Introduction

The ability to regenerate protoplasts into plantlets provides a multitude of opportunities for crop improvement, including a system for protoplast fusion (somatic hybridization), somaclonal variation, and plant transformation. In addition, plant protoplasts are an efficient experimental model for physiological and molecular studies (Vasil 1987). Despite being provided with an osmoprotectant and optimal pH in the external medium, protoplasts isolated by enzymatic methods display recalcitrance and often exhibit a change in their cellular redox environment, which may initiate a cascade leading to programmed cell death (PCD). In higher plants, PCD plays a vital role in biogenesis and morphogenesis. Similar to apoptosis in animal cells, some biotic and abiotic stimuli trigger the death pathway in plants, resulting in mitochondrial dysfunction, ROS generation, and the activation of certain proteases (Greenberg 1996; Rogers 2006). In particular, PCD has been related to aleurone and root cap cell elimination, somatic embryogenesis, leaf and petal senescence, xylogenesis, reproduction (Greenberg 1996; Pennell and Lamb 1997), and aerenchyma formation (Drew et al. 2000). Moreover, PCD is enhanced in response to biotic and abiotic stresses (Greenberg 1996; Pennell and Lamb 1997; Beers and McDowell 2001; Lam et al. 2001). In plant cells, adverse environmental stresses can increase ROS levels due to perturbations of the cellular redox balance, leading to oxidative damage and subsequent cell death (Apel and Hirt 2004; Vacca et al. 2006; Gao et al. 2008). The production of ROS, especially H2O2, and mitochondrial dysfunction are necessary components and important indicators of PCD in response to various stimuli (Gao et al. 2008; Zhang and Xing 2008; Zhang et al. 2009b). H2O2 acts in a dose-dependent manner and synergistically with nitric oxide in hypersensitive cell death (Delledonne et al. 2001; Zago et al. 2006). It has also been reported that H2O2 levels are increased during senescence of the Phalaenopsis flower (a slowly senescing flower), due to elevated activity of xanthine oxidase, an enzyme capable of O2 ·− generation (Tewari et al. 2009).

Major sources of H2O2 in plant cells include the leakage of high-potential electrons from the saturated electron transport chains of chloroplasts and mitochondria, the Mehler reaction, a wide variety of limited substrate oxidases, NADPH oxidase (NOX), and type III peroxidase (Halliwell and Gutteridge 1999; Cheeseman 2007; Doyle et al. 2010). Peroxisomes are also included among the sites of intracellular H2O2 production (Cheeseman 2007). NADPH oxidase, a plasma membrane protein, oxidizes NADPH at the cytosolic surface and reduces O2 to O2 ·− at the outer surface of the plasma membrane (Sagi and Fluhr 2006; Zhang et al. 2009a), which spontaneously, or via superoxide dismutase (SOD), is dismutated to H2O2 (Cheeseman 2007). Recent studies have indicated that RBOH A and B (respiratory burst oxidase homologue, NOX genes) from Nicotiana benthamiana function in elicitor-induced NO and H2O2 generation and stomatal closure, but not in elicitor-induced free radical generation, hypersensitive cell death, or activation of the pathogenesis-related gene, PR1 (Zhang et al. 2009a).

Brassica napus leaf protoplasts initiate senescence during isolation and senesce thereafter without cell division (Watanabe et al. 1998, 2002a). B. napus leaf protoplasts die through PCD or apoptosis, which is similar to that observed in animal cells with a few morphological differences (Watanabe et al. 2002b). H2O2 has been reported to be involved in PCD (Zhang et al. 2009b; Doyle et al. 2010). ROS generation and antioxidant enzyme activities had been analyzed before (Papadakis et al. 2001). Yasuda et al. (2007) previously tried to inhibit ROS generation by including DPI (diphenyleneiodonium chloride) in a macerating enzyme (ME) solution but such an addition had no effect on H2O2 accumulation in the chloroplasts of protoplasts. DPI seems to be degraded or inactivated in MEs during the incubation period and does not lead to cell divisions of protoplasts. In the present study, we focused on the process of H2O2 generation in B. napus leaf tissue, chloroplasts and protoplasts and presented experimental evidence for the existence of DPI-sensitive NADPH oxidase-like activity and inhibition of H2O2 accumulation in the chloroplasts. Nonetheless, we also provided flow cytometric analysis of temporal changes in mitochondrial membrane potential as an index of apoptotic cell death in B. napus leaf protoplasts.

