Research Article 4301

IntroductionPollen 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 (Stoneet al., 2004). To achieve extremely fast growth, pollen tubes havea 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 polarelongation. Recent studies showed that reactive oxygen species(ROS) were a requirement for root hair growth. Foreman andcolleagues (Foreman et al., 2003) used a loss-of-function knockoutmutant in AtrbobC/RHD2 to demonstrate that ROS wereindispensable for root hair growth and were required to stimulateCa2+ influx during root-hair elongation. AtrbobC/RHD2 encodesa superoxide (O2·–)-producing NADPH oxidase (NOX) and themutant had reduced ROS formation at the tip of very short roothairs. Subsequently, Monshausen and colleagues (Monshausen etal., 2007) detected oscillations of apoplastic ROS concentrationat the top of the flanks of root hairs, which revealed a new modelof the role of ROS in root hair growth. Otherwise, it wasdemonstrated that tip-localized ROS produced by a NOX enzymewas needed to sustain the normal rate of pollen tube growth(Potocky et al., 2007). ROS are an inevitable consequence ofaerobic metabolism, and have a variety of functions includingpathogen defence and cell signalling. In plant cells, there aremany 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 majorROS producers in animal cells and in plant cells withoutchloroplasts, such as pollen tubes. The O2·– formation inmitochondria is closely related to the coupling efficiency betweenthe respiratory chain and oxidative phosphorylation. Plasma-membrane-bound NADPH oxidase transfers electrons fromcytoplasmic NADPH to form O2·–, which undergoes an enzymaticand non-enzymatic dismutation reaction, immediately producingH2O2, the most stable form of ROS. ROS formed in the plasmamembrane 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. Usingan in vitro system, our team has identified the characteristics of S-RNase that specifically inhibit self-pollen germination and tubeelongation (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 andDNA degradation of self-generated pollen tubes (Liu et al., 2007;Wang et al., 2009). To identify whether the in vitro results mirrorwhat happens in vivo, we evaluated the nuclear DNA of pollentubes after different pollinations. ROS have an important role inpollen tube elongation as mentioned above, and the actincytoskeleton is a target for ROS in yeast (Perrone et al., 2008). Wethus speculated that tip-localized ROS disruption occurs in the SIresponse of pear.

Accepted 6 September 2010Journal of Cell Science 123, 4301-4309 © 2010. Published by The Company of Biologists Ltddoi:10.1242/jcs.075077

SummaryPear (Pyrus pyrifolia L.) has an S-RNase-based gametophytic self-incompatibility (SI) mechanism, and S-RNase has also beenimplicated in the rejection of self-pollen and genetically identical pollen. However, RNA degradation might be only the beginning ofthe SI response, not the end. Recent in vitro studies suggest that S-RNase triggers mitochondrial alteration and DNA degradation inthe incompatible pollen tube of Pyrus pyrifolia, and it seems that a relationship exists between self S-RNase, actin depolymerizationand DNA degradation. To further uncover the SI response in pear, the relationship between self S-RNase and tip-localized reactiveoxygen species (ROS) was evaluated. Our results show that S-RNase specifically disrupted tip-localized ROS of incompatible pollentubes via arrest of ROS formation in mitochondria and cell walls. The mitochondrial ROS disruption was related to mitochondrialalteration, whereas cell wall ROS disruption was related to a decrease in NADPH. Tip-localized ROS disruption not only decreasedthe Ca2+ current and depolymerized the actin cytoskeleton, but it also induced nuclear DNA degradation. These results indicate thattip-localized ROS disruption occurs in Pyrus pyrifolia SI. Importantly, we demonstrated nuclear DNA degradation in the incompatiblepollen tube after pollination in vivo. This result validates our in vitro system in vivo.

Key words: Pyrus pyrifolia, Self-incompatibility, Reactive oxygen species, NAD(P)H, Ca2+ channels, Actin cytoskeleton, Nuclear DNA

S-RNase disrupts tip-localized reactive oxygenspecies and induces nuclear DNA degradation inincompatible pollen tubes of Pyrus pyrifoliaChun-Lei Wang, Jun Wu, Guo-Hua Xu*, Yong-bin Gao, Gong Chen, Ju-You Wu, Hua-qing Wu andShao-Ling Zhang‡

College of Horticulture, State Key Laboratory of Crop Genetics and Germplasm Enhancement, Nanjing Agricultural University, Nanjing 210095,China*Present address: Department of Biology, Jiangsu Institute of Education, Nanjing 210095, China‡Author for correspondence (nnzsl@njau.edu.cn)

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ResultsS-RNase disrupts tip-localized ROS in incompatible pollentubesTo assess the effect of ROS on pollen tube growth, the pollen wasgrown in the basal medium for 1 hour at 25°C, then the NADPHoxidase inhibitor diphenylene iodonium chloride (DPI) and ROSscavenger Mn-5,10,15,20-tetrakis(1-methyl-4-pyridyl)21H,23H-porphin (TMPP) were added to the medium. As expected, DPI andTMPP obviously inhibited pollen tube growth (Fig. 1A), whichindicated that ROS are necessary for pear pollen tube growth.

