Comparative Toxicological Study of the Novel Protein Phosphatase Inhibitor 19-Epi-Okadaic Acid in Primary Cultures of Rat Cerebellar Cells

© The Author 2013. Published by Oxford University Press on behalf of the Society of Toxicology. All rights reserved. For permissions, please email: [email protected].

Comparative Toxicological Study of the Novel Protein Phosphatase Inhibitor 19-Epi-Okadaic Acid in Primary Cultures of Rat Cerebellar Cells

Maria-Teresa Fernández-Sánchez,*,1 David Cabrera-García,* Amaia Ferrero-Gutierrez,* Anabel Pérez-Gómez,* Patricia G. Cruz,† Antonio H. Daranas,† José J. Fernández,† Manuel Norte,† and Antonello Novelli*,‡

*Instituto Universitario de Biotecnología de Asturias, Departamento de Bioquímica y Biología Molecular, Universidad de Oviedo, Oviedo, Spain; †Instituto Universitario de Bio-Orgánica “Antonio González”, Universidad de la Laguna, Tenerife, Spain; and ‡Departamento de Psicología, Instituto Universitario de

Biotecnología de Asturias, Universidad de Oviedo, Oviedo, Spain

1To whom correspondence should be addressed at Departamento de Bioquímica y Biología Molecular, IUBA Edificio “Santiago Gascón”, Campus “El Cristo,” Universidad de Oviedo, 33006 Oviedo, Spain. Fax: +34 985103157. E-mail: [email protected].

Received October 22, 2012; accepted January 12, 2013

Okadaic acid (OKA) and analogues are frequent contaminants of coastal waters and seafood. Structure analysis of the isolated OKA analogue 19-epi-OKA showed important conformation differences expected to result in lower protein phosphatase (PP) inhibitory potencies than OKA. However, 19-epi-OKA and OKA inhibitory activities versus PP2A were unexpectedly found to be virtually equipotent. To investigate the toxicological relevance of these findings, we tested the effects of 19-epi-OKA on cultured cer-ebellar cells and compared them with those of OKA and its isomer dinophysistoxin-2. 19-epi-OKA caused degeneration of neurites and neuronal death with much lower potency than its conge-ners. The concentration of 19-epi-OKA that reduced after 24 h the maximum neuronal survival (EC5024) by 50% was ~300nM compared with ~2nM and ~8nM for OKA and dinophysistoxin-2, respectively. Exposure to 19-epi-OKA resulted also in less toxic-ity for cultured glial cells (EC5024,19-epi-OKA ~ 600nM; EC5024,OKA ~ 20nM). 19-epi-OKA induced apoptotic condensation and frag-mentation of chromatin, activation of caspases, and activation of ERK1/2 MAP kinases, features previously reported for OKA and dinophysistoxin-2. Also, differential sensitivity to 19-epi-OKA was observed between neuronal and glial cells, a specific characteristic shared by OKA and dinophysistoxin-2 but not by other toxins. Our results are consistent with 19-epi-OKA being included among the group of toxins of OKA and derivatives and support the suit-ability of cellular bioassays for the detection of these compounds.

Key Words: okadaic acid; diarrheic shellfish poisoning; neurotoxicity.

Diarrheic shellfish poisoning (DSP) is a severe human gas-trointestinal illness caused by the consumption of seafood contaminated with polyether toxins produced by dinoflagel-lates belonging to the genera Dinophysis or Prorocentrum (Murakami et  al., 1982; Yasumoto et  al., 1985). Okadaic acid (OKA), its isomer dinophysistoxin-2, and their methyl

homologue dinophysistoxin-1 are the main DSP toxins. In addition, many other compounds related to DSP toxins have been isolated (Domínguez et  al., 2010), and their biological and toxic activities remain in most cases undetermined. The widespread distribution of these toxins in seafood has caused increased concern due to the threat to public health and has underlined the need for suitable detection methods and toxico-logical studies to evaluate the potential risk for human health due to their presence in seafood.

OKA and its analogues share the capacity to bind and potently inhibit type 1 and type 2A ser/thr protein phosphatase (PP) activities (Bialojan and Takai, 1988), and many adverse effects of these toxins are considered to be due to this biologi-cal action. Based on this PP inhibitory activity, radioactive, colorimetric, and fluorometric functional assays for the quan-titative analysis of OKA and analogues in shellfish have been developed in the last years (See Dominguez et al., 2010 for a review; Holmes, 1991; Simon and Vermoux, 1994; Takai and Mieskes, 1991; Tubaro et al., 1996; Vieytes et al., 1997).

19-epi-okadaic acid (19-epi-OKA) is an OKA analogue isolated from cultures of Prorocentrum belizeanum (Paz et al., 2008). Structure and conformation in solution analysis of 19-epi-OKA (Cruz et al., 2007) revealed this new compound to be the C19-epimer of OKA (a planar conformation and a stereoview for 19-epi-OKA superimposed on the structure of OKA are shown in Supplementary fig. S1). Differences in structure of 19-epi-OKA with respect to OKA should result in different contact points between this new inhibitor and the protein compared with those found in the OKA-PP1/PP2A complexes and in lower inhibitory potencies of 19-epi-OKA than OKA. Consistently, 19-epi-OKA inhibited PP1 much less potently than OKA. However, the inhibitory activities of 19-epi-OKA and OKA versus PP2A were unexpectedly found to be virtually equipotent, making the interpretation of the relative

toxicological sciences 132(2), 409–418 2013doi:10.1093/toxsci/kft006Advance Access publication January 18, 2013

