicity

Palytoxin (PlTX) is one of the largest natural substances known to date, displaying high toxicity in animals (Wiles et al., 1974). From a chemical perspective, PlTX was characterized for the first time by Moore in 1981 (Moore and Bartolini, 1981) and represents one of the largest polyether-type phycotoxin. The toxin is a non-peptide substance constituted by a long, partially unsaturated, chain of 129 carbon atoms which cooperate to the lipophilic nature of this compound. Moreover, the large number of hydroxyl, amine and amide groups as side substituents participate to the hydrophilic characteristics (Moore and Bartolini, 1981; Rossini and Hess, 2010; Uemura et al., 1981a; 1981b). These mix of hydrophilic and hydrophobic features might lead to micelles formation at high concentrations of the toxin in water, behaving in a soap-like manner, and cooperate to the high ability of

* Corresponding author. E-mail address: [email protected] (M. Bellocci). 1 These authors have equally contributed to this paper. 0041-0101/$ – see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.toxicon.2010.12.013

PlTX to interact with biological systems (Rossini and Hess, 2010). A study by Wiles et al. (1974) originally brought out the ability of PlTX to exert its toxic effect in a broad range of different animal species, including rats, guinea pigs, and rabbits. In particular this work highlighted the existence of a species-specific susceptibility to PlTX and an indication that its potency is strictly related to the administration route was obtained (see recent reviews by Deeds and Schwartz, 2010; Munday, 2011; Wang, 2008; Wu, 2009). The set of observation at organismal level were later expanded by more precise and detailed investigations, to clarify the mechanistic aspects of PlTX toxicity in several systems. The latter investigations thus showed the toxin’s ability to trigger a broad variety of effects, including the contraction of vascular smooth muscle and cardiac cells (Ito et al., 1976, 1977, 1979), as well as membrane depolarization (Castle and Strichartz, 1988; Dubois and Cohen, 1977; Kudo and Shibata, 1980; Muramatsu et al., 1984; Pichon, 1982; Sheridan et al., 2005; Tosteson et al., 1991).

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Cell death is frequently recorded in isolated cells exposed to PlTX (Bignami et al., 1992; Ledreux et al., 2009; Schilling et al., 2006), where the toxin induces responses displaying uncommon characteristics. In general, cell death results in rupture of plasma membrane and the release of cellular contents in vitro. This phenomenon is observed also in the case of apoptosis, as the possible intervention of macrophages for the removal of membrane-enclosed cellular remnants (Orrenius et al., 2003) may not occur when systems comprise only a single cell line in culture vessels. Cytolytic and cytotoxic responses to a toxicant, therefore, often represent complementary aspects of the same process in isolated cells, as cell lysis is a consequence of collapsed cell functioning and represents the outcome of a toxic effect. When PlTX acts on isolated cells, however, more subtle features become apparent, as cytolysis can be induced abruptly, and a description of the process might better fit the contention that cell death is the outcome of a cytolytic response. This issue was brought to our attention by some experiments reported in the original paper by Habermann et al. (1981), when cytolysis was induced in erythrocytes that had been exposed to PlTX, after toxin removal from incubation medium and cell re-suspension in an isotonic buffer in the absence of the toxin (see, for instance, Fig. 2 in Habermann et al., 1981). We exploited those observations in our investigations, studying the effect of PlTX on MCF-7 cells, cultured as mono-layers (Bellocci et al., 2008), and the time course of a typical cytolytic effect induced by 0.3 nM PlTX, as measured by the release of cytosolic lactate dehydrogenase (LDH) in incubation medium, is shown in Fig. 1. In the first part of cell incubation, a constant level of LDH activity was measured in our samples, independently of the presence or absence of PlTX, and it was due to the enzyme contributed by the serum added to the complete culture medium used for cell treatment. The cell exposure to 0.3 nM PlTX for such a short time in culture medium, therefore, was not sufficient, by itself, to induce cell lysis in MCF-7 cells. When the culture medium was removed, replaced by isotonic buffer devoid of PlTX, and the incubation of MCF-7 cells was continued, in turn, relevant changes in the levels of LDH were recorded in the buffer bathing the cells (Fig. 1). An abrupt increase in LDH activity, in fact, was detected in the fresh incubation buffer of cells that had been previously exposed to PlTX, and this increase progressed over time in the second part of the incubation in the absence of toxin. Background levels of LDH activity, in turn, were found in the fresh incubation buffer added to control cells in paired samples (Fig. 1). Under these experimental conditions, therefore, cytolysis was induced by the addition of an isotonic buffer to isolated cells, and the effect depended on prior cell exposure to PlTX. Thus, cell lysis apparently resulted from some catastrophic event, and cell death could be viewed as the outcome of cytolysis. The apparent distinction of cytolytic and cytotoxic responses induced by PlTX can be appreciated through a comparison of the behavior of different cell lines subjected to an identical experimental procedure, as reported

in Fig. 2. In these experiments, the responses of human epithelial cells (MCF-7) and neuroblastoma cells (SH–SY5Y) to increasing concentrations of PlTX are compared. The analysis was carried out measuring two different parameters under our experimental conditions. The protein content recovered from cell cultures was measured as an indicator of the amount of cellular material existing in the samples, whereas the LDH activity was assayed to evaluate the extent of cytolysis occurring in those samples. In these experiments, the procedure was that described in Fig. 1, and consisted of a first incubation of cells with increasing concentrations of PlTX, that was terminated by removal of incubation medium, its replacement by an isotonic buffer devoid of toxin, and the incubation was continued for an additional hour (Fig. 2). The measurement of the protein content of our samples was carried out at the end of the first incubation, whereas the assay of LDH activity was carried out at the end of the second incubation, using aliquots of the buffer bathing the cells. Under those conditions, increasing toxin concentrations caused a progressive decline in the cellular material remaining in culture dishes of SH–SY5Y cells at the end of the first incubation, and the levels of total protein were almost halved after treatment with 0.5 nM PlTX (Fig. 2). MCF-7 cell cultures, in turn, did not show the same effect, as the cellular material remaining in culture dishes was not decreased by increasing PlTX concentrations under these experimental conditions. The LDH activity measured in the incubation medium at the end of the second part of the incubation was found to be increased by PlTX treatment in a dose-dependent fashion in both cell lines (Fig. 2). In the case of SH–SY5Y cells, however, the response was biphasic, and LDH activity was found to decrease in the samples where the number of cells remaining in the dish after removal of culture medium had also decreased. Overall, these data showed that both cell lines could be induced to lyse by the addition of an isotonic buffer, if they had been previously exposed to effective concentrations of PlTX, but

Fig. 1. Cytolytic effect triggered by PlTX in MCF-7 cells. MCF-7 cells grown as monolayer in complete culture medium received 0.3 nM PlTX (black dots), or vehicle (empty dots), and were incubated for 1 h at 37  C. At the end of the incubation, the medium bathing the cells was removed and replaced with an isotonic buffered solution (20 mM phosphate buffer, pH 7.4, 0.15 M NaCl) devoid of toxin. The cell incubation was then continued for 1 h at 37  C. LDH activity was measured in aliquots of incubation media which were harvested at 15 min intervals during the full incubation period. The methods used in these experiments are detailed elsewhere (Bellocci et al., 2008).

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Fig. 2. Detection of cytolysis and cell death induced by PlTX in different cellular systems. MCF-7 and SH–SY5Y cell cultures in multi-well dishes were prepared. For each cell line, paired series of cultures were used. Cells were incubated with the indicated PlTX concentrations in complete culture medium for 1 h at 37  C. At the end of the incubation, the medium bathing the cells was removed and a first series of cell samples were used for the measurement of total protein in the materials remaining in culture wells (empty dots), by the procedure of Smith et al. (1985). The second series of cell samples, received an isotonic buffered solution (20 mM phosphate buffer, pH 7.4, 0.15 M NaCl) devoid of toxin, and the incubation was continued for 1 h at 37  C. At the end of this second incubation period, LDH activity was measured in aliquots of incubation media (black dots). The methods used in these experiments are detailed elsewhere (Bellocci et al., 2008).

