负压渗透处理对花粉管加载荧光染料的影响

该研究采用负压渗透技术,以正常培养2 h的丰水梨花粉为实验材料,探索负压渗透条件下,花粉管中加载Ca²⁺荧光探针(Fluo 4/AM)的方法。结果显示：(1)将花粉及花粉管进行负压处理2 h,花粉萌发率及花粉管的活性没有受到影响。(2)对培养2 h后的花粉管进行不同条件下的负压渗透处理,辅助荧光探针Fluo 4/AM进入花粉管;激光共聚焦显微镜观察发现,在低温(4 ℃)条件下,负压(-80 kPa)渗透加载荧光探针30 min,花粉管尖端可以观察到明显的Ca²⁺梯度。(3)抑制花粉管外Ca²⁺内流或降低花粉管外Ca²⁺浓度,花粉管中荧光密度也显著降低。研究认为,负压渗透辅助加载的方法可以有效促进荧光探针进入花粉管细胞内与Ca²⁺结合。     

Propidium Iodide Competes with Ca2+ to Label Pectin in Pollen Tubes and Arabidopsis Root Hairs

We have used propidium iodide (PI) to investigate the dynamic properties of the primary cell wall at the apex of Arabidopsis (Arabidopsis thaliana) root hairs and pollen tubes and in lily (Lilium formosanum) pollen tubes. Our results show that in root hairs, as in pollen tubes, oscillatory peaks in PI fluorescence precede growth rate oscillations. Pectin forms the primary component of the cell wall at the tip of both root hairs and pollen tubes. Given the electronic structure of PI, we investigated whether PI binds to pectins in a manner analogous to Ca²⁺ binding. We first show that Ca²⁺ is able to abrogate PI growth inhibition in a dose-dependent manner. PI fluorescence itself also relies directly on the amount of Ca²⁺ in the growth solution. Exogenous pectin methyl esterase treatment of pollen tubes, which demethoxylates pectins, freeing more Ca²⁺-binding sites, leads to a dramatic increase in PI fluorescence. Treatment with pectinase leads to a corresponding decrease in fluorescence. These results are consistent with the hypothesis that PI binds to demethoxylated pectins. Unlike other pectin stains, PI at low yet useful concentration is vital and specifically does not alter the tip-focused Ca²⁺ gradient or growth oscillations. These data suggest that pectin secretion at the apex of tip-growing plant cells plays a critical role in regulating growth, and PI represents an excellent tool for examining the role of pectin and of Ca²⁺ in tip growth.The apical wall of tip-growing cells participates directly in the process of growth regulation (McKenna et al., 2009; Winship et al., 2010), yet few methods permit monitoring the wall properties of living cells. Despite this, several recent studies have enhanced our understanding of the apical cell wall. Chemical analyses of isolated pollen tube wall material have revealed a complex mixture of pectic polysaccharides with regions comprising long sequences of polygalacturonic acid. Important patterns of pectin methoxylation have been detected using immunocytochemical approaches, but these are limited to fixed cells (Dardelle et al., 2010). In a recent study, Parre and Geitmann (2005) used microindentation to show significant correlations between wall strength and growth rate. None of these techniques allow for easy investigation of the cell wall during growth.In an earlier study, we found that propidium iodide (PI) vitally stains pollen tubes of lily (Lilium formosanum) and tobacco (Nicotiana tabacum) and in particular reveals with great clarity the thickened apical cell wall (Fig. 1; McKenna et al., 2009). In addition, the apical PI fluorescence oscillates and in lily pollen tubes correlates tightly with oscillations in wall thickness measured by differential interference contrast (DIC) optics. Finally, these studies indicated that the PI fluorescence predicted cell growth rates with high confidence, suggesting that PI binding may provide useful information about the physical and chemical properties of the cell wall.Open in a separate windowFigure 1.PI fluorescence and growth rate oscillate in lily pollen tubes (A and B), Arabidopsis root hairs (C–E), and Arabidopsis pollen tubes (F and G). A, The top panel shows a DIC image of a lily pollen tube, and the bottom panel shows PI fluorescence of the same tube. The PI fluorescence