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inhibitory potencies of these compounds in structural terms difficult (Cruz et al., 2007). To test the toxicological relevance of these observations, we have studied the actions of 19-epi-OKA on primary cultures of cerebellar cells and compared the effects of this new inhibitor with other OKA derivatives, in particular with OKA and dinophysistoxin-2. Cerebellar cultures have previously proved very useful to investigate the biological action of marine toxins, including OKA derivatives. By using this experimental model, we demonstrated for the first time OKA to be a potent neurotoxin (Fernández et al., 1991, 1993) and to induce neuronal apoptosis (Fernández-Sánchez et  al., 1996). We have also used these cultures successfully to investigate both the toxic effects of dinophysistoxin-2 in neuronal and glial cells (Pérez-Gómez et al., 2004) and the effects of OKA on glial antioxidant enzyme activities (Ferrero-Gutiérrez et al., 2008). Here we show 19-epi-OKA to cause degeneration and death of cultured cerebellar cells through mechanisms similar to those reported for OKA and dinophysistoxin-2, albeit with much lower potency. The differences in neurotoxic potencies we report are consistent with the differences in structure and conformation found between these compounds and support the suitability of cellular bioassays versus enzymatic in vitro assays for the detection of these toxins.

MATERiAlS And METHOdS

Chemicals. GFAP and OX-42 antibodies were from Serotec (Oxford, UK). Okadaic acid methyl ester tetramethyl ether (OKA-TME, > 90% purity) was kindly provided by Dr. H. Fujiki (Saitama Cancer Center, Saitama, Japan). Highly pure dinophysistoxin-2 isolated from contaminated mussels was kindly provided by Dr. K. James (Cork Institute of Technology, Bishopstown, Cork, Ireland) (James et  al., 1999). Anti-phospho-p44/42 ERK was from Cell Signalling Technology, 91015, and anti-p42 ERK was from Santa Cruz Biotechnology, sc-154. All other chemicals were from Sigma.

Preparation of 19-epi-OKA. Cell cultures—A sample of 3 ml of a clonal culture of the dinoflagellate P. belizeanum containing approximately 7000 cell/ml was obtained from the IEO de Vigo collection by courtesy of Santiago Fraga. This sample was upscaled to perform large scale cultures in 80-l tanks contain-ing 40-l tanks of sea water enriched with Guillard K medium up to a final vol-ume of 1020 l. Cultures were incubated statically at 23ºC using 16:8 light:dark cycles for 3 weeks. Extraction and isolation—Due to the benthonic nature of the dinoflagellate P. belizeanum, most of the supernatant was easily separated, and finally cells were harvested by centrifugation at 3000 × g. Afterwards, cells were sonicated and extracted with acetone (4 × 1 l), and the resulting extract was filtered and concentrated to obtain 22.1 g of crude extract. The extract was subjected to successive chromatographies: first a gel filtration step using Sephadex LH-20 (Ø 6.5 cm × 60 cm) with a mixture of CHCl

3/MeOH/n-hexane

(1:1:2) yielding three fractions was carried out. The second fraction (8.11 g) was further fractionated using medium pressure reversed–phase LOBAR GRÖBE B (Ø 25 × 310 mm) LiChroprep columns eluted with MeOH:H

2O

(17:3). Finally, active fractions were subjected to HPLC purification on an XTerra (Ø 1.9 × 15 cm) column eluted with CH

3CN:H

2O (1:1). Finally, 1.0 mg

of 19-epi-OKA (> 90% purity) was obtained.

Cerebellar cell cultures. Primary cultures of rat cerebellar neurons were prepared as previously described (Novelli et al., 1988). Cytosine arabinoside (10µM) was added after 20–24 h of culture to inhibit the replication of non-neuronal cells. After 8 days in vitro, morphologically identifiable granule cells accounted for more than 95% of the neuronal population, the remaining 5%

being essentially GABAergic neurons. Astrocytes did not exceed 3% of the overall number of cells in culture. Cerebellar neurons were kept alive for more than 40 days in culture by replenishing the growth medium with glucose every 4 days and compensating for lost amounts of water due to evaporation. Mixed cerebellar cultures containing neurons and astrocytes were prepared as described (Suárez-Fernández et al., 1999). The presence of microglia in these cultures was determined by OX-42 immunostaining and did not exceed 2% of total cell popu-lation in both pure and mixed cultures and 5% in astroglial cultures.

Cell survival assessment. Cells were used between 14 and 20  days in culture. Toxins were added into the growth medium at the indicated concen-trations, and cell morphology and survival were assessed by observation of cultures under phase-contrast optics and by vital staining with fluorescein diac-etate and ethidium bromide (Fernández et al., 1991; Ferrero-Gutiérrez et al., 2008; Novelli et  al., 1988). Under fluorescent light, live neurons showed a bright green color both in the cell body and the neurites, whereas dead neurons did not retain any fluorescein and their nuclei appeared stained in red by eth-idium bromide. Live glial cells showed a bright green color and could be easily counted. For survival quantification purposes, all glial cells retaining fluores-cein were considered alive, independently of their morphology. Photographs of three randomly selected culture fields were taken and live and dead cells were counted. Total number of each type of cell per dish was calculated considering the ratio between the area of the dish and the area of the pictures (~3000).

DNA fragmentation analysis. Cells were lysed in 10mM Tris-HCl, 0.5% Triton X-100, 20mM EDTA, pH 7.4. After 20 min on ice, the lysate was cen-trifuged at 13,000 × g for 15 min at 4°C and treated with RNAse A (100 µg/ml at 37°C for 1 h). The supernatant containing degraded RNA and fragmented DNA, but not intact chromatin, was extracted with phenol chloroform. Nucleic acids were precipitated with 1 volume of ethanol and 300mM sodium acetate. Samples were electrophoresed in a 1.5% agarose gel and visualized by eth-idium bromide staining.

