The Plant Cell, Vol. 3, 637-644, June 1991 O 1991 American Society of Plant Physiologists

Microfilament Distribution in Maize Meiotic Mutants Correlates with Microtubule Organization

Christopher J. Staiger' and W. Zacheus Cande Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, California 94720

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 severa1 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.

INTRODUCTION

Relatively little is known about the function of microfila- ments in plants, despite the fact that a large number of cell types reportedly contain elaborate microfilament net- works (Staiger and Schliwa, 1987; Lloyd, 1988). It is well established that actin filaments and the mechanochemical enzyme myosin provide the motive force for cytoplasmic streaming in some lower plants and fungi (for reviews, see Williamson, 1980; Kuroda, 1990). In higher plants, there is now good evidence that actomyosin functions in cyto- plasmic streaming, especially in pollen tubes (reviewed in Staiger and Lloyd, 1991). Although less is known about its role in other processes, actin may function in tip growth, in organelle movements, and in controlling the orientation of cell wall deposition (Staiger and Schliwa, 1987). During plant cytokinesis, the phragmoplast also contains F-actin interspersed among and parallel to the prominent micro- tubule array (reviewed in Staiger and Schliwa, 1987; Lloyd, 1988). Although their presence and specific orientation are suggestive (Kakimoto and Shibaoka, 1987, 1988), no con- vincing proof exists that microfilaments play a direct role in cell plate formation. Nevertheless, it is perhaps signifi- cant that microfilaments codistribute with microtubules at Severa1 stages of the plant cell cycle. Although a direct

' To whom correspondence should be addressed. Current ad- dress: Department of Cell Biology, John lnnes Centre for Plant Science Research, Colney Lane, Norwich, NR4 7UH, United Kingdom.

functional interaction could be assumed, means for testing this are limited. Classically, microtubule or microfilament inhibitors are applied and their effect? on cytoskeletal organization examined. However, such studies are of lim- ited value because cells are rarely treated at known de- velopmental stages, and their effects are even less com- monly observed on living tissues.

Microsporogenesis, the meiotic reduction of diploid mother cells to form haploid microspores, has the potential to be an ideal system for studying cell division and the cytoskeleton in higher plants. Meiosis in maize has re- ceived considerable scrutiny, and chromosome behavior during sporogenesis has been described thoroughly (re- viewed in Carlson, 1988). The pattern of divisions during maize sporogenesis is highly controlled and yields a pre- dictable end product (reviewed in Staiger and Cande, 1990). Moreover, a large collection of mutants that disrupt meiosis at various stages has been generated (see Carl- son, 1988; Golubovskaya, 1989). Finally, in maize, it is possible to take advantage of many powerful genetic techniques, including transposon tagging, to increase our chances of identifying the genes and gene products im- portant for higher plant cell division.

Microtubule utilization and reorganization during maize sporogenesis have been analyzed using indirect immuno- fluorescence microscopy (Staiger and Cande, 1990). Inter- phase, prophase, and late telophase microtubule arrays emanate from the nuclear envelope and radiate toward the

Figure 1. Rhodamine-Phalloidin-Stained Microfilaments and 4,6-Diamidino-2-phenylindole (DAPI)-Stained Chromatin of Wild-TypeMicrosporocytes.

(a) Random cytoplasmic microfilaments in a diakinesis pollen mother cell (PMC).(b) A prometaphase PMC with F-actin interspersed among the condensed chromosomes.

Maize Meiotic Mutants 639

cortex. It has been postulated that a site closely associated with the nuclear envelope serves as a microtubule nucleat- ing site during specific stages of the plant cell cycle. Meiotic spindles, during both divisions, have highly focused poles and a predictable orientation within the cells and in the anther locule. Cytokinesis follows each meiotic division and is accomplished by a typical phragmoplast that is initiated in the spindle midzone during late anaphase and telophase. The array of parallel phragmoplast microtubules propagates centrifugally, forming a ring around the newly formed cell plate, and cytokinesis is always completed before the next division ensues. An isobilateral tetrad of coplanar microspores is the ultimate product of this con- trolled and predictable pattern of meiotic divisions. Severa1 mutants are known to alter the normal progression of meiosis and can be correlated with defects in microtubule distribution. One such mutant, dv, affects spindle pole organization (Staiger and Cande, 1990). Specifically, this lesion disrupts microtubule-organizing center structure during the transition between a prophase microtubule array and the metaphase spindle. lnstead of converging to form focused poles, the metaphase spindle poles remain broad as in prometaphase. Another mutation, ms77, has defects including excess microtubules, abnormal spindle forma- tion, improper chromosome segregation, and cytokinetic failure (C.J. Staiger and W.Z. Cande, manuscript in prep- aration). Similar abnormalities can be induced in wild-type sporocytes treated with the microtubule-stabilizing drug taxol. These results are consistent with a model in which ms 7 7 disrupts meiosis by causing microtubule hypersta- bility and an excess of spindle or cytoplasmic microtubules.