Materials and methods

Plant material, protoplast isolation, and culture

Seeds of Brassica napus L. cv. Bronowski were obtained from the National Institute of Agrobiological Resources (Tsukuba, Japan). Plants were grown in a growth chamber under controlled light flux (50 μmol m−2 s−1), photoperiod (14 h), and temperature (day/night cycle of 25/20°C) conditions. Protoplasts were prepared from the leaves of 6- to 8-week-old plants (Watanabe et al. 1992). Sterilized B. napus leaves were cut into narrow strips and incubated in a macerating enzyme (ME) solution containing 5 mM Mes (pH 5.8) with 0.6 M sorbitol, 5 mM CaCl2, 0.5% cellulase Y-C (Kyowa Kasei, Osaka, Japan) and 0.03% pectolyase Y-23 (Kyowa Kasei), for different duration (3, 6 and 12 h). Incubating leaf strips in ME for 3 or 6 h did not remove cell wall completely and exhibited oxidant and antioxidant responses similar to those isolated by 12 h exposure in ME. Thus, protoplast isolated by 12 h exposure in ME was used for further study. For corresponding controls, leaf strips were incubated in the 5 mM Mes buffer (pH 5.8) with 0.6 M sorbitol and 5 mM CaCl2 (without ME). The isolated protoplasts (5 × 105) were cultured in Petri dishes in MS medium (Murashige and Skoog 1962) supplemented with 4.5 mM 2,4-dichlorophenoxyacetic acid (2,4-D), 2.2 mM benzyl adenine (BA), 2% sucrose, 1% glucose and 0.5 M sorbitol (pH 5.8). The bottom of each dish was layered with 1.5% agar to prevent the adhesion of protoplasts. The dishes were maintained in darkness at 25°C.

Confocal microscopic and flow cytometric examinations of the mitochondrial membrane potential

For the measurement of mitochondrial membrane potential, 10 μM 3,3′-dihexyloxacarbocyanine iodide (DiOC6, Molecular Probes, Eugene, OR, USA) was added to a protoplast suspension (5 × 105). After incubation at 25°C for 30 min, the protoplasts were washed twice with 10 mM phosphate buffer containing 0.6 M sorbitol, resuspended in the same buffer, and observed under a Leica confocal laser scanning system (Leica TCS-SP2; Leica Microsystems, Tokyo, Japan) using laser excitation at 488 nm and emission at 540 nm to visualize the fluorescent probe.

Protoplasts were analyzed using an FC500 flow cytometer (Beckman Coulter, Tokyo, Japan) to monitor changes in the mitochondrial membrane potential. For all flow cytometric analyses, unstained protoplasts, collected at the time of protoplast isolation or from the same culture time, were used as autofluorescence controls. The term ‘FL1’ with the channel number refers to the quantified fluorescence at 525 nm. The fluorescence intensity was plotted on a log scale with maximum and minimum fluorescence levels of 1,000 and 1, respectively. The peak height and area in the histograms represent the number of protoplasts. Linearity of the flow cytometer was ensured using Flow Check beads with the QC 1L Flow Check protocol (Beckman Coulter). To measure the mitochondrial membrane potential, samples were stained with 500 nM DiOC6 for 30 min in the dark and analyzed immediately. DiOC6 was excited with the 488-nm line of an argon ion laser, and the fluorescence signals were detected by the FL1 detector. At least 10,000 protoplasts were analyzed using the peak height or time versus the integral of the DiOC6 fluorescence signal. Data were analyzed using Cytomics FC500 RXP software (Beckman Coulter).

Genomic DNA fragmentation

Genomic DNA of protoplasts was isolated using a Floraclean plant genomic DNA isolation kit (BIO 101, Vista, CA, USA), according to the instruction manual. DNA was separated in a 2% agarose gel, stained with ethidium bromide, and photographed under a UV-transilluminator.

