Author for correspondence: Jinxing Lin Tel: 0086 10 62836211 Email: email@example.com
• Nitric oxide (NO) plays a key role in many physiological processes in plants, including pollen tube growth. Here, effects of NO on extracellular Ca2+ flux and microfilaments during cell wall construction in Pinus bungeana pollen tubes were investigated.
• Extracellular Ca2+ influx, the intracellular Ca2+ gradient, patterns of actin organization, vesicle trafficking and cell wall deposition upon treatment with the NO donor S-nitroso-N-acetylpenicillamine (SNAP), the NO synthase (NOS) inhibitor Nω-nitro-L-arginine (L-NNA) or the NO scavenger 2-(4-carboxyphenyl)-4, 4, 5, 5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO) were analyzed.
• SNAP enhanced pollen tube growth in a dose-dependent manner, while L-NNA and cPTIO inhibited NO production and arrested pollen tube growth. Noninvasive detection and microinjection of a Ca2+ indicator revealed that SNAP promoted extracellular Ca2+ influx and increased the steepness of the tip-focused Ca2+ gradient, while cPTIO and L-NNA had the opposite effect. Fluorescence labeling indicated that SNAP, cPTIO and L-NNA altered actin organization, which subsequently affected vesicle trafficking. Finally, the configuration and/or distribution of cell wall components such as pectins and callose were significantly altered in response to L-NNA. Fourier transform infrared (FTIR) microspectroscopy confirmed the changes in the chemical composition of walls.
• Our results indicate that NO affects the configuration and distribution of cell wall components in pollen tubes by altering extracellular Ca2+ influx and F-actin organization.
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Pollen tubes are tip-growing cells that grow relatively quickly within the female gametophyte to deliver sperm nuclei for fertilization. Thus, pollen tubes are an essential part of sexual reproduction in higher plants (Hepler et al., 2001). Recent studies have indicated that the tip-focused cytoplasmic Ca2+ gradient, the regulation of actin cytoskeleton organization and polar vesicular trafficking are critical components of the tip-growth mechanism in pollen tubes (Hepler et al., 2001; Šamaj et al., 2004). Disruption of the tip-focused Ca2+ gradient in pollen tubes leads to cessation of tip growth and formation of extensive actin filament bundles in the tip region (Lancelle et al., 1997), suggesting that higher concentrations of Ca2+ at the tips of pollen tubes suppress actin filament bundle formation (Yokota et al., 2000). In addition, the construction and composition of the cell wall and the configuration of its building materials are important features that regulate pollen tube growth (Chen et al., 2007). Furthermore, secretory vesicles move to areas of higher Ca2+, where the vesicles accumulate and release cell wall precursors by fusion to the membrane, subsequently resulting in cell wall expansion and cell growth (Holdaway-Clarke et al., 1997; Camacho & Malhó, 2003).
In recent years, nitric oxide (NO) has been described as a highly active gaseous signaling molecule with multiple biological functions in plants (Courtois et al., 2008), including stimulation of seed germination (Beligni & Lamattina, 2000), root formation (Lanteri et al., 2008) and other processes of plant growth and development (Guo & Crawford, 2005), and regulation of stomatal movement (Neill et al., 2008). NO is also involved in highly polarized tip growth. For example, endogenous NO plays a positive role in root hair formation in Arabidopsis and Lactuca sativa by taking part in the auxin response (Lombardo et al., 2006). Recently, Salmi et al. (2007) reported that NO plays a signaling role in gravity-directed cell polarity in germinating Ceratopteris richardii spores. With respect to pollen tube growth, NO is involved in the growth regulation and reorientation of Lilium longiflorum and Arabidopsis pollen tubes (Prado et al., 2004, 2008). In addition, NO is thought to be involved in pollen–stigma interactions and defense in angiosperms (McInnis et al., 2006), and to play an inhibitory role in pollen germination and tube growth in Paulownia tomentosa in response to UV-B light (He et al., 2007). Previous reports have indicated that intracellular signaling via NO involves generation of cyclic guanosine monophosphate (cGMP) and cyclic ADP ribose (cADPR) and elevation of cytosolic Ca2+ (Lanteri et al., 2006). However, in many cases, NO-dependent physiological responses are governed by a complex signaling network, for which the biochemical and molecular mechanisms have not been deciphered (Neill et al., 2008). Furthermore, there are no data currently available to support the involvement of NO in cell wall construction of pollen tubes. Therefore, we investigated the regulatory roles of NO during pollen tube development in the gymnosperm Pinus bungeana. Specifically, we focused on the concentrations of NO in pollen tubes, as well as several linked cellular features that are essential for pollen tube tip growth, including extracellular Ca2+ uptake and the cytoplasmic Ca2+gradient, actin filament (AF) organization, and the composition of the cell wall during NO induction, or in response to pharmacological disturbance of NO production.
