* Biochemistry and Molecular Biology Department, University of Oviedo, Oviedo, Spain; PROTEOBIO, Mass Spectrometry Center for Proteomics and Biotoxin Research, Department of Chemistry, Cork Institute of Technology, Bishopstown, Cork, Ireland; and
Psychology Department, University of Oviedo, Oviedo, Spain
Received February 11, 2004; accepted March 26, 2004
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ABSTRACT |
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Key Words: dinophysistoxins; diarrhetic shellfish poisoning; neurotoxicity; apoptosis; cerebellar neurons; glial cells.
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INTRODUCTION |
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Diarrhetic shellfish poisoning (DSP) is a human gastrointestinal illness caused by the consumption of shellfish contaminated with polyether toxins produced by dinoflagellates belonging to the genera Dinophysis, though some DSP toxins are also produced by benthic species of Prorocentrum (Yasumoto et al., 1979, 1984
). Okadaic acid, its isomer dinophysistoxin-2 (DTX-2), and their methyl homologue dinophysistoxin-1 (DTX-1), are the main DSP toxins. Although okadaic acid is usually the principal toxin in Europe, reports concerning the presence of dinophysistoxins in coastal waters have significantly increased in the last years suggesting the spread of these toxins worldwide. DTX-1 is the dominant toxin in Japan (Yasumoto et al., 1979
), in Canada (Quilliam et al., 1993
), and in certain Norwegian fjords (Lee et al., 1989
), while DTX-2 has been shown to be the predominant toxin in Irish mussels (Carmody et al., 1996
; McMahon et al., 1996
). High amounts of DTX-2 have been also detected in shellfish from the Galician region of Spain (Blanco et al., 1995
) and in Portuguese mussels (Vale and Sampayo, 1996
, 2000
). Also, a monitoring program carried out in 1996 and 1997 confirmed for the first time the occurrence of DTX-2 in Adriatic mussels (Pavela-Vrancic et al., 2002
).
The widespread distribution of DSP toxins in seafood has caused increased concern due to the threat of public health, and has underlined the need for toxicological studies in order to evaluate the potential risk for human health due to the presence of these toxins in food. Up to now most studies have been focused on okadaic acid, while studies regarding dinophysistoxins have been very limited and very few data exist about the action of these compounds on cell survival and physiology. As for DTX-2, toxicological studies have been particularly hampered by the lack of purified material. DTX-2 is produced by phytoplankton belonging to Dinophysis sp., which cannot be maintained in culture, and therefore isolation from wild marine biological materials represents the only source of this toxin.
Cerebellar neurons in primary culture represent an experimental system that has proved very useful in a variety of toxicological studies. These cultures have been extensively used in the study of the biochemical events coupled to neurotoxicity by excitatory amino acids and the conditions controlling excitotoxicity (Fernández-Sánchez and Novelli, 1993; Lipsky et al., 2001
; Novelli et al., 1987
, 1988
). Cultured cerebellar neurons have been also identified as a very convenient model for the study of neuronal apoptosis in vitro (D'Mello et al., 1993
) and extensively used for that purpose thereafter. Moreover, we have found cultured cerebellar neurons to be very useful to investigate the action of different types of marine toxins including okadaic acid (Fernández et al., 1991
, 1993
; Fernández-Sánchez et al., 1996
; Novelli et al., 1992
). In this study we have used primary cultures of cerebellar neurons to investigate the actions of DTX-2, isolated from contaminated Irish mussels. We also used co-cultures of cerebellar neurons and astrocytes and highly enriched astroglial cultures in order to identify possible differences between the actions of this toxin on neuronal and non-neuronal cells. Our study provides the first data about the mechanisms involved in the toxic effects of DTX-2 on live cells. In view of the high sensitivity of cultured cerebellar neurons to DTX-2 and okadaic acid, this tissue culture system appears to be an appropriate model for the assessment of the potential risk for human health of DSP toxins and for the establishment of safety levels of these compounds in seafood.