Localization of H2O2 production

H2O2 production was measured by monitoring the fluorescence of dichlorofluorescein (DCF), which is the oxidation product of H2DCF (Allan and Fluhr 1997). As cell-permeant indicator of H2O2 2′,7′-dichlorodihydrofluorescein diacetate (H2DCF-DA; Molecular Probes) is non-fluorescent until the acetate groups are removed by intracellular esterases and oxidation occurs within the cell. The protoplasts were washed, resuspended in 0.6 M sorbitol in 5 mM Mes buffer (pH 5.8) and 5 mM CaCl2, and loaded with either 70 μM H2DCF-DA for 30 min in darkness. After washing the protoplasts with 0.6 M sorbitol in 5 mM Mes buffer and 5 mM CaCl2 the fluorescence signals from DCF and chlorophyll were observed at emission wavelengths of 494–559 and 680–730 nm, respectively, with an excitation wavelength of 488 nm in a Leica confocal laser scanning system (TCS-SP2; Leica). Microscope settings, for example, laser power (fixed by marking), the pinhole diameter, the objective lens, and the setting of the photomultipliers (fixed by noted digital values) were keep essentially same for individual set experiments. Excitation and emission bandwidth was saved as a file on OS for further operation with same settings. Experiment was repeated several times and relative fluorescence intensity of representative images was measured by selecting region of interest (whole cell) using ImageJ 1.42 software (http://rsb.info.nih.gov/ij/) and data was normalized against background.

Chloroplast isolation from protoplasts

Chloroplasts were isolated as described by Kieselbach et al. (1998), with minor modifications. Briefly, the protoplasts were suspended in the chloroplast isolation medium containing 0.4 M sorbitol, 50 mM Hepes (pH 7.8), 1 mM MgCl2, 10 mM KCl, 1.0 mM EDTA, and 0.15% (w/v) bovine serum albumin. To release the chloroplasts, the protoplasts were broken by passage through 20-μm nylon net that was attached over the tip of a 5.0-ml disposable plastic syringe (without needle). The protoplasts were twice drawn into the syringe through the nylon net and dispersed. Microscopic observation revealed that this treatment resulted in the complete rupture of the protoplasts and the release the chloroplasts. The chloroplasts were sedimented by centrifugation of the protoplast homogenate at 1,000g for 1.5 min, and then washed with chloroplast isolation medium and extracted in enzyme extraction buffer. For purification, chloroplast suspension was loaded on 35% Percoll and centrifuged at 2,000g for 10 min. Pellet was re-suspended in the chloroplast isolation medium and centrifuged at 1,000g for 1.5 min to sediment chloroplasts.

Enzyme extraction, protein, and plasma membrane marker determination

Young leaves strips incubated in the 5 mM Mes buffer (pH 5.8) with 0.6 M sorbitol and 5 mM CaCl2 (without ME) were homogenized in 10 ml of chilled 50 mM potassium phosphate buffer (pH 7.0) containing 0.5% (w/v) insoluble polyvinylpolypyrrolidone and 1 mM phenylmethylsulfonylfluoride, using a chilled pestle and mortar on an ice bath. The homogenate was filtered through two-fold miracloth, followed by centrifugation at 15,000g for 10 min at 4°C. The supernatant was stored at 4°C and used for enzyme assays within 4 h. For the assay of APX activity, 5.0 mM ascorbic acid was also included in the extraction medium. Protoplast and chloroplast suspensions were homogenized in the same extraction medium using a Dounce homogenizer with a tight-fitting Teflon pestle, followed by centrifugation as described above. Protein concentrations were determined according to the method of Bradford (1976). Protoplast and chloroplasts extracts were assayed for vanadate-inhibited, Mg2+-ATPase and 5′-nucleotidase activities respectively after Truitt et al. (2004) and Schimmel et al. (1973).

Enzyme assays

NADPH oxidase-like (NOX-like, EC 1.6.3.1) enzyme activity was determined as described previously (Tewari et al. 2009). This modified assay is based on the reduction of 2,3-Bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide sodium salt (XTT) by O2 ·− generated by the oxidation of NADPH. The reaction mixture contained 50 μl of enzyme extract, 0.3 mM XTT, 0.2 mM NADPH, 0.1 mM MgCl2, and 1.0 mM CaCl2 in 1 ml of 50 mM Tris–HCl (pH 7.4) except for where it stated otherwise. The reaction was initiated by the addition of NADPH, and the change in absorbance at 470 nm was recorded. NADPH oxidase-like activity was presented as unit (μmol XTT reduced min−1) mg−1 protein.

