• Chitosan, a component of the cell wall of many fungi, has been widely used to mimic pathogen attack and has been shown to induce several defence responses.
• Here we show that low concentrations (50 µg ml−1) of chitosan are able to induce an increase in cytosolic Ca2+ concentration ([Ca2+]cyt), accumulation of H2O2 in the culture medium, induction of the defence gene chalcone synthase (chs), and cell death in soybean cells (Glycine max).
• Chitosan-induced cell death occurred through cytoplasmic shrinkage, chromatin condensation and activation of caspase 3-like protease, suggesting the activation of a programmed cell death (PCD) pathway. Buffering extracellular Ca2+ with the Ca2+ chelator EGTA prevents [Ca2+]cyt elevation, H2O2 production and all downstream PCD features, but not cell death.
• Higher doses (200 µg ml−1) of chitosan evoked neither Ca2+ transient and H2O2 production nor caspase 3-like activation, but caused cell death, possibly as a result of plasma membrane disturbance.
Plants protect themselves against the invasion of a broad array of environmental microorganisms with a complex range of defence processes (Scheel, 1998; Maleck & Dietrich, 1999; Rowland & Jones, 2001). Defence responses upon pathogen infection include three steps: the initial recognition of the pathogen, a subsequent transduction of the signal, and the following execution of the defence program (Scheel, 1998). Similar to the animal counterpart, plant cells can respond to biotic and abiotic stresses by activating genetic programs for cell suicide (Beers & McDowell, 2001). This programmed cell death (PCD) has been found to be induced at the site of pathogen invasion in several plant species (Heath, 2000) as an attempt to isolate the pathogen and prevent it spreading to noninfected parts of the plant. The mechanisms that control PCD in animals have been widely investigated and well documented, whereas little is known of the regulation of plant cell death. Recent studies have reported that the hypersensitive response (HR) shares some features recognized for mammalian apoptotic process, such as elevation of cytosolic Ca2+ concentration ([Ca2+]cyt), the involvement of reactive oxygen species (ROS), nuclear condensation, protease activation, and DNA fragmentation (Dangl et al., 1996; Desikan et al., 1998; Xu & Heath, 1998; Lam et al., 1999; Mittler et al., 1999; Solomon et al., 1999). Treatments with elicitors generally make a plant more resistant to a further pathogen attack inducing HR (Bradley et al., 1992), pathogenesis-related (PR) gene expression (Doares et al., 1995; Repka, 2001), phytoalexin biosynthesis (Nurnberger et al., 1994; Jabs et al., 1997) and cell wall strengthening (Bruce & West, 1989; Bradley et al., 1992). The perception of pathogen-derived elicitors seems to be mediated by specific receptors at the plasma membrane (Zimmermann et al., 1997); however, the mechanisms involved in both elicitor perception at the plant cell surface and the following intracellular transmission of the signal have not yet been fully unravelled.
In this study we have investigated the response of aequorin-expressing soybean (Glycine max) suspension cell cultures to treatment with different concentrations of chitosan. A possible role of this elicitor in the interaction between pathogenic fungi and plant hosts has been recently proposed (El Gueddari et al., 2002). Low and high doses of the elicitor were found to induce cell death with different modalities. Either elevation of [Ca2+]cyt and consequent activation of caspase 3-like mediated PCD or induction of cell death without [Ca2+]cyt increase and caspase activation were observed, suggesting the triggering of different signal transduction pathways depending on the elicitor concentration. It is proposed that Ca2+ may participate in the switching among the different chitosan-induced cell death pathways.
Materials and Methods
Crab-shell chitosan with 85% N-deacetylation (polymerization degree: 11392) was obtained from Sigma-Aldrich (St. Louis, MO, USA). Coelenterazine was purchased from Molecular Probes Inc. (Eugene, OR, USA). Chitosan was solubilized (5 mg ml−1) in 0.5 m aqueous acetic acid and the solution was alkalized with 1 N KOH to pH 5.2.
Suspension cultured soybean cells (Glycine max L.) stably expressing cytosolic aequorin (Mithöfer et al., 1999, kindly provided by G. Neuhaus, Freiburg, Germany) were grown in Murashige and Skoog (MS) medium supplemented with 5% sucrose, 10 µg ml−1 naphthlaleneacetic acid, 2 µg ml−1 kinetin, 1 µg ml−1 thiamine-HCl and 10 µg ml−1 kanamycin. Cells were freshly reinoculated every 3 wk (10% v/v inoculum) and maintained at 24°C with 16-h photoperiod constantly shaking. Treatments with chitosan at different concentrations (25–200 µg ml−1) were performed 3 wk after reinoculation, during the exponential growth phase of the cells.
