| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Department of Botany, Stockholm University, S 106 91 Stockholm, Sweden
Received for publication June 16, 1998. Accepted for publication October 22, 1998.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
Key Words: conifer • cytoskeleton • microtubule • Picea abies; • Pinaceae • pollen • pollen tube
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
). Since these differences may be orchestrated by cytoskeletal organization, this study examines microtubule organization in germinated conifer pollen. In Norway spruce (Picea abies), pollen is released in the spring in a five-cell stage containing two degenerated prothallial cells and a body and stalk cell complex surrounded by the vegetative cell (Singh, 1978
). Pollen lands in the pollination droplet and germinates in the micropyle. The pollen tube tip secretes proteases and hydrolases (Pettitt, 1985
) as it slowly digests through integumentary tissue to reach the egg cell within the female gametophyte (Singh, 1978
). After the tube tip reaches the archegonium, the body and stalk cells separate, migrate into the tube, and the body cell undergoes mitosis to produce two sperm nuclei, but cell division is incomplete since only a partial cell plate forms in vivo (Dawkins and Owens, 1993
). One of these sperm nuclei fertilizes the egg. The gap between pollination and fertilization in P. abies is 2–3 wk (Singh, 1978
).
Angiosperms are the primary model systems for investigating the cytoskeleton in germinated pollen and pollen tubes. In angiosperms, both microfilaments and microtubules form a network in the vegetative cell within germinated pollen grains (reviewed in Pierson and Cresti, 1992
). Microfilaments are found throughout the pollen tube in a net axial array (reviewed in Pierson and Cresti, 1992;
Li et al., 1997
), but their presence in the elongating tip is controversial. Microfilaments are found in chemically fixed and cryofixed material, but recent evidence indicates that in living material microfilaments are excluded from the tip (reviewed in Taylor and Hepler, 1997
). The role of microtubules in angiosperm pollen tube growth remains controversial. Microtubules are found throughout the tube primarily in the cell cortex in either net helical or axial arrays, but are either excluded from the tip or found sparingly (Pierson and Cresti, 1992;
Li et al., 1997
). Much less is understood about the cytoskeleton in the pollination of conifers, an economically important plant group. Since P. abies pollen survives long-term storage and is easily germinated in vitro, it is a useful model system for investigating the role of the cytoskeleton in conifer pollen tube growth. Actin microfilaments in P. abies pollen tubes form an array that is parallel with the direction of elongation, ensheathes the vegetative nucleus, and is enriched at the tip (Lazzaro, 1996
). In the terminal 30 µm of the pollen tube, actin filaments form a 6-µm thick band beneath the plasma membrane, which ensheathes a microfilament depleted core (Lazzaro, 1996
). Within the pollen grain, actin microfilaments form a network in the vegetative cell that ensheathes the body and stalk cell complex. In addition, the body and stalk cells both contain actin microfilaments. The presence of microfilaments in the spermatogenous body cell is a fundamental difference from the angiosperms (Lazzaro, 1998
), where the spermatogenous generative cell reportedly lacks microfilaments but contains microtubules organized in bundles that ensheathe the nucleus (reviewed in Palevitz and Tiezzi, 1992
). Although microtubules have been identified with epifluorescence microscopy as parallel bundles in Pinus densiflora pollen tubes (Terasaka and Niitsu, 1994
), microtubule organization has not been thoroughly examined. Using primarily confocal microscopy, the organization of microtubules within the pollen grain, the body and stalk cells, the pollen tube, and the elongating pollen tube tip is described here.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
). After 24 h in a humid chamber at 30°C (Frankis and Grayson, 1990
), 95% of the pollen grains had germinated, with pollen tubes 200 µm long. The vegetative nucleus had moved into the elongating tube, while the body and stalk cells remained within the pollen grain. Germinated pollen grains were fixed directly on the agar surface for 1 h with 4% paraformaldehyde in PEM buffer (66 mmol/L PIPES, 1 mmol/L EGTA, 1 mmol/L MgSO4) at pH 7.0. Fixed pollen was then transferred to microfuge tubes and pelleted by centrifuging at 1400 x g for 10 s. Cell walls were then permeabilized using either enzymatic digestion of freeze shattering. For enzymatic digestion, pollen was then rinsed three times for 10 min each in PEM buffer at pH 5.0. Pollen tubes in the final pellet were fractured in a microfuge tube using a plastic pestle. This partially ruptured the tube cell wall and increased the enzymatic digestion of the body cell walls and the penetration of antibodies. The body cell wall was digested by agitating material for 1 h at 35°C in 1% cellulase (Onozuka R-10; Serva, Heidelberg, Federal Republic of Germany) and 1% macerase (Macerozyme R-10; Serva, Heidelberg, Federal Republic of Germany) in PEM buffer (pH 5.0) containing protease inhibitors (4 mmol/L phenylmethylsulfonyl fluoride, 20 µg/mL leupeptin, 50 µg/mL aprotinin, 10 µg/mL pepstatin).