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MATERIALS AND METHODS |
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Cell culture. Primary cultures of rat cerebellar neurons were prepared as previously described (Novelli et al., 1988). Cytosine arabinoside (10 µM) was added after 2024 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 four days and compensating for lost amounts of water, due to evaporation. Mixed cerebellar cultures containing neurons and astrocytes and highly enriched astroglial cultures 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 population in both pure and mixed cultures, and 5% in astroglial cultures. The animal procedures used were in accordance with the protocols approved by the Institutional Animal Care and Use Committee of the University of Oviedo
Neurotoxicology. Neurons were used between 1420 days in culture. Drugs were added into the growth medium at the indicated concentrations, and neuronal cultures were observed for signs of neurotoxicity thereafter by phase contrast microscopy. To quantify neuronal survival cultures were stained with fluorescein diacetate and ethidium bromide (Fernández et al., 1991; Novelli et al., 1988
), photographs of three randomly selected culture fields were taken and live and dead neurons were counted. Results were expressed as percentage of live neurons. Total number of neurons per dish was calculated considering the ratio between the area of the dish and the area of the pictures (
3000).
Confocal microscopy. Oxygen radical formation was detected with carboxy-2'7'-dichlorodihydrofluoresceine diacetate (carboxy-H2DCFDA). Following uptake, the carboxy-H2DCFDA is converted by endogenous esterases to carboxy-H2DCF, which upon exposure to oxidative species is oxidized to the fluorescent probe carboxy-DCF. Cultures were treated with DTX-2 (1050 nM) for 618 h and then loaded with 20 µM carboxy-H2DCFDA in the culture medium for 1 h. Then, the dye was removed and cultures were washed twice with a buffer containing (in mM): 154 NaCl, 5.6 KCl, 5.6 glucose, 8.6 HEPES, 1 MgCl2, 2.3 CaCl2 (pH 7.4). Carboxy-DCF fluorescence was recorded in a Bio-Rad confocal microscope with a krypton-argon laser excitation source (488 nm). Signals were digitized using Bio-Rad interface and fluorescence intensity was quantified in 1015 cell bodies per field using the software NIH Image (version 1.61).
DNA fragmentation analysis. Cells were lysed in 10 mM Tris-HCl, 0.5% Triton X-100, 20 mM ethylenediamine tetraacetic acid (EDTA), pH 7.4. After 20 min. on ice, the lysate was centrifuged at 13,000 x 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 vol. of ethanol and 300 mM sodium acetate. Samples were electrophoresed in a 1.5% agarose gel and visualized by ethidium 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.
Data presentation and analysis. For statistical analysis a one-way or a two-way analysis of variance (ANOVA) was used to identify overall treatment effects, followed by the unpaired two-tailed Student's t-test for selective comparison of individual data groups. Bonferroni's correction was applied to the significance level. Only significances relevant for the discussion of the data are indicated in each figure.
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RESULTS |
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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 induced by DTX-2 required newly synthesized proteins. No significant increase in the number of apoptotic nuclei was observed in neurons exposed to DTX-2 in the presence of actinomycin D (8 ± 2%, n = 6) compared to untreated neurons (6 ± 1%, n = 4) (Figs. 2b and 2d). Accordingly, both actinomycin D (1 µg/ml) and cycloheximide (5 µg/ml) prevented neurotoxicity by DTX-2 (Fig. 3).
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The effect of DTX-2 on astrocyte morphology and viability was further confirmed by using highly enriched astroglial cultures containing no neurons. More than 80% of the nuclei visualized by ethidium bromide staining in these cultures corresponded to GFAP-positive cells (not shown). After 10 h exposure to 50 nM DTX-2, the majority of these astrocytes had already shrunk and degenerated (Fig. 5b), and DNA staining by Hoechst 33258 revealed that glial toxicity by DTX-2 occurred via apoptosis. Thus, 30 ± 7% (n = 8) of nuclei appeared either shrunken or degraded and with aggregation of chromatin (Fig. 5d) compared to 3 ± 2% (n = 4) in control cultures in which most nuclei had regular contours and were round and large in size (Fig. 5c).