Furthermore, to evaluate the effect of S-RNase on tip-localizedROS, 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 withCM-H2DCFDA (Fig. 1B): pollen tubes with strongest fluorescencein the tip (from subapical domain to apex) were regarded as normal,whereas pollen tubes with uniform fluorescence suggested thatROS in the tip-localized pollen tube were disrupted. The samplewas stained with CM-H2DCFDA at 30 minutes after S-RNase, DPIor TMPP challenge to count the pollen tube with strongestfluorescence in tip (Fig. 1C). In the control, the proportion of pollentubes with strongest fluorescence in the tip relative to the totalnumber of tubes was 52.7±2.1% (means ± s.e.; n>100), and withthe DPI or TMPP treatments, only 32.4±3.4% (n>100) and 35.9±4.4% (n>100), respectively. In the compatible treatment, similarlyto the control, 52.0±1.4% (n>100) of pollen tubes had strongestfluorescence in the tip. However, in the incompatible treatment, thiswas only 28.8±1.7% (n>100), nearly half the control value.

Nitroblue tetrazolium (NBT) was also used for visualizing ROSdistribution. NBT is reduced by O2·– to a blue formazan precipitate,indicating the site of O2·– production. Because NBT was cytotoxicand rapidly kills pollen tubes, NBT staining provides a snapshot of

ROS formation in pollen tubes at different stages of the oscillatorygrowth cycle (Potocky et al., 2007). Fig. 1D shows typical imagesof pollen tube staining with NBT for different treatments. Thepollen tube of the control showed a tip-localized pattern of formazanstaining; there was a similar staining pattern in the compatiblepollen tubes. However, the tip-localized formazan staining of pollentubes treated with DPI or self S-RNase was disrupted. The TMPP-incubated pollen tubes were the colour of the TMPP reagent anddid not have tip-localized formazan staining. The percentage ofpollen tubes with tip-localized formazan staining pattern relativeto the total number of tubes in different treatments is shown insupplementary material Fig. S1.

Cytochemical detection of H2O2 in pollen tubesTo investigate the subcellular localization of H2O2 accumulation inthe subapical region of pollen tubes exposed to S-RNase treatment,a CeCl3 cytochemical technique, which reacts with H2O2 to produceelectron-dense deposits of cerium perhydroxides (Bestwick et al.,1997), was used. Coincidently, mitochondria were distributed inthe same subapical region. Thus the region of mitochondriaaccumulation was regarded as the subapical region of the pollentube. H2O2 accumulation in mitochondria of pollen tubes is shownin Fig. 2A–F. Control pollen tubes without CeCl3 staining areshown in Fig. 2A. The subapical region of a pollen tube grown inculture medium then stained with CeCl3 (Fig. 2B). The CeCl3deposit points were mainly in mitochondria, with some otherspresent 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 weresimilar to pollen tubes grown in culture medium, with many CeCl3deposits in mitochondria and some in cytosol (Fig. 2D). Pollentubes incubated with DPI or TMPP had no CeCl3 deposits in the

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Fig. 1. S-RNase disrupts tip-localizedROS in incompatible pollen tubes.(A)The ROS effect on pollen tube growth.The NADPH oxidase inhibitordiphenyleneiodonium chloride (DPI) orthe ROS scavenger TMPP arrests pollentube elongation. The pollen tubes wereincubated with DPI, TMPP for 60 minutesand experiments were repeated at leastthree times. Data points are means ± s.e.(B)Typical image of pollen tube stainedwith CM-H2DCFDA. Comparison offluorescence images (left) and DIC images(right) indicates the region in which ROSlocalize. (C)The percentage of pollentubes with strongest fluorescence insubapical region relative to the totalnumber of tubes in different treatments.Pollen tubes were incubated with DPI,TMPP or S-RNase from compatible(Comp) and incompatible (Incomp) stylesfor 30 minutes and experiments wererepeated at least three times. Data pointsare means ± s.e. (D)Representativeimages of pollen tubes incubated withNBT under different treatment conditions.Experiments were repeated at least threetimes with similar results.

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mitochondria or cytosol (Fig. 2E,F). H2O2 accumulation in the cellwalls of the subapical region of pollen tubes is shown in Fig. 2G-L. Typical cell walls of the subapical region of control pollen tubesgrown in culture medium without CeCl3 staining are shown in Fig.2G. When control pollen tubes were stained with CeCl3, therewere abundant CeCl3 deposits in the cell walls of the subapicalregion, facing the outside spaces (Fig. 2H). In the self S-RNasetreatment, there were no CeCl3 deposits in the cell wall of pollentubes (Fig. 2I). However, in the compatible treatment there weremany CeCl3 deposits in the cell wall, indicating great accumulationof H2O2 (Fig. 2J). There were no CeCl3 deposits in cell walls ofDPI-incubated (Fig. 2K) or TMPP-incubated (Fig. 2L) pollen tubes.These results strongly validate the methods used in this study.