Aequorin-dependent Ca2+ measurements
Reconstitution of the Ca2+-sensitive photoprotein aequorin was performed in vivo by incubating soybean cells with 5 µm coelenterazine overnight in the dark. Treatments with chitosan were carried out by injecting 50 µl of two-fold concentrated chitosan solution (dissolved in the basal cell culture medium) to the same volume of reconstituted cell-suspension culture (about 3 mg f. wt) using a light-tight-syringe. For experiments carried out in the absence of extracellular Ca2+ cells were extensively (10 vol) washed three times with Ca2+-free culture medium and then resuspended in the same medium containing 600 µm EGTA. The residual aequorin was completely discharged by adding 0.33 m CaCl2 in 10% ethanol. Luminescence data were collected and elaborated into [Ca2+]cyt by a computer algorithm based on the Ca2+ response curve of aequorin (Brini et al., 1995).
Assay for H2O2
Hydrogen peroxyde production was quantified in the culture medium of cells as described by Wolff (1994) in terms of colorimetric reaction caused by the peroxide-mediated oxidation of Fe2+ followed by the reaction of Fe3+ with xylenol orange. 0.5 ml assay solution (0.5 mm ammonium ferrous sulfate, 50 mm H2SO4, 0.2 mm xylenol orange, 200 mm sorbitol) was added to 0.5 ml of control and treated soybean cells and the absorbance at 560 nm was detected after 45 min incubation.
RNA extraction and Northern blot hybridization
Total RNA was extracted from soybean cells by using the ‘RNAeasy Plant Mini Kit’ (QIAGEN GmbH, Hilden, Germany) following manufacturer's instructions. Samples of 14 µg total RNA were separated on a 1.2% agarose gel containing 6.7% formaldehyde and transferred onto Hybond-N+ membrane (Amersham, Braunschweig, Germany). Pre-hybridization and hybridization reactions were performed at 42°C in the presence of formamide according to Sambrook et al. (1989). Filters were hybridized with a digoxigenin-labeled probe, incubated in CDP-Star (Roche, Indianapolis, IN, USA) and exposed to X-ray film (X-Omat AR, Kodak, Rochester, NY, USA). The exposed films were quantified by densitometric analysis using the ‘Quantity One’ software (Bio-Rad, Oakland, CA, USA). The probe used to detect the chalcone synthase (chs) mRNA was a cDNA fragment, labelled with digoxigenin-11-dUTP by a DIG Probe Synthesis kit (Roche), of chs gene obtained after RT-PCR on RNA extracted from soybean cells treated with 25 µg ml−1 chitosan. Briefly, RT-PCR was performed on DNase-treated RNA by ‘Ready-To-Go RT-PCR beads’ (Amersham Pharmacia, Uppsala, Sweden). Primers (forward primer: 5′-GTTTGTGCTTACATGGCACC-3′; reverse primer: 5′-ACTTAGCCTCAACTTGGTCC-3′) were designed on the basis of a published soybean chs nucleotide coding sequence (Genebank accession number X53958). First strand cDNA synthesis was at 42°C for 30 min, followed by a single step at 95°C for 5 min. PCR consisted of 35 cycles of denaturation (95°C for 1 min), annealing (57°C for 1 min) and elongation (72°C for 1 min). The amplicon of the expected size was cloned by the ‘pGEM-T Easy Vector System kit’ (Promega, Madison, WI, USA). The insert contained in the plasmid DNA isolated from a recombinant clone was confirmed to correspond to a chs cDNA by sequencing.
Detection of cell death and nuclei staining
Cell viability was determined by 10 min incubation of the cell suspension with 0.05% Evan's blue (Sigma-Aldrich). After several washings with deionized water to remove the excess of the dye, dye bound to dead cells was solubilized in 50% methanol/1% SDS and quantified spectrophotometrically by measuring the absorbance at 600 nm (Baker & Mock, 1994). Three-week-old suspension cells were incubated in the dark with 8 µg ml−1 HOECHST 33342 (HO) (Sigma-Aldrich) and 5 µg ml−1 Propidium Iodide (PI) (Sigma-Aldrich) at RT for 10 min and observed under fluorescent microscopy by using an excitation light of 350 nm and 570 nm, respectively, for the two dyes.