a dose-dependent cytotoxic response was triggered by the toxin, and measurable cell death had already occurred at the end of the first hour of toxin treatment in SH–SY5Y, but not in MCF-7 cells (Fig. 2). Under those experimental conditions, MCF-7 cell death can be detected after PlTX exposure for longer times (Sala et al., 2009). The experiments described above, show the behavior of cellular systems exposed to PlTX, and outline the apparent distinction between cytolytic and cytotoxic responses induced by the toxin. Although such a distinction between cytolytic and cytotoxic action of PlTX must be considered only in conventional terms, being referred to a single possible fate of those biological systems, the responses are actually associated with distinct features, whose further characterization could contribute to a better understanding of the molecular mechanism of cellular effects induced by PlTX. In particular, the induction of cytolysis by an isotonic buffer is unexpected, as it may not be simply explained by the rupture of plasma membrane as a consequence of some osmotic stress in cells exposed to PlTX. Within this perspective, the discussion of existing information regarding PlTX effects will be approached in the following sections by distinguishing cytolytic and cytotoxic responses, to characterize the respective features, whenever possible, and discuss their possible relationships and/or differences. Owing to the conventional nature of this distinction, cytolytic responses will comprise the detection of cytosolic material released in the medium bathing the cells, as a means to evaluate toxic effects induced by PlTX, independently of the fact that cell lysis might or might not have been deliberately induced by an operator under controlled conditions (Bellocci et al., 2008). In turn, cytotoxic responses will comprise the findings obtained when toxic effects of PlTX have been evaluated on the basis of the dimension of cell populations, assessed by either direct methods (for instance, measuring total protein in cell samples, as in Fig. 2), or indirect parameters (such as MTT assays measuring the oxidative activity of living cells). The aim of our contribution is to review existing data on cell death responses induced by PlTX in isolated cells, with

particular attention onto the molecular mechanisms responsible for cytolysis. We will not review, therefore, the mechanism of action of PlTX on Naþ,Kþ-ATPase, and the effects exerted by the toxin on mitogen-activated protein kinases, as well as the cytoskeleton, which are described in other contributions (Louzao et al., 2011; Rossini and Bigiani, 2011; Wattenberg, 2011). 1. Cytotoxic effect The powerful toxic effect recorded in various animal species after PlTX treatment in vivo (Wiles et al., 1974) has been confirmed also by in vitro studies, whose results highlighted the ability of the toxin to induce modifications to normal cell functioning. In some of these studies (e.g., Ledreux et al., 2009; Schilling et al., 2006) it was found that induced alterations led to cell death. Initial investigations were performed on excitable cells of various origin, from muscles and nervous system, and in those cases PlTX could be distinguished from other toxins on the basis of the wide range of severe effects it exerted. Changes induced by the biomolecule were phenotipically attributable to contraction, arrhythmia and loss of ability to generate action potential (Amir et al., 1997; Ito et al., 1977; Ozaki et al., 1983; Sauviat, 1989; Weidmann, 1977). Over the years, results from independent studies have highlighted the key role that alterations of ion gradients (ions fluxes) induced by PlTX at plasma membrane level played in cytotoxic and cell death events. It is known that the interaction of the toxin with Naþ,Kþ-ATPase induces the conversion of the pump into a non-selective cation channel (Artigas and Gadsby, 2003; Habermann and Chhatwal,1982; Hilgemann, 2003). As a consequence of this primary event at the cell membrane level, a loss of cation selectivity is observed, resulting in the rapid and immediate release of Kþ in the extrcellular milieu, associated with the influx on Naþ from the outside to the intracellular compartment (Habermann et al, 1981; Redondo et al., 1996; Wu, 2009). The increase in the intracellular concentration of sodium ions is pivotal in the depolarization process of the plasma membrane and for a series of secondary effects triggered by

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the rising cation concentration (Dubois and Cohen, 1977; Frelin et al., 1990a; Habermann et al, 1989; Pichon, 1982; Schilling et al., 2006; Sheridan et al., 2005). For instance the sodium influx will affect other ion transporters coupled with Naþ gradient such as Naþ/Ca2þ and Naþ/Hþ exchangers (reviewed in Rossini and Bigiani, 2011). The increase in [Naþ]i was considered responsible for the resulting changes in the distribution of Ca2þ and Hþ between the intra- and extracellular environment by some groups (Frelin and Van Renterghem, 1995; Lang et al., 2005; Yoshizumi et al., 1991). Robust data about the molecular components involved in the process, however, are not available, yet. Owing to the importance of intracellular ion homeostasis for proper cell functioning, different secondary effects induced by PlTX could be mechanistically involved in cytotoxic responses, and will be considered in this section. In particular, we will discuss major effects exerted by PlTX in cellular systems with reference to changes in the ion fluxes involving calcium, sodium and potassium, as well as protons. 1.1. PlTX and sodium/potassium As mentioned, the interaction between PlTX and Naþ,KþATPase at the membrane level converts the pump into a non-selective ion channel (Hilgemann, 2003; Ikeda et al., 1988; Muramatsu et al., 1988; Rossini and Bigiani, 2011). By this primary event the movement of sodium and potassium across the plasma membrane is no longer actively regulated, but occurs following the ion concentration gradients and results in the extrusion of potassium and intrusion of sodium, that could rise about 5 times (Pérez Gómez et al., 2010). The compromised physiological Naþ/ Kþ equilibrium results in the perturbation of different ions balance like Ca2þ (see PlTX and calcium), Hþ (see PlTX and cellular acidification) and some anions (Pérez Gómez et al., 2010). As a consequence of these events, a series of modifications of cellular proteins occurs. In particular, it is known that the action of PlTX leads to the modulation of MAP kinase cascades, although the biochemical pathway of this mechanism remains to be clarified (Wattenberg, 2011). With regard to the effect on MAP kinases, it has been established that PlTX triggers the activation of the stressactivated protein kinase/c-Jun N-terminal kinases (JNK) in a dose-dependent manner in mouse 3T3 fibroblasts (Kuroki et al., 1996). The exposure of 3T3 fibroblasts to PlTX in the nanomolar range induced activation of JNK in a time-frame of 15 min, with a maximum at 45 min for the 0.1 nM concentration. By the use of a sodium-free medium, the direct link between Naþ influx and MAPK activation was established, and it was confirmed by the use of the sodium ionophore gramicidin, that could mimic the PlTX action on the JNK system (Kuroki et al.,1996). Similar results were also obtained by a 0.3 nM PlTX treatment for 30 min in different experimental systems, such as COS7 and HeLa cells (Kuroki et al., 1997). These observations were in agreement with those obtained in Rat-1 fibroblasts under the same experimental conditions (Iordanov and Magun, 1998). In this latter study, the use of culture media rich in potassium provided evidence of the involvement of Kþ efflux in JNK activation. In fact, the blockade of Kþ extrusion, by modification of the

concentration gradient across the plasma membrane, prevented JNK activation (Iordanov and Magun, 1998). PlTX is also able to promote the activation of another protein kinase, the p38 kinase, with similar kinetics in the same in vitro systems (Li and Wattenberg, 1998, 1999). Both enzymes are classified among MAP kinases, and are considered to participate to different signalling pathways (Chang and Karin, 2001; Chen et al., 2001; Robinson and Cobb, 1997; Waskiewicz and Cooper, 1995). More recently, we have found that 8 h of 0.03 nM PlTX exposure causes the accumulation of hsp 27 isoforms phosphorylated in Ser82 (Sala et al., 2009), and this finding is in line with the above-mentioned observations. In fact, it is known that hsp 27 is phosphorylated by mitogen-activated protein kinase-acivated protein kinase 2 (MAPKAPK2), whose activity depends on kinase phosphorylation by p38 protein kinase in vivo (Dorion and Landry, 2002). Interestingly, sodium influx and potassium efflux have the same importance in determining p38 and JNK activation (Iordanov and Magun, 1998), and the impairment of potassium electrochemical gradient upon PlTX action may then represent a key feature in the activation of enzymes involved in cellular death. In fact, potassium leak from the intracellular compartment is linked to activation of caspases and nucleases, which are normally kept inactive at physiological ion concentrations, preventing the disruptive consequences of their functioning in apoptosis (Hughes et al.,1997). PlTX effects on MAP kinase cascades have raised relevant interest from investigators, because these signal transduction pathways regulate different cellular processes in response to various extracellular stimuli (Kuroki et al., 1997; Li and Wattenberg, 1999). The cell survival/cell death equilibrium is of particular relevance, as it is strongly influenced by the activation state of some MAP kinase cascades and the subsequent accumulation of different kinase isoforms (Seger and Krebs, 1995; Wada and Penninger, 2004). In keeping with the preceding observations, PlTX has been shown to promote either cell survival or cell death, perhaps depending on the concentrations used in different experimental conditions. For instance, in a study aimed at probing the tumor promoting activity of PlTX, Miura et al. (2006) showed that picomolar toxin concentrations induced an increase in the relative cell growth (18–25%) in the two-stage transformation assay of Balb/c 3T3 cells. This observation could support the notion that low amounts of PlTX exert a mild cellular damage, which could be counteracted by cell survival mechanisms. Another possible explanation for the cell behavior in presence of low PlTX concentrations could be linked to JNK activation, in terms of extent and duration: in mouse 3T3 fibroblasts, nanomolar PlTX concentrations triggered a prolonged JNK activation, detectable 15 min after the beginning of cell exposure to the toxin (fold-change of 30 at 0.1 nM PlTX) (Kuroki et al., 1996). A different cellular behavior, in turn, has been observed with high PlTX concentrations, when the toxin triggers an extensive and massive cellular damage (Sheridan et al., 2005). Extensive membrane permeability modifications and subsequent ionic equilibria alterations are not likely to be fully counteracted by homeostatic mechanisms under those conditions. Thus, the JNK activation induced by high PlTX