NAD(P)H endogenous fluorescence decreases inincompatible pollen tubesBecause NAD(P)H, but not NAD(P)+, has endogenous fluorescence,a fluorescence microscope and multimode microplate readers wereused to detect the NAD(P)H endogenous fluorescence signal ofsingle and total pollen tubes, respectively. Fig. 3A shows theNAD(P)H endogenous fluorescence signals of single pollen tube at0, 5, 10, 15 and 20 minutes after treatment. In the control, the

strongest fluorescence was in the subapical region of the pollentube. During the period 0–20 minutes, there was little change in thefluorescence intensity and distribution. In the DPI-treated pollentube, there was a similar strong fluorescence in the subapical region.However, over 0–20 minutes, the intensity of the subapical regionfluorescence increased, which is consistent with previous results(Cárdenas et al., 2006). The compatible treatment showed similarresults to the control. However, in the incompatible treatment, theintensity of the subapical region fluorescence decreased notablyduring the 0–20 minute period, in contrast to the DPI treatment.

Furthermore, the results of total pollen tubes NAD(P)Hendogenous fluorescence was consistent with the results for singlepollen tube (Fig. 3B). NAD(P)H fluorescence intensity of pollentubes in control or compatible treatments generally did not changeover 0–20 minutes. However, in the DPI treatment, the NAD(P)Hfluorescence intensity increased and NAD(P)H fluorescenceintensity of incompatible pollen tubes declined during the wholeperiod, especially during 0–5 minutes.

Tip-localized ROS disruption decreases Ca2+ currentsPatch-clamp whole-cell measurements were used to characterizeCa2+ channels in the plasma membrane of apical spheroplasts

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Fig. 2. Cytochemical localization of H2O2 accumulationin subapical region of pollen tubes with CeCl3 stainingand TEM. (A–F) Images of CeCl3 deposits in mitochondriaof pollen tubes, which directly indicate the quantity of H2O2

accumulation in the subapical regions. (A)Pollen tubes incontrols without CeCl3 staining. (B)The subapical region ofpollen tubes grown in culture medium and then stained withCeCl3. There are many CeCl3 deposits (arrows) inmitochondria. (C)In the self-S-RNase-treated pollen tube,there are no CeCl3 deposits in mitochondria. (D)Pollen tubestreated with compatible S-RNase are similar to pollen tubesgrown in culture medium. There are many CeCl3 depositpoints (arrow) in mitochondria. (E,F) Pollen tubes incubatedwith DPI or TMPP have no CeCl3 deposits in mitochondria.(G–L) CeCl3 deposits in cell walls of the subapical region ofpollen tubes. (G)Typical cell walls in the control pollen tubesubapical region, in tubes grown in culture medium withoutCeCl3 staining. (H)Pollen tubes in controls are stained withCeCl3. There are abundant CeCl3 deposits in the cell walls ofthe subapical region, facing the outside spaces (arrow). (I)Inthe incompatible treatment, there are no CeCl3 deposit pointsin the cell wall. (J)In the compatible treatment, there aremany CeCl3 deposit points (arrow) in the cell wall.(K,L)There are no CeCl3 deposits in the cell wall of DPI-incubated or TMPP-incubated pollen tubes. Pollen tubeswere incubated with DPI, TMPP or S-RNase for 30 minutesand experiments were repeated at least three times withsimilar results. Scale bars: 200 nm.

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derived from pollen tubes under NADPH oxidase inhibitor DPIand ROS scavenger TMPP challenge. Whole-cell currents froman individual spheroplast were elicited by sequential step-wisehyperpolarization of the membrane to –200 mV from a holdingpotential of 0 mV. The bathing solution contained 10 mM Ca2+.Current-voltage (I–V) relationships are shown in Fig. 4. Relativeto controls, the Ca2+ currents decreased gradually in the presenceof DPI and TMPP (Fig. 4A). For instance, the current at –200 mVin the control was approximately 200% of the level in the presenceof DPI and 400% of TMPP, respectively. Mean current at variousstep voltages under different treatments is shown in Fig. 4B. Theresults showed that DPI and TMPP clearly decreased the Ca2+

currents. At the same time, the incompatible but not thecompatible S-RNase, also decreased the Ca2+ currents(supplementary material Fig. S2). These results show the linkbetween the incompatible S-RNase, tip-localized ROS disruptionand decreased Ca2+ currents.