As an alternative to enzymatic digestion, cell walls were permeabilized with freeze shattering using a method developed by Wasteneys, Willingale-Theune, and Menzel (1997)
. Pollen tube pellets were sandwiched between two glass slides. These slides were plunged into liquid nitrogen until they reached -180°C. The slide sandwich was then placed on a precooled aluminum block (-180°C), and even downward pressure was applied with another precooled aluminum block (-180°C). The pressure created small fractures in the pollen tube walls. Slides were then separated and placed on a third aluminum block at +20°C and as the pollen tubes warmed they were bathed in extraction buffer (phosphate-buffered saline, pH 7.5 plus 1% Triton X-100) for 30 min. The freeze shattering method greatly improved the percentage of pollen tubes with optimal cell morphology and immunolabeling.
The resulting samples from enzymatic digestion or freeze shattering were rinsed three times for 5 min each in blocking buffer (phosphate-buffered saline (pH 7.5), 1% bovine serum albumin, 1% TX-100) (Wick, 1993
). The pollen sample was then divided into three aliquots for parallel treatment with primary and secondary antibody, secondary antibody alone, or no antibody. The first aliquot was incubated for 16 h at 20°C in a mouse monoclonal antibody to alpha tubulin (Amersham N356; Amersham, Buckinghamshire, UK), diluted 1:100 in blocking buffer. Pollen tubes were then rinsed four times for 10 min each in blocking buffer and then incubated for 2 h at 35°C with a Cy3-conjugated goat secondary antibody to mouse immunoglobins (Sigma C2181; Sigma, St. Louis, Missouri, USA), diluted 1:200 in blocking buffer. The pollen tubes were then rinsed three times for 10 min each in blocking buffer and mounted in Moviol. As a control, the second aliquot was processed in parallel with the first, but incubated in blocking buffer without primary antibody followed by incubation with the Cy3-conjugated secondary at 1:200 to evaluate nonspecific binding of the secondary antibody. The third aliquot was processed in parallel with the first and second, but incubated in blocking buffer without primary or secondary antibodies to measure autofluorescence. Pollen grains were examined on either a BioRad MRC600 or a Zeiss LSM510 confocal laser scanning microscope. Nomarski DIC images were collected simultaneously through the confocal scan head. Optical serial sections were collected at a thickness of 400 or 500 nm. Confocal projections were generated by digitally combining adjacent serial sections into a single image with the public domain program NIH Image on a Macintosh computer. Identical confocal settings were always used for both treated pollen tubes and controls. Confocal projections were used for the measurement of pollen tube tips, and the measurements were calibrated with an image of a stage micrometer recorded through the confocal microscope.
For electron microscopy, germinated pollen was fixed in 5% glutaraldehyde in 66 mmol/L PIPES (Piperazine-N,N'-bis[2-ethane-sulfonic acid]) buffer (pH 7.0) for 1 h directly on germination slides, and then transferred to microfuge tubes. Pollen grains were subsequently rinsed in PIPES buffer, postfixed in 1% OsO4 for 1.5 h, rinsed again in PIPES buffer, dehydrated through an acetone series, and embedded in Spurr's resin. Thin sections (90 nm) were stained with uranyl acetate and lead citrate and examined on a Zeiss EM 906.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
|
|
|
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
) so both elements may coordinate to organize the cytoplasm within the pollen grain. However, the detailed organization of microtubules within the pollen grain differs from the organization of actin microfilaments, and these differences may reflect functional distinctions between the two cytoskeletal systems. Microfilaments form an enmeshing basket in the vegetative cell that surrounds the body and stalk cells (Lazzaro, 1998
). Microfilaments are also present within the body and stalk cells and form an array throughout the cell cortex, which is enriched beneath the plasma membrane (Lazzaro, 1998
). The microtubule array was not particularly enriched beneath the plasma membrane in either the body, stalk, or vegetative cells. In the angiosperms, microtubules form a network throughout the germinated pollen grain (Tiwari and Polito, 1988,
1990
) and form helical or longitudinal arrays in the generative cell, which ensheathe the nucleus (Derksen, Pierson, and Traas, 1985;
Pierson, Derksen, and Traas, 1986;
Raudaskoski et al., 1987;
Heslop-Harrison et al., 1988
).
