the role of actin during arabidopsis trichome morphogenesis · the role of actin during arabidopsis...

1

Chapter 22

THE ROLE OF ACTIN DURING ARABIDOPSIS TRICHOME MORPHOGENESIS

Dan Szymanski Department of Agronomy, Purdue University, West Lafayette, IN 47907 USA

Key words: actin, Arabidopsis, genetics, microtubule, morphogenesis, trichome

Abstract: Arabidopsis trichome (leaf hair) development is being used as a genetic system to study actin-dependent growth. Arabidopsis trichomes are unicellular structures that are derived from the epidermis. Trichome development has been established as a powerful model system to study the control of cell fate and pattern formation. Results from several recent papers suggest that a genetic analysis of trichome morphogenesis may yield insight into the role of the cytoskeleton during cellular morphogenesis. In particular, it is found that filamentous actin (F-actin) plays an important role during Arabidopsis trichome development. Specific rearrangements of the actin cytoskeleton occur during the development of the mature trichome cell. F-actin-disrupting drugs do not affect the establishment of polarity during trichome development; however, the maintenance and coordination of the normal pattern of cell growth are very sensitive to the same agents. In contrast, inhibitors that depolymerize microtubules severely inhibit cell polarization. Disruption of the actin cytoskeleton in growing trichomes phenocopies a class of mutations that cause a distorted trichome morphology. An analysis of cell shape and microfilament structure in wild-type, mutant, and drug-treated trichomes is consistent with a role for actin microfilaments in the maintenance and co-ordination of an established growth pattern during Arabidopsis trichome formation.

1. GENETIC ANALYSIS OF THE PLANT CYTOSKELETON

The actin cytoskeleton comprises an intricate filamentous network that is essential for the trafficking of organelles and vesicles in eukaryotic cells. This transport activity is thought to be necessary for the normal growth of many

2 Szymanski

plant cell types. The actin cytoskeleton is a dynamic filamentous structure. Understanding how the organization the actin cytoskeleton is regulated in response to both intrinsic and extrinsic cues is an important and difficult question in plant biology (reviewed in Staiger, 2000). A genetic analysis of this process may lead to important insights into the regulation of the actin cytoskeleton. This chapter describes the utility of Arabidopsis trichome genetics as a model system to identify genes that control actin organization.

To date, many advances have come from studies of the role of actin during pollen tube growth. In this system, a combination of approaches relate the localization and activity of actin and actin-binding proteins to pollen tube organization and expansion during tip growth (reviewed in Heslop-Harrison et al., 1986; Staiger et al., 1997; Taylor & Hepler, 1997; see also Vidali & Hepler, this volume). Because cytoskeletal function in most plant cell types is not amenable to biochemical approaches, it is important to develop genetic strategies to study cytoskeletal organization.

The power of a genetic approach to study the actin cytoskeleton has been established in organisms as diverse as yeasts, flies and worms (Ayscough, 1998). The feasibility of using genetic techniques to study the cytoskeleton and cell growth has also been demonstrated in plants. Reverse-genetic identification of mutations in Arabidopsis actin gene family members will provide important tools to study the function of actin (McKinney et al., 1995). Actin organization is important for normal coleoptile elongation in monocots (Thimann et al., 1992; Waller & Nick, 1997). The Yin-Yang mutant of rice displays an altered actin organization in coleoptile epidermal cells during auxin-stimulted cell elongation (Wang & Nick, 1998). Mutation of the Arabidopsis kinesin-like gene ZWICHEL (ZWI) leads to defects in trichome morphogenesis (Oppenheimer et al., 1997). The presence of more than 30 genes that affect different aspects of trichome morphogenesis suggests that a detailed analyses of this process will provide a unique experimental window into cytoskeletal organization (reviewed in Marks, 1997).

2. TRICHOME STRUCTURE AND FUNCTION

Trichomes, defined as hair-like appendages that extend from the epidermis of aerial tissues, are present on the surface of almost all terrestrial plants (Johnson, 1975). Trichomes comprise a very diverse set of structures, and many plants contain several types on a single leaf. These can be divided into two basic classes: 1) complex, multicellular structures comprised of several specialized cell types that have a secretory function. The aerial epidermis of 20–30% of vascular plants contain glandular trichomes (Duke, 1994). Secreted material often accumulates in the subcuticular space of the apical cell of the

Actin and trichome morphogenesis 3

glandular trichome. And 2), simple multicellular or unicellular trichomes that extend from of the epidermal surface but lack a secretory anatomy.