Materials and Methods
Mature pollen was collected from Pinus bungeana Zucc. trees growing in the Botanical Garden of the Institute of Botany, Chinese Academy of Sciences, in May 2007 and stored at −20°C. In vitro pollen culture was performed by liquid mass culture. After 30 min of rehydration at room temperature under 100% relative humidity, pollen grains were suspended in germination medium containing 15% sucrose, 0.01% H3BO3, and 0.01% CaCl2 at pH 6.8 on a shaker (121 rpm) at 25°C in the dark. The NO synthase (NOS) inhibitor Nω-nitro-L-arginine (L-NNA) was dissolved in 0.5 N HCl, while the NO donor S-nitroso-N-acetylpenicillamine (SNAP) and the scavenger 2-(4-carboxyphenyl)-4, 4, 5, 5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO) were dissolved in double-distilled water (ddH2O) and then added to the culture medium after pollen grains had been cultured for 60 h in standard medium. The pollen tubes were subjected to further incubation for another 24 h in the presence of different concentrations of drugs. For the germination experiment, various concentrations of SNAP, cPTIO and L-NNA were added to the medium from the beginning of culture and the germination percentage was calculated after 60 h. Moreover, the control was cultured in the presence of HCl, and all working concentrations of HCl were below 10−4 mol, a level necessary to dissolve L-NNA; there were no obvious effects on pollen germination or tube growth.
Pollen tube growth determination and morphological observation
To measure the mean tube length, at least 50 pollen tubes were detected in each of five replicates at 12-h intervals. Pollen grains were not considered germinated unless the tube length was greater than the diameter of the grain. The morphology of pollen tubes was examined with a Zeiss Q500 IW light microscope, and digital images were captured with a Spot II camera (Zeiss, Göttingen, Germany).
NO assays and NO imaging
The presence of NO in pollen tubes was assayed and visualized as previously described with small modifications (Prado et al., 2004). Samples were incubated in 5 µM 4, 5-diaminofluorescein diacetate (DAF-2DA; Merck, San Diego, California, USA) for 15 min and then excess fluorophore was washed out. The specimens were examined using a 488-nm argon laser under a confocal laser-scanning microscope (CLSM Zeiss 510 META) with the same parameter settings. Emission signals were collected at 500–550 nm. The relative fluorescence intensities of at least 50 pollen tubes in each of five replicates were measured using ImageJ (NIH, Bethesda, Maryland, USA), and mean relative fluorescence intensities were calculated.
Measurement of cytoplasmic Ca2+
Pollen tubes were pressure-injected with 2.5 mM Calcium Green-1 dextran (MW = 10 000; Molecular Probes Inc., Eugene, OR, USA) in 5 mM Hepes buffer (pH 7.0). The pipette tip reached no more than 3 µm into the cytoplasm of the pollen tubes, and the agents were gently loaded into the cytoplasm within c. 5 min, then the micropipette was slowly removed from the cell within c. 30 min to allow the cell to form a plug that healed the wound, and resume normal growth, thus guaranteeing that no cytoplasm was lost. Pollen tubes that suffered severe mechanical damage or leakage of cytoplasm were discarded. The samples were then excited with a 488-nm argon laser using the CLSM and emission signals were collected at 500–550 nm. Image analysis was carried out with CLSM 510 META software. The ‘steepness’ of the tip-focused Ca2+ gradient was employed to describe the effects of NO-modulating drugs on the maintenance of the tip-focused Ca2+ gradient (Lazzaro et al., 2003) (Supporting Information, Fig. S1). To determine whether injection affected pollen tube growth, Hepes buffer was microinjected into normally growing pollen tubes as a control injection.
