S-RNase disrupts tip-localized reactive oxygen species and induces nuclear DNA degradation in incompatible pollen tubes of Pyrus pyrifolia

Pollen tubes show strictly polar cell expansion called tip growth, which is similar to that in root hairs. In some plant species, pollen tube growth rate can reach micrometers per second (Stone et al., 2004). To achieve extremely fast growth, pollen tubes have a high energy requirement that requires rapid oxygen uptake (Tadege and Kuhlemeier, 1997), and the free cytosolic calcium ([Ca²⁺]_cyt) gradient has a crucial role in modulating polar elongation. Recent studies showed that reactive oxygen species (ROS) were a requirement for root hair growth. Foreman and colleagues (Foreman et al., 2003) used a loss-of-function knockout mutant in AtrbobC/RHD2 to demonstrate that ROS were indispensable for root hair growth and were required to stimulate Ca²⁺ influx during root-hair elongation. AtrbobC/RHD2 encodes a superoxide (O₂•⁻)-producing NADPH oxidase (NOX) and the mutant had reduced ROS formation at the tip of very short root hairs. Subsequently, Monshausen and colleagues (Monshausen et al., 2007) detected oscillations of apoplastic ROS concentration at the top of the flanks of root hairs, which revealed a new model of the role of ROS in root hair growth. Otherwise, it was demonstrated that tip-localized ROS produced by a NOX enzyme was needed to sustain the normal rate of pollen tube growth (Potocky et al., 2007). ROS are an inevitable consequence of aerobic metabolism, and have a variety of functions including pathogen defence and cell signalling. In plant cells, there are many potential sources of ROS, such as chloroplasts, mitochondria and peroxisomes, and plasma membrane NADPH oxidases, cell wall peroxidases and amine oxidases (Mittler, 2002; Neill et al., 2002). Mitochondria have been considered major ROS producers in animal cells and in plant cells without chloroplasts, such as pollen tubes. The O₂•⁻ formation in mitochondria is closely related to the coupling efficiency between the respiratory chain and oxidative phosphorylation. Plasma-membrane-bound NADPH oxidase transfers electrons from cytoplasmic NADPH to form O₂•⁻, which undergoes an enzymatic and non-enzymatic dismutation reaction, immediately producing H₂O₂, the most stable form of ROS. ROS formed in the plasma membrane finally accumulate in the cell walls of the pollen tube.

Pear (Pyrus pyrifolia L.) of the family Rosaceae, has an S-RNase-based gametophytic self-incompatibility (SI) system. Using an in vitro system, our team has identified the characteristics of S-RNase that specifically inhibit self-pollen germination and tube elongation (Hiratsuka et al., 2001; Zhang and Hiratsuka, 2000; Zhang and Hiratsuka, 1999). Recently, it was confirmed that S-RNase induces the depolymerization of the actin cytoskeleton and DNA degradation of self-generated pollen tubes (Liu et al., 2007; Wang et al., 2009). To identify whether the in vitro results mirror what happens in vivo, we evaluated the nuclear DNA of pollen tubes after different pollinations. ROS have an important role in pollen tube elongation as mentioned above, and the actin cytoskeleton is a target for ROS in yeast (Perrone et al., 2008). We thus speculated that tip-localized ROS disruption occurs in the SI response of pear.

To assess the effect of ROS on pollen tube growth, the pollen was grown in the basal medium for 1 hour at 25°C, then the NADPH oxidase inhibitor diphenylene iodonium chloride (DPI) and ROS scavenger Mn-5,10,15,20-tetrakis(1-methyl-4-pyridyl)21H,23H-porphin (TMPP) were added to the medium. As expected, DPI and TMPP obviously inhibited pollen tube growth (Fig. 1A), which indicated that ROS are necessary for pear pollen tube growth.