It has been difficult to clearly demonstrate the function of plant trichomes, but several ideas have gained widespread acceptance. The presence of trichomes increases the boundary layer thickness between the epidermal tissue and the environment, and can protect against heat and water loss. In many species, trichomes are thought to protect the plant against insect or pathogen attack. These defensive qualities can be due to either chemical components that are secreted from trichomes or via the physical structure of the cell that limits insect access to or mobility on vegetative tissues. Figure 1 illustrates a glandular alfalfa trichome (Fig. 1A) and a Mentzelia pumila hooked trichome (Figs. 1B–E), both of which have the capacity to non-specifically immobilize insects at the leaf surface.

Figure 1. Scanning electron micrograph of trichomes. (A) An alfala weevil larvae attached to the sticky glandular exudate of an alfalfa trichome. Copyright (1975) RE Shade, reproduced with permission. (B) Type 1 Mentzelia pumila trichome. (C) Type 3 M. pumila trichome. (D) An agromyzid fly entangled in trichomes on the surface of a M. pumila plant. (E) High magnification of (D) showing the antennae of the fly wedged between type 1 and type 3 trichomes; white arrowhead, type 3 trichome; white arrow, type 1 trichome; black arrow, fly leg. (B–E) Copyright (1998) National Academy of Sciences USA, reproduced with permission.

Based on the potential economic importance of trichomes for the pharmaceutical, insecticidal, flavor and textile industries, considerable effort has been spent to understand the interplay between the cellular organization and function of plant trichomes. For example, the ultrastructure and vacuolar organization of the multicellular acid-secreting trichomes of cowpea have been

4 Szymanski

examined in fixed specimens and in living cells (Lazzaro & Thomson, 1992, 1996). Cotton fibers are trichomes that develop from the outer integument cells of the ovule, and are an excellent model system to study cytoskeletal organization and cell wall synthesis. The microtubule and microfilament organization of developing cotton fibers has been examined in detail, (Seagull, 1986, 1990, 1992; Tiwari & Wilkins, 1995). The fine structure of trichomes from dozens of species of economic significance also has been documented (Rodriguiez et al., 1984). However, understanding how trichome structure and function is determined requires the use of molecular genetic techniques in conjunction with cell biological tools.

3. ARABIDOPSIS TRICHOME DEVELOPMENT

3.1 Trichome Initiation

Arabidopsis leaf trichomes are unicellular stellate trichomes that usually contain three branches. Historically, Arabidopsis trichome development has been used to address the question of how cell fate and pattern formation are regulated in the plane of the epidermis (reviewed in Larkin et al., 1997; Marks, 1997). Cell fate decision-making is complex and involves the concerted activity of at least seven genes that positively and negatively regulate trichome initiation. Many of the early acting genes encode putative transcriptional regulators (reviewed in Szymanski et al., 2000).

The ability of an epidermal cell to receive trichome differentiation signals is strictly regulated, and does not appear to be determined solely by the presence or absence of transcription factors. Cells that are mitotically cycling appear incapable of adopting the trichome cell fate (Pyke et al., 1991; Lloyd et al., 1994; Larkin et al., 1996). As leaf development progresses, leaf epidermal cells exit the mitotic cycle and undergo variable rounds of endoreduplication (DNA synthesis without cell division) (Szymanski & Marks, 1998). The ability of cells to receive trichome differentiation signals is limited to a narrow developmental window during the transition from the mitotic to the endomitotic cycle (Lloyd et al., 1994). Genes that alter the cell cycle also affect the spacing and tissue distribution of trichome formation, but the relationship between cell cycle status and the ability to receive trichome differentiation signals is not clear (Schnittger et al., 1998; Szymanski & Marks, 1998). Trichome development represents a powerful experimental system to understand how transcription factors and cell cycle parameters interact with cytoskeletal components to alter the fate and morphogenesis of a cell.


3.2 Trichome Morphogenesis

Once an epidermal precursor enters the trichome pathway, it undergoes a complex morphogenetic program (Hülskamp et al., 1994; reviewed in Hülskamp et al., 1998). Based on morphological landmarks, trichome growth can be divided into discrete stages (Szymanski et al., 1998), as shown in Figure 2. The first sign of trichome formation is isodiametric expansion within the plane of the epidermis (stage 1). In cross section, the trichome precursor appears as an enlarged spherical cell. The cell nucleus is greatly enlarged, and it has been proposed that the trichome has undergone additional endoreduplication cycles relative to neighboring cells (Hülskamp et al., 1994). However, DNA content has not been measured in developing trichomes. During stage 1, neighboring socket cells form a tight association with the spherical trichome precursor. Stage 2 trichomes display obvious polarized elongation perpendicular to the leaf plane until the cell reaches a length of ~30 µm. The changes in cellular organization and shape that occur during stage 2 resemble tip-growing cells, but the nature of stage 2 growth has not been examined directly. During stage 3, branch initiation occurs sequentially on the developing stalk, often yielding a cell with three branches. The elongating branch buds initially have a blunt tip morphology (stage 4), but as the branches elongate, the tip morphology becomes more pointed (stage 5). The transition to stage 5 occurs very early in trichome development, and the vast majority of the cell volume is generated by diffuse growth during this phase. Once cell expansion has ceased, the cell wall acquires a papillate surface (stage 6). Although there is some variation between trichomes in the shape changes that occur during morphogenesis, each cell executes a similar developmental program.