Measurement of pollen tube tip extracellular Ca2+ fluxes
Net Ca2+ fluxes were measured using the scanning ion-selective electrode technique (SIET) as described previously (Holdaway-Clarke et al., 1997) with small modifications in Xu-Yue company (Sci. & Tech. Co. Ltd, Beijing, China; http://www.xuyue.net). Ca2+-selective microelectrodes with an external tip diameter of c. 3 µm were manufactured and salinized with tributylchlorosilane, and the tips were backfilled with commercially available ion-selective cocktails (Calcium Ionophore I – Cocktail A, 21048; Fluka, Busch, Switzerland). The self-referencing vibrating probe oscillated with an excursion of 10 µm, completing a whole cycle in c. 5.72 s. Extracellular Ca2+ fluxes at the tip of a pollen tube were measured by positioning the electrode tip on the normal to the tangent at the very tip and setting the direction of vibration to be parallel to the long axis of the pollen tube (Video S1). Pollen tubes selected for measurement were c. 100 µm in length and were growing normally. All experiments were repeated three times, and the Ca2+ fluxes of at least 10 pollen tubes were measured in each treatment each time. Pollen tubes that showed stable fluctuations in the preliminary detection were selected for the subsequent net Ca2+ flux measurements with SNAP, cPTIO or L-NNA, and representative changes upon different treatments were plotted against time. Furthermore, the mean values for different treatments were calculated from at least five pollen tubes to illustrate the Ca2+ flux variations upon different pharmacological applications. The data obtained were analyzed using an Excel spreadsheet to convert data from the background-mV estimation of concentration and the microvolt-difference estimation of the local gradient into specific ion influx (pmol cm−2 s−1) (Franklin-Tong et al., 2002).
Fluorescence labeling of F-actin
Labeling of F-actin was performed as previously described with small modifications (Chen et al., 2007). Samples were observed under CLSM with excitation at 488 nm and emission at 500–550 nm. All images were projected along the z-axis.
FM4-64 staining to analyze vesicle trafficking in the tube apex
Loading of cells with FM4-64 dye was generally achieved by direct application to the growing pollen tubes as described previously with slight modifications (Chen et al., 2007). The samples were excited at 514 nm with a 25-mW argon laser. Serial optical sections were performed every 20 s for c. 60 images 1 min after dye application until the fluorescence finally reached saturation, and the images were processed with CLSM 510 META software.
Fluorescent immunolabeling of pectins in the pollen tube cell wall
Immunolabeling of pectins in pollen tube cell walls was carried out according to Chen et al. (2007) with slight modifications. All the samples were excited at 488 nm under CLSM and emission signals were collected at 500–550 nm. Controls were prepared by omitting the primary antibody.
Fluorescence labeling of callose
Callose in the pollen tube cell wall was labeled with decolorized aniline blue as described by Chen et al. (2007). The stained samples were examined with differential interference contrast (DIC) followed by epifluorescence (ultraviolet excitation), and then photographed using a Zeiss Axioskop 40 microscope (excitation filter BP395-440; chromatic beam splitter FT460; barrier filter LP 470).
Fourier transform infrared (FTIR) analysis of the pollen tube walls
Pollen tubes were repeatedly washed with ddH2O four times and then dried in a layer on a barium fluoride window (13 mm diameter × 2 mm). FTIR spectra were recorded on a Perkin-Elmer Cetus MAGNA 750 FTIR spectrometer (Nicolet Corp., Tokyo, Japan) equipped with a mercury–cadmium–telluride (MCT) detector, and the Perkin-Elmer Cetus microscope was interfaced to a personal computer. An area of c. 100 × 100 µm was selected for FTIR analysis. The acquisition parameters were 8 cm−1 resolution, 128 co-added interferograms, and spectra were normalized to obtain relative absorbance (Chen et al., 2007).