In this paper we describe microfilament distribution dur- ing wild-type and mutant (dv and ms77) maize microspo- rogenesis. By comparing microfilament distribution with our prior observations on microtubule distribution (Staiger and Cande, 1990), we find that these two cytoskeletal elements codistribute in the spindle during metaphase and anaphase. Analysis of meiotic mutants that alter microtu- bule distribution in a stage-specific and site-specific man- ner demonstrated the interaction of microfilaments and

microtubules in the meiotic spindle and phragmoplast but not in other subcellular locations.

R E SU LTS

Methodology

The technique used to visualize F-actin differs significantly from that used to observe microtubules in maize sporo- cytes (Staiger and Cande, 1990). Attempts to localize microfilaments in aldehyde-fixed cells using actin antibod- ies or rhodamine-phalloidin were only partially successful (data not shown). Other researchers have reported similar difficulties in preserving plant microfilaments (Parthasara- thy et al., 1985; Seagull et al., 1987), and, consequently, severa1 new methods have been developed (reviewed in Lloyd, 1988). The most common alternative technique, and the one used in this study, involves treating unfixed, permeabilized cells with rhodamine-phalloidin (Traas et al., 1987, 1989). Because phalloidin binds to and stabilizes F-actin (Schliwa, 1986), it could be suspected of causing actin polymerization. Permeabilization of cells before rho- damine-phalloidin treatment should eliminate most G-actin, thereby reducing the chances of artificial polymerization. Another concern is the potential for phalloidin (or DMSO) to cause microfilament rearrangement in the unfixed cy- toplasm. To address these concerns, a third technique was explored (Sonobe and Shibaoka, 1989). In preliminary experiments on wild-type cells, the mild protein cross- linking reagent m-maleimidobenzoyl N-hydroxysuccinimide ester (MBS, 100 pM) was used to stabilize F-actin fila- ments. Only minor amounts of DMSO (0.1%) were present during this treatment, and cells were post-fixed in 4% paraformaldehyde before application of a rhodamine-phal- loidin solution. Microfilament distribution in MBS-treated cells was indistinguishable from the results described be- low and indicates, therefore, that microfilament arrays observed in unfixed, permeabilized cells are representative of actin distribution in the living cell.

Figure 1. (continued).

(c) DAPl staining of the cell shown in (b). (d) Spindle and other cytoplasmic microfilaments in a metaphase I cell. (e) DAPI-stained metaphase chromosomes of the cell shown in (d). (f) Spindle F-actin becomes concentrated between chromosomes and poles during anaphase, but is also present in the midzone. (9) A late anaphase PMC with dramatic accumulation of F-actin in the spindle midzone. (h) A telophase PMC with phragmoplast microfilaments normal to cell plate orientation, random cytoplasmic filaments, and an aggregation of rhodamine-phalloidin at the former spindle pole (arrowhead). (i) A dyad of secondary sporocytes with random cytoplasmic array. (i) Phragmoplast microfilaments during early telophase II. (k) A tetrad of microspores with reticulate cytoplasmic F-actin array. (I) A young microspore after release from the tetrad containing ordered cortical microfilaments. Magnification x700.

Figure 2. Comparison of Microtubule and Microfilament Distribution at Key Stages in Two Meiotic Mutants, dv and ms17.

(a) Spindle microtubules in a wild-type, metaphase I pollen mother cell (PMC) detected with antitubulin.(b) A similar metaphase PMC stained with rhodamine-phalloidin contains both cytoplasmic and spindle microfilaments.(c) DAPI-stained chromosomes of cell shown in (b).

