应用水稻花粉离体萌发体系观察花粉管内微丝分布

采用非固定、DMSO渗透和异硫氰酸标记的鬼笔环肽（FITC—Ph）染色方法,观察水稻花粉离体萌发过程中花粉管内肌动蛋白微丝的形态和分布。结果表明：（1）水稻花粉水合2min后即可萌发,花粉管生长速度在600～1500μm／h之间。（2）水合而未萌发的花粉粒中,大量较短的梭形微丝束构成微丝网络结构,萌发过程中花粉粒内的梭形微丝束松解,部分微丝转移至萌发的花粉管内沿花粉管纵轴呈束状结构;随着花粉管的伸长,微丝束主要分布在花粉管中前端,但在花粉管顶端区域始终未见明显的微丝束。（3）水合后不能正常萌发的花粉粒内肌动蛋白微丝呈弥散不规则分布,在相同萌发时间生长迟缓的花粉管中,微丝束较少,且主要位于花粉管近萌发孔的部位。表明微丝骨架的形态和分布影响水稻花粉管的萌发和生长。     

杜仲花粉离体萌发特征及花粉管微丝骨架分布

以干燥的杜仲成熟花粉为材料,对杜仲花粉离体萌发的适宜液体培养基配方进行了筛选,并对花粉萌发特征及花粉管微丝骨架分布规律进行了研究。结果表明:(1)适宜杜仲花粉离体萌发的液体培养基组成为200g/L蔗糖+30mg/L硼酸+10mg/L Ca(NO3)2,于26℃条件下离体培养18h的花粉萌发率可达46.29%±3.75%。(2)在适宜液体培养条件下,杜仲花粉萌发率在培养6h内急剧增长,随后趋于平稳;而花粉管在培养8h内伸长较快,之后有放缓趋势,至培养48h时,花粉管长度可达363.14±30.51μm。(3)杜仲花粉属于2胞花粉,花粉萌发过程中,营养核和生殖核的移动存在一定的时序性,通常营养核先于生殖核进入到花粉管;杜仲花粉生殖核的有丝分裂发生在花粉管中,离体培养12h可逐渐观察到有丝分裂行为。(4)花粉萌发过程,微丝骨架形成束状,与花粉管伸长方向平行排布,与较为稀疏的网状微丝阵列组成连续系统,引导细胞核的运动。     

微丝骨架的构成及其对花粉管极性生长的调控作用

微丝骨架是细胞骨架的重要组成部分,它由肌动蛋白和肌动蛋白结合蛋白组成,广泛存在于真核细胞中。近年来,大量研究表明植物花粉及花粉管中存在丰富的微丝骨架。目前,在微丝骨架作为信号转导途径的靶标参与对花粉管极性生长的调控、微丝骨架在花粉和花粉管中的分布及其在花粉管生长过程中与其他信号分子之间的相互作用等方面取得了一系列突破性进展。     

Staining Germinating Pollen and Pollen Tubes

Germinating pollen on stigmas and pollen tubes in styles of Antirrhinum, Brassica, Oenothera, Raphanus, Rosa, solatium and Tagetes spp. were prepared for examination as follows: The styles were fixed in ethyl alcohol-acetic acid 3:1 for 1 hr, and hydrolyzed at 60°C for 5 to 60 min (depending on the species) in 45% acetic acid. The stigma with its attached strand(s) of stigmatoid tissue was then dissected out under a stereoscopic microscope, placed in a few drops of a staining solution made by dissolving 150 mg of safranin O and 20 mg of aniline blue in 25 ml of hot 45% acetic acid. After 5-15 min in this stain, the tissue was placed in a fresh drop of stain on a microscope slide and gently squashed under a cover glass. Because of a gradual precipitation of the aniline blue component, the stain had to be filtered regularly before use. However, a staining solution could be kept at room temperature for several weeks.     

Keto Acids in Pollen and Pollen Tubes of Sesbania aegyptica

Pollen and pollen tubes of Sesbania aegyptica Pers. contain α-ketoglutaric acid, oxaloacetic acid and pyruvic acid. Changes in the keto acids have been correlated with their corresponding amino acids during different phases of germination. It is suggested that keto acids were readily turned over during the elongation of pollen tubes.     