Superoxide dismutase (SOD, EC 1.15.1.1) activity was assayed by measuring its ability to inhibit the photochemical reduction of nitro-blue tetrazolium (NBT) at 560 nm. The reaction mixture (5.0 ml) contained 25 mM phosphate buffer (pH 7.8), 65 μM NBT, 2 μM riboflavin, enzyme extract, and 15 μl of N,N,N′,N′-tetramethylethylenediamine (TEMED; modified from Beauchamp and Fridovich (1971). The reaction mixture was exposed to light (350 μmol m−2 s−1) for 15 min. The amount of SOD corresponding to 50% inhibition of the reaction was defined as one unit of enzyme.

Catalase (CAT, EC 1.11.1.6) activity was measured after Beers and Sizer (1952). Briefly, 25 μl of the extract was added to 3 ml of 5 mM H2O2 in 50 mM potassium phosphate (pH 7.0). The decrease in absorbance at 240 nm was monitored for 3 min, and the linear part of the curve was used to quantitate the rate of decrease by using an extinction coefficient of H2O2 at 240 nm of 0.0435 mM−1 cm−1. CAT activity was presented as unit (μmol H2O2 decomposed min−1) mg−1 protein.

Ascorbate peroxidase (APX, EC 1.11.1.11) was measured in a 3.0 ml reaction mixture containing 50 mM phosphate buffer (pH 7.0), 0.5 mM AsA, 0.1 mM H2O2, 0.1 mM EDTA, and a suitable quantity of enzyme extract. The blanks included reactions without tissue extract and without H2O2. The change in absorbance at 290 nm was measured every 15 s (Nakano and Asada 1981). The activity of APX was expressed as unit (μmol AsA oxidized min−1) mg−1 protein).

Native PAGE and visualization of enzymes on native gels

Enzyme isoforms were separated by polyacrylamide gel electrophoresis in a discontinuous gel under non-denaturing conditions, as described by Hames (1990). Native gels were run at constant current (30 mA) at 4°C. NOX, SOD, and guaiacol peroxidase (POD) were visualized on native gels as described previously (Tewari et al. 2008). NADPH oxidase-like activity was visualized on a 12.5% native gel by soaking the gel in a reaction mixture containing 50 mM Tris–HCl (pH 7.4), 0.2 mM NBT, 0.1 mM MgCl2, 1 mM CaCl2, and 0.2 mM NADPH in the absence and presence of 200 U of SOD (O2 ·− scavenger) and 100 μM DPI (NOX inhibitor).

Superoxide dismutase (SOD) activity was visualized after Beauchamp and Fridovich (1971) by soaking the gel in 2.45 mM NBT for 20 min, followed by immersion in a solution containing 28.0 mM TEMED, 0.028 mM riboflavin, and 36 mM potassium phosphate buffer (pH 7.8). The gel was then placed on a clean glass plate and illuminated under cool fluorescent light for 2–3 min.

Guaiacol POD was separated in a 12.5% native gel and was visualized by soaking the gel in a reaction mixture containing 20 ml of 0.1 M phosphate buffer (pH 7.0), 4 ml of 0.01% (v/v) H2O2, and 4.0 ml of 0.5% (w/v) guaiacol for 20 min. For the detection of DAB-POD activity gel was stained in a reaction mixture containing 0.1 M sodium acetate buffer (pH 5.5) containing 1 mM 3,3-diaminobenzidine (DAB) and 0.03% H2O2 (Wang et al. 2009). All gels were rinsed with distilled water and then imaged.

Ascorbate, dehydroascorbate and lipid peroxidation

Fresh leaf tissue (250 mg) was homogenized in 2.0 ml of 10% (w/v) trichloroacetic acid, followed by centrifugation at 10,000g for 5 min. Total ascorbate, which was determined after the reduction of dehydroascorbate (DHA) to AsA by dithiothreitol (DTT) and then the neutralization of the excess DTT by N-ethylmaleimide, and AsA in the supernatant were measured as the Fe+2-bipyridyl complex (A max at 525 nm) formed from the reduction of Fe+3 by AsA, according to the method of Law et al. (1983). The DHA content was calculated from the difference between total ascorbate and AsA. Lipid peroxidation was determined by method of Heath and Packer (1968) in terms of malondialdehyde (MDA) content by thiobarbituric acid (TBA) reaction. The amount of TBA reactive substance (TBARS) was calculated from the difference in absorbance at 532 and 600 nm using extinction coefficient of 155 mM−1 cm−1.