Assessment of nuclear morphology. Cells were labeled with Hoechst 33258 (5 µg/ml) for 15 min, washed in PBS, and fixed in 4% formaldehyde. Fixed cells were washed and viewed on an Olympus IMT-2 inverted research microscope using the filter for 340 nm.

Immunoblotting analysis. Neurons were washed thrice with ice-cold PBS and quickly harvested in a cell lysis buffer (Tris 50mM, pH 7, 2mM EDTA, 0.5mM EGTA, 0.5mM dithiothreitol, 1% phosphatase inhibitor cocktail, 1% protease inhibitor cocktail). Equal amounts of lysate proteins (50 µg/lane) were subjected to SDS-PAGE and transferred to PDVF membranes that were then incubated with anti-phospho-p44/42 and anti-p42 ERK antibodies. Membranes were rinsed with TBS-T (50mM Tris-HCl, pH 7.6, 150mM NaCl, and 0.05% Tween 20) and incubated with horseradish peroxidase–conjugated anti-rabbit IgG (1:2000), and immunoreactivity was detected by the enhanced chemilu-minescence technique. Densitometric quantification of the immunoblot bands was performed from scanned images using NIH Image J (version 1.31 v) den-sitometry software.

Determination of caspase-3 activity. Caspase-3 activity was determined using a fluorimetric assay as described (Smith et al., 2003). After exposure to toxins, culture medium was aspirated, cells were washed twice with 1 ml PBS, and centrifuged at 1000 rpm for 5 min. Cell pellets were then resuspended in 1 ml lysis buffer containing 25mM HEPES, pH 7.5, 0.1%Tritón X-100, 5mM MgCl

2, 2mM DTT, 1.3mM EDTA, 1mM EGTA, 20µM leupeptin, and 0.15µM

pepstatin. After incubation on ice for 30 min, the lysates were centrifuged at 16,000 × g (25 min, 4ºC), and the supernatants were stored at −70ºC till use. Aliquots of 100 µg total protein were assayed in triplicate in caspase buffer (100mM HEPES, pH 7.5; 2mM DTT). The reaction was started in dark condi-tions with the addition of 40µM caspase-3 substrate acetyl-Asp-Glu-Val-Asp-7-amino-4-methylcoumarin (Ac-DEVD-AMC), and the reaction was followed at 37ºC for up to 24 h. Fluorescence was measured at excitation 360 nm and emission 460 nm in a spectrofluoromether (Cyofluor 2300). Data are presented as fluorescence arbitrary units per micrograms of protein. Total protein was determined by the Bradford method (Bradford, 1976).

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Data presentation and analysis. For statistical analysis, we have used the software InStat for Macintosh, version 2.01. An unpaired two-tailed one-way ANOVA or a Student t-test was used to identify overall significant differences. A Tuckey-Kramer multiple comparison test was used for selective compari-son of individual data groups. Homogeneity of variances was tested using a Bartlett’s test. Only significances relevant for the discussion of the data are indicated in each figure.

RESulTS

Exposure to 19-epi-OKA Causes Degeneration of Neurites and Neuronal Death With Much Lower Toxic Potency Than OKA and Dinophysistoxin-2

Dose-response experiments showed 19-epi-OKA to result in significant neurotoxicity at concentrations above 200nM (see Fig.  1). Similarly to OKA and dinophysistoxin-2 (Fernández et  al., 1991, 1993; Pérez-Gómez et  al., 2004), 19-epi-OKA neurotoxicity was a long-term process because neither cell death nor morphological signs of toxicity were observed in neurons exposed to the toxin for less than 15 h, and toxicity was characterized first by neurite weakness and fragmentation (15–18 h) and later (> 24 h) by swelling of cell bodies and cel-lular death (see Fig. 2). However, in comparison with OKA and dinophysistoxin-2, 19-epi-OKA showed a much lower toxic potency for cultured cerebellar neurons. Degeneration of the neuronal network started to become evident at 19-epi-OKA concentrations of approximately 200nM, whereas after expo-sure to 500nM 19-epi-OKA, cell bodies appeared rounded and swollen and all neurites were very weak or completely frag-mented (Fig.  2). The concentration of 19-epi-OKA that pro-duced a 50% reduction in maximum neuronal survival (EC50) after 24 h was estimated at approximately 300nM. It should be noted that despite the strong signs of toxicity observed in neu-rons exposed to 500nM 19-epi-OKA for 24 h (see Fig.  2), a

considerable number of cells (~60%) were still able to retain the vital dye fluorescein diacetate, and therefore, they were con-sidered alive under this criterion (data not shown, see Materials and Methods section). In parallel experiments, the EC50 for OKA and dinophysistoxin-2 was ~2nM and ~8nM respectively, whereas OKA-TME, a derivative lacking PP inhibitory activity (Nishiwaki et al., 1990), caused no morphological changes (not shown) nor reduction in neuronal survival at concentrations up to 1µM OKA-TME (Fig. 1; Fernández-Sánchez et al., 1996).