Distribution of Transglutaminase in Pear Pollen Tubes in Relation to Cytoskeleton and Membrane Dynamics

Transglutaminases (TGases) are ubiquitous enzymes that take part in a variety of cellular functions. In the pollen tube, cytoplasmic TGases are likely to be involved in the incorporation of primary amines at selected peptide-bound glutamine residues of cytosolic proteins (including actin and tubulin), while cell wall-associated TGases are believed to regulate pollen tube growth. Using immunological probes, we identified TGases associated with different subcellular compartments (cytosol, membranes, and cell walls). Binding of cytosolic TGase to actin filaments was shown to be Ca²⁺ dependent. The membrane TGase is likely associated with both Golgi-derived structures and the plasma membrane, suggesting a Golgi-based exocytotic delivery of TGase. Association of TGase with the plasma membrane was also confirmed by immunogold transmission electron microscopy. Immunolocalization of TGase indicated that the enzyme was present in the growing region of pollen tubes and that the enzyme colocalizes with cell wall markers. Bidimensional electrophoresis indicated that different TGase isoforms were present in distinct subcellular compartments, suggesting either different roles or different regulatory mechanisms of enzyme activity. The application of specific inhibitors showed that the distribution of TGase in different subcellular compartments was regulated by both membrane dynamics and cytoskeleton integrity, suggesting that delivery of TGase to the cell wall requires the transport of membranes along cytoskeleton filaments. Taken together, these data indicate that a cytoplasmic TGase interacts with the cytoskeleton, while a different TGase isoform, probably delivered via a membrane/cytoskeleton-based transport system, is secreted in the cell wall of pear (Pyrus communis) pollen tubes, where it might play a role in the regulation of apical growth.Transglutaminases (TGases [EC 2.3.2.13]; protein-Gln γ-glutamyltransferase) are a family of ubiquitous Ca²⁺-activated enzymes that are involved in animal cell morphogenesis and differentiation, apoptosis, cell death, inflammation, cell migration, and wound healing (Griffin et al., 2002; Mehta et al., 2006; Beninati et al., 2009). TGases are associated with different subcellular compartments, such as cytosol, plasma membrane, nucleus, mitochondria, and extracellular matrix. The specific localization of TGases is likely to determine both the biochemical activity and the type of proteins and/or substrates with which TGases react (Park et al., 2010). The distribution profile of TGase is affected by Ca²⁺, since the enzyme is preferentially associated with the lysosome compartment of liver cells in the absence of Ca²⁺ (Juprelle-Soret et al., 1984).TGase was initially detected in association with the cytosol, with the particulate (probably the microsomal) fraction (Birckbichler et al., 1976), and with the nucleus of animal cells (Remington and Russell, 1982). The association of TGase with the plasma membrane was related to its activity in promoting cell adhesion and to the interaction of cells with the extracellular matrix, while the presence of TGase in the nucleus is likely related to cell apoptosis (Griffin et al., 2002). How TGase is delivered to its final destination in animal cells remains to be clarified. Since the cytoskeleton is essential for the correct positioning of proteins in the cells, this interplay has often been studied in terms of potential substrates of TGase activity (Griffin et al., 2002). For example, the TGase-mediated incorporation of polyamines (PAs) stimulates actin polymerization (Takashi, 1988; Griffin et al., 2002). TGase was also found to associate with myosin in stress fibers of vascular smooth cells (Chowdhury et al., 1997). The association between TGase and microtubules (MTs) was initially studied in view of the importance of MTs in Alzheimer’s disease (Griffin et al., 2002), whereas the dynamics of MTs is also likely to be controlled by TGase (Al-Jallad et al., 2011). Interestingly, MTs are also a substrate of TGase activity in cells committed to apoptosis (Piredda et al., 1999). TGase was also shown to posttranslationally modify MT-associated proteins such as tau (Griffin et al., 2002).Information about the localization and function of TGases in plant cells is limited. Following the early evidence of an enzyme-based incorporation of PAs in plants (Serafini-Fracassini et al., 1988), a number of reports described the presence and role of TGase in nonphotosynthetic/photosynthetic tissues and in isolated chloroplasts (Serafini-Fracassini and Del Duca, 2008, and refs. therein). Attempts have also been made to examine the differences and similarities between plant and animal TGases. For example, a tobacco (Nicotiana tabacum) TGase was proposed to be involved in the programmed cell death (PCD) of the flower corolla (Della Mea et al., 2007); in such a case, TGase is likely to be released into the cell wall by a Golgi vesicle-based transport. Plant TGases might also be involved in protection against viruses (Del Duca et al., 2007) and in the self-incompatibility (SI) response involving pollen and stigma during sexual reproduction (Del Duca et al., 2010). Recently, different TGase isoforms were detected in meristematic apices of Jerusalem artichoke (Helianthus tuberosus) tuber sprouts (Beninati et al., 2013).The pollen tube is a widely investigated tip-growing plant cell (Lee and Yang, 2008). Studies are generally aimed at clarifying the many aspects related either to its growth or to rejection by the stigma/style. Early evidence for a role of PAs during pollen tube emergence (Bagni et al., 1981) was confirmed through the detection of PA binding via a Ca²⁺-activated TGase activity (Del Duca et al., 1997) and later by the identification of actin and tubulin as substrates of purified pollen TGase (Del Duca et al., 2009). In pollen, the enzyme affected the polymerization state and activity of actin filaments (AFs) and MTs (Del Duca et al., 2009) and existed as both soluble and cell wall associated (Di Sandro et al., 2010). Visualization of fluorescently labeled TGase products indicated that the cross-linking activity of TGase occurred at the apex of pollen tubes, in a basal region close to the pollen grain and within the pollen grain itself (Iorio et al., 2008). The enzyme was found as a soluble cytoplasmic form likely involved in the regulation of unspecified physiological processes (possibly associated with the cytoskeleton; Del Duca et al., 2009).Although the association of pollen TGases with organelles/vesicles has not been reported, an extracellular form of a Ca²⁺-dependent TGase was shown to be involved in pollen tube growth (likely as a modulator of cell wall building and strengthening). Moreover, pollen TGase was secreted in the incubation medium during germination, where it might catalyze the cross linking of PAs with secreted proteins (Di Sandro et al., 2010). This suggests that pollen TGase may be secreted through a vesicle-based mechanism. Finally, a TGase activity was also observed in planta, consistent with a possible role of TGase during tube migration through the style (Di Sandro et al., 2010) or in the SI response of pollen tubes (Del Duca et al., 2010).The pollen tube is an excellent model to study how a given plant protein is either secreted or delivered to its final destination. Although we know that actin and tubulin are substrates of TGase activity, and that the active enzyme is located in the cell wall and released outside, how TGase is distributed in the cells and how this process is dependent on cytoskeleton and membrane dynamics remain unknown. Here, we wanted to study in detail the localization and distribution of TGase in growing pollen tubes of pear (Pyrus communis) in relation to both cytoskeleton and membrane dynamics. The aim was to shed light on the mechanism by which TGase is transported and secreted, a process that is still not well understood even in animal cells. Specific antibodies that cross react with the TGase of pollen tubes were used to localize the enzyme in different membrane compartments and in the cell wall. The use of specific inhibitors indicated that the delivery of extracellular TGase is dependent on both AFs and membrane dynamics. Analysis by bidimensional electrophoresis (2-DE) showed that distinct TGase isoforms are associated with different cell compartments, suggesting that TGase might be differently regulated according to its position in the cell. Together, these data may contribute to our understanding of the mechanisms underlying pollen tube growth, an essential aspect of fertilization processes.     