Tip-localized ROS disruption depolymerizes the actincytoskeleton of pollen tubesWe previously demonstrated that S-RNase inducesdepolymerization of the actin cytoskeleton of self-generated pollentubes (Liu et al., 2007). In the present paper, the percentage of thepollen tubes with actin cytoskeleton depolymerization relative tototal number of tubes in the S-RNase treatments was determined.At the same time, the effect of tip-localized ROS disruption onactin cytoskeletons of pollen tubes was evaluated. A typical pollentube with normal or depolymerized actin cytoskeleton is shown in

Fig. 5A. The percentage of pollen tubes with actin cytoskeletondepolymerization relative to the total number pollen tubes underdifferent treatments are shown in Fig. 5B. In controls, 13.2±1.6%(means ± s.e.) of pollen tubes had actin cytoskeletondepolymerization relative to the total number; in the compatibletreatment, this was 17.8±2.5%. There were no obvious differencesbetween controls and the compatible treatment; however, in theincompatible treatment, the value was 73.9±6.7%. Thus thepercentage in the incompatible treatment was approximately sixtimes higher than the controls. Coincidentally, the DPI and TMPPtreatments were similar to the incompatible treatment, the valueswere 54.7±5.6% and 60.4±5.1%, respectively.

Tip-localized ROS disruption induces nuclear DNAdegradationWe demonstrated that S-RNase triggered DNA degradation inincompatible pollen tubes. Since we discovered that S-RNasedisrupted tip-localized ROS in incompatible pollen tubes, wespeculated that tip-localized ROS disruption would induce nuclearDNA degradation. The pollen tubes were stained with DAPI afterdifferent treatments, and the percentage of pollen tubes with DNAdegradation relative to total number in a certain treatment wascounted. Staining with DAPI revealed three types of pollen tubesin each treatment: binucleate tubes were regarded as normal; asingle nucleus suggested that vegetative nuclear DNA had degradedentirely; whereas complete absence of nuclei indicated that allnuclear DNA had degraded (Fig. 6). The percentage of binucleate,single nucleate, and tubes with no nucleus relative to the total

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Fig. 3. NAD(P)H endogenous fluorescence signal ofpollen tubes with different treatments over time.(A)Typical images of NAD(P)H endogenous fluorescencesignal of single pollen tubes at 0, 5, 10, 15 and 20 minutesafter different treatment. Experiments were repeated at leastthree times with similar results. Scale bar: 20m. (B)TheNAD(P)H endogenous fluorescence intensity of total pollentubes. The fluorescence of pure culture medium is shown asthe datum line. ‘Comp’ indicates compatible treatment, and‘Incomp’, incompatible treatment.

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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 were69.9±4.7%, 16.2±3.1% and 13.9±3.5%, respectively. In theincompatible treatment, the percentages were 13.5±4.0%,27.3±4.9% and 59.2±5.8%, respectively. The results with S-RNasetreatment were similar to those in our previous paper (Wang et al.,

2009). In the DPI treatment, however, the percentage of binucleatetubes relative to other types decreased markedly to only 42.7±8.9%,whereas single nucleus tubes and those lacking a nucleus reached25.2±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 nuclearDNA to be degraded.

To further check the relationship between the actin cytoskeletonand nuclear DNA, pollen tubes were incubated with an actin-depolymerizing agent, cytochalasin B (CB) or the actin-stabilization

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Fig. 4. Whole-cell currents from an individualspheroplast derived from pollen tubes. Tubeswere treated with the NADPH oxidase inhibitorDPI (300M) or the ROS scavenger TMPP(300M) and whole-cell currents were elicited bysequential step-wise hyperpolarization of themembrane to –200 mV from a holding potential of0 mV. (A)Relative to controls, the Ca2+ currentsdecrease gradually in the presence of DPI orTMPP. (B)Mean current at various step voltagesfor different treatments. Data points are means ±s.e.

Fig. 5. The actin cytoskeleton is depolymerized in the SI of pear.(A)Typical pollen tubes with normal or depolymerized actin cytoskeletons.(B)Pollen tubes with actin cytoskeleton depolymerization relative to the totalnumber of tubes in different treatments (%). Pollen tubes were incubated withDPI, TMPP or S-RNase for 30 minutes. ‘Comp’ indicates compatibletreatment, and ‘Incomp’, incompatible treatment. Data points are means ± s.e.

Fig. 6. Tip-localized ROS disruption induces nuclear DNA degradation.There was no difference in the percentage of pollen tubes with degraded DNAbetween the control and compatible treatment (Comp). Incompatible S-RNase(Incomp), NADPH oxidase inhibitor DPI, ROS scavenger TMPP, actin-depolymerizing agent, cytochalasin B (CB) and actin-stabilization agent,phalloidin, caused nuclear DNA to degrade. Pollen tubes were incubated withDPI, TMPP, CB, phalloidin or S-RNase for 30 minutes. Binucleate tubes wereregarded as normal; pollen tubes with a single nucleus implied that thevegetative nuclear DNA had degraded completely, whereas those lacking anucleus suggested that both vegetative and generative DNA had degraded.Data points are means ± s.e.