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concentrations (equal or higher than 1 nM) (Kuroki et al., 1996) might switch cellular events towards cell death, rather than cell survival. The conversion of the Naþ, Kþ-ATPase into a non-selective channel is reflected on the balances of ions other than sodium and potassium, and could involve also anion equilibria. A recent study, focused onto the ability of PlTX to enhance the vulnerability of neurons to domoic acid provided evidence of the crucial role played by anions in this phenomenon (Pérez Gómez et al., 2010). In particular, an unspecific blocker of anion transporter (4,40 -diisothiocyanatostilbene-2,20 -disulfonic acid, DIDS) was able to totally prevent the potentiating effect of PlTX. The use of a second blocker more specific for the Cl/HCO 3 exchanger, in turn, could not inhibit PlTX action, demonstrating that other anion transporters could be involved (Pérez Gómez et al., 2010). 1.2. PlTX and calcium Bivalent calcium cation plays multiple roles as a chemical messenger. Under physiological conditions, and depending on the cellular district, the ion can be involved in a variety of processes, such as, for instance, contraction of muscle cells and nervous signal generation in neurons (Baylor and Hollingworth, 2010; Gover et al., 2009). Due to the crucial physiological roles of calcium, its cytosolic concentration is finely tuned by numerous molecular components. Under physiological conditions, a four-way balance between the calcium pools of the extracellular space, the cytosol, the mitochondria and the endoplasmic reticulum (RE) is maintained (Nicotera and Orrenius, 2006; Orrenius et al., 1989). Studies from several research groups have shown that PlTX determines the increase of [Ca2þ]i in many types of excitable cells (reviewed in Frelin and Van Renterghem, 1995; Pérez Gómez et al., 2010) as well as non-excitable systems (Amano et al., 1997; Monroe and Tashjian, 1995; Schilling et al., 2006; Wattenberg et al., 1989). The identification of the pathway/s responsible for this process is/are

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still uncertain, despite numerous studies have been conducted (Frelin et al., 1990b; Satoh et al., 2003; Sheridan et al., 2005). In some system the increase in intracellular Ca2þ has been related to ion influx from the extracellular environment, rather than its release from the endoplasmic reticulum (Satoh et al., 2003). Changes in calcium concentrations in one of the cellular districts has an impact on the overall ion balance inducing alterations in cell homeostasis (Berridge et al., 2000; Giorgi et al., 2008; Pinton et al., 2008; Rizzuto and Pozzan, 2003). Consequent responses are generated, according to the magnitude in the ion concentration sensed by its cellular targets. For instance, a rising in cytosolic calcium concentration leads to activation of calcium-dependent proteases that exert their function on a-actinin filaments, disrupting the actin based cytoskeleton, and on the actin binding proteins at membrane level compromising the anchor points of the cytoskeleton (McConkey and Orrenius, 1996; Nicotera and Orrenius, 2006; Orrenius et al., 1989). Furthermore, the activation of phospholipases (including phospholipases A2 and C) is another consequence of the [Ca2þ]i rising, resulting in increased hydrolysis of membrane phospholipids and the release of intracellular signalling mediators (Berridge et al., 2003; Giorgi et al., 2008; Jouaville et al., 1999; Nicotera and Orrenius, 2006; Orrenius et al., 1989, 2003; Schinder et al., 1996). Moreover, endonucleases responsible for DNA fragmentation are activated by increasing calcium concentrations at the nuclear level (Jones et al., 1989; Orrenius et al., 1989), and the production of ATP is altered by hyper- or depolarization of the mitochondrial membrane (Giorgi et al., 2008; Jouaville et al., 1999; Orrenius et al., 1989). Some of these phenomena correspond to those observed in different cell types after exposure to PlTX (Table 1). In particular, when rabbit enterocytes (Ares et al., 2005) were treated with 75 nM PlTX for 4 h, damages to the actin cytoskeleton were recorded, without obvious morphological changes of enterocytes. The measured

Table 1 Effects of intracellular calcium overload induced by PlTX treatment. PlTX-induced [Ca2þ]i overload effect

Cell type

References

Morphological changes

Rat aortic muscle Rabbit enterocytes Rat neurons

Sheridan et al., 2005 Ares et al., 2005 Pérez-Gómez et al., 2010

y n n

Protease activation and disruption of actin cytoskeleton

Rabbit enterocytes Human colon Human neuroblastoma

Ares et al., 2005 Valverde et al., 2008a Valverde et al., 2008b

y y y

Phospholipase activation and release of mediator

Mouse macrophages Rat brain Rat liver

Aizu et al., 1990 Habermann and Laux 1986 Lavine and Fujiki 1985

n y y

Endonuclease activation and DNA fragmentation

Human neuroblastoma

Valverde et al., 2008b

n

Muscle contraction

Rat muscle Rabbit aorta Guinea pig aorta Frog heart Porcine coronary artery

Amir et al., 1997 Ito et al., 1977 Ozaki et al., 1983 Sauviat, 1989 Ishii et al., 1997

y y y y y

Neurotransmitter release

Rat aorta Guinea pig vas deferens Rat pheochromocytoma

Nagase and Karaki 1987 Ishida et al., 1985 Tatsumi et al. 1984

y y y

Effects analyzed in different experimental systems are indicated with reference to their detection (y) or lack of detection (n).

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reduction of F-actin levels related to toxin exposure was found to be lower in Ca2þ free solution than in a medium containing Ca2þ, confirming the involvement of Ca2þ influx in this process in enterocyte response to PlTX. Similarly, Valverde et al. (2008a, 2008b), investigating PlTX effects on F-actin in intestinal (Caco2) (Valverde et al., 2008a) and neuronal (BE(2)-M17) cells (Valverde et al., 2008b), confirmed its action on actin based cytoskeleton (Table 1). In the same study they also evaluated the possible damage exerted by the toxin on DNA, that could be due to endonucluase activation coupled to [Ca2þ]i rising. DNA fragmentation or laddering of the nucleic acid were not detected, however, in cells exposed to PlTX concentrations as high as 100 nM (Table 1). On the one hand, some of these results are still uncertain and ambiguous as, for example, the action triggered by PlTX on cell morphology was found modified in rat aortic muscle cells (Sheridan et al., 2005), but not in rabbit enterocytes (Ares et al., 2005), indicating a possible cellspecific susceptibility (Table 1). Similar uncertainties are found with regard to PlTX effects on phospholipids and mediator release (Table 1). On the other hand, muscle contraction and neurotransmitter release were detected in different types of animal cells (Table 1). At present, neither the mechanisms through which PlTX induces the recorded increase in [Ca2þ]i, nor the role of this response in PlTX-induced effects on cellular functions, are fully clarified. Several studies are still ongoing, and further information on doses, time frame and molecular pathways involved in the alteration of intracellular calcium concentrations by PlTX’s action will help to better characterize and clarify the mechanism of its cytotoxic effects. Overall, the cellular responses to PlTX described above could contribute to an oncotic necrosis process triggered by an increase in intracellular Ca2þ concentrations (Giorgi et al., 2008), as a consequence of cell exposure to PlTX. 1.3. PlTX and cellular acidification Proper cellular functioning results from a fine tuning of a wide array of factors, among which ion equilibria play key roles. The loss of selectivity of Naþ,Kþ-ATPase pump after PlTX interaction was found to affect also the cytosolic pH (pHi) (Monroe and Tashjian, 1996). The pathways of Hþ entry into the cells affected by PlTX have not been identified and several mechanisms have been proposed in different studies (Frelin et al., 1990a; Monroe and Tashjian, 1996; Vale-González et al., 2007). In human osteoblast-like Saos-2 cells (Monroe and Tashjian, 1996) treatment with 8 nM PlTX-induced a decrease in pHi of about 0.2 units, as a consequence of PlTX-induced changes in [Naþ]i and [Ca2þ]i. The cytosolic acidification was related to a reduction of basal activity of Naþ/Hþ antiport and an incentive of Ca2þ/Hþ exchange through Ca2þ-ATPase in plasma membrane. The involvement of Ca2þ-ATPase in the pHi decrease could occur also in primary cultures of cerebellar granule calls (CGC) (Vale-González et al., 2007). Higher cytosolic acidification, corresponding to 0.6 units of pH reduction, was observed in the tested range of PlTX concentrations (10–50 nM). The intracellular acidification was detected