Pollen tube growth and morphological observations
As shown in Tables 1–3, SNAP stimulated pollen germination and pollen tube growth in a dose-dependent manner (Table 1). By contrast, cPTIO and L-NNA significantly delayed pollen germination and pollen tube growth in a dose-dependent manner (Tables 2, 3). Furthermore, under control conditions, pollen tubes had a uniform diameter and a clear zone at the tip (Fig. S2a,e). However, pollen tube elongation was stimulated by 15 µM SNAP and the morphology of SNAP-treated pollen tubes did not show significant changes (Fig. S2b,f). In contrast, pollen tubes were shorter and many of them exhibited obvious abnormalities, including swollen tips, loss of the clear zone at the tube tips, and even bipolar growth, in the presence of 100 µM cPTIO (Fig. S2c,g) or 45 µM L-NNA (Fig. S2d,h) for 24 h.
Table 1. Effects of S-nitroso-N-acetylpenicillamine (SNAP) on pollen germination and mean tube length in Pinus bungeana
SNAP (µmol l−1)
Germination percentage (%)
Pollen tube length (µm)
Only pollen tubes that were longer than the diameter of the pollen grain were considered to have germinated. Values are the mean ± SD.
81.3 ± 3.12
75.6 ± 2.93
92.7 ± 4.17
102 ± 3.93
84.4 ± 2.58
74.9 ± 3.22
97.3 ± 3.79
110 ± 4.37
86.9 ± 3.02
75.1 ± 3.49
104 ± 4.68
117 ± 4.01
90.1 ± 2.76
74.7 ± 3.61
112 ± 5.22
124 ± 3.55
Table 2. Effects of 2-(4-carboxyphenyl)-4, 4, 5, 5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO) on pollen germination and mean tube length in Pinus bungeana
cPTIO (µmol l−1)
Germination percentage (%)
Pollen tube length (µm)
Only pollen tubes that were longer than the diameter of pollen grain were considered to have germinated. Values are the mean ± SD.
82.7 ± 3.87
74.4 ± 3.44
93.1 ± 3.48
100 ± 4.55
67.9 ± 3.07
75.2 ± 2.91
88.9 ± 4.02
95.8 ± 4.29
48.7 ± 2.95
74.6 ± 3.65
85.2 ± 4.73
89.0 ± 3.12
45.3 ± 2.33
73.7 ± 2.87
80.3 ± 4.59
83.4 ± 3.51
74.1 ± 3.05
76.2 ± 3.88
78.7 ± 3.43
Table 3. Effects of Nω-nitro-L-arginine (L-NNA) on pollen germination and mean tube length in Pinus bungeana
L-NNA (µmol l−1)
Germination percentage (%)
Pollen tube length (µm)
Only pollen tubes that were longer than the diameter of the pollen grain were considered to have germinated. Values are the mean ± SD.
80.9 ± 3.30
74.9 ± 3.12
93.3 ± 4.16
104 ± 3.14
73.8 ± 2.77
75.5 ± 3.26
86.6 ± 3.64
95.3 ± 3.66
67.5 ± 3.15
74.8 ± 3.44
82.8 ± 4.08
88.9 ± 3.05
47.6 ± 2.06
73.7 ± 3.67
78.1 ± 4.96
82.5 ± 4.81
74.3 ± 4.01
75.9 ± 5.19
77.3 ± 4.73
SNAP, cPTIO and L-NNA affect intracellular NO production
In the control, NO was distributed throughout nearly the entire tube in a compartmentalized fashion, but almost no fluorescence was detected in the tip region (Fig. 1a). In comparison, NO was distributed throughout almost the entire tube in cells treated with 100 µM cPTIO or 45 µM L-NNA, including the tip region (Fig. 1b,c), but fluorescence was weaker compared with that in control cells (Fig. 1b,c,e). In pollen tubes treated with 15 µM SNAP, significant fluorescence was detected almost along the entire tubes, except at the very tip (Fig. 1d,e).