Statistical analysis

All results are means of experimental replicates (at least n = 6) of protoplast/chloroplast images or protoplast/chloroplast preparations as specified in captions. The data were analyzed by analysis of variance (ANOVA) and tested for significance by Bonferroni t test using Sigma-stat software.

Results

Temporal effect of exposure of ME on oxidative metabolism

Incubating leaf strips in ME for 3 or 6 h did not dissolve cell wall completely as revealed by non-circular cell shape (Fig. 1a). Moreover, irrespective of exposure duration (3, 6, or 12) in ME, cells/protoplasts showed an increase in H2O2 (Fig. 1a, b) and MDA accumulation (Fig. 1c), higher NOX-like enzyme activity and lower SOD and APX (Fig. 2a, b, d) activities. Macerating enzyme did not cause any significant change in CAT activity (Fig. 2c). Therefore, we performed experiments on B. napus leaf protoplasts isolated by 12 h exposure in macerating enzyme.

Fig. 1
figure 1

Hydrogen peroxide (H2O2) generation and lipid peroxidation in B. napus leaf protoplasts/cells obtained by differential exposure duration in macerating enzyme (ME). Protoplasts were prepared and stained as described in “Materials and methods”. a Cell/Protoplasts were isolated at different time intervals and either not stained (Blank) or loaded with 70 μM H2DCF-DA. Images obtained in green, red and transmission channels were merged and presented. b Quantitative analysis of the relative DCF fluorescence intensity from representative images. c Lipid peroxidation in B. napus leaf strips (−Macerating enzyme) or isolated cell/protoplasts (+Macerating enzyme) obtained by differential exposure duration with ME. Bars represent mean ± SE of ROIs (region of interest i.e. whole cell) of different protoplasts. Bars having different letters are significantly different by Bonferroni t test (P < 0.05)

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Fig. 2
figure 2

NADPH oxidase, NOX (a), superoxide dismutase, SOD (b), catalase (c) and ascorbate peroxidase, APX (d) activities in the B. napus leaf strips (−Macerating enzyme) or protoplasts/cells (+Macerating enzyme) obtained by differential exposure duration (0, 3, 6, 12 h). Vertical bars represent the mean ± SE (n = 6). Bars having different letters are significantly different by Bonferroni t test (P < 0.05)

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B. napus leaf protoplasts undergo apoptotic-like cell death

B. napus leaf protoplasts undergo apoptotic cell death as indicated by decreased mitochondrial membrane potential and DNA fragmentation. DiOC6 (a carbocyanine dye accumulates in active mitochondria) (Fig. 3a) was used to monitor the mitochondrial membrane potential by flow cytometry (Fig. 3b). The mitochondrial membrane potential decreased with culture time. Only 8.4% of protoplasts were detected in the lower fluorescence intensity region (T) after 10 h of culture, whereas more than 82% of protoplasts were detected in the same region after 60 h. No DNA fragmentation was detected in the protoplasts cultured for 10 h; however, DNA fragmentation was induced during 10–24 h of culture, and advanced further with culture period, 48 h (Fig. 3c).

Fig. 3
figure 3

Mitochondrial membrane potential of B. napus leaf protoplasts. a DiOC6-stained protoplasts showing DiOC6-specific fluorescence image, chlorophyll fluorescence image, and transmission image under a confocal laser scanning microscope. b Flow cytometric analysis of the changes in mitochondrial membrane potential of B. napus leaf protoplasts during culture. T and U regions represent the percentages of cells showing non-specific fluorescence and DiOC6-specific fluorescence (mitochondrial membrane potential), respectively, at the time points indicated. Non-stained protoplasts served as the blank. c Agarose gel analysis for nuclear DNA fragmentation in B. napus leaf protoplasts cultured for 10, 24, and 48 h

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Hydrogen peroxide generation in protoplasts

H2O2 was generated excessively in the protoplasts during isolation. H2O2 signal was very weak in the transverse sections of leaf tissue used for protoplasts isolation (Supplementary Fig. S1). The highest DCF-fluorescence, an index of H2O2 accumulation, was detected in the protoplasts isolated by 6 or 12 h exposure in ME on the day of protoplast isolation (Fig. 1a, b). The level of DCF-fluorescence although declined gradually with culture time but decrease was non-significant (Fig. 4a, b). When 100 μM DPI was added to the protoplast or chloroplast suspension before the addition of 70 μM H2DCF-DA, an inhibition of DCF fluorescence was observed in both chloroplasts and protoplasts (Fig. 5 a, b).