rapidly after PlTX addition, and was coupled to the [Ca2þ]i rise. The increase of [Naþ]i, other than [Ca2þ]i, was related to cytosolic acidification event recorded in chick cardiac cells exposed to nanomolar PlTX concentration (Frelin et al., 1990a). Sodium influx triggered by toxin action on Naþ,KþATPase would stimulate a reverse functioning of the Naþ/Hþ antiporter, thus inducing a pHi reduction. Although available data show a direct relationship between PlTX action and intracellular environment acidification, a full characterization of the mechanisms through which PlTX-induced acidification would affect cellular systems is still lacking. Biochemical evidence would suggest that intracellular pH reduction is involved in many processes that could drastically determine cellular fate. A quick cytoplasmatic acidification in neuronal cells was proposed to be responsible for programmed cell death, as it is associated to activation of endonucleases (Vincent et al., 1999), a phenomenon that accompanies apoptosis in many experimental systems (Ellis et al., 1991). More recently, other studies have approached this issue, to identify the causeeffect relationship between intracellular acidification and establish mechanisms involved in cell death. For instance, it has been shown that an increase in the proton concentration in the cytosol could lead to activation of hydrolases, such as IL-1b Converting Enzyme (ICE)-like protease in BAF3 cells (Furlong et al., 1997), as well as DNAse II in HL60 cells (Famulski et al., 1999). Furthermore, one cannot rule out the possible involvement of acidification in BAX translocation to mitochondria (Yang et al., 2008), that would favor cell death (Pinton et al., 2008; Pucci et al., 2008). Cellular processes set in motion by cytosol acidification would then result in many different responses, such as DNA degradation, activation of proteolitic enzymes and mitochondrial membrane depolarization, this latter as a result of the membrane interaction with BAX. Taken individually, these events can be viewed as part of the programmed cell death (apoptosis) cascade. Among all the consequences related to intracellular acidification event described above, the possible relation between PlTX acidification and some typical markers of apoptotic cell death has been investigated (Valverde et al., 2008a, 2008b). Chromatin condensation, DNA fragmentation and activation of caspases have not been recorded in neuroblastoma cell line after treatment with 100 nM PlTX for 20 h. Under the same conditions, Caco2 cells showed neither DNA degradation nor activation of caspases (Valverde et al., 2008a, 2008b). Hence, a scenario for PlTXinduced cell death that might not include apoptosis is conceivable. 2. Cytolytic effect The first evidence regarding cytolytic effects of PlTX was obtained by Habermann and his co-workers in 1981, studying the effects triggered by PlTX using non-excitable cells (Habermann et al., 1981). Habermann’s first observations were based on the hypothesis, subsequently proved erroneous, that the toxin was a pure haemolysin (Habermann et al., 1981). Accordingly, the experimental

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system they chose consisted in rat erythrocytes. This investigation then provided proof that PlTX is an extremely potent haemolysin, displaying a non-conventional mode of lytic action. The peculiarity of PlTX mode of action is related from one hand with the relatively long time needed to induce the lytic process that is recorded after a lag phase of one or more hours (Habermann et al., 1981). On the other hand, the lytic effect was found to be causally linked to the release of potassium ions (Kþ) in the external milieu (Habermann et al., 1981). This event was recorded in a time frame of minutes after erythrocyte exposure to the toxin and was completed before the hemolytic event took place, leading to the recognition that cell lysis is delayed with reference to loss of intracellular Kþ ions (Habermann et al., 1981). Since the lytic process depended on both the osmolarity of the solution bathing the cells and the intracellular ion composition, the process was defined as osmotic haemolysis (Ahnert-Hilger et al., 1982). In order to better characterize the PlTX-induced cytolytic process, and to elucidate the details concerning the influence of the composition of the cellular environment, numerous studies took place during the following decades. Useful information on these two aspects of the toxicity emerged from further investigations conducted by Habermann’s group, by using erythrocytes of different origins (Habermann et al., 1981), substances that alter the ion flux (Habermann and Chhatwal, 1982) and resealed ghosts (Chhatwal et al., 1983). The operative choice of monitoring the cytolytic effect in erythrocytes from various animal species was matched by the intracellular ion compositions of these cells, and it was found that the susceptibility of the system to the hemolytic action of PlTX was related to intracellular ion composition (Habermann et al., 1981). Most of the species susceptible to the lytic action of PlTX had intracellular sodium and potassium concentrations in the 103 M and 101 M ranges, respectively. In the case of cattle and sheep, in turn, the orders of magnitude of concentrations of the two monovalent ions were reversed, with Naþ as the main internal cation, and, conversely, the concentrations of Kþ ions were much lower (Habermann et al., 1981). In the case of rat, mouse and rabbit, the PlTX doses that caused half-maximal lytic effects (ED50) after a 4 h treatment were in the ng/ml range. Hemolysys of cattle and sheep erythrocytes, instead, required PlTX doses higher by three or more orders of magnitude (Habermann et al., 1981). These data were particularly interesting in the light of the fact that the erythrocytes from these two animals are particularly rich in sodium, showing high [Naþ]i, in the 102–101 M range (Habermann et al., 1981) Habermann and co-workers continued their studies attempting to clarify the key role ascribed to ionic fluxes in the cytolytic process induced by PlTX. In subsequent investigations, the attention was shifted from the evaluation of the hemolytic process as such, to the assessment of potassium release from erythrocytes, as an indicator of the ongoing process. Indeed, this choice was supported by the apparently indissoluble nature of cytolysis and potassium release from sensitive erythrocytes, that emerged in the case of PlTX-induced responses. Among the compounds that could affect ion fluxes at the cell membrane level,

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ouabain was found particularly effective in preventing the action of PlTX. More precisely, in these studies it was observed that ouabain inhibited the increased permeability to cations of erythrocytes treated with PlTX, in a dose- and time-dependent manner (Habermann and Chhatwal, 1982). Thus, the existence of some interference between the mechanisms of action of ouabain and PlTX was revealed, and the Naþ,Kþ-ATPase, the recognized target of ouabain action, could be proposed as the receptor of PlTX (Habermann and Chhatwal, 1982). In this scenario, a useful contribution arose from the study conducted in 1983 by Chhatwal et al. Resealed ghosts from erythrocytes were used in this investigation, representing a particular experimental system which made it possible to modify the intracellular ion composition. More precisely, erythrocytes were subjected to a mild lysis under controlled conditions, to lose their internal content, and the subsequent resealing of the cell membranes was obtained in a solution whose composition was tightly controlled by the operator, and would then represent the environment enclosed by the plasma membrane of resealed ghosts. Thus, it was possible to obtain an experimental system with a high degree of control that allowed to study the effects of PlTX in different sets of experimental conditions. In particular, the effect of ATP, Naþ and Kþ inside the ghosts, and the influence of ouabain in the medium bathing the cells were evaluated. From these analyses the capacity of PlTX to promote the free passage of Naþ and Kþ across plasma membrane following their concentration gradients was shown (Chhatwal et al., 1983). The outward movement of Kþ ions was favored by the presence of intracellular ATP under those conditions (Chhatwal et al., 1983). Finally, it was shown that ouabain treatment inhibited the ion exchange induced by PlTX involving monovalent cations (Chhatwal et al., 1983). This evidence confirmed the involvement of the Naþ,KþATPase in the toxic response of erythrocytes to PlTX treatment, and showed that the interaction between the toxin and the membrane protein was promoted by the availability of ATP and was inhibited by the presence of ouabain (Chhatwal et al., 1983; Habermann and Chhatwal, 1982). The confirmation that PlTX was not able to act as an ionophore but rather transformed the sodium pump into a non-selective pore permeable to small molecules (<0.36 nm) was equally important (Ahnert-Hilger et al., 1982). This last conclusion was reached using ghosts loaded with radio-labeled molecules of different sizes. The quantitation of these molecules in the extracellular medium, following cell treatment with PlTX, revealed that only molecules smaller than 180 Da were allowed to cross the membrane of the resealed ghosts. The results obtained in these studies proved particularly useful for planning subsequent investigations devoted to get further knowledge about the molecular target of PlTX and its mechanism of action (Rossini and Bigiani, 2011). Subsequent studies in non-excitable cells other than erythrocytes, and in excitable systems as well, have then provided support to the contention that the cytolytic effect of PlTX could be a common response induced by the toxin under in vitro conditions. Sheridan et al. (2005) and our group (Bellocci et al., 2008) have shown that PlTX cause