Maize Meiotic Mutants 641

Microfilament Arrays in Wild-Type and Mutant Cells

Microfilament distribution during wild-type maize sporo- genesis is depicted in Figure 1. During prophase I, an extensive, random network fills the cytoplasm (Figure 1 a). Perinuclear staining increases during diplotene and diaki- nesis, but unlike microtubule distribution, the cytoplasmic array does not decrease in extent (Figure Ia). At nuclear envelope breakdown, microfilaments invade the nucleus and are interspersed with the chromosomes (Figures 1 b and 1 c). However, unlike microtubule distribution (Staiger and Cande, 1990), there is no bipolar organization to the microfilament network in prometaphase. The first obvious colocalization of microfilaments and microtubules occurs in the metaphase spindle (Figures I d and le). Spindle microfilaments are predominately aligned parallel to the long axis, but do not show any obvious association with chromosomes (Figure Id). Figures 2a and 2b show that the number of microfilaments in the spindle is relatively limited compared with the wealth of microtubules. In ad- dition to spindle staining, a complex array of cytoplasmic F-actin remains throughout division (Figures I d to 1 h). At anaphase, microfilaments between the chromosomes and poles appear to shorten, and prominent interzone fibers are obvious (Figure I f ) . Microfilaments accumulate in the spindle midzone coincident with phragmoplast formation. These may be organized in an irregular fashion (data not shown) or as severa1 bundles on each side of the equatorial region (Figure 19). As the phragmoplast propagates cen- trifugally, parallel microfilaments codistribute with micro- tubules; however, an unstained region corresponding to the cell plate is not always obvious (Figure 1 h). During telophase, a brightly stained focus is often visible on the dista1 side of each daughter nucleus approximately in the location of the former spindle pole (arrow; Figure 1 h).

A random cytoplasmic microfilament array is present in secondary sporocytes (Figure 1 i) and throughout the sec- ond meiotic division. Similar to division I, colocalization of F-actin with microtubules is observed in meiosis II spindles and phragmoplasts (Figure 1 j). Tetrads of microspores

have a reticulate array of microfilaments (Figure 1 k). This distribution differs from the radial microtubule organization observed in similarly staged microspores (see Figure 2r in Staiger and Cande, 1990). After microspore release from the tetrad, actin redistributes to the cell’s cortex (Figure 11). Similar to microtubule distribution at this stage (see Figure 2t in Staiger and Cande, 1990), the cortical micro- filaments converge on foci at opposite ends of the cell (Figure 1 I).

Two mutants that cause stage-specific disruptions to the microtubule arrays have been examined. The first, ms77, is initially perturbed during late prophase I (C.J. Staiger and W.Z. Cande, manuscript in preparation). Un- usual and abundant accumulations of microtubules are observed in the cortical cytoplasm during diplotene and diakinesis, and often remain throughout karyokinesis (data not shown). Spindle formation is also abnormal, with mul- tiple poles, and extra spindles occurring quite frequently (Figure 2d). Changes in F-actin distribution accompany only a subset of the aforementioned microtubule re- arrangements in ms77 cells. We have not observed ab- normal cortical F-actin arrays, even though atypical cortical microtubules are commonly found (data not shown). How- ever, microfilaments do accurately represent the abnormal arrangement of spindle microtubules. For example, individ- ual minispindles contain parallel arrays of F-actin that reflect the size and shape of the spindle (Figures 2e and 2f). A second mutant, dv, affects spindle formation, spe- cifically during the transition from a prophase microtubule array to the metaphase spindle (Staiger and Cande, 1990). lnstead of converging to form highly focused poles, the metaphase spindle remains divergent (Figure 29). Rhoda- mine-phalloidin staining of metaphase dv microsporocytes reflects this abnormal spindle structure (Figures 2h and 2i). Moreover, the focus of rhodamine-phalloidin staining observed during late telophase in wild-type cells (Figure 2j) is spread out in a linear fashion in mutant cells (Figures 2k and 21). Multiple spindles and phragmoplasts are formed during the second division in dv sporocytes, and these also contain F-actin (not shown).

Figure 2. (continued).