To rule out the possibility that the much lower neurotoxic potency observed for 19-epi-OKA was attributable to a lower uptake of this toxin into neurons compared with OKA, we did experiments in which extracellular medium (EM) from cul-tures treated for 24 h with 5nM OKA (EM-OKA) or 500nM 19-epi-OKA (EM-19-epi-OKA) was collected and used to replace medium in untreated sister cultures. Cultures receiving EM-OKA or EM-19-epi-OKA were then evaluated for toxicity 24 and 48 h after medium replacement, as an indirect meas-urement of the concentration of toxins not entering into the cells during the first round of exposure (24 h) and remaining in the EM (EM-OKA (1) and EM-19-epi-OKA (1) in Table 1). Medium exchange followed by evaluation of EM-induced tox-icity was repeated two more times, 48 and 72 h after the ini-tial exposure to the toxin, and the results obtained (EM-OKA (2), EM-19-epi-OKA (2) and EM-OKA (3), EM-19-epi-OKA (3), respectively) are summarized in Table 1. Extensive toxic-ity was observed after 24-h exposure to OKA or 19-epi-OKA. However, no toxicity was observed in cultures exposed for 24 h to EM-OKA or EM-19-epi-OKA after the first medium replacement experiment. Toxicity was observed at longer expo-sure times (48 h) in medium exchange experiments (1) and (2), but no significant differences could be observed in either case between neuronal survival in cultures exposed to EM-OKA

Fig. 1. Concentration-dependent neurotoxicity of 19-epi-OKA on cultured cerebellar neurons. Percentage of live neurons per dish (mean ± SD, n = 6–8) after exposure of cultures to the indicated concentrations (0.1–1000nM) of 19-epi-OKA for 24 h. Dose-response curves obtained for OKA and dinophysistoxin-2 (DTX-2) are also shown for comparison. OKA-TME, a derivative of OKA lacking PP inhibitory activity, was also used as a negative control.

19-Epi-OKA TOXICITY IN CULTURED CEREBELLAR CELLS 411

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or EM-19-epi-OKA (see Table  1). No toxicity was observed in cultures exposed for 24 or 48 h to EM-OKA or EM-19-epi-OKA after the third medium replacement (EM-OKA (3) and EM-19-epi-OKA (3) in Table 1).

To exclude the possible degradation of OKA and of 19-epi-OKA in the culture medium, we did the following experiments: solutions of OKA (5nM) and of 19-epi-OKA (500nM) were prepared in culture medium and then kept in the same incubator as cell cultures (5% CO

2, 37°C). Aliquots (2 ml) from these

solutions were taken after 0, 24, 48, and 72 h and used to replace medium in cultures. Toxicity was then evaluated after 24 and 48 h to determine possible changes in the toxicity of the compounds in the culture medium. No decrease/delay in neurotoxicity by either 19-epi-OKA or OKA maintained in the incubator for up to 72 h (neuronal survival

19-epi-OKA,72h  =  15 ± 7%; neuronal

survivalOKA,72h

 = 10 ± 7% live neurons) was observed compared with freshly made solutions of toxins (neuronal survival

19-epi-

OKA,0h = 12 ± 8% live neurons; neuronal survival

OKA,0h = 13 ± 5%),

demonstrating that no relevant degradation of 19-epi-OKA or OKA is taking place in the culture medium during a 72-h period.

Neurotoxicity by 19-epi-OKA Is Due to Gene Expression-Dependent Apoptosis and Involves Caspase Activity Activation

To test whether, similar to OKA and dinophysistoxin-2, neurotoxicity by 19-epi-OKA also involved the activation of biochemical pathways leading to apoptosis, we examined the DNA from cultured neurons exposed to 19-epi-OKA. Agarose gel electrophoresis of soluble DNA extracted from neurons treated with 19-epi-OKA revealed large DNA fragmentation characteristic of apoptotic cells (Fig.  3), resulting from cleavage of nuclear DNA in internucleosomal regions (Wyllie, 1980). Fragmentation of DNA was an early event in the death process. It was evident in DNA extracted from neurons that had been exposed to 19-epi-OKA for 13 h, whereas 24-h exposure was necessary to observe clear morphological signs of toxicity and the significant decrease in neuronal survival reported in Figure 1. Based on the intensity of ethidium bromide staining, DNA from neurons exposed to 500nM 19-epi-OKA for 18 h showed quantitatively similar fragmentation as DNA from neurons exposed for the same time to 5nM OKA or 25nM dinophysistoxin-2, whereas no soluble fragmented DNA was obtained from control neurons (Fig. 3A). Changes in the nuclear morphology of affected neurons, typical of occurrence of apoptosis, were also observed in cultures exposed to 19-epi-OKA. Staining with the DNA-binding fluorochrome Hoechst 33258 revealed a significantly higher number of apoptotic nuclei showing condensation and fragmentation of chromatin in 19-epi-OKA–treated cultures (38 ± 4%, n  =  6) compared with control cultures (8 ± 2%) (Fig.  3B). As apoptosis can generally be inhibited by the suppression of gene expression, we used the transcriptional inhibitor actinomycin D and the protein synthesis inhibitor cycloheximide to examine whether neuronal apoptosis by 19-epi-OKA required newly synthesized

Fig. 2. Time dependence of the neurotoxic effects of 19-epi-OKA. Phase contrast photomicrographs of neurons exposed to 200nM or 500nM 19-epi-OKA for the indicated times. Arrows indicate granulation and fragmentation of neuronal processes. Fragmentation of neurites was observed approximately 15–16 h upon addition of 19-epi-OKA and was almost complete after 24 h. Extensive degeneration of neuronal somas occurred after 24-h exposure to 19-epi-OKA. Results from a parallel experiment using 5nM OKA are also shown for comparison. Very similar effects were observed in neurons exposed to 500nM 19-epi-OKA and 5nM OKA at each time.