Furthermore, to evaluate the effect of S-RNase on tip-localized ROS, pollen tubes were stained with the ROS fluorescence probe, 5-(and 6-)chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate (CM-H₂DCFDA). Two types of pollen tubes were stained with CM-H₂DCFDA (Fig. 1B): pollen tubes with strongest fluorescence in the tip (from subapical domain to apex) were regarded as normal, whereas pollen tubes with uniform fluorescence suggested that ROS in the tip-localized pollen tube were disrupted. The sample was stained with CM-H₂DCFDA at 30 minutes after S-RNase, DPI or TMPP challenge to count the pollen tube with strongest fluorescence in tip (Fig. 1C). In the control, the proportion of pollen tubes with strongest fluorescence in the tip relative to the total number of tubes was 52.7±2.1% (means ± s.e.; n>100), and with the DPI or TMPP treatments, only 32.4±3.4% (n>100) and 35.9± 4.4% (n>100), respectively. In the compatible treatment, similarly to the control, 52.0±1.4% (n>100) of pollen tubes had strongest fluorescence in the tip. However, in the incompatible treatment, this was only 28.8±1.7% (n>100), nearly half the control value.

Nitroblue tetrazolium (NBT) was also used for visualizing ROS distribution. NBT is reduced by O₂•⁻ to a blue formazan precipitate, indicating the site of O₂•⁻ production. Because NBT was cytotoxic and rapidly kills pollen tubes, NBT staining provides a snapshot of ROS formation in pollen tubes at different stages of the oscillatory growth cycle (Potocky et al., 2007). Fig. 1D shows typical images of pollen tube staining with NBT for different treatments. The pollen tube of the control showed a tip-localized pattern of formazan staining; there was a similar staining pattern in the compatible pollen tubes. However, the tip-localized formazan staining of pollen tubes treated with DPI or self S-RNase was disrupted. The TMPP-incubated pollen tubes were the colour of the TMPP reagent and did not have tip-localized formazan staining. The percentage of pollen tubes with tip-localized formazan staining pattern relative to the total number of tubes in different treatments is shown in supplementary material Fig. S1.

To investigate the subcellular localization of H₂O₂ accumulation in the subapical region of pollen tubes exposed to S-RNase treatment, a CeCl₃ cytochemical technique, which reacts with H₂O₂ to produce electron-dense deposits of cerium perhydroxides (Bestwick et al., 1997), was used. Coincidently, mitochondria were distributed in the same subapical region. Thus the region of mitochondria accumulation was regarded as the subapical region of the pollen tube. H₂O₂ accumulation in mitochondria of pollen tubes is shown in Fig. 2A–F. Control pollen tubes without CeCl₃ staining are shown in Fig. 2A. The subapical region of a pollen tube grown in culture medium then stained with CeCl₃ (Fig. 2B). The CeCl₃ deposit points were mainly in mitochondria, with some others present in the cytosol. In the self-S-RNase-treated pollen tube, however, there was no CeCl₃ deposited in mitochondria or cytosol (Fig. 2C). Compatible pollen tubes treated with S-RNase were similar to pollen tubes grown in culture medium, with many CeCl₃ deposits in mitochondria and some in cytosol (Fig. 2D). Pollen tubes incubated with DPI or TMPP had no CeCl₃ deposits in the mitochondria or cytosol (Fig. 2E,F). H₂O₂ accumulation in the cell walls of the subapical region of pollen tubes is shown in Fig. 2G-L. Typical cell walls of the subapical region of control pollen tubes grown in culture medium without CeCl₃ staining are shown in Fig. 2G. When control pollen tubes were stained with CeCl₃, there were abundant CeCl₃ deposits in the cell walls of the subapical region, facing the outside spaces (Fig. 2H). In the self S-RNase treatment, there were no CeCl₃ deposits in the cell wall of pollen tubes (Fig. 2I). However, in the compatible treatment there were many CeCl₃ deposits in the cell wall, indicating great accumulation of H₂O₂ (Fig. 2J). There were no CeCl₃ deposits in cell walls of DPI-incubated (Fig. 2K) or TMPP-incubated (Fig. 2L) pollen tubes. These results strongly validate the methods used in this study.

Because NAD(P)H, but not NAD(P)⁺, has endogenous fluorescence, a fluorescence microscope and multimode microplate readers were used to detect the NAD(P)H endogenous fluorescence signal of single and total pollen tubes, respectively. Fig. 3A shows the NAD(P)H endogenous fluorescence signals of single pollen tube at 0, 5, 10, 15 and 20 minutes after treatment. In the control, the strongest fluorescence was in the subapical region of the pollen tube. During the period 0–20 minutes, there was little change in the fluorescence intensity and distribution. In the DPI-treated pollen tube, there was a similar strong fluorescence in the subapical region. However, over 0–20 minutes, the intensity of the subapical region fluorescence increased, which is consistent with previous results (Cárdenas et al., 2006). The compatible treatment showed similar results to the control. However, in the incompatible treatment, the intensity of the subapical region fluorescence decreased notably during the 0–20 minute period, in contrast to the DPI treatment.