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agent phalloidin. The percentage of pollen tubes with nuclear DNAdegradation to total number tubes was determined. Similarly toDPI and TMPP treatments, the binucleate tubes relative to theother tube types with CB and phalloidin treatment decreased to25.4±3.8% and 22.8±8.0%, respectively; whereas the values insingle nucleus tubes were 27.0±3.5% and 33.6±4.5%; and tubeswith no nucleus 47.7±4.2% and 43.5±9.0%, respectively.

Nuclear DNA of pollen tubes is degraded afterincompatible pollinationIn a previous paper, we demonstrated that S-RNase triggersnuclear DNA degradation in vitro. To demonstrate that the invitro results mirrored those in vivo, the nuclear DNA of thepollen tubes concealed in the style after different pollinationswere evaluated. Pollen tube growth started to arrest after 9 hourswith incompatible pollination (data not shown). Thus, the 9 hourtime point after pollination was chosen to evaluate the nuclearDNA of pollen tubes embedded in the style. Typical images ofnuclear DNA in the pollen tubes embedded in the style are shownat 9 hours after pollination in Fig. 7A. The pollen tubes werestained with aniline blue, and nuclear-DNA stained with DAPI.The results show that the nuclei of pollen tubes are smaller thanthose of the style organization cells, and the DAPI fluorescenceintensity of pollen tube nuclei is stronger than that of nuclei ofthe style cells.

The terminal deoxynucleotidyl transferase-mediated dUTP nick-end labelling (TUNEL) method was adopted to further assess andconfirm the nuclear DNA degradation of pollen tubes in theincompatible pollination. The TUNEL assay was performed 9hours after pollination, and DAPI staining showed that the TUNEL-positive signal corresponded to nuclear DNA (Fig. 7B). In theincompatible 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).

DiscussionS-RNase disrupts tip-localized ROS in incompatible pollentubesRecent studies show that tip-localized ROS are a requirement fortip growth (Foreman et al., 2003; Potocky et al., 2007). The presentresults supported this conclusion (Fig. 1A). The percentage ofpollen tubes with the strongest fluorescence in the tip followingCM-H2DCFDA staining decreased in the incompatible, but notcompatible treatment (Fig. 1C), indicating that S-RNase particularlydisrupts tip-localized ROS of incompatible pollen tubes in vitro.The NBT staining experiment presented similar results. Asmentioned earlier, there are two central potential sources of ROSin pollen tubes, one is mitochondria and the other is plasmamembrane NADPH oxidases (Mittler, 2002; Neill et al., 2002). Soaccumulation of ROS in these two parts was assessed bycytochemical detection after S-RNase challenge. In the normal orcompatible pollen tube, there were many CeCl3 deposits in the cellwall or mitochondria, but the deposits could not be observed inthese two regions of incompatible tubes, which suggested thatROS in mitochondria or cell walls are disrupted in incompatibletubes. It was shown that S-RNase induced mitochondrial changes,including membrane potential collapse, cytochrome c leakage andswelling (Wang et al., 2009). These mitochondrial alterationsinduced loss of mitochondrial function, resulting in disruption ofmitochondrial ROS.

Our results also showed that S-RNase particularly decreased theautofluorescence signal of NAD(P)H. The autofluorescence signaltest in our protocol was from both mitochondrial and cytosolicNADH and NADPH, because the autofluorescence spectra overlap;

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Fig. 7. The extent of pollen tube nuclear DNA degradationafter pollination. (A)After pollination, pollen tubes wereembedded in the style. The pollen tube (pt) is stained withAniline Blue (green), and nuclear DNA with DAPI (blue).(B)In incompatible pollination, the nucleus appears positive inthe TUNEL assay (arrows), unlike compatible pollination(arrows). DAPI staining shows that the TUNEL-positive signalcorresponds to nuclear DNA. (C)The percentage of TUNEL-positive nuclei in incompatible (Incomp) or compatible (Comp)pollination. Data points are means ± s.e.

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thus it is not possible to distinguish between the two signals.Previously published studies indicate that decreased NAD(P)Hfluorescence is due to oxidation of NAD(P)H, where the oxidizedform, NAD(P)+, is nonfluorescent, rather than to a shift frombound (stronger fluorescence) to free NAD(P)H (weakerfluorescence) (Wakita et al., 1995; Cárdenas et al., 2006; Kasimovaet al., 2006). Therefore, increased autofluorescence indicates anincrease in the reduced form of the pyridine nucleotide, anddecreased autofluorescence indicates an increase in the oxidizedform (Schuchmann et al., 2001). Because NADH and NADPHtend to have opposite redox states, NAD mostly oxidized andNADP mostly reduced, the decreased autofluorescence signal inthe incompatible treatment mainly resulted from a decrease in thereduced form of NADP. As an NADPH is an electron provider, adecrease in its concentration blocked plasma membrane ROSformation, and finally resulted in disruption of cell wall ROS.There was a similar absence of CeCl3 deposits in DPI-incubated orTMPP-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 ROSformation in the mitochondria or cell wall.