Transmission electron microscopy
For transmission electron microscopy, soybean cells were collected after 4 h of treatment and fixed with 3% glutaraldehyde in 0.1 m cacodylate buffer for 24 h at 4°C. Cells were postfixed for 2 h at 4°C with 1% osmium tetroxide in 0.1 m cacodylate buffer, dehydrated in a graded ethanol series and then embedded in araldite resin. Ultra-thin sections (0.05 µm) were cut with an ultramicrotome, stained with uranyl acetate in ethanol for 30 min and observed at 75 KV in a Hitachi 300 TEM. About 200 cells were analysed in each condition.
Caspase 3-like activity
15 µg of total protein extracts were incubated for 2 h at 37°C with the synthetic tetrapeptide DEVD-p-nitroaniline (pNA), in the presence or absence of the caspase 3 inhibitor Ac-DEVD-CHO (20 µm), as described by the manufacturer (‘caspase-3 colorimetric activity assay kit’, Chemicon International, Temecula, CA, USA). Addition of the substrate resulted in a signal caused by the caspase 3-dependent cleavage of the chromophore pNA from the labelled substrate. Caspase 3-like activity was determined by spectrophotometric quantification of the free pNA (λem = 405 nm).
Data are presented as mean ± SD and the differences between groups were assessed with Student's t-test.
Effect of chitosan on cytosolic Ca2+ elevations and H2O2 production in soybean cell cultures
Changes in [Ca2+]cyt and extracellular H2O2 accumulation were monitored after administration of increasing concentrations of the exogenous elicitor chitosan (25–200 µg ml−1) to aequorin-expressing soybean suspension cells. A transient increase in [Ca2+]cyt from a basal level of about 0.20 µm to 0.77 µm ± 0.16 µm and 0.68 µm ± 0.14 µm was recorded after cell treatment with 25 and 50 µg ml−1 chitosan, respectively (Fig. 1a). The maximal [Ca2+]cyt peak was reached after about 3 min upon chitosan administration and fell back to the basal level after about 5 min (Fig. 1a). A gradual attenuation of the Ca2+ rise was observed using increasing concentrations of chitosan, with an almost complete annulment of the Ca2+ transient following treatment with the 200 µg ml−1 dose (Fig. 1a).
A similar reverse dose-dependent trend was observed for H2O2 accumulation in the culture medium (Fig. 1b). The concentration of 25 µg ml−1 chitosan caused a maximum production of H2O2 (167 µm ± 8 µm reached after about 45 min), whereas the 200 µg ml−1 dose did not induce any detectable H2O2 accumulation.
Cells challenged with culture medium containing the same percentage of aqueous acetic acid, pH 5.2, as chitosan-treated cells, did not show any [Ca2+]cyt elevation or extracellular H2O2 accumulation (Fig. 1a,b).
In soybean cells a maximum accumulation of chs transcripts was observed after 4 h of incubation with 25 µg ml−1 chitosan (Fig. 2). The elicitor-induced increase of chs mRNA was remarkably reduced with 100 µg ml−1 and with the highest concentration of 200 µg ml−1 (Fig. 2).
Chitosan triggers a cell death program
We analysed the effect of chitosan in the induction of cell death. Administration of 25–200 µg ml−1 chitosan to soybean cells for 4 h and 24 h caused cell death, as measured by Evan's blue staining (Turner & Novacky, 1974; Levine et al., 1996). The percentage of cell death rose with the increase in elicitor concentration and length of treatment (Fig. 3). The highest value was observed with 200 µg ml−1 chitosan for 24 h (about 78% ± 5%), whereas 25 µg ml−1 caused about 27% ± 2% cell death after the same time of elicitor incubation (Fig. 3). Control soybean cell cultures exhibited about 5% ± 0.5% of Evan's blue stained cells (Fig. 3).
To further determine the nature of the cell death induced by chitosan we analysed the occurrence of the main PCD hallmarks recognized for plant cells such as morphological changes, nuclear morphology, caspase 3-like activity, and DNA fragmentation (Chen et al., 2000; McCabe & Leaver, 2000), focusing our attention on the two concentrations of 50 µg ml−1 and 200 µg ml−1.