Furthermore, the results of total pollen tubes NAD(P)H endogenous fluorescence was consistent with the results for single pollen tube (Fig. 3B). NAD(P)H fluorescence intensity of pollen tubes in control or compatible treatments generally did not change over 0–20 minutes. However, in the DPI treatment, the NAD(P)H fluorescence intensity increased and NAD(P)H fluorescence intensity of incompatible pollen tubes declined during the whole period, especially during 0–5 minutes.

Patch-clamp whole-cell measurements were used to characterize Ca²⁺ channels in the plasma membrane of apical spheroplasts derived from pollen tubes under NADPH oxidase inhibitor DPI and ROS scavenger TMPP challenge. Whole-cell currents from an individual spheroplast were elicited by sequential step-wise hyperpolarization of the membrane to −200 mV from a holding potential of 0 mV. The bathing solution contained 10 mM Ca²⁺. Current-voltage (I–V) relationships are shown in Fig. 4. Relative to controls, the Ca²⁺ currents decreased gradually in the presence of DPI and TMPP (Fig. 4A). For instance, the current at −200 mV in the control was approximately 200% of the level in the presence of DPI and 400% of TMPP, respectively. Mean current at various step voltages under different treatments is shown in Fig. 4B. The results showed that DPI and TMPP clearly decreased the Ca²⁺ currents. At the same time, the incompatible but not the compatible S-RNase, also decreased the Ca²⁺ currents (supplementary material Fig. S2). These results show the link between the incompatible S-RNase, tip-localized ROS disruption and decreased Ca²⁺ currents.

We previously demonstrated that S-RNase induces depolymerization of the actin cytoskeleton of self-generated pollen tubes (Liu et al., 2007). In the present paper, the percentage of the pollen tubes with actin cytoskeleton depolymerization relative to total number of tubes in the S-RNase treatments was determined. At the same time, the effect of tip-localized ROS disruption on actin cytoskeletons of pollen tubes was evaluated. A typical pollen tube with normal or depolymerized actin cytoskeleton is shown in Fig. 5A. The percentage of pollen tubes with actin cytoskeleton depolymerization relative to the total number pollen tubes under different treatments are shown in Fig. 5B. In controls, 13.2±1.6% (means ± s.e.) of pollen tubes had actin cytoskeleton depolymerization relative to the total number; in the compatible treatment, this was 17.8±2.5%. There were no obvious differences between controls and the compatible treatment; however, in the incompatible treatment, the value was 73.9±6.7%. Thus the percentage in the incompatible treatment was approximately six times higher than the controls. Coincidentally, the DPI and TMPP treatments were similar to the incompatible treatment, the values were 54.7±5.6% and 60.4±5.1%, respectively.

We demonstrated that S-RNase triggered DNA degradation in incompatible pollen tubes. Since we discovered that S-RNase disrupted tip-localized ROS in incompatible pollen tubes, we speculated that tip-localized ROS disruption would induce nuclear DNA degradation. The pollen tubes were stained with DAPI after different treatments, and the percentage of pollen tubes with DNA degradation relative to total number in a certain treatment was counted. Staining with DAPI revealed three types of pollen tubes in each treatment: binucleate tubes were regarded as normal; a single nucleus suggested that vegetative nuclear DNA had degraded entirely; whereas complete absence of nuclei indicated that all nuclear DNA had degraded (Fig. 6). The percentage of binucleate, single nucleate, and tubes with no nucleus relative to the total number of control tubes were 71.6±3.5%, 18.6±4.5% and 9.8±6.7%, respectively. In the compatible treatment, the percentages were 69.9±4.7%, 16.2±3.1% and 13.9±3.5%, respectively. In the incompatible treatment, the percentages were 13.5±4.0%, 27.3±4.9% and 59.2±5.8%, respectively. The results with S-RNase treatment were similar to those in our previous paper (Wang et al., 2009). In the DPI treatment, however, the percentage of binucleate tubes relative to other types decreased markedly to only 42.7±8.9%, whereas single nucleus tubes and those lacking a nucleus reached 25.2±6.1% and 32.1±9.5%, respectively; in the TMPP treatment, values were 28.2±3.8%, 25.5±4.8% and 46.2±3.6%, respectively. The results indicate that tip-localized ROS disruption causes nuclear DNA to be degraded.