is pseudocolored, with white representing high signal and blue representing low signal. Bar = 10 μm. B, Growth rate (blue) and PI fluorescence (red) are plotted on a line graph. Both oscillate with the same period but with different phases. C, DIC image (top panel) and PI fluorescence image (bottom panel) of an Arabidopsis root hair. Bar = 10 μm. D, Two PI fluorescence images of the same root hair focused on the apex representing peak (top) and trough (bottom) PI signals. Bar = 5 μm. E, A line graph showing the growth rate (blue) and peak PI fluorescence at the apex (red) for the same root hair shown in C and D. F, The top panel shows a DIC image of an Arabidopsis pollen tube, and the bottom panel shows PI fluorescence of the same tube. The PI fluorescence is pseudocolored, with white representing high signal and blue representing low signal. Bar = 5 μm. G, Growth rate (blue) and PI fluorescence (red) are plotted on a line graph. Both oscillate with the same period but with different phases. The growth rate between individual 3-s frames was smaller than the pixel size for our optics in both Arabidopsis cell types; to remove the noise this generated, a four-image (pollen) or five-image (root hair) running average is shown. A.U., Arbitrary units.PI is commonly used to visualize plant cell walls by wide-field fluorescence and confocal microscopy (Fiers et al., 2005; Tian et al., 2006; Estevez et al., 2008) and to select viable cells during cell sorting (Deitch et al., 1982; Jones and Senft, 1985). A positively charged phenanthridine derivative, the propidium ion stains cell walls but does not pass through the intact cell membranes of living cells. It readily diffuses into dead cells and forms highly fluorescent complexes by intercalation between base pairs of double-stranded nucleic acids, thus acting as an excellent indicator for cell vitality (Hudson et al., 1969). Binding to cell walls presumably occurs by a different mechanism, since it is not accompanied by the dramatic increase in fluorescence and shift in absorption and emission maxima observed when PI binds to nucleic acids. The mechanism of PI binding needs further exploration, as does the potential for broader use in other tip-growing plant cells.In this report, we test two hypotheses: first, that PI stains other tip-growing cells with pectin-containing cell walls; and second, that PI and Ca²⁺ bind to the same sites in these walls. This binding would occur through the interaction of partial positive charges caused by localized deficits in π-orbital electrons associated with three of the four nitrogen atoms of PI (Luedtke et al., 2005) coordinating with negatively charged carboxyl and hydroxyl groups on homogalacturonans (HGs), as has been suggested in Oedogonium bharuchae (Estevez et al., 2008).Our findings indicate that both hypotheses are satisfied. Notably, oscillatory changes in apical PI fluorescence occur and are observed to anticipate oscillations in growth rate in Arabidopsis (Arabidopsis thaliana) root hairs and Arabidopsis pollen tubes. In addition, competition studies indicate that PI and Ca²⁺ bind to the same sites in cell walls. Supporting these studies, we demonstrate that pectin methyl esterase (PME) creates more sites for PI binding, presumably by demethoxylating HGs as they are secreted, and that pectinase reduces PI fluorescence dramatically. However, unlike other pectin-binding dyes, PI does not block Ca²⁺ channels at the concentration used in live cell studies, nor does it alter oscillatory growth characteristics. Our findings provide evidence that PI may be employed as a quantitative measure of Ca²⁺-binding sites and thus may have use as an indicator of the degree of cross-linking of HGs and of cell wall extensibility.     