Growing Pollen Tubes Possess a Constitutive Alkaline Band in the Clear Zone and a Growth-dependent Acidic Tip

Using both the proton selective vibrating electrode to probe the extracellular currents and ratiometric wide-field fluorescence microscopy with the indicator 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF)-dextran to image the intracellular pH, we have examined the distribution and activity of protons (H⁺) associated with pollen tube growth. The intracellular images reveal that lily pollen tubes possess a constitutive alkaline band at the base of the clear zone and an acidic domain at the extreme apex. The extracellular observations, in close agreement, show a proton influx at the extreme apex of the pollen tube and an efflux in the region that corresponds to the position of the alkaline band. The ability to detect the intracellular pH gradient is strongly dependent on the concentration of exogenous buffers in the cytoplasm. Thus, even the indicator dye, if introduced at levels estimated to be of 1.0 μM or greater, will dissipate the gradient, possibly through shuttle buffering. The apical acidic domain correlates closely with the process of growth, and thus may play a direct role, possibly in facilitating vesicle movement and exocytosis. The alkaline band correlates with the position of the reverse fountain streaming at the base of the clear zone, and may participate in the regulation of actin filament formation through the modulation of pH-sensitive actin binding proteins. These studies not only demonstrate that proton gradients exist, but that they may be intimately associated with polarized pollen tube growth.     