Tip-localized ROS disruption induces nuclear DNAdegradationTo assess the influence of ROS disruption in pollen tubes, theNADPH oxidase inhibitor DPI or ROS scavenger TMPP wereused. DPI and TMPP have been widely used together to assess theinfluence of ROS disruption in pollen tube growth (Coelho et al.,2008; Potocky et al., 2007). Although DPI is not a specific inhibitorof NADPH oxidase, it is a more suitable inhibitor for ROSproduction (Majander et al., 1994; Møller, 2001). On the one hand,DPI arrested ROS production via inhibiting NAD(P)H oxidases orNAD(P)H dehydrogenases. On the other hand, when the NAD(P)Hdehydrogenases were inhibited, complex I of the mitochondrialelectron transport chain will still produce ROS in the presence ofrotenone because rotenone blocks electron transfer downstream ofthe FMN-containing active site; DPI could arrest this pathway ofROS production via inhibiting flavoproteins; DPI also inhibitsother enzymes (e.g. peroxidases). In our experiments, the sampleswere also treated with TMPP, which showed similar results as DPItreatment. This combination ensured that the alterations were aresult of ROS disruption. DPI or TMPP decreased Ca2+ currents,depolymerized actin cytoskeletons and induced nuclear DNAdegradation at same time, which suggests that ROS disruptioninduces arrest of Ca2+ currents, depolymerization of actincytoskeletons and degradation of nuclear DNA. Foreman and co-workers (Foreman et al., 2003) observed activation ofhyperpolarization-activated Ca2+ channels by ROS in root hairs,which is similar to our results in pollen tubes. As the self S-RNasewas added to the medium, the NAD(P)H fluorescence decreasedimmediately, which directly arrested cell wall ROS formation.These results indicate that tip-localized ROS disruption is theupstream event of SI in P. pyrifolia. Furthermore, our resultsshowed that tip-localized ROS disruption induced actindepolymerization and nuclear DNA degradation. Severalresearchers have reported that either stabilization ordepolymerization of the actin cytoskeleton is adequate to induceprogrammed 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 (Papaverrhoeas). Moreover, in our experiments, degradation of nuclearDNA of pollen tubes, a hallmark feature of apoptosis, was inducedby the actin-depolymerizing agent, cytochalasin B (CB), or theactin-stabilization agent, phalloidin. Phalloidin-induced nuclearDNA degradation might be caused by toxins that bind polymericF-actin, which interfere with the function of actin-rich structuresand destroy the oscillation of tip F-actin. More importantly, it wasdemonstrated that nuclear DNA degradation also occurred in thepollen tube after incompatible pollination in vivo, which suggestedour in vitro system mirrored the in vivo situation. It was alsoshown that ROS might have a role in pollen–stigma interaction(McInnis et al., 2006). Accordingly, our results showed inhibitionof ROS is involved in SI, which is also involved in pollen–stigmainteraction in many plants.

Do different SI systems use common mechanisms toreject incompatible pollen?It is widely accepted that the growth of pollen tubes is inhibited byS-RNase degradation of RNA of incompatible pollen tubes in thepear. However, RNA degradation is not the only event in SI. It wasdemonstrated that S-RNase specifically induces tip-localized ROSdisruption, actin cytoskeleton depolymerization and nuclear DNAdegradation in incompatible pollen tubes of the pear, and alterationsin actin and DNA degradation were the cause, not the result, ofpollen tube growth arrest (Liu et al., 2007; Wang et al., 2009). Thisis the first report to show that tip-localized ROS have a role in SI.The mechanisms involved in inhibition of pollen by the pear SIresponse shared some features with the mechanistically differentSI in the poppy, where S-protein (recently renamed PrsS) triggersa Ca2+-dependent signal cascade, including protein phosphorylation,depolymerization of actin cytoskeleton and microtubules, and PCD(Geitmann et al., 2000; Staiger and Franklin-Tong, 2003; Thomasand Franklin-Tong, 2004; McClure and Franklin-Tong, 2006;Thomas et al., 2006; Poulter et al., 2008). Endomembranebreakdown in incompatible pollen tubes was found in Nicotiana(Goldraij et al., 2006) and also in Brassica, where vacuolardisruption in incompatible stigmatic papilla cells and rejection istriggered 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 thedifferent SI systems.