Fig. 4
figure 4

Hydrogen peroxide (H2O2) generation in B. napus leaf protoplasts during culture. Protoplasts were prepared, cultured, and stained as described in “Materials and methods”. a Protoplasts were harvested at different time intervals and either not stained (Blank) or loaded with 70 μM H2DCF-DA. Images obtained in green, red and transmission channels were merged and presented. b Quantitative analysis of the relative DCF fluorescence intensity from representative images. Bars represent mean ± SE of ROIs (region of interest i.e. whole cell) of different protoplasts. Bars having different letters are significantly different by Bonferroni t test (P<0.05)

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Fig. 5
figure 5

Effect of a NADPH oxidase inhibitor (100 μM DPI) on hydrogen peroxide (H2O2) generation in B. napus leaf chloroplasts and protoplasts. Chloroplasts and protoplasts were prepared and stained as described in “Materials and methods”. a Chloroplasts and protoplasts were either not stained (Blank) or loaded with 70 μM H2DCF-DA in the absence or presence of 100 μM DPI. b Quantitative analysis of the relative DCF-fluorescence intensity (an index of in vivo H2O2 generation) from representative images. Bars represent mean ± SE of ROIs (region of interest i.e. whole chloroplast or cell) of different chloroplasts/protoplasts. Bars having different letters are significantly different by Bonferroni t test (P < 0.05)

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NADPH oxidizing, DPI-sensitive NOX-like enzyme is involved in H2O2 generation

The inhibition of DCF fluorescence (Fig. 5) and NOX-like enzyme activity (Fig. 6a, d) in the presence of 100 μM DPI (NOX inhibitor) revealed NOX-like enzyme-mediated generation of O2 ·−, which subsequently dismutated and accumulated as H2O2 in the chloroplasts and protoplasts. The NOX-like enzyme-mediated generation of O2 ·− in chloroplasts was confirmed by the visualization of NOX-like enzyme activity on a native gel in the presence of 200 U of SOD (Fig. 6c). The activity of the NOX-like enzyme isoforms disappeared almost completely in the presence of SOD and 100 μM DPI (Fig. 6c, d). To rule out the possibility of plasma membrane contamination in chloroplasts fractions, we prepared chloroplasts as described in the “Materials and methods”, also purified with 30% Percoll gradient and monitored for marker enzymes of plasma membrane; vanadate sensitive ATPase and 5′-nucleotidase. None of these plasma membrane enzymes activities were detected in chloroplast fraction but it showed presence of NOX-like enzyme activity (Fig. 6e).

Fig. 6
figure 6

NADPH oxidase-like (NOX-like) activity of leaf (L), protoplasts (P) and chloroplasts isolated from protoplasts (PC) measured in cuvette (a) or in native gels in the absence (b) or presence of 200 U of SOD (c) or 100 μM DPI (d). NOX-like activity in leaf (L) and Percoll purified chloroplasts (PPC) (e). Superoxide dismutase (SOD) (f) and guaiacol peroxidase (G-POD) (g) and 3,3-diaminobenzidine peroxidase (DAB-POD) (h) activities in native gels. Extracts of leaf (L), protoplasts (P), and chloroplasts isolated from protoplasts (PC) of B. napus were separated by native PAGE. Leaf extract of Petunia hydrida was considered as positive control for G-POD. Enzyme activities were visualized using standard protocols as described in “Materials and methods”

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Ineffective cellular antioxidant system provides poor protection against oxidative stress