DNA Is Taken Up by Root Hairs and Pollen,and Stimulates Root and Pollen Tube Growth

Phosphorus (P) enters roots as inorganic phosphate (P_i) derived from organic and inorganic P compounds in the soil. Nucleic acids can support plant growth as the sole source of P in axenic culture but are thought to be converted into P_i by plant-derived nucleases and phosphatases prior to uptake. Here, we show that a nuclease-resistant analog of DNA is taken up by plant cells. Fluorescently labeled S-DNA of 25 bp, which is protected against enzymatic breakdown by its phosphorothioate backbone, was taken up and detected in root cells including root hairs and pollen tubes. These results indicate that current views of plant P acquisition may have to be revised to include uptake of DNA into cells. We further show that addition of DNA to P_i-containing growth medium enhanced the growth of lateral roots and root hairs even though plants were P replete and had similar biomass as plants supplied with P_i only. Exogenously supplied DNA increased length growth of pollen tubes, which were studied because they have similar elongated and polarized growth as root hairs. Our results indicate that DNA is not only taken up and used as a P source by plants, but ironically and independent of P_i supply, DNA also induces morphological changes in roots similar to those observed with P limitation. This study provides, to our knowledge, first evidence that exogenous DNA could act nonspecifically as signaling molecules for root development.Phosphorus (P) is an essential macronutrient that limits plant growth in many situations due to a low availability in soils (for review, see Schachtman et al., 1998; Raghothama, 1999; Vance et al., 2003; Lambers et al., 2008). P enters plant roots as orthophosphates (P_i) via active transport across the plasma membrane (Smith et al., 2003; Park et al., 2007; Xu et al., 2007). Concentrations of P_i in soil solution are generally very low (<10 μm; Bieleski, 1973) and plants have evolved root specializations to access P from inorganic and organic sources (Raghothama, 1999; Hinsinger, 2001; López-Bucio et al., 2003; Vance et al., 2003; Lambers et al., 2008). Roots exude enzymes and chemicals to mobilize P directly from soil compounds or indirectly via enhanced activity of soil microbes, and form symbioses with P-mobilizing mycorrhizal fungi (Schachtman et al., 1998; Raghothama, 1999; Bucher, 2007).However, similar to other nutrients, notably nitrogen, research on P nutrition of plants has focused on inorganic sources although organic P (P_org) in soil can account for 40% to 80% of the total P pool of mineral and organic soils, respectively (Bower, 1945; Raghothama, 1999; Vance et al., 2003). P_org compounds in soils are derived from plant residues, soil biota, and from synthesis by soil microbes (Jencks et al., 1964). Soil P_org is composed primarily of phospholipids, nucleic acids, and phytin (Dyer and Wrenshall, 1941). Phytic acid (inositol hexaphosphate) and its salts phytate, account for a large proportion of the P_org pool of soils (Anderson, 1980). Nucleic acids (RNA, DNA) represent approximately 1% to 2% of the soil P_org pool (Dalal, 1977). It can be released from prokaryotic and eukaryotic cells after death and protected against nuclease degradation by its adsorption on soil colloids and sand particles (Pietramellara et al., 2009).Although P_org can be a substantial constituent of the soil P pool, its contribution to the P nutrition of plants is poorly understood. P_org can be converted to P_i via root-exuded enzymes (Tarafdar and Claassen, 1988; Marschner, 1995; Vance et al., 2003). Secretion of nucleolytic enzymes and breakdown of nucleic acid were considered the reason for the observed growth of axenic Arabidopsis (Arabidopsis thaliana) and wheat (Triticum aestivum) on nucleic acid substrates as the sole P source (Chen et al., 2000; Richardson et al., 2000).Whether plants take up intact DNA has not been reported. We recently showed that roots take up protein, possibly via endocytosis (Paungfoo-Lonhienne et al., 2008). We hypothesized that roots may take up DNA by a similar process and grew Arabidopsis in the presence of phosphorothioate oligonucleotides (S-DNA) labeled with Cy3-fluorescent dye. S-DNA has a sulfur backbone and cannot be digested by plant nucleases, allowing tracking DNA of known size into cells (Spitzer and Eckstein, 1988). We examined if S-DNA of 25 nucleotides in length enters root hairs and pollen tubes as both types of cells are strongly elongated and have similar polarized growth (Schiefelbein et al., 1993; Hepler et al., 2001). We also assessed if addition of DNA to the growth medium affects the morphology of roots and pollen tubes. Here, we present evidence that plants take up DNA and demonstrate that the presence of DNA in the growth medium enhances lateral branching of roots, and the length of root hairs and pollen tubes, irrespective of P_i supply.     