TAblE 1Evaluation of Toxicity by 19-epi-OKA and OKA Remaining in

the EM After 24 h in Medium Replacement Experiments Shows no differences between 19-epi-OKA and OKA neuronal uptake

Treatment

Neuronal survival (%)

Time

24 h 48 h

None 98 ± 2 91 ± 619-epi-OKA 11 ± 8* n.d.OKA 8 ± 5* n.d.EM-19-epi-OKA (1) 95 ± 3 20 ± 6*EM-OKA (1) 96 ± 4 24 ± 5*EM-19-epi-OKA (2) 96 ± 2 53 ± 7*EM-OKA (2) 97 ± 3 49 ± 8*EM-19-epi-OKA (3) 98 ± 1 92 ± 5EM-OKA (3) 97 ± 2 95 ± 5

Notes. Neurons were exposed to 19-epi-OKA (500nM) or OKA (5nM), and toxicity was determined after 24 h. EM from cultures exposed for 24 h to 19-epi-OKA or OKA was collected and used to replace medium in sister cultures, and toxicity by EM-OKA and EM-19-epi-OKA was determined 24 and 48 h after medium replacement (EM-19-epi-OKA(1) and EM-OKA(1)). Medium exchange followed by evaluation of EM-induced toxicity was repeated 48 h (EM-19-epi-OKA(2) and EM-OKA(2)) and 72 h (EM-19-epi-OKA(3) and EM-OKA (3)) after the initial exposure to each toxin. Data are the mean ± SD (n = 4–6).

*p < 0.05 versus none in the same column.

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proteins, as previously reported for OKA and dinophysistoxin-2 (Fernandez-Sánchez et  al., 1996; Pérez-Gómez et  al., 2004). The presence of actinomycin D (1  µg/ml) or cycloheximide (5 µg/ml) prevented both neurotoxicity and the appearance of apoptotic nuclei by 19-epi-OKA (see Table 2). To determine whether 19-epi-OKA–induced apoptosis was a caspase-dependent process, we measured the caspase activity in extracts from 19-epi-OKA–treated cells. Measurements were carried out by using the caspase-3 fluorogenic substrate Ac-DEVD-AMC and were compared to those found after neuronal treatment with OKA. As shown in Figure  4, extracts from both 19-epi-OKA– and OKA-treated neurons were able to

hydrolyze Ac-DEVD-AMC significantly. Caspase-3 activation by 19-epi-OKA became evident after 3-h exposure to the toxin, and maximum activation levels approaching a twofold increase compared with control cells were observed after 15 h. As for OKA, approximately a 3.5-fold increase in caspase-3 activity compared with controls was consistently found in neurons exposed to the toxin for 1 h, whereas activation levels similar to those observed for 19-epi-OKA occurred in neurons exposed to OKA for 3 h or longer. It should be noted that induction of caspase activity occurred long before the first slight signs of toxicity, affecting mostly neurite strength and integrity, were morphologically evident in neurons exposed to either 19-epi-OKA or OKA for ≥ 17 h (see Fig. 2), and no significant decrease in the number of surviving neurons was observed up to 20-h exposure. A second caspase-3 activation peak was consistently found in cultures exposed to 19-epi-OKA or OKA for 24 h. This treatment was associated to a significant decrease in neuronal survival of approximately 30–40% (see Fig. 1).

Effects of 19-epi-OKA on the Activation of Extracellular Signal-Regulated Protein Kinase1/2 Mitogen-Activated Protein Kinase

Exposure to 19-epi-OKA (500nM) caused a time-dependent increase in the phosphorylation of ERK 1/2, with a maximal of 5.5- ± 0.3-fold induction at 3 h that then declined to 3- ± 0.3-fold at 16 h (see Supplementary fig. S3). In parallel experi-ments with OKA, a time-dependent increase in ERK 1/2 phos-phorylation was also observed in neurons exposed to OKA (5nM). ERK1/2 phosphorylation induction by OKA peaked at 1 h with a 7.1- ± 0.2-fold increase was 5- ± 0.2-fold after 3-h exposure to the toxin and then declined until 2- ± 0.8-fold acti-vation at 16 h.

Fig. 3. 19-epi-OKA neurotoxicity involves DNA fragmentation and condensation and fragmentation of chromatin typical of apoptosis. (A) Agarose gel electrophoresis of soluble DNA revealed considerable DNA fragmentation in neurons exposed to 19-epi-OKA (500nM) for 13 h (lane 2) or 18 h (lane 3) but not in untreated neurons (lane 1). DNA fragmentation in neurons exposed for 18 h to okadaic acid (5nM, lane 4) and dinophysistoxin-2 (25nM, lane 5) is also shown. Sizes of the DNA molecular size markers (lane M) are indicated in base pairs (bp). (B) Percentage of apoptotic nuclei in control neurons and after exposure to 19-epi-OKA (500nM), OKA (5nM), or dinophysistoxin-2 (DTX-2, 25nM) for 18 h was determined from photomicrographs stained with the DNA-binding fluo-rochrome Hoechst 33258 in which nuclei in control cultures appeared large in size and weakly stained and apoptotic nuclei appeared smaller and brightly stained due to condensed chromatin. Values are the mean ± SD, n = 5. *Significantly different from control (p < 0.001).