Effects of NO on the intracellular Ca2+ gradient and extracellular Ca2+ flux at the pollen tube tip
Pollen tubes grown under normal conditions exhibited the typical tip-to-base cytoplasmic Ca2+ concentration gradient (Fig. 2a); whereas pollen tubes treated with 100 µM cPTIO showed a negligible cytosolic Ca2+ gradient (Fig. 2b). Similarly, treatment with 45 µM L-NNA also led to a narrow cytosolic Ca2+ gradient from tip to base (Fig. 2c). However, pollen tubes exhibited a sharper tip-focused Ca2+ gradient after treatment with 15 µM SNAP in comparison to that of control pollen tubes (Fig. 2d). Furthermore, the pollen tubes retained normal morphology and a well-characterized clear zone in the tip region after control injection, and recovered normal fountain-like cytoplasmic streaming (Video S2), indicating that injection did not affect pollen tube growth.
A consistent Ca2+ influx near the very tip and stable fluctuations were observed in normally growing pollen tubes within the first 360 s, and then 15 µM SNAP, 100 µM cPTIO or 45 µM L-NNA was added to the controls. Ca2+ influx significantly increased after addition of 15 µM SNAP, and then became relatively stable after c. 180 s (Fig. 3a). In addition, the range of influx in SNAP-treated pollen tubes was c. 77 to 130 pmol cm−2 s−1 (with a mean value of 100 ± 5.7 pmol cm−2 s−1; n = 15), whereas the influx in controls ranged from c. 48 to 82 pmol cm−2 s−1 (with a mean value of 63 ± 3.5 pmol cm−2 s−1; n = 15) (Fig. 3a). By contrast, treatment with 100 µM cPTIO led to a decrease in Ca2+ influx and the amplitude of influx ranged from 10 to 30 pmol cm−2 s−1 (with a mean value of 22 ± 2.3 pmol cm−2 s−1; n = 15), which was narrower than that of control pollen tubes (ranging from 41 to 78 pmol cm−2 s−1; the mean value was 59 ± 2.7 pmol cm−2 s−1; n = 15) (Fig. 3b). Similarly, the Ca2+ influx dramatically decreased upon treatment with 45 µM L-NNA and also showed a narrow flux range (from 15 to 36 pmol cm−2 s−1; the mean value was 27 ± 3.1 pmol cm−2 s−1; n = 15) in comparison to that of control pollen tubes (from 49 to 80 pmol cm−2 s−1; the mean value was 64 ± 3.2 pmol cm−2 s−1; n = 15) (Fig. 3c).
Stimulatory effects of L-NNA on actin polymerization and organization
Microscopic analysis revealed that the AFs were distributed throughout the entire pollen tube in a net axial array that was largely parallel to the direction of elongation in control tubes, except in the elongation tip region (Fig. 4a), where only a dense array of fine AFs was detected (Fig. 4b). Furthermore, a net axial array of AFs parallel to the direction of elongation was also observed in pollen tubes treated with 15 µM SNAP (Fig. 4c), but AFs at the tips of pollen tubes were depolymerized and even thinner (Fig. 4c,d). By contrast, thick actin bundles were detected along the entire length of pollen tubes treated with 100 µM cPTIO (Fig. 4e) or 45 µM L-NNA (Fig. 4g). Moreover, these thick actin bundles extended into the extreme tip of cPTIO- (Fig. 4f) or L-NNA-treated (Fig. 4h) pollen tubes.
FM4-64 staining following NO manipulation
Uptake of FM4-64 into pollen tubes was strictly time-dependent (Fig. 5). The plasma membrane was stained immediately after application of the dye. Bright spherical structures were observed in apical and subapical regions within 5 min (Fig. 5a). A typical reverse V-like staining pattern became apparent after c. 16 min (Fig. 5a). In SNAP-treated pollen tubes, the same staining pattern was observed in the cytoplasm within c. 10 min (Fig. 5b).