Under light microscopy control cells showed a well defined structure with round nuclei, a central vacuole and cortical chloroplasts (Fig. 4a), while chitosan-treated cells were found to undergo various progressive morphological changes. Cells treated for 4 h with 50 µg ml−1 chitosan showed a cell disorganization with a gradual condensation of the cytoplasm and a consequent detaching of the plasma membrane from the cell wall (Fig. 4d,g). Different degrees of cell disorganization were found to coexist, indicating a different sensitivity to the elicitor molecule in an asynchronized cell population. Treatment with 200 µg ml−1 led to highly collapsed cells already after 4 h (Fig. 4l). After 24 h treatment an increased percentage of cells showing cellular alterations was observed with both chitosan concentrations (data not shown).
To further investigate the cellular changes induced by chitosan a double staining of cells with HO/PI dyes was carried out. This staining allows the simultaneous detection of the early stages of apoptotic cells (HO+ and PI− nuclei) and of late apoptotic (HO+ and PI+ nuclei) or necrotic cells (mainly HO− and PI+ nuclei) (Hamatake et al., 2000).
As shown in Fig. 4, fluorescence from HO/PI of soybean control cells was faint or even not detectable (Fig. 4b,c), while cells treated for 4 h with 50 µg ml−1 chitosan showed an enhanced HO fluorescence (Fig. 4e,h), sometimes occurring with PI positive nuclei (Fig. 4i). The positivity in HO fluorescence in treated cells compared with the control indicates the presence of a highly condensed chromatin that sometimes appears to be localized at the periphery of the nucleus (Fig. 4e,h). On the whole the data reveal the presence of both early (Fig. 4e,f) and late (Fig. 4h,i) stages of PCD in the 4 h chitosan-treated cell population, resembling those observed during apoptosis in animal cells (Hamatake et al., 2000; Maruyama et al., 2000). After 24 h treatment the percentage of PI positive cells increased confirming the results obtained with the Evan's blue method (data not shown).
The double-staining of cells treated with 200 µg ml−1 chitosan showed that about 60–70% of cells were HO−/PI+ after 4 h chitosan incubation and about 80% after 24 h (Fig. 4m,n).
Transmission electron microscopy confirmed the induction of PCD-like morphological changes in soybean cells treated for 4 h with 50 µg ml−1 chitosan. Figure 5 shows various stages of the ultrastructural alterations caused by the elicitor (b–h) compared with the control (a). In about 25% of the cell population chitosan was found to induce detachment of the plasma membrane from the cell wall with shrinkage of the cell content (Fig. 5e,g). A condensation of chromatin is also evident in about 50% of chitosan-treated nuclei (Fig. 5c,g,h) compared with the control (Fig. 5a) and it seems to rise progressively with the advancing of the PCD-induced undoing of the cell. Intriguingly, chloroplast morphology was also affected by chitosan. In comparison with the control, a significant swelling of the stroma occurred (in about 50% of the cell population) after 4 h of treatment (Fig. 5b,d,f) (Fig. 5a).
The activation of caspase 3-like proteases has been shown to occur during PCD in several plant systems (Chen et al., 2000; Korthout et al., 2000) suggesting that some forms of plant PCD may have a caspase triggering pathway similar to the animal counterpart. To measure caspase 3-like activity, total protein extracts from soybean suspension cells treated with chitosan were incubated with the synthetic tetrapeptide DEVD-pNA, a specific substrate of caspase 3. Addition of the substrate resulted in a signal caused by the cleavage of the chromophore pNA from the labelled substrate. The caspase 3-like activity was monitored after 4 h of cell treatment with increasing concentrations of chitosan (50–200 µg ml−1). Only the 50 µg ml−1 dose was found to induce a statistically significant increase in protease activity, compared with untreated cells (Fig. 6a). Caspase 3-like activation was observed to occur after 1 h of 50 µg ml−1 chitosan treatment and to gradually rise with the incubation time (Fig. 6b). By contrast, 200 µg ml−1 chitosan did not induce any significant caspase 3-like activation throughout the considered time lapse (Fig. 6c). The use of the caspase 3 inhibitor Ac-DEVD-CHO completely abolished the caspase 3-like activity (Fig. 7a) and cell death (Fig. 7b) induced by 50 µg ml−1 chitosan.
Taken together the present data are suggestive of the induction by 50 µg ml−1 chitosan of a cell death pathway showing some PCD-like features recognized for animal apoptosis. Nevertheless, it was not possible to detect any typical DNA laddering either with agarose gel electrophoresis or in situ TUNEL assay within a 24-h time lapse (data not shown). Higher doses of chitosan seem to cause a different pathway of cell death occurring without involvement of caspase 3-like protease and chromatin condensation.