TAblE 2 Protein Synthesis inhibitors inhibit Apoptotic neurotoxicity by

19-epi-OKA

Neuronal survival (%) Apoptotic nuclei (%)

Treatment None +AcD +CHX None +AcD +CHX

None 83 ± 12 82 ± 6 98 ± 1 5 ± 1 6 ± 2 8 ± 119-epi-OKA 32 ± 3** 88 ± 5* 87 ± 4* 48 ± 4** 8 ± 3* 11 ± 4*OKA 20 ± 5** 80 ± 2* 89 ± 1* 62 ± 4** 18 ± 2* 20 ± 3*DTX-2 30 ± 1** 85 ± 5* 90 ± 2* 52 ± 7** 9 ± 1* 12 ± 3*

Notes. Neurons were exposed to 19-epi-OKA (500nM), OKA (5nM), or dinophysistoxin-2 (DTX-2, 25nM) in the absence (None) or in the presence of the transcriptional inhibitor actinomycin D (AcD, 1 µg/ml) or the protein synthesis inhibitor cyclohexymide (CHX, 5 µg/ml). AcD and CHX were added 1 h before toxins. Percentage of apoptotic nuclei and surviving neurons were determined after 20- and 24-h exposure to each toxin, respectively. Each value represents the mean ± SD (n = 4).

*p < 0.005 versus same treatment in the absence of inhibitors. **p < 0.01 versus none in the same column.

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Differential Toxicity of 19-epi-OKA in Cultured Cerebellum Neurons and Astrocytes

We have previously reported DSP toxins OKA and dinophy-sistoxin-2 to induce differential toxicity in cultured cerebellum neurons and astrocytes (Fernández et al., 1991; Pérez-Gómez et  al., 2004), a feature that is not shared by other polyether toxins. To evaluate the selectivity of the neurotoxic effect of 19-epi-OKA, we used mixed neuroglial cultures containing neurons and astrocytes (see Materials and Methods section). In these cultures, nonneuronal cells, confirmed to be astrocytes by immunostaining with glial fibrillary acidic protein GFAP (not shown), accounted for about 10% of the overall number of cells in culture and could be easily distinguished from neurons and counted. Dose-response experiments for 19-epi-OKA and

OKA were performed in neuroglial cultures, and survival of neurons and glial cells was determined after different times of exposure to each toxin. The general pattern of glial toxicity was similar for both 19-epi-OKA and OKA, although with differ-ent dose responses, 19-epi-OKA showing in all cases a much lower potency than OKA (see Fig. 5). Morphological damage (not shown) by 19-epi-OKA or OKA was observed in these experiments much earlier in glial cells than in neurons although glial cell damage required significantly higher concentrations of the toxins. Thus, 7-h exposure to 19-epi-OKA (> 500nM) or OKA (> 20nM) caused marked morphological changes (not shown) and significant reduction in the survival (Fig. 5A) of glial cells present in mixed cultures, whereas morphology (not shown) and survival (Fig. 5B) of cocultured neurons remained

Fig. 4. 19-epi-OKA increases caspase-3 activity. Extracts from neurons treated with 19-epi-OKA (500nM) or OKA (5nM) for the indicated times were incu-bated with the caspase-3 fluorogenic substrate Ac-DEVD-AMC (DEVD-7-amino-4-methylcoumarine) for 24 h. Fluorescence from released AMC due to substrate fragmentation by caspase-3 was measured at λ

ex = 360 nm and at λ

em = 460 nm. Caspase-3 activity was determined in controls (vehicle-treated cells) at 0, 6, 15,

and 24 h with similar results. Values represent the mean ± SD of three independent experiments (n = 9). *p < 0.005 versus 0 h.

Fig. 5. 19-epi-OKA induced differential toxicity in cultured cerebellum neurons and glial cells. (A and B) Dose-response experiments for toxicity of neurons and glial cells in mixed neuroglial cultures after exposure to 19-epi-OKA. Percentages of live glial cells (A) or live neurons (B) after treatment of mixed neuroglial cultures with 19-epi-OKA for 7 or 24 h at the indicated concentrations are represented. Dose-response curves obtained for OKA are also shown for comparison. Compared to neurotoxicity, glial cell toxicity by 19-epi-OKA was observed much earlier (7 h instead of 24 h) and required higher concentrations of toxins (EC50

7h

≈ 920 ± 78nM 19-epi-OKA for glial cells and EC5024h

≈ 285 ± 5nM 19-epi-OKA for neurons).

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unaltered after 7-h exposure to much higher concentrations (up to 2.5µM 19-epi-OKA or 100nM OKA) of the toxins (Fig. 5B). Longer exposures (24 h) to low concentrations (up to 250nM 19-epi-OKA or 5nM OKA) affected morphology and viability of neurons but not glial cells present in mixed cultures, whereas extensive toxicity of both types of cells was observed after 24-h exposure to high concentrations of the toxins (Figs. 5A and B). Concentrations of toxin producing 50% reduction of sur-viving glial cells after 7-h exposure (EC50

7) were estimated

at EC507EpiOKA,Glia

≈ 920 ± 78nM 19-epi-OKA and EC507OKA,Glia

≈ 33 ± 6nM OKA, respectively, and longer exposures (24 h) to the toxins shifted these values to 582 ± 26nM 19-epi-OKA and 22 ± 5nM OKA (Fig. 5A), respectively. In the case of neu-rons, no toxicity was observed at 7 h, and the estimated EC50

24

values were EC5024EpiOKA,Neu

  =  285 ± 5nM 19-epi-OKA and EC50

24OKA,Neu  =  2.5 ± 1nM OKA. As for 24-h exposure, con-

centration dependence, as well as maximal toxicity and EC50 values, was similar to those observed in pure neuronal cultures (see Fig. 1).