Microtubule bundles can be traced between the body and stalk cells from the cytoplasm of the body cell to the adjoining cell wall and into the cytoplasm of the stalk cell. These cells are mitotic sisters (Singh, 1978
) and are connected by plasmodesmata. The organization of microtubules and the presence of plasmodesmata suggest that microtubules form a path for intercellular communication by projecting from the cytoplasm to interconnecting plasmodesmata. There is evidence that the cytoskeleton is involved in intercellular traffic via plasmodesmata. The movement protein from tobacco mosaic virus colocalizes with regions of the cytoplasm containing microtubules and microfilaments suggesting that protein transport through plasmodesmata may use the cytoskeleton (Heinlein et al., 1995
; McLean, Zupan, and Zambryski, 1995
). Microfilaments may control the size exclusion limit of plasmodesmata since cytochalasins and profilin, which depolymerize microfilaments, increase the size exclusion limit and diameter of plasmodesmata (White et al., 1994
; Ding, Kwon, and Warnberg, 1996
). However, since microtubules were not specifically detected within plasmodesmata in electron micrographs, one can only conclude from the confocal micrographs that the organization of microtubules provides a direct path up to and away from the plasmodesmata between the body and stalk cells. Since there was no continuity between microtubules in the body and stalk cells and the surrounding vegetative cell, this pathway appears specific for the plasmodesmata between body and stalk cells and not vegetative cells. In Picea glauca pollen germinated in vitro, plasmodesmata were observed between all three cells, but in pollen germinated in vivo, they were only observed between the body and stalk cells and not to the surrounding vegetative cell (Dawkins and Owens, 1993
). Although actin microfilaments have been detected in plasmodesmata in other plant material (White et al., 1994
), they were not seen in continuity between the body and stalk cells in Picea abies pollen (Lazzaro, 1998
).
Microtubule bundles extend through the pollen grain aperture and down the length of the tube in a generally axial orientation. This is similar to the overall organization of microtubules in Pinus densiflora (Terasaka and Niitsu, 1994
), Pinus sylvestris (de Win et al., 1996
), and in angiosperm pollen tubes (Derksen, Pierson, and Traas, 1985
; Pierson, Derksen, and Traas, 1986
; Raudaskoski et al., 1987
). Microtubules are enriched around the vegetative nucleus and are closely associated with the nuclear envelope. Actin microfilaments also form a net axial organization in these pollen tubes and are enriched around the vegetative nucleus (Lazzaro, 1996
). The possibility therefore exists that both microtubule and microfilament systems may be involved in the transport of the tube nucleus. In Pinus sylvestris, microtubules and microfilaments are closely associated in cryofixed material (de Win et al., 1996
). In angiosperms, treatment with cytoskeletal inhibitors indicates that microfilaments (Franke et al., 1972
; Heslop-Harrison et al., 1988
) and microtubules (Joos, van Aken and Kristen, 1994
; Åström, Sorri, and Raudaskoski, 1995
; Heslop-Harrison and Heslop-Harrison, 1996
) direct the movement of the vegetative nucleus. In Pinus densiflora, the microfilament motor protein myosin is found on the nuclear surface (Terasaka and Niitsu, 1994
). Myosin is also detected on the surface of the vegetative nucleus in angiosperms (Tang, Hepler, and Scordilis, 1989
; Heslop-Harrison and Heslop-Harrison, 1989
; Miller, Scordilis, and Hepler, 1995
), but the microtubule motor proteins kinesin and dynein are not (Cai et al., 1993
; Liu and Palevitz, 1996
; Moscatelli et al., 1998
).