The activity of the antioxidant enzymes SOD was lower in the protoplasts compared with the activities in the leaf extract of B. napus (Fig. 2b). Seven SOD isoforms were detected in the leaf and protoplast extracts on native gels; however, the intensities of the SOD isoforms in the protoplasts were lower than those in the leaf extract (Fig. 6f). The activity of APX was lower in protoplasts compared to leaf (Fig. 2d). No G-POD isoforms were detected in the leaf extract, protoplasts, or chloroplasts of B. napus on a native gel (Fig. 6g), although two isoforms of G-POD were detected in the leaf extract of Petunia hybrida (which considered to be a positive control for G-POD) on the same gel (Fig. 6g). Activity of DAB-POD was reduced in protoplasts and chloroplasts did not show DAB-POD activity (Fig. 6h). These reductions in the activities of APX and DAB-POD and activation of NOX-like enzyme resulted in the accumulation of H2O2 and lipid peroxidation. While protoplasts isolated by 3 h exposure in ME did not show any significant change in ascorbate concentration, the protoplasts isolated by 6 or 12 h exposure in ME enhanced AsA concentration compared to the corresponding control leaves (Fig. 7a, b). The AsA/DHA ratio was higher in the protoplasts isolated by 12 h exposure in ME solution (Fig. 7b).

Fig. 7
figure 7

Concentrations of ascorbic acid (AsA) and total ascorbate (TAsc) (a), and the AsA/DHA ratio (b) in the leaf strips (−Macerating enzyme) or cells/protoplasts obtained by differential exposure period with ME (+Macerating enzyme) of B. napus plants. Vertical bars represent the mean ± SE (n = 6). Bars for an individual parameter having different letters are significantly different by Bonferroni t test (P < 0.05)

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Discussion

Mitochondrial dysfunction is characteristic of apoptotic cells (Zhang et al. 2009b). The ROS generated initially in chloroplasts may induce mitochondrial dysfunction. The loss of mitochondrial membrane potential (mitochondrial dysfunction) accompanied by DNA fragmentation in the B. napus leaf protoplasts suggests that the protoplasts underwent apoptosis during culture.

The occurrence of the highest DCF fluorescence, an index of H2O2 generation (Kristiansen et al. 2009), in B. napus leaf protoplasts on the first day of their isolation suggests that the ME solution probably induced a form of metabolic redox disturbance during protoplast isolation, and this probably induced excessive ROS generation and oxidative stress. The observed increase in the generation of O2 ·− and H2O2 in B. napus leaf protoplasts is consistent with the findings of Ishii (1987) and Papadakis et al. (2001). H2DCF-DA has also been reported to react with O2 ·− (Allan and Fluhr 1997; Kristiansen et al. 2009); however, it should also be taken into account that observed intense fluorescence in B. napus protoplasts might be an outcome of reactivity of both O2 ·− and H2O2 with fluorescence probe (H2DCF-DA). Superoxide itself is highly reactive and rapidly dismutated to H2O2. Therefore, it generally leads to the low persistence of O2 ·− in chloroplast. The inhibition of DCF fluorescence in chloroplasts or protoplasts upon incorporation of 100 μM DPI, an inhibitor of NADPH oxidase (NOX), before the addition of H2DCF-DA suggests NOX-like enzyme-mediated generation of O2 ·− in chloroplasts, which subsequently dismutated to H2O2 and accumulated in the chloroplasts. Thus, H2O2 is produced by spontaneous or SOD-mediated dismutation (to a lower extent) in B. napus leaf protoplasts. Extent of H2O2 accumulation in the protoplasts isolated by 6 or 12 h exposure in the ME was almost similar. They also exhibited a similar response for the activities of NOX-like enzyme, APX and SOD. Excessive accumulation H2O2 in protoplasts suggests a response similar to wounding or pathogenic infection leading to apoptosis-like cell death (Zago et al. 2006; Chaki et al. 2009). Commercially available ME (cellulase and pectolyase) used for the isolation of protoplasts either contain some elicitor (because they obtained from fungal sources) or that generated in the process of cell wall dissolution which might have triggered oxidative burst (ROS generation) in the protoplasts.