Concomitant Exposure to PP1 Specific Inhibitor Tautomycetin Increases Vulnerability of Neurons to 19-epi-OKA

19-epi-OKA showed a PP2A/PP1 specificity 10-fold greater than OKA or dinophysistoxin-2 (Cruz et al., 2007), making it one of the most selective PP2A inhibitors known. To test the possible role of the lack of inhibition of PP1 in the much lower toxic potency we observed for 19-epi-OKA compared with OKA or dinophysisitoxin-2, we used the highly specific PP1 inhibitor tautomycetin (Mitsuhashi et al., 2001). Dose-response experiments showed no morphological signs of toxicity or sig-nificant reduction in neuronal survival at concentrations of tau-tomycetin up to 500nM for 40 h (see Supplementary fig. S2). Based on both these results and those shown in Figure  1, in further experiments we combined 200nM tautomycetin and 100nM 19-epi-OKA, concentrations that did not compromise neuronal viability for at least 48 h when applied separately. Widespread neurotoxicity was observed in neurons exposed to both tautomycetin and 19-epi-OKA. Interestingly, OKA (1nM) neurotoxicity was not enhanced in the presence of the PP1-specific inhibitor tautomycetin (see Table 3).

diSCuSSiOn

In this study, we report for the first time data about the action on cell survival and physiology of 19-epi-OKA, a new PP inhibitor isolated from cultures of P.  belizeanum (Cruz et al., 2007; Paz et al., 2008). We show 19-epi-OKA to cause degeneration and death of cerebellar cells in primary culture. Concentrations of 19-epi-OKA in the nanomolar range caused visible signs of neurotoxicity, including degeneration of neur-ites and swelling of cell bodies, leading to widespread neuronal death. Neurotoxicity by 19-epi-OKA was accompanied by the

laddering-like fragmentation of DNA, a hallmark of apoptosis (Raff, 1992), and chromatin condensation and fragmentation. This neurotoxicity pattern is similar to that reported previ-ously for OKA and dinophysistoxin-2 (Fernández et al., 1991, 1993; Pérez-Gómez et al., 2004) and is consistent with 19-epi-OKA being included among the toxins responsible for DSP. Further evidence supporting this idea includes the observation that both 19-epi-OKA and OKA induced the activation of the extracellular signal-regulated protein kinase1/2 mitogen-acti-vated protein kinase (ERK1/2 MAPK) to a similar extent and with similar time dependencies (see Supplementary fig. S3) and induced caspase-3 activity. Caspase-3 activation appears to be required for apoptotic chromatin condensation and DNA fragmentation and may also function before or at the stage when commitment to loss of cell viability is made (Porter and Jänicke, 1999). The early caspase-3 activation observed in neu-rons exposed to 19-epi-OKA or OKA preceding the appearance of apoptotic hallmarks is therefore consistent with the implica-tion of caspase-3 activity in the apoptotic neurotoxicity induced by these OKA derivatives. In addition, differential sensitivity to 19-epi-OKA was observed between neuronal and nonneuronal cells, a quite specific characteristic shared by OKA and dino-physistoxin-2 (see Fig. 5 and Pérez-Gómez et al. (2004) ) but not by other toxins including yessotoxin and palytoxin (data not shown).

Compared with OKA and dinophysistoxin-2, the neurotoxic actions of 19-epi-OKA occurred at much higher concentrations of the toxin. Thus, the minimum concentration of 19-epi-OKA producing morphological signs of neurotoxicity was approxi-mately 150nM compared with 0.5nM for OKA and 2.5nM for dinophysistoxin-2 (Fernández et al., 1991; Pérez-Gómez et al., 2004), and the EC50

24 value for 19-epi-OKA (~300nM) was

approximately 150 and 40 times higher than the correspond-ing EC50

24 values for OKA (2nM) and dinophysistoxin-2

(8nM), respectively. The lower toxicity of 19-epi-OKA we observed is consistent with the structural features found in this new compound (Cruz et al., 2007). A key difference between 19-epi-OKA and OKA is that the carboxylic acid (C1) and the hydroxyl groups located at C24 are situated in 19-epi-OKA on different faces of the molecule, hindering the possibility

TAblE 3 Tautomycetin Enhances Vulnerability of neurons to 19-epi-OKA

Treatment

Neuronal survival (%)

None Tautomycetin

None 96 ± 2 92 ± 419-epi-OKA 95 ± 2 46 ± 12*OKA 80 ± 7 70 ± 10

Notes. Neurons were exposed to 19-epi-OKA (100nM) or OKA (1nM) in the absence (None) or in the presence of the PP1 inhibitor tautomycetin (200nM). Percentage of surviving neurons was determined after 24-h exposure to 19-epi-OKA or OKA. Each value represents the mean ± SD (n = 4).

*p < 0.05 versus none in the same row.