By contrast, a completely distinct staining pattern was observed in the cytoplasm of L-NNA-treated pollen tubes (Fig. 5c,d). In pollen tubes with a significantly swollen tip, FM dye uptake took place all along these swollen regions (Fig. 5c). In addition, FM dye uptake into the L-NNA-treated pollen tubes without balloon tips took place not only in the apical and subapical regions but also in the basal part of the pollen tubes, and finally the fluorescence was distributed almost evenly throughout the pollen tube (Fig. 5d). Furthermore, the uptake of FM4-64 in L-NNA-treated cells took c. 20 min, which was longer than in the controls (Fig. 5c,d), indicating possible variations in either cell wall construction or the pattern of dye internalization.
Effects of L-NNA on pectin distribution
In control pollen tubes, the distribution of JIM5-reactive (de-esterified) pectins was relatively uniform, except at the tip (Fig. 6a), whereas localization of JIM7-reactive (esterified) pectins was limited to the very tip of the growing tubes (Fig. 6c). By contrast, de-esterified pectins were detected across the entire surface of the pollen tubes treated with 45 µM L-NNA, including the tips (Fig. 6b), and esterified pectins were detected only in basal sites (Fig. 6d).
Effects of L-NNA on callose deposition
Callose was distributed almost evenly along the long tube shank in control pollen tubes, as shown by aniline blue staining (Fig. 6e). By contrast, strong fluorescence was detected in the tip region of the pollen tubes treated with 45 µM L-NNA, suggesting enhanced callose synthesis and deposition at the tip in response to L-NNA treatment (Fig. 6f).
FTIR analysis of wall components in pollen tubes
Typical FTIR spectra obtained from the tip region of control and 15 µM SNAP- or 45 µM L-NNA-treated pollen tubes are shown in Fig. 7(a). A saturated ester peak (representative of esterified pectins) was detected at 1743 cm−1 (Morikawa et al., 1978). The carboxylic acid peak (representative of de-esterified pectins) at c. 1419 cm−1 (Morikawa et al., 1978) was not distinct. To identify possible changes in pectins, differential spectra were generated by the digital subtraction of spectra for the tip region of control pollen tubes from those of 15 µM SNAP- or 45 µM L-NNA-treated pollen tubes. Based on these spectra, a carboxylic acid peak with a negative value appeared at c. 1419 cm−1, and a saturated ester peak at c. 1743 cm−1 exhibited increased absorbance intensity (Fig. 7b), indicating that exposure to 15 µM SNAP lead to decrease in the de-esterified pectin and increase in the esterified pectin content. By contrast, treatment with 45 µM L-NNA resulted in an increase in de-esterified pectin content and a decrease in esterified pectin content (Fig. 7c).
In recent years, NO has emerged as an important endogenous signaling molecule in plants with regulatory roles in many developmental and physiological processes, including tip growth (Prado et al., 2008). Recent in vitro studies have indicated that, as lily pollen tubes move into the NO gradient produced by SNAP, their growth is reduced or abrogated while the pollen tube reorients, and is then subsequently resumed (Prado et al., 2004). Our data show that pollen tube growth is stimulated by SNAP in a dose-dependent manner, while cPTIO and L-NNA inhibit pollen germination and pollen tube growth, accompanied by significant morphological alterations, indicating that cPTIO and L-NNA inhibit pollen germination and tube elongation in a dose-dependent manner, whereas He et al. (2007) reported that cPTIO and L-NAME (a NOS inhibitor) had no effect on pollen germination and tube growth in P. tomentosa. These differences in sensitivity to NO-modulating drugs may result in the general differences in responsiveness to SNAP, cPTIO or L-NNA that have been observed in several plant species. These differences may come about because P. bungeana pollen tubes grow far more slowly than angiosperm pollen tubes.