Fluorescence Microscopy in the Study of Pollen Grains and Pollen Tubes

Fresh pollen gains were either crushed directly in a 0.01% solution of acridine orange (0.1 M phosphate-citrate buffer, pH 5.2-5.4) or they were germinated previously in 5-25% sucrose solution (glass-distilled water of pH 5.0-6.0 with 100 ppm H₃BO₃) inside moist incubating chambers at 24-30° C. Observations and records were made by using ultraviolet or blue-violet light with suitably coupled exciter and barrier filters. When the pollen grains, tube walls or cytoplasm interfered with observation of a particular cell content, the same was either pressed or dissected out of the gain or the tube. The vegetative, generative or sperm cells as well as other protoplasmic contents, such as plastid-like bodies, lipid granules and mitochondria could be differentiated.     

Cytoskeleton in Pollen and Pollen Tubes of Ginkgo biloba L.

The distribution of F-actin and microtubules was investigated in pollen and pollen tubes of Ginkgo biloba L. using a confocal laser scanning microscope after fluorescence and immunofluorescence labeling. A dense F-actin network was found in hydrated Ginkgo pollen. When Ginkgo pollen was germinating,F-actin mesh was found under the plasma membrane from which the pollen tube would emerge. After pollen germination, F-actin bundles were distributed axially in long pollen tubes of G. biloba. Thick F-actin bundles and network were found in the tip of the Ginkgo pollen tube, which is opposite to the results reported for the pollen tubes of some angiosperms and conifers. In addition, a few circular F-actin bundles were found in Ginkgo pollen tubes. Using immunofluorescence labeling, a dense microtubule network was found in hydrated Ginkgo pollen under confocal microscope. In the Ginkgo pollen tube, the microtubules were distributed along the longitudinal axis and extended to the tip. These results suggest that the cytoskeleton may have an essential role in the germination of Ginkgo pollen and tube growth.     

Haustorial Pollen Tubes in Fuchsia boliviana

Persistent pollen tubes have been observed in Fuchsia boliviana.It is proposed that after fertilization the pollen tube becomesan important channel conveying food materials from the surroundingtissues to the embryo. Fuchsia, pollen tubes, callosic channel, Onagraceae     

Persistent Symmetry Frustration in Pollen Tubes

Pollen tubes are extremely rapidly growing plant cells whose morphogenesis is determined by spatial gradients in the biochemical composition of the cell wall. We investigate the hypothesis (MP) that the distribution of the local mechanical properties of the wall, corresponding to the change of the radial symmetry along the axial direction, may lead to growth oscillations in pollen tubes. We claim that the experimentally observed oscillations originate from the symmetry change at the transition zone, where both intervening symmetries (cylindrical and spherical) meet. The characteristic oscillations between resonating symmetries at a given (constant) turgor pressure and a gradient of wall material constants may be identified with the observed growth-cycles in pollen tubes.     

Spatial Scales of Pollen and Seed-Mediated Gene Flow in Tropical Rain Forest Trees

Gene flow via seed and pollen is a primary determinant of genetic and species diversity in plant communities at different spatial scales. This paper reviews studies of gene flow and population genetic structure in tropical rain forest trees and places them in ecological and biogeographic context. Although much pollination is among nearest neighbors, an increasing number of genetic studies report pollination ranging from 0.5–14 km for canopy tree species, resulting in extensive breeding areas in disturbed and undisturbed rain forest. Direct genetic measures of seed dispersal are still rare; however, studies of fine scale spatial genetic structure (SGS) indicate that the bulk of effective seed dispersal occurs at local scales, and we found no difference in SGS (Sp statistic) between temperate (N?=?24 species) and tropical forest trees (N?=?15). Our analysis did find significantly higher genetic differentiation in tropical trees (F _ST?=?0.177; N?=?42) than in temperate forest trees (F _ST?=?0.116; N?=?82). This may be due to the fact that tropical trees experience low but significant rates of self-fertilization and bi-parental inbreeding, whereas half of the temperate tree species in our survey are wind pollinated and are more strictly allogamous. Genetic drift may also be more pronounced in tropical trees due to the low population densities of most species.     