Materials and MethodsPlant materialsAdult pear (Pyrus pyrifolia L.) trees planted in the orchards of Nanjing AgriculturalUniversity, Jiangsu, China were used. The cultivars and their S-genotypes wereKosui (S4S5) and Imamuraaki (S1S6). Flowers from each cultivar were collected afew days before anthesis, and the styles detached, weighed, and stored in liquidnitrogen. Anthers of cultivar Kosui were collected, dehisced, dried in bottlescontaining desiccants, and stored at –20°C.

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

Pollen culture and SI challengePollen grains of cultivar Kosui were precultured for 2 hours at 25°C in a basalmedium and in darkness, based on published methods (Hiratsuka et al., 2001). Thebasal 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-RNasewas added to the medium as an SI challenge (incompatible treatment), Imamuraaki

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stylar S-RNase was added to the medium as a compatible treatment, and mediumwithout S-RNase was used as a control. The final activity of the S-RNases in thebasal medium was 0.15 U.

Detection of ROS in pollen tubesTo evaluate the ROS effect on pollen tube growth, pollen was grown in the basalmedium for 2 hours at 25°C, and then diphenylene iodonium chloride (DPI, 300 Mfinal concentration) or Mn-5,10,15,20-tetrakis(1-methyl-4-pyridyl) 21H,23H-porphin(TMPP, 300 M final concentration) was added to basal medium and incubated for60 minutes. Lengths of ≥100 pollen tubes were measured per treatment, with eachtreatment 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 M) or nitrobluetetrazolium (NBT; 1 mg/ml). Before CM-H2DCFDA or NBT staining, pollen tubeswere incubated with S-RNase, DPI or TMPP for 30 minutes. The stained sampleswere investigated with a Zeiss Axio Imager A1 fluorescence microscope and thepercentage of all pollen tubes with strongest fluorescence in the tip was counted ineach treatment. When tip-localized fluorescence strength was ≥threefold tubefluorescence strength, the pollen tube was counted as having strongest fluorescence.Per treatment, ≥100 pollen tubes were observed, with each treatment repeated threetimes. Images were processed using Adobe Photoshop CS.

Cytochemical detection of H2O2

H2O2 was visualized at the subcellular level using CeCl3 for localization (Bestwicket al., 1997). Electron-dense CeCl3 deposits are formed in the presence of H2O2 andare visible by transmission electron microscopy (TEM). Pollen was grown in thebasal medium for 2 hours at 25°C and then incubated with S-RNase, DPI or TMPPfor 30 minutes. After treatment, pollen tubes were incubated in freshly prepared 5mM 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 for45 minutes in 1% (v/v) osmium tetroxide, and then dehydrated in a graded ethanolseries (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, thesamples 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 (300mesh). Sections were examined with a Hitachi H-7650 TEM.

NAD(P)H endogenous fluorescence detectionNAD(P)H is fluorescent and can be visualized by exciting at 340–365 nm anddetecting emission at 400–500 nm (Cárdenas et al., 2006; Hu et al., 2002; Lakowiczet al., 1992). The NAD(P)H endogenous fluorescence signal of single pollen tubewas assessed as described (Cárdenas et al., 2006). The fluorescence signal wasvisualized with a fluorescence microscope (Zeiss Axio Imager A1), equipped with aCCD camera and a Xenon excitatory light source. The filters were a 360 nm (10 nmbandpass) excitation filter and a 420–470 nm emission filter. The images werecaptured 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 totalpollen tubes. Pollens were precultured in a 5 ml centrifugal tube, and 2 hours laterthe tube was gently oscillated to ensure uniform pollen tube density throughout thetube. A pipette was used to add an equal volume of medium with suspended pollentubes into each well of a 96-well microplate. Thus there was an equal quantity ofpollen tubes in each well. When S-RNase or DPI was added to the well, pollen tubeswere immediately examined for NAD(P)H endogenous fluorescence and repetitivelyexcited by 345 nm light and emissions detected at 420 nm at intervals of 30 secondsover a total period of 20 minutes. The experiment was repeated three times.

Ca2+ currents of pollen tubeThe method used was previous published (Qu et al., 2007). After preculture, pollentubes were washed twice with deionized water and incubated in an enzyme solutionfor 2.5 hours at 32°C to release spheroplasts. The enzyme solution was composedof 1% (w/v) macerozyme R-10, 2.0% (w/v) cellulase RS-10, 0.7% (w/v) pectolyaseY-23 and 1% (w/v) bovine serum albumin. The enzyme solution was then exchangedwith 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 coatedwith Sylgard (184 silicone elastomer kit; Dow Corning, Midland, MI). The range ofpipette resistance was 15–35 in 10 mM CaCl2. The pipette solution comprised 1mM 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, adjustedto pH 7.3 and an osmolality of 1100 mOsM by Tris and D-sorbitol, respectively. ATPwas incorporated to delay rundown of currents (Forscher and Oxford, 1985) andGTP was incorporated to sustain possible G-protein-related activity (Edwards et al.,1989; Yawo and Momiyama, 1993). The free Ca2+ concentration in the pipettesolution was ~10 nM, calculated with the chemical speciation program GEOCHEM(Parker et al., 1987). Whole-cell plasma membrane currents were measured using anAxon 200B amplifier (Axon Instruments, Foster City, CA). The whole-cell