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plasma membrane lysis in smooth muscle cells from aorta (A7r5), as well as in human breast cancer cells (MCF-7), respectively. In both systems the lytic event was evaluated by monitoring the release of LDH into the media bathing the cells. If the treatment with the toxin was preceded by cell exposure to the Naþ,Kþ-ATPase inhibitor ouabain, a significant reduction of the effect of PlTX was recorded in both cases. Under these conditions, the release of LDH in the extracellular medium could be reduced by 50% or more. The cytolytic response of MCF-7 cells, as recorded in Fig. 1, is a consequence of cell exposure to PlTX, but is actually induced by the addition of an isotonic buffer to toxin-treated cells in the absence of PlTX, a condition stressing the complexity of a process which should then include osmolyte-dependent and independent components. 3. Conclusions and perspectives The results obtained over the years have provided increasing support to the identification of the Naþ,KþATPase as the main molecular target of PlTX (Artigas and Gadsby, 2003; Habermann and Chhatwal, 1982; Hilgemann, 2003). If the attention is focused onto the direct action that PlTX exerts on the Naþ/Kþ pump and on its conversion into a non-specific ion channel, the PlTX’s effect might be limited to the trans-membrane distribution of Naþ and Kþ ions. However, extending the view on processes triggered by the primary molecular event, a wide range of secondary effectors could be called into action by the toxin. Owing to the complex array of responses, some of which appear cell-specific, the distinction between cytolytic and cytotoxic events becomes conventional, and mostly related to the parameters measured by the operator. In particular the cytolytic response is detected when cytosolic material is found in the media bathing the cells, independently of the fact that cell lysis might or might not have been deliberately induced by an operator under controlled conditions (Bellocci et al., 2008; Sheridan et al., 2005). In turn, cytotoxic responses are detected when the size of the cell population is measured. The early observation of PlTX’s hemolytic effect (Habermann et al., 1981) led to recognition of the lytic event as an erythrocyte-specific response. The identification of PlTX as an atypic hemolysin, however, shifted the attention of scientists to its more general cytotoxic activity. The following years were then characterized by several studies aimed at obtaining a better characterization of cytotoxic responses induced by PlTX in experimental systems other than erythrocytes (e.g., Bignami et al., 1992; Habermann et al., 1989; Hori et al., 1988; Kuroki et al., 1996; Lauffer et al., 1985; Satoh et al., 2003). Thus cytolysis has been considered the end-point of a cascade of events that originates from the interaction between PlTX and Naþ/Kþ pump, and the effect could be extended to include many cellular systems. For instance, Sheridan et al. (2005) used the determination of the LDH released into culture medium as indicator of the final cytotoxic effect induced by PlTX in smooth muscle cells, following morphological changes of that biological system: “These alterations culminated in a loss of viability as indicated by marked increases in the release of lactate dehydrogenase”. The presence of this

cytosolic protein in the medium bathing the cells, in fact, is recognized as a marker of cytolysis, in keeping with the lack of distinction between PlTX-induced cytotoxicity and cytolysis. In fact, the two responses have been exploited without a net mechanistic distinction in most of the cellbased assays for the detection and quantification of PlTXgroup toxins developed so far (Bignami et al., 1992; Cañete and Diogène, 2008; Ledreux et al., 2009; Riobó et al., 2008). Subtle differences, however, can be found when the cellular responses to PlTX are analyzed with reference to cell death, representing a common end-point of its action in cell culture systems. This condition can be exemplified by the features of two distinct assays developed for a selective identification and quantification of PlTX in biological matrices: a cytolytic assay based on the use of cultured MCF-7 breast cancer cells (Bellocci et al., 2008), and a cytotoxic assay performed on Neuro-2a cells (Ledreux et al., 2009). Both methods can selectively detect PlTX, by inclusion of ouabain-treated samples, to check for the involvement of the Naþ,KþATPase in the recorded cell death responses. However, the two procedures can be distinguished according to the parameter used to monitor cell death, and the way cell death ensues in PlTX-treated cells. In the first assay, based on the use of MCF-7 cells, cell lysis is induced by an operator through a modification of the extracellular environment composition, after 1 h PlTX exposure (0–3 nM), and the extent of cell disruption is monitored by measuring LDH activity (Bellocci et al., 2008). In the Neuro-2a cellbased method, in turn, cell death is assessed by measuring cell viability, by monitoring the overall metabolic activity remaining in cell cultures (using the MTT assay), following their exposure to PlTX (Ledreux et al., 2009). The cell lysis induced by medium change (Fig. 1) in the MCF-7 cell-based assay was shown to be a PlTX-specific response, that is not induced by other cytolytic components under those experimental conditions (Bellocci et al., 2008). In turn, measurement of PlTX effect in the mouse neuroblastoma cell assay is based on evaluation of a general toxic effect, consisting in the reduction of cell viability (vs control), detected by the rate of oxidative metabolism in biological system (Ledreux et al., 2009). In this case, the selective detection of PlTX is ascertained by the introduction of the pretreatment with ouabain. Thus, cytolysis represents a PlTX-specific response that can be part, but might not necessarily represent a unique aspect, of its cytotoxic effects. The distinction between PlTX-induced cell lysis and cell death can be also appreciated by comparing the characteristics of the two responses in the same cell line under different experimental conditions (Bellocci et al., 2008; Sala et al., 2009) and in different cellular systems under identical experimental conditions (Fig. 2). It seems likely, therefore, that a detailed characterization of the molecular mechanism of PlTX-induced cell death might provide a better understanding of the modes of action of this toxin in animal systems. Over the years different and independent studies have been performed, to expand the knowledge concerning the cytotoxic effect of PlTX. By an overall evaluation of available information, the existence of a complex and still in complete landscape in PlTX action has been pointed out. On

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one hand some observations obtained in different systems, including both excitable and non-excitable cells show the ability of PlTX to induce programmed cell death. On the other hand, existing data would indicate that cell death consequent to PlTX treatment involves a different type of non-programmed cell death with many features of oncotic necrosis (Majno and Joris, 1995). Majno and Joris (1995) in “Apoptosis, Oncosis, and Necrosis: an overview of cell death” pointed out a list of characteristics associated with oncotic necrosis events. Putative markers of this kind of death included: swelling of cells and their organelles, an increase of membrane permeability related to impairment of ionic pumps, the alteration of ATP metabolism, blebbing of plasma membrane, vacuolization and protein denaturation. Comparing the body of knowledge collected during these years about oncotic necrosis and apoptosis, on the one hand, and the information gathered on PlTX-induced effects, such as cell rounding (Valverde et al., 2008a, 2008b) and swelling (Amir et al.,1997; Falciola et al.,1994; Mullin et al.,1991), cell lysis and loss of membrane integrity resulting in leakage of cytosolic enzymes (Bellocci et al., 2008; Sheridan et al., 2005) together with the lack of chromatin condensation, DNA fragmentation and activation of caspases (Valverde et al., 2008a, 2008b), the support to mechanisms of oncotic necrosis in PlTX-induced cell death would appear substantial. However, the studies on activation of JNK and p38 MAP kinase phosphorylation cascades (Li and Wattenberg, 1998), the accumulation of phosphorylated hsp 27 and oxidized isoforms of DJ1 cell stress protein (Sala et al., 2009), as well as the activation of proteases and nucleases (Famulski et al., 1999; Furlong et al.,1997), and the breakdown of mitochondrial membrane potential (Ledreux et al., 2009; Valverde et al., 2008a, 2008b) might provide support to the contention that some form of apoptosis might be induced by PlTX in some systems. If the findings collected during the years are put into perspective, they provide a contribution to the characterization of the complex toxicity of PlTX, although the picture seems far from being complete. Our working hypothesis regarding the molecular mechanisms of PlTX-induced cell death includes a first osmolyte-sensitive step triggered by the interaction between PlTX and Naþ,Kþ-ATPase, followed by a loss of normal cellular functions, which culminates with the destruction of the system and, finally, cell death. Fig. 3 summarizes our views and includes the major effects triggered by PlTX in biological systems. The interaction of the toxin with the Naþ/Kþ pump at the membrane level leads to a perturbation of cellular physiological conditions. The change of this pump into a non-selective ion channel allows the movement of Naþ and Kþ according to their concentrations gradients. The potassium efflux is accompanied by the cytosolic loading of sodium. The increased [Naþ]i is counterbalanced by the action of two different processes. On the one hand, the sodium is extruded from the cell by the Naþ/Hþ antiporter with a consequent intracellular acidification. On the other hand, the sodium is exchanged for calcium through the Naþ, Ca2þ exchanger. Moreover, membrane depolarization due to the rising in [Naþ]i would lead to Ca2þ entry into the cells through voltage gated calcium channels (VGCC) (Lopez