To further check the relationship between the actin cytoskeleton and nuclear DNA, pollen tubes were incubated with an actin-depolymerizing agent, cytochalasin B (CB) or the actin-stabilization agent phalloidin. The percentage of pollen tubes with nuclear DNA degradation to total number tubes was determined. Similarly to DPI and TMPP treatments, the binucleate tubes relative to the other tube types with CB and phalloidin treatment decreased to 25.4±3.8% and 22.8±8.0%, respectively; whereas the values in single nucleus tubes were 27.0±3.5% and 33.6±4.5%; and tubes with no nucleus 47.7±4.2% and 43.5±9.0%, respectively.

In a previous paper, we demonstrated that S-RNase triggers nuclear DNA degradation in vitro. To demonstrate that the in vitro results mirrored those in vivo, the nuclear DNA of the pollen tubes concealed in the style after different pollinations were evaluated. Pollen tube growth started to arrest after 9 hours with incompatible pollination (data not shown). Thus, the 9 hour time point after pollination was chosen to evaluate the nuclear DNA of pollen tubes embedded in the style. Typical images of nuclear DNA in the pollen tubes embedded in the style are shown at 9 hours after pollination in Fig. 7A. The pollen tubes were stained with aniline blue, and nuclear-DNA stained with DAPI. The results show that the nuclei of pollen tubes are smaller than those of the style organization cells, and the DAPI fluorescence intensity of pollen tube nuclei is stronger than that of nuclei of the style cells.

The terminal deoxynucleotidyl transferase-mediated dUTP nick-end labelling (TUNEL) method was adopted to further assess and confirm the nuclear DNA degradation of pollen tubes in the incompatible pollination. The TUNEL assay was performed 9 hours after pollination, and DAPI staining showed that the TUNEL-positive signal corresponded to nuclear DNA (Fig. 7B). In the incompatible pollination, 62.7±8.1% of visible nuclei appeared positive in the TUNEL assay; whereas in the compatible pollination, this number was only 8.8±2.7% (Fig. 7C).

Recent studies show that tip-localized ROS are a requirement for tip growth (Foreman et al., 2003; Potocky et al., 2007). The present results supported this conclusion (Fig. 1A). The percentage of pollen tubes with the strongest fluorescence in the tip following CM-H₂DCFDA staining decreased in the incompatible, but not compatible treatment (Fig. 1C), indicating that S-RNase particularly disrupts tip-localized ROS of incompatible pollen tubes in vitro. The NBT staining experiment presented similar results. As mentioned earlier, there are two central potential sources of ROS in pollen tubes, one is mitochondria and the other is plasma membrane NADPH oxidases (Mittler, 2002; Neill et al., 2002). So accumulation of ROS in these two parts was assessed by cytochemical detection after S-RNase challenge. In the normal or compatible pollen tube, there were many CeCl₃ deposits in the cell wall or mitochondria, but the deposits could not be observed in these two regions of incompatible tubes, which suggested that ROS in mitochondria or cell walls are disrupted in incompatible tubes. It was shown that S-RNase induced mitochondrial changes, including membrane potential collapse, cytochrome c leakage and swelling (Wang et al., 2009). These mitochondrial alterations induced loss of mitochondrial function, resulting in disruption of mitochondrial ROS.