Genetic and pharmacological experiments suggest that precise cytoskeletal function is required throughout trichome development. The ZWI gene encodes a minus-end directed kinesin-like motor protein that is required for normal branch formation and stalk elongation (Oppenheimer et al., 1997; Song et al., 1997). Further genetic analysis has identified many additional genes that are required for normal branch formation (Folkers et al., 1997; Krishnakumar & Oppenheimer, 1999; Luo & Oppenheimer, 1999; Perazza et al., 1999). An analysis of double mutant combinations for apparent loss-of-function branching mutants has uncovered a highly redundant branch initiation control pathway (Luo & Oppenheimer, 1999). Pharmacological data suggest that microtubule-dependent function is required for stalk and branch initiation (see below). Drugs that affect actin organization have very distinct effects on trichome morphogenesis; polarized elongation and branch initiation are not obviously affected, while the coordinated expansion of branches and the stalk following stage 3 is severely inhibited (see below).

6 Szymanski

Figure 2. Stages of trichome development. Scanning electron micrographs of the adaxial surface of Arabidopsis leaves that illustrate aspects of trichome spacing and morphogenesis. (a) An example of secondary trichomes (white arrows) initiating around an existing central trichome (black arrows). Numbers to the left of each labeled trichome indicate the developmental stage. (b) An illustration of additional stages of trichome development: 1, isodiametric expansion in the plane of the epidermis; 2, stalk emergence and polar expansion; 3, branch initiation; 4, expansion of the stalk and branches with a blunt tip morphology; 5, trichome expansion with pointed branch tips; and 6, mature trichome with a papillate cell wall. Reproduced from Szymanski et al., 2000.

4. ACTIN FUNCTION AND CELL POLARITY IN OTHER SYSTEMS

Like all other polarized cell types, developing trichomes must define a specific cortical position for cell expansion and reinforce and maintain the asymmetry during morphogenesis (Drubin & Nelson, 1996). This hierarchical cascade of functional requirements has been studied in detail in many other experimental systems. In many cases, the actin cytoskeleton plays an essential role in the polarity establishment and maintenance.

The role of actin in cell polarity has been studied most thoroughly in the budding yeast Saccharomyces cerevisiae. Genetic analysis of bud formation has identified a clear hierarchy of regulation (Pringle et al., 1995). The GTPase cdc42 acts near the top of the regulatory cascade to locally regulate actin organization and the subsequent assembly of bud site-specific factors (Ayscough et al., 1997). F-actin is also essential for normal polarity establishment in the embryos of the brown algae, Fucus and Pelvetia. Pharmacological experiments with cytochalasins indicate that an unperturbed F-actin cytoskeleton is required for the normal onset of cell polarization in Fucus embryos (Quatrano, 1973; Brawley & Robinson, 1985). In Pelvetia, an F-actin patch in living embryos has been reported as an early marker of cell polarization and predicts the position of rhizoid emergence (Alessa & Kropf, 1999). Pharmacological evidence and F-actin localization also suggest that actin plays an important role during pollen germination (Mascarenhas &


LaFountain, 1972; Tiwari & Polito, 1990). In the germinated pollen tube, a complex longitudinal array of actin bundles that terminate distal to the pollen tube tip likely plays an important role in vesicle delivery to the apical region of the cell (Picton and Steer, 1981; Lancelle et al., 1987, Miller et al., 1996). However, recent studies suggest that organization of specific membrane domains and actin-dependent function in the tip region of the tube are essential for normal pollen tube growth (Gibbon et al., 1999; Kost et al., 1999)

The cytoskeletal requirements for sensory bristle and hair formation in Drosophila are similar to those for Arabidopsis trichomes. In developing sensory bristles, a dense population of individual microtubules is surrounded by longitudinal hexagonal cortical bundles of microfilaments (Tilney et al., 1995). During early developmental stages of bristle development, the assembly of the core microtubules appears to precede microfilament bundle assembly (Tilney et al., 1996). Rearrangement of the microtubule and microfilament cytoskeletons also has been documented during hair formation on the Drosophila wing (Eaton et al., 1996; Turner & Adler, 1998). A pharmacological analysis suggests that microtubules are essential for wing hair initiation and morphogenesis, and the primary role of the actin cytoskeleton is to maintain the established growth pattern (Turner & Adler, 1998).