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Fig. 3. Palytoxin effects on major cellular functions in vitro. Normal cellular conditions (A), are perturbed after the interaction of PlTX with the Naþ,KþATPase on plasma membrane, converts the pump into a non-selective cation channel, resulting in extensive perturbations of intracellular ion homeostasis (B). This alteration causes changes in cellular structures and functions (C), and, after a lag phase sensitive to experimental conditions, cell destruction is observed (D). See the text for details.

et al., 1995). The alteration of ion distribution across the plasma membrane, consequent to PlTX action would be the driving force of secondary effects as, for instance, MAPK activation, cytoskeletal reorganization, changes in cell morphology and/or shape, as well as the alteration of the mitochondrial membrane potential (ΔJm). In the processes recorded in vitro, cellular conditions and experimental parameters play key roles in the cellular fate of the biological system after PlTX treatment. Thus, the identification of the mechanistic links among the series of events triggered by PlTX, and further studies filling existing knowledge gaps will be essential for a full understanding of the molecular mechanisms of PlTX-induced cell death and its toxicity pathway.

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Conflict of interest None. Acknowledgements We greatly thank Prof. Gian Paolo Rossini for insightful discussions and all his support. Our investigations on palytoxin effects have been supported by the Italian MIUR (Grant 2007FXSCL2). References Ahnert-Hilger, G., Chhatwal, G.S., Hessler, H.J., Habermann, E., 1982. Changes in erythrocyte permeability due to palytoxin as compared to amphotericin B. Biochim. Biophys. Acta 688 (2), 486–494. Aizu, E., Yamamoto, S., Nakadate, T., Kato, R., 1990. Differential effects of various skin tumor-promoting agents on prostaglandin E2 release from primary cultures of mouse epidermal cells. Eur. J. Pharmacol. 182 (1), 19–28. Amano, K., Sato, K., Hori, M., Ozaki, H., Karaki, H., 1997. Palytoxin-induced increase in endothelial Ca2þ concentration in the rabbit aortic valve. Naunyn Schmiedebergs Arch. Pharmacol. 355 (6), 751–758. Amir, I., Harris, J.B., Zar, M.A., 1997. The effect of palytoxin on neuromuscular junctions in the anococcygeus muscle of the rat. J. Neurocytol 26, 367–376. Ares, I.R., Louzao, M.C., Vieytes, M.R., Yasumoto, T., Botana, L.M., 2005. Actin cytoskeleton of rabbit intestinal cells is a target for potent marine phycotoxins. J. Exp. Biol. 208 (Pt 22), 4345–4354. Artigas, P., Gadsby, D.C., 2003. Ion occlusion/deocclusion partial reactions in individual palytoxin-modified Na/K pumps. Ann. N. Y Acad. Sci. 986, 116–126. Baylor, S.M., Hollingworth, S., 2010. Calcium indicators and calcium signalling in skeletal muscle fibres during excitation-contraction coupling. Prog. Biophys. Mol. Biol.. doi:10.1016/j.pbiomolbio.2010.06.001. Bellocci, M., Ronzitti, G., Milandri, A., Melchiorre, N., Grillo, C., Poletti, R., Yasumoto, T., Rossini, G.P., 2008. A cytolytic assay for the measurement of palytoxin based on a cultured monolayer cell line. Anal. Biochem. 374, 48–55. Berridge, M.J., Lipp, P., Bootman, M.D., 2000. The versatility and universality of calcium signalling. Nat. Rev. Mol. Cell Biol. 1 (1), 11–21. Berridge, M.J., Bootman, M.D., RodericK, H.L., 2003. Calcium signalling: dynamics, homeostasis and remodelling. Nat. Rev. Mol. Cell Biol. 4 (7), 517–529. Bignami, G.S., Senter, P.D., Grothaus, P.G., Fischer, K.J., Humphreys, T., Wallace, P.M., 1992. N-(4’-hydroxyphenylacetyl)palytoxin: a palytoxin prodrug that can be activated by a monoclonal antibody-penicillin G amidase conjugate. Cancer Res. 52, 5759–5764. Cañete, E., Diogène, J., 2008. Comparative study of the use of neuroblastoma cells (Neuro-2a) and neuroblastomaxglioma hybrid cells (NG108-15) for the toxic effect quantification of marine toxins. Toxicon 52 (4), 541–550. Castle, N.A., Strichartz, G.R., 1988. Palytoxin induces a relatively nonselective cation permeability in frog sciatic nerve which can be inhibited by cardiac glycosides. Toxicon 26 (10), 941–951. Chang, L., Karin, M., 2001. Mammalian MAP kinase signalling cascades. Nature 410, 37–40. Chen, Z., Gibson, T.B., Robinson, F., Silvestro, L., Pearson, G., Xu, B., Wright, A., Vanderbilt, C., Cobb, M.H., 2001. MAP kinases. Chem. Rev. 101, 2449–2476. Chhatwal, G.S., Hessler, H.J., Habermann, E., 1983. The action of palytoxin on erythrocytes and resealed ghosts. Formation of small, nonselective pores linked with Naþ, Kþ-ATPase. Naunyn Schmiedebergs Arch. Pharmacol. 323 (3), 261–268. Deeds, J.R., Schwartz, M.D., 2010. Human risk associated with palytoxin exposure. Toxicon 56 (2), 150–162. Dorion, S., Landry, J., 2002. Activation of the mitogen-activated protein kinase pathways by heat shock. Cell Stress Chaperones 7, 200–206. Dubois, J.M., Cohen, J.B., 1977. Effect of palytoxin on membrane and potential and current of frog myelinated fibers. J. Pharmacol. Exp. Ther. 201 (1), 148–155. Ellis, R.E., Yuan, J.Y., Horvitz, H.R., 1991. Mechanisms and functions of cell death. Annu. Rev. Cell Biol. 7, 663–698. Falciola, J., Volet, B., Anner, R.M., Moosmayer, M., Lacotte, D., Anner, B.M., 1994. Role of cell membrane Na, K-ATPase for survival of human lymphocytes in vitro. Biosci. Rep. 14, 189–204.