(d) An ms77/ms77 PMC with major spindle and microspindle (arrow) in common cytoplasm. (e) Microfilament distribution in a different ms77/ms77 PMC with multiple spindles demonstrates that minispindles contain F-actin (arrow). (f) The minispindle in (e) appears to contain only one chromosome (arrowhead). (9) The metaphase I spindle in this dv/dv sporocyte has broad, unfocused poles. (h) A similarly staged dv cell stained with rhodamine-phalloidin mirrors the divergent spindle structure. (i) Chromosome distribution of the cell shown in (h). (i) A wild-type telophase PMC with phragmoplast F-actin and a focus of staining at the former spindle poles (arrowhead). (k) lnstead of being concentrated in a small spot, the polar staining is broad (arrowhead), and nearly linear in a telophase dv meiocyte (outlined). (I) Chromosome distribution (arrows) of the cell shown in (k). Magnification x700.

642 The Plant Cell

DlSCUSSlON

Comparison of Microfilament Distribution in Wild-Type and Mutant Cells

Four classes of qctin microfilaments are observed during maize sporogenesis. A complex network of microfilaments is present during the earliest meiotic stages and persists throughout both divisions. During chromosome segrega- tion and cytokinesis, microfilaments codistribute with mi- crotubules in the spindle and phragmoplast. Following the meiotic divisions, microfilaments redistribute to the cortical cytoplasm. Similar arrays of microfilaments in the inter- phase cytoplasm, spindle, and phragmoplast have been observed by Traas et al. (1989) in a study of eggplant meiosis, as well as by VanLammeren and coworkers (1 989) in Gasteria. Lilium microsporocytes reportedly con- tain phragmoplast F-actin; however, they were found to lack a significant population of spindle microfilaments (Sheldon and Hawes, 1988). Instead, small stellate foci were associated with the metaphase chromosomes but did not invade the spindle (Sheldon and Hawes, 1988). An extensive cortical microfilament array like the one seen in maize microspores has only been reported in Gasteria (VanLammeren et al., 1989).

A dependence of microfilament distribution on micro- tubule arrays has been assumed based on colocaliza- tion and use of specific inhibitors (Traas et al., 1989; VanLammeren et al., 1989). In this study, we demon- strated that microfilament distribution is changed by mu- tations that alter microtubule organization. Specifically, F-actin codistributes with the multiple spindles and phrag- moplasts associated with two meiotic mutations. In the case of dv, F-actin staining reveals a picture similar to that obtained with antitubulin. Spindle form is altered, with poles remaining broad and divergent. The focused spot of rhodamine-phalloidin stain observed in late telophase is similarly disturbed by dv. Evidence that this pattern reflects the former spindle pole is substantiated by the observation that the staining is broad and spread out in dv sporocytes. These results are consistent with other findings suggesting an interaction of microtubules and microfilaments in the spindle and phragmoplast. For example, when microtu- bules are abolished by colchicine or cold treatment, F-actin is not found in spindle-like or phragmoplast-like arrays (Traas et al., 1989; VanLammeren et al., 1989). However, it is important to note that this dependence of microfila- ments on microtubules is apparently limited to certain stages and arrays. In ms77 sporocytes, a new microtubule array has been observed in the cortical cytoplasm during late prophase and metaphase (C.J. Staiger and W.Z. Cande, unpublished data). That microfilament distribution is not obligatorily coupled to microtubules is indicated by the fact that such cells do not have a similar accumulation of F-actin at this site. Therefore, it can be concluded that

microfilament distribution is linked to microtubule-contain- ing structures only during specific stages of the meiotic cell cycle, namely karyokinesis and cytokinesis. These results do not imply that perturbations in microfilament organization cause the meiotic defects and male sterility associated with these two mutants. On the contrary, in both examples, defects in F-actin distribution lag behind the most obvious lesion, which involves abnormalities in microtubule arrays.