configuration was obtained using a short burst of suction applied to the pipetteinterior to rupture the membrane, resulting in a substantial increase in capacitance.Series resistance and capacitance were compensated accordingly. The membranewas held at a holding potential of 0 mV and then the voltage was either clamped atdiscrete 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. Datawere sampled at 2 kHz and filtered at 0.5 kHz, and then analysed using PCLAMP9.0 (Axon Instrument). Junction potentials were corrected according to Amtmannand Sanders (Amtmann and Sanders, 1997). All experiments were conducted atroom temperature (20–22°C). The permeability of the channels to Ca2+ relative tochlorine (Cl–) (PCa/PCl) was estimated using an equation derived from the Goldman–Hodgkin–Katz equation (Goldman, 1943; Hodgkin and Katz, 1949).

Actin cytoskeleton fluorescent labellingPollen was grown in the basal medium for 2 hours at 25°C and then incubated withS-RNase, DPI or TMPP for 30 minutes. Samples were treated with freshly preparedm-maleimidobenzoyl-N-hydroxy-succinimide ester at a final concentration of 200M in basal medium for 7 minutes to stabilize the actin filaments according topublished methods (Liu et al., 2007). They were subsequently fixed for 1 hour with4% formaldehyde in phosphate-buffered saline buffer. The samples were washedthree times in phosphate-buffered saline and then placed on a cover-slide previouslysmeared with poly-L-lysine. Surplus buffer was removed with filter paper, and thesamples stained with 5 g/ml fluorescein isothiocyanate-phalloidin in 5% dimethylsulfoxide, 5 mM EGTA and 10% (w/v) sucrose, pH 6.9. Specimens were washedwith buffer before mounting in glycerol. The stained samples were investigated witha Zeiss fluorescence microscope and the pollen tubes with actin cytoskeletondepolymerization were counted. In each treatment, ≥100 pollen tubes were countedand the experiment repeated three times.

Pollen tube nuclear DNA stainPollen was grown in the basal medium for 2 hours at 25°C and then incubated withDPI, TMPP, CB or phalloidin for 30 minutes. To assess the extent of nuclear DNAdegradation, pollen tubes were fixed in 95% ethanol:glacial acetic acid (3:1) for 1hour at 4°C, transferred into 70% ethanol at –20°C for at least 4 hours, and thenstained with 0.05 g/ml DAPI in citrate buffer at pH 4.1 for 2 hours. The stainedsamples were examined with a Zeiss Axioskop40 fluorescence microscope. Ineach treatment, ≥100 pollen tubes were counted and the experiment repeated threetimes.

Pollen tube nuclear DNA degradation detection in vivoTo check nuclear DNA of pollen tubes after compatible or incompatible pollination,the full-bloom flowers were emasculated. The Kosui pollen grains were transferredto the stigma of emasculated Imamuraaki flowers as a compatible pollination,whereas Kosui pollen grains were transferred to stigmas of Kosui flowers as anincompatible pollination. At 9 hours after pollination, the styles were collected andfixed in FAA (37% formaldehyde:glacial acetic acid:50% ethanol; 5:5:9) and storedat 4°C until used. The styles were washed thoroughly under running tap water andincubated 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 styleswere washed with citrate buffer (pH 4.1) and stained with 0.05 g/ml DAPI in citratebuffer at pH 4.1 for 2 hours. Specimens were washed with citrate buffer again, andthen squashed on glass slides, and mounted in glycerol. The stained samples wereinvestigated with a Zeiss Axioskop40 fluorescence microscope. The DeadEndColorimetric TUNEL System (Promega, http://www.promega.com) was used for theindependent assessment of nuclear DNA degradation in pollen tubes after compatibleor incompatible pollination. After tissues were softened, the styles were processedfollowing the manufacturer’s instructions. Then the styles were stained with DAPIas previously mentioned. DAPI staining showed that the TUNEL-positive signalcorresponded to nuclear DNA. The samples were investigated with an OLYMPUSBX51 fluorescence microscope. The percentage of positive-TUNEL-reactive nucleito all visible nuclei was counted in each treatment. At least ten styles were observedper treatment and the experiment was repeated three times. Images were processedusing Adobe Photoshop CS.

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

Supplementary material available online athttp://jcs.biologists.org/cgi/content/full/123/24/4301/DC1

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