Our results also showed that S-RNase particularly decreased the autofluorescence signal of NAD(P)H. The autofluorescence signal test in our protocol was from both mitochondrial and cytosolic NADH and NADPH, because the autofluorescence spectra overlap; thus it is not possible to distinguish between the two signals. Previously published studies indicate that decreased NAD(P)H fluorescence is due to oxidation of NAD(P)H, where the oxidized form, NAD(P)⁺, is nonfluorescent, rather than to a shift from bound (stronger fluorescence) to free NAD(P)H (weaker fluorescence) (Wakita et al., 1995; Cárdenas et al., 2006; Kasimova et al., 2006). Therefore, increased autofluorescence indicates an increase in the reduced form of the pyridine nucleotide, and decreased autofluorescence indicates an increase in the oxidized form (Schuchmann et al., 2001). Because NADH and NADPH tend to have opposite redox states, NAD mostly oxidized and NADP mostly reduced, the decreased autofluorescence signal in the incompatible treatment mainly resulted from a decrease in the reduced form of NADP. As an NADPH is an electron provider, a decrease in its concentration blocked plasma membrane ROS formation, and finally resulted in disruption of cell wall ROS. There was a similar absence of CeCl₃ deposits in DPI-incubated or TMPP-incubated pollen tubes, which validated the method used. These results suggest that S-RNase particularly disrupts tip-localized ROS of incompatible pollen tubes by arresting ROS formation in the mitochondria or cell wall.

To assess the influence of ROS disruption in pollen tubes, the NADPH oxidase inhibitor DPI or ROS scavenger TMPP were used. DPI and TMPP have been widely used together to assess the influence of ROS disruption in pollen tube growth (Coelho et al., 2008; Potocky et al., 2007). Although DPI is not a specific inhibitor of NADPH oxidase, it is a more suitable inhibitor for ROS production (Majander et al., 1994; Møller, 2001). On the one hand, DPI arrested ROS production via inhibiting NAD(P)H oxidases or NAD(P)H dehydrogenases. On the other hand, when the NAD(P)H dehydrogenases were inhibited, complex I of the mitochondrial electron transport chain will still produce ROS in the presence of rotenone because rotenone blocks electron transfer downstream of the FMN-containing active site; DPI could arrest this pathway of ROS production via inhibiting flavoproteins; DPI also inhibits other enzymes (e.g. peroxidases). In our experiments, the samples were also treated with TMPP, which showed similar results as DPI treatment. This combination ensured that the alterations were a result of ROS disruption. DPI or TMPP decreased Ca²⁺ currents, depolymerized actin cytoskeletons and induced nuclear DNA degradation at same time, which suggests that ROS disruption induces arrest of Ca²⁺ currents, depolymerization of actin cytoskeletons and degradation of nuclear DNA. Foreman and co-workers (Foreman et al., 2003) observed activation of hyperpolarization-activated Ca²⁺ channels by ROS in root hairs, which is similar to our results in pollen tubes. As the self S-RNase was added to the medium, the NAD(P)H fluorescence decreased immediately, which directly arrested cell wall ROS formation. These results indicate that tip-localized ROS disruption is the upstream event of SI in P. pyrifolia. Furthermore, our results showed that tip-localized ROS disruption induced actin depolymerization and nuclear DNA degradation. Several researchers have reported that either stabilization or depolymerization of the actin cytoskeleton is adequate to induce programmed cell death (PCD) in yeast and some animal cells (Celeste Morley et al., 2003; Gourlay and Ayscough, 2005; Janmey, 1998; Levee et al., 1996). Supporting this, Thomas and colleagues (Thomas et al., 2006) found that actin depolymerization was sufficient to induce PCD in the SI pollen of the poppy (Papaver rhoeas). Moreover, in our experiments, degradation of nuclear DNA of pollen tubes, a hallmark feature of apoptosis, was induced by the actin-depolymerizing agent, cytochalasin B (CB), or the actin-stabilization agent, phalloidin. Phalloidin-induced nuclear DNA degradation might be caused by toxins that bind polymeric F-actin, which interfere with the function of actin-rich structures and destroy the oscillation of tip F-actin. More importantly, it was demonstrated that nuclear DNA degradation also occurred in the pollen tube after incompatible pollination in vivo, which suggested our in vitro system mirrored the in vivo situation. It was also shown that ROS might have a role in pollen–stigma interaction (McInnis et al., 2006). Accordingly, our results showed inhibition of ROS is involved in SI, which is also involved in pollen–stigma interaction in many plants.