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to form a hydrogen bond between them, which is considered to play a crucial role in the formation of the OKA-PP1/PP2A complexes. As a result, the contact points between this new inhibitor and the protein should be very different than those found for OKA and eventually result in lower relative PP1/PP2A inhibitory potencies and cell toxicity by 19-epi-OKA. Previous data from in vitro assays (Cruz et al., 2007) do not allow, however, for a simple correlation between the differ-ences in the toxic potency of 19-epi-OKA and OKA we report and the PP1/PP2A inhibitory potencies of both compounds. In those experiments, 19-epi-OKA and OKA were found to be equipotent in inhibiting the catalytic subunit of PP2A (PP2Ac), whereas 19-epi-OKA inhibited the catalytic subunit of PP1 (PP1c) much less potently (~7.5-fold) than OKA (Cruz et al., 2007). Different possibilities might be taken in consideration in order to explain this apparent discrepancy. First, it might be attributed to differences in the plasma membrane perme-ability of the compounds. Because of the absence of the intra-molecular C24/C1 hydrogen bond, 19-epi-OKA appears to be indeed much more polar than OKA, as demonstrated also by the fact that it was soluble in methanol but not in chloroform (Cruz et al., 2007). However, medium exchange experiments demonstrated that the amount of 19-epi-OKA that had not entered into the cells after 24-h exposure and remained in the culture medium was too low to induce neurotoxicity in sister cultures and showed also no significant neurotoxicity differ-ences between 19-epi-OKA and OKA, thus arguing against a different cellular uptake of both compounds. A second possible explanation could be that the ability of 19-epi-OKA to interact and inhibit PPs, and in particular PP2A, differed significantly in vitro from the much more complex situation in vivo. PP2A is an important regulator of numerous target proteins within diverse signaling pathways and consequently its activity in vivo must be tightly regulated. The predominant form of PP2A within the cell appears to be a heterotrimeric holoenzyme including scaffolding (A), regulatory (B), and catalytic (C) subunits. Distinct mutually exclusive families of regulatory B subunits have been identified that control phosphatase activity, substrate selectivity, and target PP2A holoenzymes to distinct cellular compartments and to specific substrates (Janssens and Goris, 2001; McCright and Virshup, 1995; McCright et  al., 1996; Strack et  al., 1998; Virshup and Shenolikar, 2009; Virshup et al., 2000). B subunits showed tissue-specific locali-zation, and, interestingly, some members of the B subunit family were detectable only in brain regions including cer-ebellum (Strack et  al., 1998). PP inhibitors including OKA and microcystin-LR have been shown to be able to disrupt the interaction of PP2A with some of these subunits (Kloeker et  al., 2003). However, the differences in structure and con-formation in solution occurring in 19-epi-OKA (Cruz et  al., 2007; Fernández et al., 2003) may prevent a similar disruption by this inhibitor, thus explaining the much lower inhibitory potency of this compound compared with OKA or dinophysis-toxin-2. A third possibility could be related to posttranslational

modifications, i.e., carboxymethylation, ubiquitination, and phosphorylation of the PP2A subunits, that have been previ-ously shown to influence the activity, oligomeric composition, and intracellular levels of this enzyme (Bryant et  al., 1999; Chen et al., 1992; Trockenbacher et al., 2001). Such regulatory mechanisms are not present in in vitro paradigms and may be also affected by 19-epi-OKA. Finally, it could be also argued that the observed effects of these compounds on neurons and glial cells might not be related to their PP inhibitory activity. However, previous studies have demonstrated the actions of OKA on neurons and glial cells to be effectively mimicked by other PP inhibitors (Fernández-Sánchez et al., 1996; Ferrero-Gutiérrez et al., 2008), with greater correlation with inhibition of PP2A rather than PP1 activity. Moreover, the closely related structural analogue of OKA, OKA-TME, lacking PP inhibitory activity, had no effect on neurons or glial cells, thus confirming the role of PP inhibition in the toxic action of OKA on these cells. These observations, together with the strong similarities in the morphological toxicity pattern induced by 19-epi-OKA, OKA and dinophysistoxin-2, strongly support the involvement of PP inhibition in the observed effects of 19-epi-OKA.

The differential contribution of PP activities other than PP2A to the cellular effects exerted by OKA and 19-epi-OKA may play also an important role in the observed difference between the toxicological and biochemical potencies of these com-pounds. Although inhibition of PP2A appears to have a major contribution in OKA-induced apoptotic death in cerebellar neu-rons (Fernández-Sánchez et al., 1996), OKA potently inhibits also the activity of PP1 and the activity of other PPs including PP4, PP5, and likely PP6A (see Swingle and Honkanen, 2007). In contrast, 19-epi-OKA is a very selective PP2A inhibitor, with a PP2A/PP1 specificity 10-fold higher than OKA (Cruz et  al., 2007). Inhibition of PP1 activity, which together with PP2A represents the most abundant PP in mammalian neu-rons, could therefore represent a prominent contributing factor to OKA but not 19-epi-OKA–induced neurotoxicity, and our observations that concomitant exposure to PP1 specific inhibi-tor tautomycetin increases vulnerability of neurons to 19-epi-OKA would be consistent with this possibility. Also, inhibition of PP1 has been previously demonstrated to induce apoptotic cell death associated with upregulated expression of caspase-3 at the mRNA, protein, and enzyme activity levels (Wang-Chen et al., 2001), suggesting a role for PP1 inhibition in the increase in caspase-3 activity observed after 1-h exposure to OKA but not 19-epi-OKA.

We have previously proposed a neuronal bioassay (García-Rodríguez et al., 1998) for the detection of OKA derivatives based in the high sensitivity of cultured cerebellar neurons to OKA and dinophysistoxin-2 (Fernández et  al., 1991, 1993; Pérez-Gómez et  al., 2004). Although the enzyme inhibition assay remains a rapid, simple and very convenient method for the current detection of OKA derivatives in monitoring pro-grammes, the results described herein further support the suit-ability of cultured neurons as an alive experimental system

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sufficiently sensitive to provide useful information about the rel-ative toxicological potency of these molecules. It is worth not-ing that the differences in the neurotoxic potency we observed for OKA and 19-epi-OKA are consistent with the differences in structure and conformation between these compounds. The observed neurotoxic effects of these molecules also validate the suitability of these neurons in culture for the development of neuronal networks on multielectrode arrays as biosensors for marine biotoxins (Hogberg et  al., 2011), including OKA derivatives. In view of the observed effects of 19-epi-OKA on glial cells, previously described also for OKA and dinophysis-toxin-2 (Perez-Gómez et al., 2004), cultured astrocytes, which can be of much easier handling and commercialization than cultured neurons, appear also as a very convenient model for complementing the biological analysis of these toxins.

Funding

Spanish Ministry for Science and Technology, grants CTQ2008-06754-C04-03/PPQ, SAF2011-28883-C03-03.

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