A Method for Staining Pollen Tubes in Pistil

A quadruple staining procedure has been developed for staining pollen tubes in pistil. The staining mixture is made by adding the following in the order given: lactic acid, 80 ml; 1% aqueous malachite green, 4 ml; 1% aqueous acid fuchsia, 6 ml; 1% aqueous aniline blue, 4 ml; 1 % orange G in 50% alcohol, 2 ml; and chloral hydrate, 5 g. Pistils are fixed for 6 hr in modified Carnoy's fluid (absolute alcohol:chloroform:glacial acetic acid 6:4:1), hydrated in descending alcohols, transferred to stain and held there for 24 hr at 45±2 C They were then transferred to a clearing and softening fluid containing 78 ml lactic acid, 10 g phenol, 10 g chloral hydrate and 2 ml 1% orange G. The pistils were held there for 24 hr at 45±2 C, hydrolyzed in the clearing and softening fluid at 58±1 C for SO min, then stored in lactic acid for later use or immediately mounted in a drop of medium containing equal parts of lactic acid and glycerol for examination. Pollen tubes are stained dark blue to bluish red and stylar tissue light green to light greenish blue. This stain permits pollen tubes to be traced even up to their entry into the micropyle.     

Fertilizing Pollen Tubes Parasitise Ovary Wall in Grevillea

Tissue-invasive pollen tube branching occurs in an Australianshrub, Grevillea banksii R.Br. (Proteaceae). Parenchyma andphloem of the ovary wall are soon infiltrated after fertilization,by intercellular branches and the lumina of xylem elements areentered. Ovular tissues are not affected. Grevillea banksii, pollen, haustoria, ovary wall     

Microfilament Distribution in Maize Meiotic Mutants Correlates with Microtubule Organization

Microtubules and microfilaments often codistribute in plants; their presumed interaction can be tested with drugs although it is not always clear that these are without side effects. In this study, we exploited mutants defective in meiotic cell division to investigate in a noninvasive way the relationship between the two cytoskeletal elements. By staining unfixed, permeabilized cells with rhodamine-phalloidin, spatial and temporal changes in microfilament distribution during maize meiosis were examined. In wild-type microsporocytes, a microtubule array that radiates from the nucleus disappeared during spindle formation and returned at late telophase. This result differed from the complex cytoplasmic microfilament array that is present at all stages, including karyokinesis and cytokinesis. During division, a second class of microfilaments also was observed in the spindle and phragmoplast. To analyze this apparent association of microtubules and microfilaments, we examined several meiotic mutants known to have stage-specific disruptions in their microtubule arrays. Two mutations that altered the number or form of meiotic spindles also led to a dramatic reorganization of F-actin. In contrast, rearrangement of nonspindle, cytoplasmic microtubules did not lead to concomitant changes in F-actin distribution. These results suggested that microtubules and microfilaments interact in a cell cycle-specific and site-specific fashion during higher plant meiosis.     

Hexose Transport in Growing Petunia Pollen Tubes and Characterization of a Pollen-Specific, Putative Monosaccharide Transporter