Simple,Rapid and Inexpensive Quantitative Fluorescent PCR Method for Detection of Microdeletion and Microduplication Syndromes

Because of economic limitations, the cost-effective diagnosis of patients affected with rare microdeletion or microduplication syndromes is a challenge in developing countries. Here we report a sensitive, rapid, and affordable detection method that we have called Microdeletion/Microduplication Quantitative Fluorescent PCR (MQF-PCR). Our procedure is based on the finding of genomic regions with high homology to segments of the critical microdeletion/microduplication region. PCR amplification of both using the same primer pair, establishes competitive kinetics and relative quantification of amplicons, as happens in microsatellite-based Quantitative Fluorescence PCR. We used patients with two common microdeletion syndromes, the Williams-Beuren syndrome (7q11.23 microdeletion) and the 22q11.2 microdeletion syndromes and discovered that MQF-PCR could detect both with 100% sensitivity and 100% specificity. Additionally, we demonstrated that the same principle could be reliably used for detection of microduplication syndromes, by using patients with the Lubs (MECP2 duplication) syndrome and the 17q11.2 microduplication involving the NF1 gene. We propose that MQF-PCR is a useful procedure for laboratory confirmation of the clinical diagnosis of microdeletion/microduplication syndromes, ideally suited for use in developing countries, but having general applicability as well.     

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.     

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.     

荧光标记在植物花粉管构造及生长特性研究中的应用

花粉(管)构造及花粉管生长特性长期以来一直是人们研究的焦点,近年来,随着生物学和新材料不断发展,新荧光标记技术的运用,植物花粉管生长特性的研究取得巨大进步.本文介绍了花粉管生长特性等研究中用到的主要荧光标记物质,包括:苯胺蓝、Flou-3、罗丹明鬼笔环肽、TUNEL标记以及荧光蛋白等的理化性质、作用原理,并对应用荧光标记探索植物花粉管构造及生长特性的研究进展进行了综述.     

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.     

Haustorial Role of Pollen Tubes

In angiosperms, the pollen tube is siphonogamous and its mainfunction is to carry the male gametes for double fertilization.In some taxa, as in Cucurbitaceae, the tube branches after enteringthe ovule, prior to fertilization. The tube may even swell andform a bulla. During post-fertilization development of the ovule,a portion of the tube may persist in the micropyle, or in theembryo sac, or in both, sometimes even in the micropyle of themature seed. Haustorial function has been presumed in a numberof taxa. In Grevillea, following fertilization, the pollen tube branchesat the micropyle, and the branches grow intercellularly intothe ovarian tissue where further branching occurs. A haustorialrole of the pollen tube is presumed from circumstantial evidence.In gymnosperms (for example, Cycas, Zamia and Ginkgo) the pollentube is nonsiphonogamous, arises from the distal (upper) poleof pollen grain, and grows laterally in the apical region ofthe nucellus. The tube branches in Cycas and Ginkgo but remainsunbranched in Zamia. These pollen tube branches are enucleate,and are not concerned with the transport of male gametes forfertilization. However, the haustorial role has been well documented.In Podocarpus, the pollen tube is siphonogamous and arises fromthe proximal (lower) pole of pollen grain. After traversingthe nucellus, the tube forms a bulla at the point of contactwith the female gametophyte, and several branches originatefrom the bulla. The pollen tube branches grow along the innersurface of the nucellus and the outer surface of the femalegametophyte. The haustorial role of the pollen tube branchesis uncertain. Procedures for convincingly demonstrating thehaustorial role of pollen tubes are discussed. Angiosperms, gymnosperms, pollen tube, bulla, fertilization, haustorial role     