Famulski, K.S., Macdonald, D., Paterson, M.C., Sikora, E.,1999. Activation of a low pH-dependent nuclease by apoptotic agents. Cell Death Differ. 6, 281–289. Frelin, C., Van Renterghem, C., 1995. Palytoxin. Recent electrophysiological and pharmacological evidence for several mechanisms of action. Gen. Pharmacol. 26 (1), 33–37 (Review). Frelin, C., Vigne, P., Breittmayer, J.P., 1990a. Palytoxin acidifies chick cardiac cells and activates the Naþ/Hþ antiporter. FEBS Lett. 264, 63–66. Frelin, C., Vigne, P., Breittmayer, J.P., 1990b. Mechanism of the cardiotoxic action of palytoxin. Mol. Pharmacol. 38 (6), 904–909. Furlong, I.J., Ascaso, R., Lopez Rivas, A., Collins, M.K., 1997. Intracellular acidification induces apoptosis by stimulating ICE-like protease activity. J. Cell. Sci. 110, 653–661. Giorgi, C., Romagnoli, A., Pinton, P., Rizzuto, R., 2008. Ca2þ signaling, mitochondria and cell death. Curr. Mol. Med. 8 (2), 119–130. Gover, T.D., Moreira, T.H., Weinreich, D., 2009. Role of calcium in regulating primary sensory neuronal excitability. Handb Exp. Pharmacol., 563–587. Habermann, E., Ahnert-Hilger, G., Chhatwal, G.S., Beress, L., 1981. Delayed haemolytic action of palytoxin. General characteristics. Biochim. Biophys. Acta 649, 481–486. Habermann, E., Chhatwal, G.S., 1982. Ouabain inhibits the increase due to palytoxin of cation permeability of erythrocytes. Naunyn Schmiedebergs Arch. Pharmacol. 319 (2), 101–107. Habermann, E., Hudel, M., Dauzenroth, M.E., 1989. Palytoxin promotes potassium outflow from erythrocytes, HeLa and bovine adrenomedullary cells through its interaction with Naþ, Kþ -ATPase. Toxicon 27, 419–430. Habermann, E., Laux, M., 1986. Depolarization increases inositolphosphate production in a particulate preparation from rat brain. NaunynSchmiedebergs Arch. Pharmacol. 334 (1), 1–9. Hilgemann, D.W., 2003. From a pump to a pore: how palytoxin opens the gates. Proc. Natl. Acad. Sci. U S A 100 (2), 386–388. Hori, M., Shimizu, K., Nakajyo, S., Urakawa, N., 1988. Palytoxin-induced Kþ efflux from ileal longitudinal smooth muscle of the guinea-pig. Jpn. J. Pharmacol. 46, 285–292. Hughes Jr., F.M., Bortner, C.D., Purdy, G.D., Cidlowski, J.A., 1997. Intracellular Kþ suppresses the activation of apoptosis in lymphocytes. J. Biol. Chem. 272, 30567–30576. Ikeda, M., Mitani, K., Ito, K., 1988. Palytoxin induces a nonselective cation channel in single ventricular cells of rat. Naunyn-Schmeideberg’s Arch. Pharmacol 337, 591–593. Iordanov, M.S., Magun, B.E., 1998. Loss of cellular Kþ mimics ribotoxic stress. Inhibition of protein synthesis and activation of the stress kinases SEK1/ MKK4, stress-activated protein kinase/c-Jun NH2-terminal kinase 1, and p38/HOG1 by palytoxin. J. Biol. Chem. 273, 3528–3534. Ishii, K., Ito, K.M., Uemura, D., Ito, K., 1997. Possible mechanism of palytoxin-induced Caþþ mobilization in porcine coronary artery. J. Pharmacol. Exp. Ther 281 (3), 1077–1084. Ishida, Y., Kajiwara, A., Takagi, K., Ohizumi, Y., Shibata, S., 1985. Dual effect of ouabain on the palytoxin-induced contraction and norepinephrine releas in the guinea-pig vas deferens. J. Pharmac exp Ther. 232, 551–556. Ito, K., Karaki, H., Ishida, Y., Urakawa, N., Deguchi, T., 1976. Effects of palytoxin on isolated intestinal and vascular smooth muscles. Jpn. J. Pharmacol., 683–692. Ito, K., Karaki, H., Urakawa, N., 1977. The mode of contractile action of palytoxin on vascular smooth muscle. Eur. J. Pharmacol. 46 (1), 9–14. Ito, K., Karaki, H., Urakawa, N., 1979. Effects of palytoxin on mechanical and electrical activities of guinea pig papillary muscle. Jpn. J. Pharmacol. 29 (3), 467–476. Jones, D.P., McConkey, D.J., Nicotera, P., Orrenius, S., 1989. Calcium-activated DNA fragmentation in rat liver nuclei. J. Biol. Chem. 264 (11), 6398–6403. Jouaville, L.S., Pinton, P., Bastianutto, C., Rutter, G.A., Rizzuto, R., 1999. Regulation of mitochondrial ATP synthesis by calcium: evidence for a longterm metabolic priming. Proc. Natl. Acad. Sci. U S A 96 (24), 13807–13812. Kudo, Y., Shibata, S., 1980. The potent depolarizing action of palytoxin isolated from Palythoa tuberculosa on the isolated spinal cord of the frog. Br. J. Pharmacol. 71, 575–579. Kuroki, D.W., Bignami, G.S., Wattenberg, E.V., 1996. Activation of stressactivator protein kinase/c-Jun N-terminal kinase by the non-TPA-type tumor promoter palytoxin. Cancer Res. 56 (3), 637–644. Kuroki, D.W., Minden, A., Sánchez, I., Wattenberg, E.V., 1997. Regulation of a c-Jun amino-terminal kinase/stress-activated protein kinase cascade by a sodium-dependent signal transduction pathway. J. Biol. Chem. 272 (38), 23905–23911. Lang, F., Föller, M., Lang, K.S., Lang, P.A., Ritter, M., Gulbins, E., Vereninov, A., Huber, S.M., 2005. Ion channels in cell proliferation and apoptotic cell death. J. Membr. Biol. 205 (3), 147–157. Lauffer, L., Stengelin, S., Béress, L., Hucho, F., 1985. Palytoxin-induced permeability changes in excitable membranes. Biochim. Biophys. Acta 818, 55–60.

M. Bellocci et al. / Toxicon 57 (2011) 449–459 Ledreux, A., Krys, S., Bernard, C., 2009. Suitability of the Neuro-2a cell line for the detection of palytoxin and analogues (neurotoxic phycotoxins). Toxicon 53, 300–308. Levine, L., Fujiki, H., 1985. Stimulation of arachidonic acid metabolism by different types of tumor promoters. Carcinogenesis 6 (11), 1631–1634. Li, S., Wattenberg, E.V., 1998. Differential activation of mitogen-activated protein kinases by palytoxin and ouabain, two ligands for the Naþ, Kþ-ATPase. Toxicol. Appl. Pharmacol. 151, 377–384. Li, S., Wattenberg, E.V., 1999. Cell-type-specific activation of p38 protein kinase cascades by the novel tumor promoter palytoxin. Toxicol. Appl. Pharmacol. 160 (2), 109–119. Lopez, M.G., Artalejo, A.R., Garcia, A.G., Neher, E., Garcia-Sancho, J., 1995. Veratridine-induced oscillations of cytosolic calcium and membrane potential in bovine chromaffin cells. J. Physiol. 482 (1), 15–27. Louzao, M.C., Ares, I.R., Cagide, E., Espiña, B., Vilariño, N., Alfonso, A., Vieytes, M.R., Botana, L.M., 2011. Palytoxins and cytoskeleton: an overview. Toxicon 57 (3), 460–469. Majno, G., Joris, I., 1995. Apoptosis, oncosis, and necrosis. An overview of cell death. Am. J. Pathol. 146, 3–15. McConkey, D., Orrenius, S., 1996. The role of calcium in the regulation of apoptosis. J. Leukoc. Biol. 59, 775–783. Miura, D., Kobayashi, M., Kakiuchi, S., Kasahara, Y., Kondo, S., 2006. Enhancement of transformed foci and induction of prostaglandins in Balb/c 3T3 cells by palytoxin: in vitro model reproduces carcinogenic responses in animal models regarding the inhibitory effect of indomethacin and reversal of indomethacin’s effect by exogenous prostaglandins. Toxicol. Sci. 89, 154–163. Monroe, J.J., Tashjian Jr., A.H., 1995. Actions of palytoxin on Naþ and Ca2þ homeostasis in human osteoblast-like Saos-2 cells. Am. J. Physiol. 269 (3 Pt 1), C582–C589. Monroe, J.J., Tashjian Jr., A.H., 1996. Palytoxin modulates cytosolic pH in human osteoblast-like Saos-2 cells via an interaction with Na(þ)-K (þ)-ATPase. Am. J. Physiol. 270, C1277–C1283. Moore, R.E., Bartolini, G., 1981. Structure of palytoxin. J. Am. Chem. Soc. 103, 2491–2494. Mullin, J.M., Snock, K.V., McGinn, M.T., 1991. Effects of apical vs. basolateral palytoxin on LLC-PK1 renal epithelia. Am. J. Physiol. 260, C1201–C1211. Munday, R., 2011. Palytoxin toxicology: animal studies. Toxicon 57 (3), 470–477. Muramatsu, I., Uemura, D., Fujiwara, M., Narahashi, T., 1984. Characteristics of palytoxin-induced depolarization in squid axons. J. Pharmacol. Exper. Ther. 231, 488–494. Muramatsu, I., Nishio, M., Kigoshi, S., Uemura, D., 1988. Single ionic channels induced by palytoxin in guinea-pig ventricular myocytes. Br. J. Pharmacol. 93, 811–816. Nagase, H., Karaki, H., 1987. Palytoxin-induced contraction and release of prostaglandins and norepinephrine in the aorta. J. Pharmacol. Exp. Ther. 242 (3), 1120–1125. Nicotera, P., Orrenius, S., 2006. Ca2þ and cell death. Ann. New York Acad. Sci. 648, 17–27. Neurotoxins and Neurodegenerative Disease. Orrenius, S., McConkey, D.J., Bellomo, G., Nicotera, P., 1989. Role of Ca2þ in toxic cell killing. Trends Pharmacol. Sci. 10 (7), 281–285. Orrenius, S., Zhivotovsky, B., Nicotera, P., 2003. Regulation of cell death: the calcium-aptoptosis link. Nat. Rev. Mol. Cell Biol. 4 (7), 552–565. Ozaki, H., Tomono, J., Nagase, H., Urakawa, N., 1983. The mechanism of contractile action of palytoxin on vascular smooth muscle of guineapig aorta. Jpn. J. Pharmacol. 33 (6), 1155–1162. Pérez-Gómez, A., Novelli, A., Fernandez-Sanchez, T., 2010. Naþ/Kþ-ATPase inhibitor palytoxin enhances vulnerability of cultured cerebellar neurons to domoic acid via sodium-dependent machanisms. J. Neurochem. 114, 28–38. Pichon, Y., 1982. Effects of palytoxin on sodium and potassium permeabilities in unmyelinated axons. Toxicon 20 (1), 41–47. Pinton, P., Giorgi, C., Siviero, R., Zecchini, E., Rizzuto, R., 2008. Calcium and apoptosis: ER-mitochondria Ca2þ transfer in the control of apoptosis. Oncogene 27 (50), 6407–6418. Pucci, B., Bertani, F., Karpinich, N.O., Indelicato, M., Russo, M.A., Farber, J.L., Tafani, M., 2008. Detailing the role of Bax translocation, cytochrome c release, and perinuclear clustering of the mitochondria in the killing of HeLa cells by TNF. J. Cell Physiol. 217 (2), 442–449. Redondo, J., Fiedler, B., Scheiner-Bobis, G., 1996. Palytoxin-induced Naþ influx into yeast cells expressing the mammalian sodium pump is due to the formation of a channel within the enzyme. Mol. Pharmacol 49 (1), 49–57. Riobó, P., Paz, B., Franco, J.M., Vázquez, J.A., Murado, M.A., 2008. Proposal for a simple and sensitive haemolytic assay for palytoxin. Toxicological dynamics, kinetics, ouabain inhibition and thermal stability. Harmful Algae 7, 415–429.