The annulment of the Ca2+ signal leads to different pathways of cell death
Incubation of cells with Ca2+-free medium containing 600 µm EGTA before challenge with the elicitor (50 µg ml−1) caused a complete inhibition of both the Ca2+ rise and H2O2 elevation (Fig. 8a). To investigate the outcome of the Ca2+ transient annulment on the cell death program we evaluated the induction of caspase 3-like activity and cell death in the absence of extracellular Ca2+. Incubation of cells with 50 µg ml−1 chitosan in Ca2+-free medium containing 600 µm EGTA did not induce any significant caspase 3-like activity compared with the control after 1 h of treatment (Fig. 8b). Nevertheless, the percentages of cell death in cells treated with chitosan in standard culture conditions and in Ca2+-free medium containing 600 µm EGTA were comparable (Fig. 8c). Moreover, the double staining of cells with HO and PI showed a staining of nuclei only by PI, suggesting the occurrence of cell death without chromatin condensation (Fig. 8d).
We showed that high doses of chitosan (200 µg ml−1) do not evoke [Ca2+]cyt elevation (Fig. 1a), as well as all the downstream PCD events (Figs 1b, 4l–n, 6a,c) but lead to a high percentage of dead cells (Fig. 3). Thus, the annulment of the Ca2+ signal either by deprivation of the extracellular Ca2+ pool or by treatment with high doses of elicitor, seems to have a key role in determining the type of cell death induced by chitosan.
High doses of chitosan (200 µg ml−1) induce alterations in plasma membrane permeability
To check if the lack of Ca2+ signalling upon high doses of chitosan may be caused by extensive plasma membrane damage, pretreated cells were incubated in Ca2+-free medium and subsequently challenged with a relatively high Ca2+ concentration (3 mm). Cells under conditions that alter membrane integrity are expected to face the sudden availability of extracellular Ca2+ with a major passive flow of the ion, resulting in altered sustained basal Ca2+ level. Upon CaCl2 addition, a rapid and steep Ca2+ increase was induced in both 200 µg ml−1 and 50 µg ml−1 chitosan-treated cells (Fig. 9). Following this rapid increase, the 50 µg ml−1 chitosan-treated cells rapidly recovered the Ca2+ basal level (about 110 nm), whereas in the 200 µg ml−1-treated cells a higher steady state of [Ca2+]cyt (about 440 nm) was reached, suggesting a remarkable alteration of the plasma membrane Ca2+ permeability. The observed Ca2+ transient increase seemed to be specifically induced by Ca2+ re-addition, as the supplying of Ca2+-free medium did not evoke any Ca2+ increase (Fig. 9, inset).
In this paper, soybean suspension cell cultures were treated with exogenous chitosan to investigate its effectiveness in eliciting both early cell responses, such as cytosolic Ca2+ elevation and extracellular H2O2 accumulation, and later events like defence-gene expression and cell death. Administration of increasing chitosan concentrations to aequorin-expressing soybean cells results in a transient increase of [Ca2+]cyt with a reverse dependence on the dose (Fig. 1). The [Ca2+]cyt elevation occurs with unique parameters, which determine the specificity of the Ca2+ transient. Hence, as observed for other elicitors (Clayton et al., 1999; Lecourieux et al., 2002; Navazio et al., 2002), these data provide evidence for the involvement of Ca2+ in the chitosan-mediated signalling.
Incubation with 50 µg ml−1 chitosan gradually induces distinctive changes in cell and nuclear morphology, well-known hallmarks of PCD in plant and animal systems (Danon et al., 2000; McCabe & Leaver, 2000): cellular disorganization, cytoplasmic shrinkage and chromatin condensation. By contrast, when higher doses of chitosan were applied, cells appeared completely disrupted already after 4 h of treatment.