It is widely accepted that the growth of pollen tubes is inhibited by S-RNase degradation of RNA of incompatible pollen tubes in the pear. However, RNA degradation is not the only event in SI. It was demonstrated that S-RNase specifically induces tip-localized ROS disruption, actin cytoskeleton depolymerization and nuclear DNA degradation in incompatible pollen tubes of the pear, and alterations in actin and DNA degradation were the cause, not the result, of pollen tube growth arrest (Liu et al., 2007; Wang et al., 2009). This is the first report to show that tip-localized ROS have a role in SI. The mechanisms involved in inhibition of pollen by the pear SI response shared some features with the mechanistically different SI in the poppy, where S-protein (recently renamed PrsS) triggers a Ca²⁺-dependent signal cascade, including protein phosphorylation, depolymerization of actin cytoskeleton and microtubules, and PCD (Geitmann et al., 2000; Staiger and Franklin-Tong, 2003; Thomas and Franklin-Tong, 2004; McClure and Franklin-Tong, 2006; Thomas et al., 2006; Poulter et al., 2008). Endomembrane breakdown in incompatible pollen tubes was found in Nicotiana (Goldraij et al., 2006) and also in Brassica, where vacuolar disruption in incompatible stigmatic papilla cells and rejection is triggered by SRK (S-locus receptor kinase) (Iwano et al., 2007). Actin depolymerization was also detected in stigmatic papilla cells (Iwano et al., 2007). These data indicate a potential link among the different SI systems.

Adult pear (Pyrus pyrifolia L.) trees planted in the orchards of Nanjing Agricultural University, Jiangsu, China were used. The cultivars and their S-genotypes were Kosui (S₄S₅) and Imamuraaki (S₁S₆). Flowers from each cultivar were collected a few days before anthesis, and the styles detached, weighed, and stored in liquid nitrogen. Anthers of cultivar Kosui were collected, dehisced, dried in bottles containing desiccants, and stored at −20°C.

To isolate S-RNase, 4 g of styles were prepared following our previously described method (Hiratsuka et al., 2001; Zhang and Hiratsuka, 2000). The isolated S-RNase sample was stored in Eppendorf tubes at −80°C. S-RNase concentration and activity were determined by the methods of Bradford (Bradford, 1976) and Brown and Ho (Brown and Ho, 1986), respectively.

Pollen grains of cultivar Kosui were precultured for 2 hours at 25°C in a basal medium and in darkness, based on published methods (Hiratsuka et al., 2001). The basal medium consisted of a MES NaOH buffer supplemented with 10% sucrose, 15% polyethylene glycol 4000, 0.01% H₃BO₃, 0.07% Ca(NO₃)₂.4H₂O, 0.02% MgSO₄.7H₂O and 0.01% KNO₃, pH 6.0–6.5. After preculture, Kosui stylar S-RNase was added to the medium as an SI challenge (incompatible treatment), Imamuraaki stylar S-RNase was added to the medium as a compatible treatment, and medium without S-RNase was used as a control. The final activity of the S-RNases in the basal medium was 0.15 U.

To evaluate the ROS effect on pollen tube growth, pollen was grown in the basal medium for 2 hours at 25°C, and then diphenylene iodonium chloride (DPI, 300 μM final concentration) or Mn-5,10,15,20-tetrakis(1-methyl-4-pyridyl) 21H,23H-porphin (TMPP, 300 μM final concentration) was added to basal medium and incubated for 60 minutes. Lengths of ≥100 pollen tubes were measured per treatment, with each treatment repeated three times. To detect ROS formation in the pollen tube, precultured pollen tubes were stained with 5-(and 6-)chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate (CM-H₂DCFDA; 20 μM) or nitroblue tetrazolium (NBT; 1 mg/ml). Before CM-H₂DCFDA or NBT staining, pollen tubes were incubated with S-RNase, DPI or TMPP for 30 minutes. The stained samples were investigated with a Zeiss Axio Imager A1 fluorescence microscope and the percentage of all pollen tubes with strongest fluorescence in the tip was counted in each treatment. When tip-localized fluorescence strength was ≥threefold tube fluorescence strength, the pollen tube was counted as having strongest fluorescence. Per treatment, ≥100 pollen tubes were observed, with each treatment repeated three times. Images were processed using Adobe Photoshop CS.