We investigated the molecular and physiological processes of sugar uptake and metabolism during pollen tube growth and plant fertilization. In vitro germination assays showed that petunia (Petunia hybrida) pollen can germinate and grow not only in medium containing sucrose (Suc) as a carbon source, but also in medium containing the monosaccharides glucose (Glc) or fructose (Fru). Furthermore, high-performance liquid chromatography analysis demonstrated a rapid and complete conversion of Suc into equimolar amounts of Glc and Fru when pollen was cultured in a medium containing 2% Suc. This indicates the presence of wall-bound invertase activity and uptake of sugars in the form of monosaccharides by the growing pollen tube. A cDNA designated pmt1 (petunia monosaccharide transporter 1), which is highly homologous to plant monosaccharide transporters, was isolated from petunia. Pmt1 belongs to a small gene family and is expressed specifically in the male gametophyte, but not in any other vegetative or floral tissues. Pmt1 is activated after the first pollen mitosis, and high levels of mRNA accumulate in mature and germinating pollen. A model describing the transport of sugars to the style, the conversion of Suc into Glc and Fru, and the active uptake by a monosaccharide transporter into the pollen tube is presented.The meiotic division of a pollen mother cell early in the development of the anther generates four immature male gametophytes. The gametophyte undergoes one mitotic division to generate one vegetative and one generative cell, after which the generative cell further divides to form two sperm cells (for review, see McCormick, 1993). The sperm cells are delivered to the female reproductive cells by unidirectional growth of the vegetative cell. This pollen tube grows through the stigma and style toward the ovules in the pistil. Much of the recent molecular research on the physiology of pollen tube growth focuses on specialized processes such as the incompatibility reaction, or on substances such as kinases, pectinases, polygalacturonases, and flavonols (Mascarenhas, 1993; McCormick, 1993). In contrast to this, one of the most striking phenomena of plant fertilization, the extreme speed and long-range capacity of pollen tube growth, has been poorly investigated on a molecular level. To enable the fast growth of the pollen tube, a rapid synthesis of wall material (Derksen et al., 1995) and a high energy supply is necessary. Therefore, a high level of sugar import is required (Schlüpmann et al., 1994).During maturation in the anther, pollen accumulates high levels of carbohydrates that represent the major part of the mature grain''s dry weight (Stanley and Linskens, 1974a; Pacini, 1996). After germination on a compatible stigma, the fast growth of the pollen tube is supported by the pistil. In the stylar fluids of petunia (Petunia hybrida) pistils the free sugars Suc, Glc, and Fru are available to the pollen tube (Konar and Linskens, 1966). After absorption by the pollen, sugars are utilized as an energy source and are converted to wall material as pectins, cellulose, and callose (Mascarenhas, 1993; Derksen et al., 1995). Constant de novo synthesis of cell wall material is essential because many of the carbohydrates used for wall synthesis are dissipated for participation in further pollen tube formation.The primary source of carbohydrates in the pistil and pollen lies in the photosynthesizing mature leaves, where assimilation takes place. After assimilation sugars are transported through the phloem to sink tissues such as the anther and stylar apoplast, mostly in the form of Suc (Bush, 1993). Nonetheless, it should be noted that the transmitting tract cells of the petunia pistils contain chloroplasts (B. Ylstra, personal observations) and might therefore also contribute directly to sugars in the stylar apoplast (Jansen et al., 1992). The final destination of the sugars, however, is the pollen, which requires translocation from the anther, stigma, and stylar apoplast over the pollen (tube) membrane.Several sugar transporters were identified in plants (Sauer and Tanner, 1989, 1993; Bush, 1993) and their genes were cloned and characterized by transgenic expression in yeast (Sauer and Tanner, 1993; Sauer and Stolz, 1994). The gene products were designated mono- and dissaccharide transmembrane symporters, and actively translocate sugars across plasma membranes driven by a proton-electrochemical potential (Stadler et al., 1995). The dissaccharide-symporter genes isolated are especially transcribed in mature leaves. The monosaccharide transporters reported so far are primarily transcribed in sink tissues such as young leaves and in storage and floral organs (Sauer and Tanner, 1993). In terms of sink-source relations pollen should be regarded as a strong sink, since it is not able to assimilate but requires high levels of starch and carbohydrates during maturation, germination, and growth.In vitro germination assays are helpful in the study of the growth requirements of pollen tubes. Different chemical constituents, pH, and viscosity could be related to the quality of pollen development and quantity of tube growth (Stanley and Linskens, 1974a, 1974b; Jahnen et al., 1989; Derksen et al., 1995). Suc was most commonly included in the in vitro germination medium as a carbon source. However, sugar import into germinating and growing pollen has only been studied to a limited extent in lily (Deshusses et al., 1981). We present physiological and biochemical experiments that enable us to describe the uptake of carbohydrates by pollen tubes in molecular terms.