The most intriguing observation in this study is the organization of microtubules in the pollen tube tip, where they are enriched in the terminal 20 µm of the pollen tube, coincident with the enriched actin microfilament zone (Lazzaro, 1996
). Microtubules in the cell cortex at the tip are oriented perpendicular to the growing axis. This orientation changes abruptly 20 µm from the tip to a net axial organization. The microtubules perpendicular to the elongating axis are coincident with the 6-µm thick microfilament band in the tip that ensheathes a microfilament-depleted core (Lazzaro, 1996
). In medial sections through the tip, microtubules are present in a net axial orientation and in the same place as the microfilament-depleted core (Lazzaro, 1996
), so this core region, which contains small organelles, does have a cytoskeletal (microtubule) network. The cytoskeletal organization of the tip is very different from that in angiosperm pollen tubes (reviewed in Pierson and Cresti, 1992
; Li et al., 1997
), where microtubules are either not observed in the tips or form short elements that are not organized in a radial array (Derksen, Pierson, and Traas, 1985
; Raudaskoski et al., 1987
; Del Casino et al., 1993
). However, in fern protonemal cells, microtubules form a specific radial array at the growing tip, which shifts to a net axial array farther back in the tube (Kadota and Wada, 1992
), similar to the organization seen in Picea abies.
Why are microtubules enriched in the growing tip and oriented perpendicular to the growing axis? Conifer pollen tube tips secrete proteases and hydrolases (Pettitt, 1985
) as they slowly digest their way through integumentary tissue to reach the egg cell within the female gametophyte (Singh, 1978
). Pollen tubes grow by the deposition of cell wall components at the tip (reviewed in Derksen et al., 1995
; Taylor and Hepler, 1997
). My working model is that the radial microtubules at the tip may orient the deposition of cellulose microfibrils at the tip region. The cell wall may develop in the terminal 20 µm but may be mature in older parts of the tube. Farther back in the tube where the microtubules are axial, their function may change to direct the migration of the nucleus and other organelles, rather than the growth of the cellulose microfibrils. There is an abrupt transition in microtubule organization from radial to axial in the pollen tube cortex 20 µm back from the tip. As the tube elongates and this transition region moves down the tube, microtubules are presumably probably broken down and reorganized. Microtubules are dynamic structures where tubulin dimers are continuously added and removed, and plant microtubules have a high degree of dynamic instability (Moore et al., 1997
) and can rapidly reorganize from transverse to longitudinal in living cells (Yuan et al., 1994
).
In conclusion, the organization of microtubules in the pollen grain and elongating tube of this conifer has some similarities to microtubule organization in angiosperm pollen, but there are important differences, especially in the elongating tip where they are enriched in a radial array. This work provides additional evidence that cytoskeletal organization in conifer pollen is unique.
|
FOOTNOTES |
|---|
2 E-mail:lazzaro{at}botan.su.se
|
LITERATURE CITED |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
Cai, G., A. Bartalesi, C. Del Casino, A. Moscatelli, A. Tiezzi, and M. Cresti. 1993 The kinesin immunoreactive homolog from Nicotiana tabacum pollen tubes: biochemical properties and subcellular localization. Planta 191: 496–506.[ISI]
Dawkins, M. D., and J. N. Owens. 1993 In vitro and in vivo pollen hydration, germination, and pollen tube growth in white spruce, Picea glauca (Moench) Voss. International Journal of Plant Science. 154: 506–521.[CrossRef][ISI]
Del Casino, C., Y. Li, A. Moscatelli, M. Scali, A. Tiezzi, and M. Cresti. 1993 Distribution of microtubules during the growth of tobacco pollen tubes. Biology of the Cell 79: 125–132.[CrossRef][ISI]
Derksen, J., E. S. Pierson, and J. A. Trass. 1985 Microtubules in vegetative and generative cells of pollen tubes. European Journal of Cell Biology 38: 142–148.
———, T. Rutten, T. van Amstel, A. de Win, F. Doris, and M. Steer. 1995 Regulation of pollen tube growth. Acta Botanica Neerlandica 44: 93–119.