Role of Actin in Division

Although localization studies similar to this one could be used to argue for the function of F-actin in cell division and cytokinesis, severa1 authors hold the view that the role of actin in cell division is limited (Mole-Bajer et al., 1988; McCurdy and Gunning, 1990). They cite as evidence the fact that cytochalasin treatment or actin antibody microin- jection does not inhibit chromosome to pole movement or cytokinesis in plant cells (Palevitz, 1980; Bajer et al., 1987a, 1987b; Schmit and Lambert, 1987, 1988; Lloyd and Traas, 1988). There is also a large body of literature citing work on animal and lower eukaryotic cells where no role for actin in anaphase chromosome movement has been found (reviewed in Cande, 1989). However, these results are contradicted by a recent study in which microin- jected phalloidin slowed anaphase chromosome to pole movement and reduced the extent of spindle elongation in Haemanthus endosperm (Schmit and Lambert, 1990). It should be emphasized that cytochalasins sometimes are not very effective in destroying plant microfilaments or only affect certain classes of microfilaments (Cande et al., 1973; Lancelle and Hepler, 1988; Tang et al., 1989). For example, VanLammeren and coworkers (1 989) report that cytochal- asin B preferentially destroyed cytoplasmic microfilaments, whereas spindle staining persisted. One possibility is that actin microfilaments serve as a structural component nec- essary for maintaining spindle integrity and are, therefore, only indirectly involved in chromosome movement. Alter- natively, microinjected phalloidin may affect overall cyto- plasmic viscosity and thereby indirectly affect chromosome separation. Spindle microfilaments may also function in nonchromosomal spindle transport. However, no inhibitor studies have addressed this issue, and such movements are an order of magnitude slower than cytoplasmic stream- ing (Bajer et al., 1987a).

The existence of microfilaments in the phragmoplast is not disputed, but their role in cytokinesis remains uneluci- dated. lsolated phragmoplasts contain both microtubules and microfilaments, with a large percentage of the micro- filaments arranged in parallel with the spindle long axis (Kakimoto and Shibaoka, 1988). Heavy meromyosin dec- oration, a technique used to study F-actin orientation, shows that most of these microfilaments are of uniform

Maize Meiotic Mutants 643

polarity, with their barbed ends facing the cell plate (Kaki- moto and Shibaoka, 1988). This orientation is as one would predict if microfilaments directed myosin-coated vesicles toward the cell plate. Eggplant sporocytes treated with cytochalasin failed to form phragmoplasts and yielded multinucleate products, leading the authors to conclude that F-actin is necessary for phragmoplast formation and function. However, similar treatment of other cell types disrupts neither phragmoplast integrity nor cell plate for- mation (Palevitz and Hepler, 1974; Palevitz, 1980). In addition to microtubule-associated F-actin, a raft of micro- filaments between the leading edge of the phragmoplast and the cortical division site has been observed in vacu- olate suspension culture cells (Kakimoto and Shibaoka, 1987; Traas et al., 1987; Lloyd and Traas, 1988). This observation has led to the conclusion that actin plays a role in guiding the phragmoplast to the cortical division site (Lloyd and Traas, 1988). However, sporocytes, including those of maize, do not exhibit a concentration of microfil- aments between phragmoplast and cortex, yet division planes are under strict control (this study; Traas et al., 1989; VanLammeren et al., 1989; Staiger and Cande, 1990). Perhaps the obvious endoplasmic microfilament arrays in maize meiocytes function in positioning and/or anchoring the telophase nuclei, thereby influencing division plane orientation. Additional careful studies will be neces- sary to define unequivocally the role of microfilaments during plant mitosis and cytokinesis.

To study the role of the cytoskeleton during cell division, we initiated a study of maize meiotic mutants. In this report we exploited mutants known to alter microtubule distri- bution and spindle organization to establish that F-actin organization depends on microtubules. Specifically, micro- filaments were found to codistribute with spindle and phragmoplast microtubules. However, the pattern of other cytoplasmic microfilament arrays is apparently unaffected by perturbations to microtubule organization. To charac- terize further the role of actin in cell division, it will be necessary to identify mutations whose first cytological defect involves an alteration in F-actin organization and to characterize these mutations at the molecular level.

METHODS

Maize (Zea mays) inbred strains A344 and W23 were grown in a greenhouse at the University of California, Berkeley, and staged as described previously (Staiger and Cande, 1990). Sporocyte microfilament arrays were visualized using rhodamine-phalloidin on unfixed, permeabilized cells as described by Traas et al. (1 987, 1989). Sporocytes from cut anthers were extruded into permea- bilization solution containing 50 mM Pipes (pH 7.0), 10 mM MgCI2, 10 mM EGTA, 0.35 M sucrose, 5% (v/v) DMSO, and 0.1% (v/v) Nonidet P-40. After 10 min of permeabilization, the solution was

replaced with an identical one containing 0.33 fiM rhodamine- phalloidin (Molecular Probes, Eugene, OR) and 1 pg/mL 4,6- diamidino-2-phenylindole (Sigma) to stain microfilaments and chro- matin, respectively. Staining was allowed to progress for 1 O min to 15 min. Cells were mounted in 2 mg/mL 1,4-diazobicyclo- (2,2,2)-octane (Aldrich Chemical Co.) to reduce fluorescence fad- ing and viewed immediately on a Zeiss Photoscope III equipped with epifluorescence optics and a rhodamine filter set. lmages were recorded on Kodak TMAX 400 film exposed at 1600 ASA and developed in Kodak 0-76.