H₂O₂ was visualized at the subcellular level using CeCl₃ for localization (Bestwick et al., 1997). Electron-dense CeCl₃ deposits are formed in the presence of H₂O₂ and are visible by transmission electron microscopy (TEM). Pollen was grown in the basal medium for 2 hours at 25°C and then incubated with S-RNase, DPI or TMPP for 30 minutes. After treatment, pollen tubes were incubated in freshly prepared 5 mM CeCl₃ in 50 mM 3-3-(N-morpholino) propanesulfonic acid at pH 7.2 for 1 hour. Pollen tubes were then fixed in 1.25% (v/v) glutaraldehyde and 1.25% (v/v) formaldehyde in 50 mM sodium cacodylate buffer, pH 7.2, for 1 hour. After fixation, pollen tubes were washed twice for 10 minutes in the same buffer and postfixed for 45 minutes in 1% (v/v) osmium tetroxide, and then dehydrated in a graded ethanol series (30–100%; v/v) and embedded in Eponaraldite (Agar Aids, Bishops, UK). After 12 hours in pure resin, followed by a change of fresh resin for 4 hours, the samples were polymerized at 60°C for 48 hours. Blocks were sectioned (70–90 nm) on a Reichert-Ultracut E microtome, and mounted on uncoated copper grids (300 mesh). Sections were examined with a Hitachi H-7650 TEM.

NAD(P)H is fluorescent and can be visualized by exciting at 340–365 nm and detecting emission at 400–500 nm (Cárdenas et al., 2006; Hu et al., 2002; Lakowicz et al., 1992). The NAD(P)H endogenous fluorescence signal of single pollen tube was assessed as described (Cárdenas et al., 2006). The fluorescence signal was visualized with a fluorescence microscope (Zeiss Axio Imager A1), equipped with a CCD camera and a Xenon excitatory light source. The filters were a 360 nm (10 nm bandpass) excitation filter and a 420–470 nm emission filter. The images were captured at 0, 5, 10, 15 and 20 minutes, respectively.

To validate the results, multimode microplate readers (Infinite M200, TECAN; http://www.tecan.com) were used to measure the fluorescence intensity of total pollen tubes. Pollens were precultured in a 5 ml centrifugal tube, and 2 hours later the tube was gently oscillated to ensure uniform pollen tube density throughout the tube. A pipette was used to add an equal volume of medium with suspended pollen tubes into each well of a 96-well microplate. Thus there was an equal quantity of pollen tubes in each well. When S-RNase or DPI was added to the well, pollen tubes were immediately examined for NAD(P)H endogenous fluorescence and repetitively excited by 345 nm light and emissions detected at 420 nm at intervals of 30 seconds over a total period of 20 minutes. The experiment was repeated three times.

The method used was previous published (Qu et al., 2007). After preculture, pollen tubes were washed twice with deionized water and incubated in an enzyme solution for 2.5 hours at 32°C to release spheroplasts. The enzyme solution was composed of 1% (w/v) macerozyme R-10, 2.0% (w/v) cellulase RS-10, 0.7% (w/v) pectolyase Y-23 and 1% (w/v) bovine serum albumin. The enzyme solution was then exchanged with the control bathing solution (0.2 mM glucose, 10 mM CaCl₂ and 5 mM MES, adjusted to an osmolality of 800 mOsM and pH 5.8 with D-sorbitol and Tris, respectively). The pipettes were pulled from borosilicate glass blanks and coated with Sylgard (184 silicone elastomer kit; Dow Corning, Midland, MI). The range of pipette resistance was 15–35 Ω in 10 mM CaCl₂. The pipette solution comprised 1 mM MgCl₂, 0.1 mM CaCl₂, 4 mM Ca(OH)₂, 10 mM ethyleneglycoltetraacetic acid (EGTA), 2 mM MgATP, 10 mM HEPES, 100 mM CsCl and 0.1 mM GTP, adjusted to pH 7.3 and an osmolality of 1100 mOsM by Tris and D-sorbitol, respectively. ATP was incorporated to delay rundown of currents (Forscher and Oxford, 1985) and GTP was incorporated to sustain possible G-protein-related activity (Edwards et al., 1989; Yawo and Momiyama, 1993). The free Ca²⁺ concentration in the pipette solution was ~10 nM, calculated with the chemical speciation program GEOCHEM (Parker et al., 1987). Whole-cell plasma membrane currents were measured using an Axon 200B amplifier (Axon Instruments, Foster City, CA). The whole-cell configuration was obtained using a short burst of suction applied to the pipette interior to rupture the membrane, resulting in a substantial increase in capacitance. Series resistance and capacitance were compensated accordingly. The membrane was held at a holding potential of 0 mV and then the voltage was either clamped at discrete values for 2.5 seconds or changed rapidly and continuously in a ‘ramp’. Voltage protocols for tail current analysis are described in the figure legends. Data were sampled at 2 kHz and filtered at 0.5 kHz, and then analysed using PCLAMP 9.0 (Axon Instrument). Junction potentials were corrected according to Amtmann and Sanders (Amtmann and Sanders, 1997). All experiments were conducted at room temperature (20–22°C). The permeability of the channels to Ca²⁺ relative to chlorine (Cl⁻) (P_Ca/P_Cl) was estimated using an equation derived from the Goldman–Hodgkin–Katz equation (Goldman, 1943; Hodgkin and Katz, 1949).