De Win, A. H. N., B. Knuiman, E. S. Pierson, H. Geurts, H. M. P. Kengen, and J. Derksen. 1996 Development and cellular organization of Pinus sylvestris pollen tubes. Sexual Plant Reproduction 9: 93–101.
Ding, B., M. O. Kwon, and L. Warnberg. 1996 Evidence that actin filaments are involved in controlling the permeability of plasmodesmata in tobacco mesophyll. Plant Journal 10: 157–164.[CrossRef][ISI]
Franke, W. W., W. Herth, J. van der Woude, and D. J. Morré. 1972 Tubular and filamentous structures in pollen tubes: possible involvement as guide elements in protoplasmic streaming and vectorial migration of secretory vesicles. Planta 105: 317–341.[CrossRef][ISI]
Frankis, R. C., and G. K. Grayson. 1990 Heat-shock response in germinating pine pollen. Sexual Plant Reproduction 3: 195–199.[ISI]
Heinlein, M., B. L. Epel, H. S. Padgett, and R. N. Beachy. 1995 Interaction of tobamovirus movement proteins with the plant cytoskeleton. Science 270: 1983–1985.
Heslop-Harrison, J., and Y. Heslop-Harrison. 1989 Myosin associated with the surfaces of organelles, vegetative nuclei and generative cells in angiosperm pollen grains and tubes. Journal of Cell Science 94: 319–325.
———, and ———. 1996 Microtubules and the positioning of the vegetative nucleus and generative cell in the angiosperm pollen tube: a quantitative study. Proceedings of the Royal Society of London 263: 1299–1304.[CrossRef]
———, ———, M. Cresti, A. Tiezzi, and A. Moscatelli. 1988 Cytoskeletal elements, cell shaping and movement in the angiosperm pollen tube. Journal of Cell Science 91: 49–60.
Joos, U., J. van Aken, and U. Kristen. 1994 Microtubules are involved in maintaining the cellular polarity in pollen tubes of Nicotiana sylvestris. Protoplasma 179: 5–15.
Kadota, A., and M. Wada. 1992 Reorganization of the cortical cytoskeleton in tip growing fern protonemal cells during phytochrome mediated phototropism and blue light induced apical swelling. Protoplasma 166: 35–41.[CrossRef][ISI]
Lazzaro, M. D. 1996 The actin microfilament network within elongating pollen tubes of the gymnosperm Picea abies (Norway spruce). Protoplasma 194: 186–194.[CrossRef][ISI]
———. 1998 The spermatogenous body cell of the conifer Picea abies (Norway spruce) contains actin microfilaments. Protoplasma 201: 194–201.[CrossRef][ISI]
Li, Y., Moscatelli, G. Cai, and M. Cresti. 1997 Functional interactions among cytoskeleton, membranes, and cell wall in the pollen tube of flowering plants. International Review of Cytology 176: 133–199.[ISI][Medline]
Liu, B., and B. A. Palevitz. 1996 Localization of a kinesin-like protein in generative cells of tobacco. Protoplasma 195: 78–89.[CrossRef][ISI]
McLean, B. G., J. Zupan, and P. C. Zambryski. 1995 Tobacco mosaic virus movement protein associates with the cytoskeleton in tobacco cells. Plant Cell 7: 2101–2114.[Abstract]
Miller, D. D., S. P. Scordilis, and P. K. Hepler. 1995 Identification and localization of three classes of myosins in pollen tubes of Lilium longiflorum and Nicotiana alata. Journal of Cell Science 108: 2549–2653.
Moore, R. C., M. Zhang, L. Cassimeris, and R. J. Cyr. 1997 In vitro assembled plant microtubules exhibit a high state of dynamic instability. Cell Motility and Cytoskeleton 38: 278–286.[CrossRef][ISI][Medline]
Moscatelli, A., G. Cai, F. Ciampolini, and M. Cresti. 1998 Dynein heavy chain-related polypeptides are associated with organelles in pollen tubes of Nicotiana tabacum. Sexual Plant Reproduction 11: 31–40.