ACKNOWLEDGMENTS

We thank members of the Cande lab and Clive Lloyd for their helpful comments on this manuscript. This work was supported by U.S. Department of Agriculture Grant 8901117 to W.Z.C. C.J.S. was supported by U.S. Department of Agriculture Predoc- toral Fellowship 84-GRAD-9-0015.

Received March 11, 1991 ; accepted April 19, 1991.

REFERENCES

Bajer, A.S., Vantard, M., and Mole-Bajer, J. (1 987a). Multiple mitotic transports expressed by chromosome and particle movement. Fortschr. Zool. 34, 171-186.

Bajer, AS., Vantard, M., Schmit, C., Cypher, C., Hewitt, P.C., Huynh, T.T., and Mole-Bajer, J. (1987b). Dynamics of micro- tubules and F-actin in higher plant endosperm mitosis, analyzed with immuno-gold and video microscopy. In The Cytoskeleton in Cell Differentiation and Development. ICSU Symposium Se- ries, R.B. Maccioni and J. Arechaga, eds (Washington, DC: IRL Press), pp. 25-36.

Cande, W.Z. (1989). Mitosis in vitro. In Mitosis: Molecules and Mechanisms, J.S. Hyams and B.R. Brinkley, eds (San Diego: Academic Press Inc.), pp. 303-326.

Cande, W.Z., Goldsmith, M.H.M., and Ray, P.M. (1973). Polar auxin transport and auxin-induced elongation in the absence of cytoplasmic streaming. Planta 111, 279-296.

Carlson, W.R. (1988). The cytogenetics of com. In Corn and Corn Improvement. Agronomy Series, No. 18, 3rd ed., G.F. Sprague and J.W. Dudley, eds (Madison, WI: American Society of Agron- omy), pp. 259-343.

Golubovskaya, I.N. (1989). Meiosis in maize: mei genes and conception of genetic control of meiosis. Adv. Genet. 26,

Kakimoto, T., and Shibaoka, H. (1987). Actin filaments and microtubules in the preprophase band and phragmoplast of tobacco cells. Protoplasma 140, 151 -1 56.

Kakimoto, T., and Shibaoka, H. (1 988). Cytoskeletal ultrastruc- ture of phragmoplast-nuclei complexes isolated from cultured tobacco cells. Protoplasma (suppl. 2), 95-1 03.

149-1 92.

644 The Plant Cell

Kuroda, K. (1990). Cytoplasmic streaming in plant cells. Int. Rev.

Lancelle, S.A., and Hepler, P.K. (1 988). Cytochalasin-induced ultrastructural alterations in Nicotiana pollen tubes. Protoplasma

Cytol. 121, 267-307.

(SUPP~. 2), 65-75. Lloyd, C.W. (1988). Actin in plants. J. Cell Sci. 90, 185-188.

Lloyd, C.W., and Traas, J.A. (1988). The role of F-actin in determining the division plane of carrot suspension cells. Drug studies. Development 102, 21 1-221.

McCurdy, D.W., and Gunning, B.E.S. (1 990). Reorganization of cortical actin microfilaments and microtubules at preprophase and mitosis in wheat root-tip cells: A double label immunofluo- rescence study. Cell Motil. Cytoskel. 15, 76-87.

Mole-Bajer, J., Bajer, AS., and Inoue, S. (1 988). Three-dimen- sional localization and redistribution of F-actin in higher plant mitosis and cell plate formation. Cell Motil. Cytoskel. 10,

Palevitz, B.A. (1 980). Comparative effects of phalloidin and cy- tochalasin B on motility and morphogenesis in Allium. Can. J.

Palevitz, B.A., and Hepler, P.K. (1974). The control of the plane of division during stomatal differentiation in Allium. II. Drug studies. Chromosoma 46,327-341.