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DISCUSSION |
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Neurotoxicity by DTX-2 was accompanied by the laddering-like fragmentation of DNA, a hallmark of apoptosis (Raff, 1992), similarly to what has been previously observed for okadaic acid (Fernández-Sánchez et al., 1996
). Assessment of chromatin condensation by staining of nuclei with the Hoechst dye further confirmed that neuronal degeneration by DTX-2 was attributable to apoptosis. DNA fragmentation occurred early in the death process, as it could be detected long before neuronal degeneration and death could be observed. Also, the observation that it could be prevented by inhibitors of RNA and protein synthesis indicates that apoptosis by DTX-2 is a genetically controlled process. These observations are consistent with a regulation by DTX-2 of expression of genes coding for proteins playing active roles in the control of apoptosis. Among possibilities, p53 and members of the bcl-2 family, Bcl-2, BAX, and bcl-x, appear to be plausible candidates as they have been suggested to be involved in the apoptotic response induced by okadaic acid in other cell types (Benito et al., 1997
; Sheikh et al., 1996
; Yan et al., 1997
). Experiments are in progress to investigate the possible role for the expression of these genes in the neuronal apoptosis induced by DTX-2.
The rescue of neurons by the transcription inhibitor actinomycin D also raises the question of whether aberrant activation of cell cycle regulatory proteins could play a role in neurotoxicity by DTX-2, as demonstrated for apoptosis by other stimuli in post-mitotic neurons including cerebellar granule neurons (Martin-Romero et al., 2000; Padmanabhan et al., 1999
; Park et al., 1996
). We are currently investigating the possible involvement of cyclin-dependent kinases in the effects of DTX-2. Interestingly, preliminary data showed that in glial cells apoptosis by DTX-2 could not be inhibited by actinomycin D (data not shown). Thus, comparison between neuronal and glial cells may provide new insights about the mechanisms controlling apoptosis in neuronal versus non-neuronal cells and these cerebellar cultures appear to be an appropriate model for these types of studies.
Recent data suggest that toxic insults to glial cells might cause degeneration of surrounding neurons both in experimental systems and in human pathologies (Akiyama et al., 2000; Ekdahl et al., 2003
). Consistently, we have previously reported that apoptotic degeneration of cultured cerebellar glial cells due to long exposures to aluminum induced neuronal degeneration and death (Suárez-Fernández et al., 1999
). In contrast, DTX-2 toxicity appeared to occur independently in neurons and glial cells. The relative resistance of neurons to short exposures of high concentrations of DTX-2 compared to the co-cultured astrocytes, suggests that DTX-2-induced glial toxicity is not likely to be associated either with increased glial reactivity toward neuronal cells or with the release of neurotoxic factors from glial cells. Preliminary experiments indicate a similar pattern of neuro-glial toxicity for okadaic acid (data not shown). Thus, these DSP toxins may represent very interesting biochemical tools for elucidating how glial cells may die without affecting neighboring neurons.
We have previously proposed a neuronal bioassay (García-Rodríguez et al., 1998) for the detection of DSP toxins, based in the high sensitivity of cultured cerebellar neurons to okadaic acid (Fernández et al., 1991
, 1993
). By using purified okadaic acid as reference standard, neuronal bioassay proved to be a suitable method for the detection and quantification of DSP toxicity in toxic mussel extracts which were demonstrated by HPLC to contain both okadaic acid and DTX-2, and allowing for a detection limit as low as 20 µg okadaic acid equivalents per kg of fresh animal tissue. Results described herein further support the suitability of cultured neurons for the analysis of total DSP toxicity. It is worth noting that DTX-2 appeared to be a less potent neurotoxin than okadaic acid (Fig. 1). Thus, cultured cerebellar neurons provide a sensitive model for the evaluation of DSP toxins that may give also useful information about the relative potency of toxins. In view of the marked effects of DTX-2 on astrocytes we report, and given that these cells can be of much more easily handling and commercialization than cultured neurons, cultured astrocytes also represent a very promising model for the biological detection of DSP toxins.
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed at Antonello Novelli or María-Teresa Fernández-Sánchez, Departamento de Bioquímica y Biología Molecular, Edificio "Santiago Gascón". Campus "El Cristo", Universidad de Oviedo, 33006 Oviedo, Spain. Fax: (+34) 985103157. E-mail: neurolab{at}bioquimica.uniovi.es.
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