The involvement of NO in tip growth has been investigated using pharmacological agents to alter endogenous NO concentrations (Prado et al., 2004; He et al., 2007; Salmi et al., 2007). In the present study, SNAP was selected as the NO donor instead of sodium nitroprusside (SNP), because application of SNP as an NO donor has side effects (Planchet & Kaiser, 2006; Schröder, 2006). The results showed that endogenously generated NO was almost absent at the tip of control pollen tubes but was at higher concentrations behind the clear zone of the tip region, which are in accordance with the findings of Prado et al. (2004) in lily pollen tubes. These results indicate that an appropriate concentration of NO behind the tip region is a positive growth regulator, at least in gymnosperm pollen tubes. NO was found in the tip region of pollen tubes treated with cPTIO or L-NNA, because cPTIO or L-NNA disrupted the clear zone of pollen tubes. This result might give rise to potential changes in the regulation of Ca2+ influx as indicated by the results from the net Ca2+ flux measurements.
Ca2+ serves as a second messenger in a variety of plant physiological processes. There is a close coupling between the intracellular tip-focused Ca2+ gradient, extracellular tip-directed Ca2+ influx, and elongation of the pollen tube (Holdaway-Clarke & Hepler, 2003). Increasing evidence suggests that there is a connection between NO and Ca2+ signaling pathways. Pharmacological and biochemical studies have shown that NO signaling in plants is mediated by cGMP (Prado et al., 2004; Salmi et al., 2007) and cyclic nucleotide-gated ion channels (CNGCs), which are permeable to both monovalent and divalent cations (typically K+, Na+, and Ca2+) and are directly activated by cGMP and/or cAMP (Lanteri et al., 2006). Recently, Prado et al. (2008) reported that activation of Ca2+ influx in pollen tubes partially eliminated the balloon tips induced by cPTIO, indicating that a putative NO-cGMP signaling pathway is dependent on Ca2+ signaling, through effects on cytosolic Ca2+ concentrations during NO-induced pollen tube reorientation. On the basis of the results obtained regarding the Ca2+ gradient and Ca2+ influx in the present study, we conclude that NO regulates the cytoplasmic Ca2+ gradient largely by mediating Ca2+ influx, which is probably dependent on cGMP-activated channels in pollen tubes (Frietsch et al., 2007), to regulate pollen tube development. Furthermore, the 2-fold tip-focused Ca2+ gradient in conifer pollen tubes is much lower than the 10-fold gradient within 20 µm in angiosperm pollen tubes (Lazzaro et al., 2005). Considering the slow growth rate of gymnosperm pollen tubes in comparison to angiosperm pollen tubes and the previous results showing that the Ca2+ gradient was related to the polarized growth rate (Silverman-Gavrila & Lew, 2003; Cárdenas et al., 2008), we assume that the increase in growth rate upon SNAP treatment may partly result from the greater steepness of the tip-focused Ca2+ gradient in comparison to that of control pollen tubes, and that the slower growth rate of the pollen tube in response to cPTIO/L-NNA probably results from the lower steepness of the Ca2+ gradient.
In addition, accumulating evidence indicates that AFs control cytoplasmic streaming and hence the transport of secretory vesicles, and that actin polymerization itself also contributes to pollen tube growth (Chen et al., 2007). The dynamic state of the cytoskeleton is controlled via numerous regulatory factors, including several actin-binding proteins activated in response to Ca2+ (Cárdenas et al., 2008). Our results show that NO stimulation induced depolymerization of F-actin accompanied by a sharper Ca2+ gradient, and a reduction of NO induced the polymerization of F-actin in the pollen tube tip region accompanied by a negligible Ca2+ gradient. This suggests that F-actin organization in the tip region of pollen tubes sensitive to NO is partly dependent on the Ca2+ gradient during NO signaling in pollen tubes. Furthermore, the effects of cPTIO and L-NNA are reminiscent of previous studies which reported that polymerization of actin in pollen tube tips blocks tube growth (Cárdenas et al., 2005). Time-lapse images of L-NNA-treated pollen tubes showed a distinct FM dye staining pattern from that of controls, suggesting that vesicular trafficking was perturbed by the reduction of NO. This is consistent with a previous report showing that FM is a marker for polar growth (Camacho & Malhó, 2003). Interestingly, internalization of the dye was faster in SNAP-treated cells and slower in L-NNA-treated cells than in control cells. This is supported by results obtained by Lowenstein (2007) showing that NO accelerates endocytosis in cardiovascular cells. Taking these findings together, we speculate that NO stimulates endocytosis, indicating that changes in cell wall modeling may be partly dependent on the polymerization status of F-actin in pollen tubes.