Electrotropism of Nicotiana Pollen Tubes

Electrotropism of tobacco pollen tubes towards the anode wasanalysed. The threshold and saturation values for the electrotropismwere less than 50 mV mm^–1 and 200 mV mm^–1, respectively.The tropic response gradually increased with increasing durationto exposure, but no further increase in the tropic responsewas observed when exposure of the electric field was terminated.Pollen tubes growing towards the cathode had a tendency to burstin a strong electric field. These results suggest that an externallyapplied electric field acts as a motive force for electrotropismbut not as a trigger and that endogenous currents play a rolein tip growth of pollen tubes. Possible mechanisms responsiblefor the electrotropism of pollen tubes are discussed. (Received July 9, 1993; Accepted September 18, 1993)     

Microinjection of Guanine NucleotideAnalogues into Lily Pollen Tubes Results in Isodiametric TipExpansion

Abstract: Heterotrimeric and small G-proteins aresupposed to participate in tip growth of plant cells. Quantitative changes intip growth rate after introduction of non-hydrolysable guanine nucleotideanalogues (NA) into lily pollen tubes have been interpreted as support for thehypothesis that heterotrimeric G-proteins regulate tube elongation (Ma et al.,1999 IDREF="R243-11">11). Here, we report that microinjection of guanineNA into lily pollen tubes causes loss of growth polarity, resulting inisodiametric tip swelling. Our results are compatible with current modelssuggesting an involvement of plant Rho-related small G-proteins (Rop) in themaintenance of pollen tube polarity and in tip morphogenesis.     

Colonization of Wheat Root Hairs and Roots by Agrobacteria

Formation of extracellular structures in pure culture and in interaction with wheat root surface was studied by scanning and transmission electron microscopy. The effects of various factors (growth temperature as well as pretreatment of agrobacteria with kalanchoe extract, acetosyringone, and centrifugation) on formation of extracellular structures was tested. The data on Agrobacterium tumefaciens (wild-type strain C58 and mutants LBA2525 (virB2::lacZ) and LBA288 (without the Ti plasmid)) adhesion to wheat root surface and root hairs after pretreatment of agrobacteria with inducer of virulence genes (vir) acetosyringone were obtained. Formation of agrobacterial cell aggregates on wheat root hair tips was demonstrated. The proportion of root hairs with agrobacterial aggregates on the root hair tip insignificantly changed after pretreatment with acetosyringone but considerably increased after treatment of A. tumefaciens C58 and LBA2525 with kalanchoe leaf extract. The most active colonization of root hairs and formation of agrobacterial aggregates on hair root tips was observed at 22°C. The capacity of agrobacteria for adhesion on monocotyledon surface could be changed by pretreatment of bacteria with various surface-active substances. Bacterial cells subjected to centrifugation had a decreased capacity for attachment to both wheat root surface and root hairs. The relationship between the capacity for adhesion and pilus production in agrobacteria was considered.     

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.     

Use of Fluorescent Brighteners to Visualise the Sites of Cellulose Synthesis in the Root Hairs of Species of Peperomia

Branched and unbranched root hairs of Peperomia fraseri and P. blanda were examined by bright field transmission and fluorescence microscopy in the presence of the optical brightener Photine HV. Many highly fluorescent bands and spots were detected along the length of the root hairs, in addition to those in the expected site of tip growth at the apex of the hairs.     

Rapid Dye Decolorization Method for Screening Potential Wood Preservatives

We developed a new screening method for potential wood preservatives based on decolorization of the dye Remazol Brilliant Blue R by extracellular oxidative agents produced by wood decay fungi. Oxidative biodegradation of lignin yielded decolorized zones around and under fungal cultures on a dyed agar medium. Inhibitory effects were detected by direct observation and measurement of the decolorized zones.