Palevitz, B. A., and A. Tiezzi. 1992 Organization, composition, and function of the generative cell and sperm cytoskeleton. International Review of Cytology 140: 149–185.[CrossRef]
Pettitt, J. M. 1985 Pollen tube development and characteristics of the protein emission in conifers. Annals of Botany 56: 379–397.
Pierson, E. S., and M. Cresti. 1992 Cytoskeleton and cytoplasmic organization of pollen and pollen tubes. International Review of Cytology 140: 73–125.[CrossRef]
———, J. Derksen, and J. A. Traas. 1986 Organization of microfilaments and microtubules in pollen tubes grown in vitro or in vivo in various angiosperms. European Journal of Cell Biology 41: 14–18.
Raudaskoski, M., H. Åström, K. Pertillä, I. Virtanen, and J. Louhelian. 1987 Role of the microtubule cytoskeleton in pollen tubes: an immunocytochemical and ultrastructural approach. Biology of the Cell 61: 177–188.[ISI]
Singh, H. 1978 Embryology of gymnosperms. In W. Zimmermann, S. Carlquist, P. Ozenda, and H. D. Wulff [eds.], Handbuch der Pflanzenanatomie, vol. 10, part 2. Gebrüder Borntraeger, Berlin.
Tang, X., P. K. Hepler, and S. P. Scordilis. 1989 Immunochemical and immunocytochemical identification of a myosin heavy chain polypeptide in Nicotiana pollen tubes. Journal of Cell Science 92: 569–574.
Taylor, L. P., and P. K. Hepler. 1997 Pollen germination and tube growth. Annual Review of Plant Physiology and Plant Molecular Biology 48: 461–491.[CrossRef][ISI]
Terasaka, O., and T. Niitsu. 1994 Differential roles of microtubule and actin-myosin cytoskeleton in the growth of Pinus pollen tubes. Sexual Plant Reproduction 7: 264–272.[ISI]
Tiwari, S. C., and V. S. Polito. 1988 Organization of the cytoskeleton in pollen tubes of Pyrus communis: a study employing conventional and freeze-substitution electron microscopy, immunofluorescence, and rhodamine-phalloidin. Protoplasma 147: 100–112.[CrossRef][ISI]
———, and ———. 1990 The initiation and organization of microtubules in germinating pear (Pyris communis L.) pollen. European Journal of Cell Biology 53: 384–389.[ISI][Medline]
Wasteneys, G. O., J. Willingale-Theune, and D. Menzel. 1997 Freeze shattering: a simple and effective method for permeabilizing higher plant cell walls. Journal of Microscopy 188: 51–61.[ISI][Medline]
White, R. G., K. Badelt, R. L. Overall, and M. Vesk. 1994 Actin associated with plasmodesmata. Protoplasma 180: 169–184.[CrossRef][ISI]
Wick, S. M. 1993 Immunolabeling of antigens in plant cells. Methods in Cell Biology 37: 171–200.[ISI][Medline]
Yuan, M., P. J. Shaw, R. M. Warn, and C. W. Lloyd. 1994 Dynamic reorientation of cortical microtubules, from transverse to longitudinal, in living plant cells. Proceedings of the National Academy of Sciences, USA 91: 6050–6053.
This article has been cited by other articles:
|
|
 |
|
  X. Sheng, Z. Hu, H. Lu, X. Wang, F. Baluska, J. Samaj, and J. Lin Roles of the Ubiquitin/Proteasome Pathway in Pollen Tube Growth with Emphasis on MG132-Induced Alterations in Ultrastructure, Cytoskeleton, and Cell Wall Components Plant Physiology, August 1, 2006; 141(4): 1578 - 1590. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
X. Wang, Y. Teng, Q. Wang, X. Li, X. Sheng, M. Zheng, J. Samaj, F. Baluska, and J. Lin Imaging of Dynamic Secretory Vesicles in Living Pollen Tubes of Picea meyeri Using Evanescent Wave Microscopy Plant Physiology, August 1, 2006; 141(4): 1591 - 1603. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. D. Lazzaro, L. Cardenas, A. P. Bhatt, C. D. Justus, M. S. Phillips, T. L. Holdaway-Clarke, and P. K. Hepler Calcium gradients in conifer pollen tubes; dynamic properties differ from those seen in angiosperms J. Exp. Bot., October 1, 2005; 56(420): 2619 - 2628. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