Parthasarathy, M.V., Perdue, T.D., Witztum, A., and Alvernaz, J. (1985). Actin network as a normal component of the cytoskeleton in many vascular plant cells. Am. J. Bot. 72,

Schliwa, M. (1 986). The Cytoskeleton. An lntroductory Survey. Cell Biology Monographs. Vol. 13, (Wien: Springer-Verlag).

Schmit, A.-C., and Lambert, A.-M. (1 987). Characterization and dynamics of cytoplasmic F-actin in higher plant endosperm cells during interphase, mitosis, and cytokinesis. J. Cell Biol. 105,

Schmit, A.-C., and Lambert, A.-M. (1988). Plant actin filament and microtubule interactions during anaphase-telophase tran- sition: Effects of antagonist drugs. Biol. Cell 64, 309-31 9.

217-228.

Bot. 58, 773-785.

131 8-1 323.

2157-2166.

Schmit, A.-C., and Lambert, A.-M. (1990). Microinjected fluores- cent phalloidin in vivo reveals the F-actin dynamics and assem- bly in higher plant mitotic cells. Plant Cell 2, 129-138.

Seagull, R.W., Falconer, M.M., and Weerdenburg, C.A. (1987). Microfilaments: Dynamic arrays in higher plant cells. J. Cell Biol.

Sheldon, J.M., and Hawes, C. (1988). The actin cytoskeleton during male meiosis in Lilium. Cell Biol. Int. Rep. 12, 471-476.

Sonobe, S., and Shibaoka, H. (1989). Cortical fine filaments in higher plant cells visualized by rhodamine-phalloidin after pre- treatment with m-maleimidobenzoyl N-hydroxysuccinimide es- ter. Protoplasma 148, 80-86.

Staiger, C.J., and Schliwa, M. (1987). Actin localization and function in higher plant cells. Protoplasma 141, 1-12.

Staiger, C.J., and Cande, W.Z. (1 990). Microtubule distribution in dv, a maize meiotic mutant defective in the prophase to metaphase transition. Dev. Biol. 139, 231 -242.

Staiger, C.J., and Lloyd, C.W. (1991). The plant cytoskeleton. Curr. Opin. Cell Biol. 3, 33-42.

Tang, X., Lancelle, S.A., and Hepler, P.K. (1 989). Fluorescence microscopic localization of actin in pollen tubes: Comparison of actin antibody and phalloidin staining. Cell Motil. Cytoskel. 12,

Traas, J.A., Doonan, J.H., Rawlins, D.J., Shaw, P.J., Watts, J., and Lloyd, C.W. (1987). An actin network is present in the cytoplasm throughout the cell cycle of carrot cells and associ- ates with the dividing nucleus. J. Cell Biol. 105, 387-395.

Traas, J.A., Burgain, S., and Dumas de Vaulx, R. (1989). The organization of the cytoskeleton during meiosis in eggplant (Solanum melongena (L.)): Microtubules and F-actin are both necessary for coordinated meiotic division. J. Cell Sci. 92,

VanLammeren, A.A.M., Bednara, J., and Willemse, M.T.M. (1 989). Organization of the actin cytoskeleton during pollen development in Gasferia verrucosa (Mill.) H., Duval visualized with rhodamine-phalloidin. Planta 178, 531 -539.

Williamson, R.E. (1980). Actin in motile and other processes in plant cells. Can. J. Bot. 58, 766-772.

104,995-1004.

21 6-224.

541 -550.

DOI 10.1105/tpc.3.6.637 1991;3;637-644Plant Cell

C. J. Staiger and W. Z. CandeOrganization.

Microfilament Distribution in Maize Meiotic Mutants Correlates with Microtubule

 This information is current as of February 4, 2021

 

Permissions 8X

https://www.copyright.com/ccc/openurl.do?sid=pd_hw1532298X&issn=1532298X&WT.mc_id=pd_hw153229

eTOCs http://www.plantcell.org/cgi/alerts/ctmain

Sign up for eTOCs at:

CiteTrack Alerts http://www.plantcell.org/cgi/alerts/ctmain

Sign up for CiteTrack Alerts at:

Subscription Information http://www.aspb.org/publications/subscriptions.cfm

is available at:Plant Physiology and The Plant CellSubscription Information for

ADVANCING THE SCIENCE OF PLANT BIOLOGY © American Society of Plant Biologists