Pollen was grown in the basal medium for 2 hours at 25°C and then incubated with S-RNase, DPI or TMPP for 30 minutes. Samples were treated with freshly prepared m-maleimidobenzoyl-N-hydroxy-succinimide ester at a final concentration of 200 μM in basal medium for 7 minutes to stabilize the actin filaments according to published methods (Liu et al., 2007). They were subsequently fixed for 1 hour with 4% formaldehyde in phosphate-buffered saline buffer. The samples were washed three times in phosphate-buffered saline and then placed on a cover-slide previously smeared with poly-L-lysine. Surplus buffer was removed with filter paper, and the samples stained with 5 μg/ml fluorescein isothiocyanate-phalloidin in 5% dimethyl sulfoxide, 5 mM EGTA and 10% (w/v) sucrose, pH 6.9. Specimens were washed with buffer before mounting in glycerol. The stained samples were investigated with a Zeiss fluorescence microscope and the pollen tubes with actin cytoskeleton depolymerization were counted. In each treatment, ≥100 pollen tubes were counted and the experiment repeated three times.

Pollen was grown in the basal medium for 2 hours at 25°C and then incubated with DPI, TMPP, CB or phalloidin for 30 minutes. To assess the extent of nuclear DNA degradation, pollen tubes were fixed in 95% ethanol:glacial acetic acid (3:1) for 1 hour at 4°C, transferred into 70% ethanol at −20°C for at least 4 hours, and then stained with 0.05 μg/ml DAPI in citrate buffer at pH 4.1 for 2 hours. The stained samples were examined with a Zeiss Axioskop40 fluorescence microscope. In each treatment, ≥100 pollen tubes were counted and the experiment repeated three times.

To check nuclear DNA of pollen tubes after compatible or incompatible pollination, the full-bloom flowers were emasculated. The Kosui pollen grains were transferred to the stigma of emasculated Imamuraaki flowers as a compatible pollination, whereas Kosui pollen grains were transferred to stigmas of Kosui flowers as an incompatible pollination. At 9 hours after pollination, the styles were collected and fixed in FAA (37% formaldehyde:glacial acetic acid:50% ethanol; 5:5:9) and stored at 4°C until used. The styles were washed thoroughly under running tap water and incubated in 1 M NaOH for 2 hours to soften the tissues and then soaked in 0.1% aniline blue solution with 0.1 M K₃PO₄ for 2 hours at 60°C in darkness. The styles were washed with citrate buffer (pH 4.1) and stained with 0.05 μg/ml DAPI in citrate buffer at pH 4.1 for 2 hours. Specimens were washed with citrate buffer again, and then squashed on glass slides, and mounted in glycerol. The stained samples were investigated with a Zeiss Axioskop40 fluorescence microscope. The DeadEnd Colorimetric TUNEL System (Promega, http://www.promega.com) was used for the independent assessment of nuclear DNA degradation in pollen tubes after compatible or incompatible pollination. After tissues were softened, the styles were processed following the manufacturer's instructions. Then the styles were stained with DAPI as previously mentioned. DAPI staining showed that the TUNEL-positive signal corresponded to nuclear DNA. The samples were investigated with an OLYMPUS BX51 fluorescence microscope. The percentage of positive-TUNEL-reactive nuclei to all visible nuclei was counted in each treatment. At least ten styles were observed per treatment and the experiment was repeated three times. Images were processed using Adobe Photoshop CS.

This work was supported by the earmarked fund for Modern Agro-industry Technology Research System (nycytx-29). The language in the manuscript was improved by International Science Editing.