459

Rizzuto, R., Pozzan, T., 2003. When calcium goes wrong: genetic alterations of a ubiquitous signaling route. Nat. Genet. 34 (2), 135–141. Robinson, M.J., Cobb, M.H., 1997. Mitogen-activated protein kinase pathways. Curr. Opin. Cell Biol. 9 (2), 180–186. Rossini, G.P., Bigiani, A., 2011. Palytoxin action on the Na+, K+-ATPase and the disruption of ion equilibria in biological systems. Toxicon 57 (3), 429–439. Rossini, G.P., Hess, P., 2010. Phycotoxins: chemistry, mechanisms of action and shellfish poisoning. EXS 100, 65–122. Sala, G.L., Bellocci, M., Rossini, G.P., 2009. The cytotoxic pathway triggered by palytoxin involves a change in the cellular pool of stress response proteins. Chem. Res. Toxicol. 22, 2009–2016. Satoh, E., Ishii, T., Nishimura, M., 2003. Palytoxin-induced increase in cytosolic-free Ca(2þ) in mouse spleen cells. Eur. J. Pharmacol. 465, 9–13. Sauviat, M.P., 1989. Effect of palytoxin on the calcium current and the mechanical activity of frog heart muscle. Br. J. Pharmacol. 98 (3), 773– 780. Seger, R., Krebs, E.C.,1995. The MAPK signaling cascade. FASEB J. 9, 726–735. Schilling, W.P., Snyder, D., Sinkins, W.G., Estacion, M., 2006. Palytoxininduced cell death cascade in bovine aortic endothelial cells. Am. J. Physiol. Cell Physiol. 291 (4), C657–C667. Schinder, A.F., Olson, E.C., Spitzer, N.C., Montal, M, 1996. Mitochondrial dysfunction is a primary event in glutamate neurotoxicity. J. Neurosci. 16 (19), 6125–6133. Sheridan, R.E., Deshpande, S.S., Adler, M., 2005. Cytotoxic actions of palytoxin on aortic smooth muscle cells in culture. J. Appl. Toxicol. 25, 365–373. Smith, P.K., Krohn, R.I., Hermanson, G.T., Mallia, A.K., Gartner, F.H., Provenzano, M.D., Fujimoto, E.K., Goeke, N.M., Olson, B.J., Klenk, D.C., 1985. Measurement of protein using bicinchoninic acid. Anal. Biochem. 150 (1), 76–85. Erratum in: Anal. Biochem., 1987 163(1), 279. Tatsumi, M., Takahashi, M., Ohizumi, Y., 1984. Mechanism of palytoxininduced [3H] norepinephrine release from a rat pheochromocytoma cell line. Molec Pharmac 25 (3), 379–383. Tosteson, M.T., Halperin, J.A., Kishi, Y., Tosteson, D.C., 1991. Palytoxin induces an increase in the cation conductance of red cells. J. Gen. Physiol. 98 (5), 969–985. Uemura, D., Ueda, K., Hirata, Y., Naoki, H., Iwashita, T., 1981a. Further studies on palytoxin. 1. Tetrahedron Lett. 22, 1909–1912. Uemura, D., Ueda, K., Hirata, Y., Naoki, H., Iwashita, T., 1981b. Further studies on palytoxin. 2. Structure of palytoxin. Tetrahedron Lett. 22, 2781–2784. Vale-González, C., Gómez-Limia, B., Vieytes, M.R., Botana, L.M., 2007. Effects of the marine phycotoxin palytoxin on neuronal pH in primary cultures of cerebellar granule cells. J. Neurosci. Res. 85 (1), 90–98. Valverde, I., Lago, J., Vieites, J.M., Cabado, A.G., 2008a. In vitro approaches to evaluate palytoxin-induced toxicity and cell death in intestinal cells. J. Appl. Toxicol. 28, 294–302. Valverde, I., Lago, J., Reboreda, A., Vieites, J.M., Cabado, A.G., 2008b. Characteristics of palytoxin-induced cytotoxicity in neuroblastoma cells. Toxicol. In Vitro 22, 1432–1439. Vincent, A.M., TenBroeke, M., Maiese, K., 1999. Neuronal intracellular pH directly mediates nitric oxide-induced programmed cell death. J. Neurobiol. 40 (2), 171–184. Wada, T., Penninger, J.M., 2004. Mitogen-activated protein kinases in apoptosis regulation. Oncogene 23, 2838–2849. Wang, D.Z., 2008. Neurotoxins from marine dinoflagellates: a brief review. Mar Drugs 6 (2), 349–371. Waskiewicz, A.J., Cooper, J.A., 1995. Mitogen and stress response pathways: MAP kinase cascades and phosphatase regulation in mammals and yeast. Curr. Opin. Cell Biol. 7 (6), 798–805. Wattenberg, E.V., 2011. Modulation of protein kinase signaling cascades by palytoxin. Toxicon 57 (3), 440–448. Wattenberg, E.V., McNeil, P.L., Fujiki, H., Rosner, M.R., 1989. Palytoxin down-modulates the epidermal growth factor receptor through a sodium-dependent pathway. J. Biol. Chem. 264 (1), 213–219. Weidmann, S., 1977. Effects of palytoxin on the electrical activity of dog and rabbit heart. Experientia 33, 1487–1489. Wiles, J.S., Vick, J.A., Christensen, M.K., 1974. Toxicological evaluation of palytoxin in several animal species. Toxicon 12 (4), 427–433. Wu, C.H., 2009. Palytoxin: membrane mechanisms of action. Toxicon 54 (8), 1183–1189. Yang, L., Mei, Y., Xie, Q., Han, X., Zhang, F., Gu, L., Zhang, Y., Chen, Y., Li, G., Gao, Z., 2008. Acidification induces Bax translocation to the mitochondria and promotes ultraviolet light-induced apoptosis. Cell. Mol. Biol. Lett. 13, 119–129. Yoshizumi, M., Houchi, H., Ishimura, Y., Masuda, Y., Morita, K., Oka, M., 1991. Mechanism of palytoxin-induced Naþ influx into cultured bovine adrenal chromaffin cells: possible involvement of Naþ/Hþ exchange system. Neurosci. Lett. 130, 103–106.