Antisense knockouts and gene silencing experiments in tobacco as well as an analysis of Arabidopsis mutants have confirmed the role of NOX as a source of O2 ·− (Simon-Plas et al. 2002) in the oxidative burst that regulates cell death (Torres et al. 2002; Yoshioka et al. 2003). H2O2 is a relatively stable product of spontaneous or SOD-catalyzed dismutation of O2 ·−. Therefore, this study focused on H2O2 instead of O2 ·−. In gel localization revealed higher NOX-like activity, particularly in the B. napus chloroplasts and protoplasts compared with the leaf, suggests the involvement of a NOX-like enzyme in O2 ·− generation in the chloroplasts. The activity of NOX-like enzyme isoforms disappeared completely in the presence of SOD (200 U) or DPI (50 μM). These observations suggest the existence of a NOX-like protein in B. napus leaf chloroplasts. The most promising candidate in the chloroplasts, which may exhibit NOX-like enzyme activity is small ferredoxin:NADP+ oxidoreductase (FNRs). It has been reported previously that small FNR isoforms of cyanobacterium Synechocystis sp. strain PCC6803, (FNRs ≈ 34 kDa) which corresponds to plastid FNR, is a NADPH oxidase (Thomas et al. 2006; Korn et al. 2009). Moreover, there is a report that supports the existence of NOX in chloroplasts: the hydrogen peroxide generated by a Mn2+-dependent enzyme in the chloroplasts of Euglena has been shown to be involved in the decarboxylation of glyoxylate (Yokota et al. 1983). However, no report is yet available on NADPH oxidase-like activity in the chloroplasts.

The reduced activities of the antioxidant enzymes SOD and POD in the protoplasts and chloroplasts compared with the leaf extract suggest a weakening of antioxidant protection in B. napus protoplasts. The decreased DAB-POD activity and lack of G-POD probably caused an over-accumulation of H2O2 in the B. napus leaf protoplasts due to a lower antioxidant potential of the cells. SOD is an O2 ·−-dismutating antioxidant enzyme and is present in various cellular compartments such as chloroplasts, mitochondria, and cytosol (Mittler et al. 2004). SOD generally scavenges the O2 ·− generated under normal conditions and dismutates O2 ·− into H2O2 (Mittler et al. 2004). H2O2 can diffuse across the membrane or move through water channels (Bienert et al. 2007) and is scavenged generally by the participation of various antioxidant enzymes such as catalase, non-specific peroxidase, ascorbate peroxidase, and glutathione peroxidase (Cheeseman 2007). The superoxide anion, generated by excessive NOX-like activity in the chloroplasts, was dismutated to H2O2, spontaneously and/or via chloroplastic SOD activity. However, H2O2 was not scavenged by the available cellular antioxidants because of the undetectable G-POD and reduced DAB-POD activities in B. napus protoplasts. An ineffective antioxidant system as seen in B. napus protoplasts, may be characteristic of apoptosis of protoplasts. Previously, Watanabe et al. (2002a) have reported a high protease activity and excessive production of ethylene gas in apoptotic protoplasts of B. napus. Papadakis et al. (2001) have been reported reduced activity of the cellular antioxidants in the apoptotic protoplasts. They also reported a lower AsA/AsA+DHA ratio in non-totipotent protoplasts compared with the leaf, indicating the accumulation DHA. However, our observations with B. napus protoplasts are not in agreement with the findings of Papadakis et al. (2001), as B. napus protoplasts displayed enhanced AsA, lower DHA, and a higher AsA/DHA ratio. Generally, AsA has been considered as an efficient free radical scavenger (Arrigoni and De Tullio 2002); however, AsA exerts pro-oxidant effects as well (Podmore et al. 1998; Arrigoni and De Tullio 2002). AsA exerts pro-oxidant effects in vitro, usually by interactions with transition metal ions to promote its oxidation, accompanied by H2O2 production (Halliwell 2001). Moreover, an enhanced apoptotic-like cell death has been reported in the cell culture of Arabidopsis exposed to heat stress by ascorbic acid or sodium ascorbate treatment (Doyle et al. 2010).

In summary, the isolation of protoplasts using macerating enzymes is a stress-inducing process and results in the accumulation of H2O2 in chloroplasts due to the activation of NOX-like activity. The inactivation of DAB-POD and lack of guaiacol POD activity appears to be another cause of sustained H2O2.accumulation. The sustained generation of H2O2 induced the apoptotic cell death of B. napus leaf protoplasts. Therefore, promising approach would be to suppress NOX-mediated ROS generation to control apoptotic cell death of protoplasts. Inhibition of NOX-like enzyme-mediated generation of H2O2 may be implied in the regulation of apoptotic-like death of B. napus protoplasts. However, further experiments are required to reach on this conclusion.