The results of the caspase 3 assay highlight the induction of this enzyme activity by the lowest chitosan dose already after 1 h of treatment, with the caspase 3 inhibitor Ac-DEVD-CHO preventing this increase. In animal cells the cysteinyl aspartate-specific proteases are the major physiological executors of apoptosis (Chang & Yang, 2000). Although previous data have suggested an involvement of serine and cysteinyl proteases in plant PCD, only recently has a definite role for caspase-like machinery in plant PCD been recognized (Elbaz et al., 2002; Woltering et al., 2003). During xylogenesis cysteine and serine protease genes are induced (Ye & Yarner, 1996) and the differentiation of tracheary elements in Zinnia elegans involves a 40-kDa serine protease (Groover & Jones, 1999). Moreover, cysteine proteases have been shown to be involved in oxidative stress-induced PCD in soybean cells (Solomon et al., 1999) and caspase 3-like activation has been observed in tobacco (Tian et al., 2000), barley (Korthout et al., 2000) and tomato (De Jong et al., 2000) suspension cell cultures in response to different elicitors. Chitosan has been widely used in the medical field as a causative agent of apoptosis through activation of caspase 3-like activity in bladder tumour cells (Hasegawa et al., 2001). In keeping with all these findings, our data highlight a role for caspase 3-like protease in chitosan-induced PCD and suggest the possible activation by chitosan of similar biochemical pathways in both animal and plant cells.
By contrast to the degradation of DNA to nucleosomal fragments observed in several plant PCD processes (Ryerson & Heath, 1996; Xu & Heath, 1998), no detectable DNA fragmentation was observed in soybean cells undergoing chitosan-induced PCD in the considered time interval. Nevertheless, the need for internucleosomal DNA fragmentation for the complete accomplishment of apoptosis is still controversial and PCD occurring without DNA-laddering has been noticed (Mittler & Lam, 1995; Dangl et al., 1996; Fukuda, 2000).
Interestingly, chloroplast morphology is seriously compromised by chitosan treatment. Similar to what has been found during chilling stress (Kratsch & Wise, 2000), chitosan induces a stroma swelling, which seems to precede nuclear morphological changes and to increase with the length of the treatment. An open question is whether chloroplasts participate in the PCD pathway induced by chitosan or are just one of the targets of the PCD program. Different alterations in the chloroplast morphology have been previously shown to occur during plant responses to biotic and abiotic stresses (Mittler et al., 1997; Pedroso & Durzan, 2000).
Our results suggest that the signal transduction pathway leading to caspase 3-like dependent cell death following administration of low doses of chitosan is likely to be Ca2+-mediated. Indeed, the complete block of the chitosan-induced Ca2+ transient by chelation of Ca2+ in the culture medium with EGTA prevents both early and late downstream responses, such as extracellular H2O2 accumulation and caspase 3-like activation, but not cell death. A similar behaviour is obtained also with 200 µg ml−1 dose: in this case a prominent disturbance of the plasma membrane integrity, rather than a differential stimulus perception and transduction, seems to be the trigger of cell death. It has been reported that treatments that disrupt plasma membrane integrity are often accompanied by alterations of cell Ca2+ signalling (Pizzo et al., 2002). It is also known that the polycationic nature of chitosan may lead to membrane disturbance through its interaction with negatively charged membrane phospholipids (Shibuya & Minami, 2001). Notably, the experimental device that we used to demonstrate plasma membrane alterations in 200 µg ml−1 chitosan-treated cells highlighted the occurrence of a rapid and steep Ca2+ transient in both control and treated cells upon Ca2+ re-addition to cells deprived of external Ca2+. Considering that removal of Ca2+ from the culture medium causes depletion of intracellular Ca2+ pools in tobacco cells (Cessna & Low, 2001), it can be hypothesized that the intracellular Ca2+ increase observed when Ca2+ was re-added is likely to be caused by a store-operated Ca2+ influx. This capacitative Ca2+ entry has been well characterized in animal cells (Parekh, 2003) but is still poorly considered in plant cells.
On the whole our data suggest that chitosan may induce different signalling pathways in soybean cells depending on its concentration and on the induction of cytosolic Ca2+ elevations. Thus, Ca2+ may play a key role in determining the switching among different chitosan-induced cell death pathways.
The authors are grateful to G. Neuhaus (Center for Applied Biosciences, University of Freiburg, Freiburg, Germany) for kindly providing the transgenic soybean cells expressing cytosolic aequorin, to M. Terbojevich (Dipartimento di Chimica Organica, Padova, Italy) for helpful advice concerning the determination of the chitosan polymerization degree, and to P. Pizzo (Dipartimento di Scienze Biomediche, Padova, Italy) for fruitful suggestions and discussion on EGTA experiments. This work was supported by COFIN 2001 prot. 2001055885 and FIRB 2002 prot. RBNE01K2E7.