The dynamic balance between cell wall extensibility and rigidity is another key factor that regulates tip growth in pollen tubes (Rockel et al., 2008). Immunolabeling with JIM5 and JIM7 showed that de-esterified pectins along the longitudinal axis seemed to decline gradually from the distal end towards the tube tip, whereas esterified pectins were present only in the tube tip region in control pollen tubes. This is consistent with data showing that the gradient in cell wall composition from apical esterified to distal de-esterified pectins, which is correlated with an increase in the degree of cell wall rigidity and a decrease in visco-elasticity, influences pollen tube growth and architecture (Parre & Geitmann, 2005). Compared with the controls, de-esterified pectins accumulated whereas esterified pectins decreased in concentration or disappeared completely in the tip region of L-NNA-treated pollen tubes. We propose that the conversion of esterified pectins to de-esterified pectins and/or the biosynthesis of esterified pectins is under the control of NO, and that the reduction in NO leads to excess wall rigidity at the tip of the pollen tube, which may partly account for the inhibition of pollen tube elongation in the presence of L-NNA. In addition, both types of pectin are sensitive to NO inhibitors because they are transported via either Golgi apparatus-based (esterified pectins) constitutive secretion (Geitmann et al., 1996) or endosomal recycling (de-esterified pectins) pathways (Baluška et al., 2002). Furthermore, in the present study, callose was deposited in the apical region of L-NNA-treated pollen tubes, indicating that NO spatially and developmentally mediates the synthesis and distribution of callose, thereby contributing to the mechanical properties of the cell wall and thus affecting pollen tube growth. This result confirms the findings that massive accumulation of callose in pollen tube tips is an important manifestation of abnormally growing tubes (Chen et al., 2007) and a common indicator of incompatible pollen (Guyon et al., 2004).
FTIR analyses showed distinct peaks corresponding to saturated esters (1743 cm−1) and other polysaccharides (1200–900 cm−1) in control pollen tubes. However, the saturated ester peak was relatively weak in inhibitor-treated tubes and relatively strong in donor-treated tubes. Peaks designated as pectins were observed in a difference spectrum, showing that there were fewer esterified pectins in the inhibitor-treated pollen tube tips than in the control tube tips and that there were more esterified pectins in the donor-treated tube tips. The FTIR results suggest that the NO signaling pathway significantly influences deposition of cell wall components such as carboxylic acids and pectins in the pollen tubes, further confirming the results obtained by immunolabeling with JIM5/JIM7.
In summary, our investigation into the effects of SNAP, cPTIO and L-NNA on P. bungeana pollen tubes provides a more global view of the role of NO in polarized tip growth in pollen tubes. We found that NO promotes extracellular Ca2+ influx, which may play a role in the maintenance of the tip-focused Ca2+ gradient and which may alter AF organization. The resulting differences in vesicle trafficking and cell wall construction lead to variations in tip growth (Fig. 8). This study produced two novel findings. Firstly, NO is indispensable for maintenance of the typical tip-focused cytosolic Ca2+ concentration gradient, partially through extracellular Ca2+ influx, in P. bungeana pollen tubes. Secondly, inhibitor treatment can cause changes in the main cell wall components of pollen tubes, such as pectins and callose, especially in the apical region. This combined cytological and biochemical study provides new insights into the multifaceted mechanistic framework for the functions of NO in polarized tip growth of pollen tubes.
This work was funded by the National Key Basic Research Program (2009CB119105) from MOST, the Knowledge Innovation Program of the Chinese Academy of Sciences, Grant No. KJCX2-YW-L08 and a key project from NSFC (30730009). We are indebted to Professor Yuan Ming and Professor Ren Hai-Yun for their expert advice concerning the microinjection.