The specialized requirements for F-actin assembly during bristle development have been exploited to identify several genes that regulate microfilament organization. For example, removal of profilin or actin-capping protein genes is lethal in Drosophila. However, weak alleles of both genes yield bristle defects (Verheyen & Cooley, 1994; Hopmann et al., 1996). The forked and singed mutations cause bristle shape defects (Bender, 1960; Hoover et al., 1993). The predicted protein product of the SINGED gene shares amino acid similarity with the actin-bundling protein fascin (Cant et al., 1994). FORKED and SINGED appear to act sequentially to regulate actin bundle formation in developing bristles (Tilney et al., 1996; Wulfkuhle et al., 1998).

5. CYTOSKELETAL FUNCTION IN ARABIDOPSIS TRICHOME DEVELOPMENT

5.1 Pharmacology

Two recent publications provide preliminary data that Arabidopsis trichome formation is a useful genetic system to study actin-dependent growth (Mathur et al., 1999; Szymanski et al., 1999). Pharmacological data demonstrate that organized actin is not required for the initiation of polarized growth. Polarized growth during stage 2 and branch initiation during stage 3 are not noticeably

8 Szymanski

affected by F-actin disrupting agents (Fig. 3A). However, subsequent cell expansion during stages 4 and 5 is disordered, and cell shape defects are apparent (Fig. 3B). F-actin-dependent function is required to coordinate cell expansion following the formation of a polarized stalk and branch structures.

The apparent unimportance of F-actin during the establishment of polarized growth contrasts with many other developmental processes in which F-actin is required. This is not due to an absence of F-actin during these stages of development. Using conventional fixation techniques coupled with the freeze-shattering permeabilization approach described by Wasteneys et al. (1997), anti-actin antibodies label intricate networks of F-actin in both stage 1 and late stage 2 cells (Szymanski et al., 1999). Perhaps the role of F-actin during stage 2 is to facilitate apical transport, but in its absence, diffusion or microtubule-dependent function are sufficient. It should be emphasized that while disruption of actin organization does not appear to affect the rate of cell elongation through stage 3, drug-treated cells at early stages of development are slightly swollen, and subtle kinetic effects on growth have not been examined (Szymanski et al., 1999).

Polar growth during stages 2 and 3 requires microtubule-dependent activities (Figs. 3C, 3D). Agents that either stabilize or depolymerize microtubules can inhibit the establishment of polarity (Mathur et al., 1999). This was clearly shown using a dexamethose-regulated form of the maize R gene to induce trichome formation in the presence or absence the microtubule-depolymerizing agent oryzalin (Szymanski et al., 1999). Application of high concentrations of oryzalin prior to initiation causes isotropic cell expansion without branch formation. However, at lower drug concentrations some polar expansion is observed (Mathur et al., 1999). Because staining for microtubule organization in Arabidopsis trichomes has not been conducted, the extent to which the microtubule cytoskeleton is disrupted in drug-treated cells is not known. Residual polar expansion of treated cells may be due to a drug-resistant population of microtubules. Cotton fiber elongation displays a similar sensitivity to microtubule-destabilizing agents (Tiwari & Wilkins, 1995), and a population of apparently functional drug-resistant microtubules have been described (Seagull, 1990).

Agents that disrupt F-actin organization severely disrupt trichome morphogenesis following branch initiation in stage 3 (Fig. 3B). In untreated cells, cell expansion during stages 4 and 5 is strictly regulated and includes diffuse growth of the stalk and branches, as well as potential tip-directed growth in the elongating branches. Drug-treated stage 4 and 5 trichomes display swollen stalks, twisted and swollen branches, and aborted branches. High concentrations of cytochalasin D (CD) do not stop cell growth. Leaves exposed to the drug for six days give rise to enlarged cells with a striking, distorted morphology (Fig. 3B). Interestingly, Mathur et al. (1999) show that


the effects of F-actin stabilizing agents phalloidin and jasplakinolide are indistinguishable from those of CD and Latrunculin B, which tend to destabilize microfilaments. It was shown that each actin-binding drug severely disrupts cellular organization (Mathur et al., 1999). It is possible that the actin-dependent reinforcement of an organized growth pattern requires dynamic actin filaments or the activity of F-actin-binding proteins, both of which can be affected by different classes of actin-binding agents.

Figure 3. Effects of cytoskeletal inhibitors on trichome morphogenesis. Scanning electron micrographs of the adaxial leaf surface of drug-treated leaves. (A) Trichome morphology after 48 h of treatment with 50 µM Latrunculin B. (B) Trichome morphology after treatment with 50 µM cytochalasin D for 6 d. (C) Shape defects of cells treated with 100 µM oryzalin for 48 h. (D) High magnification of (C) demonstrating the swollen or distorted morphology of stage 3 and 4 oryzalin-treated cells. Numbers printed below each trichome indicate the developmental stage. Arrows indicate bulges that may represent defective trichome branches.

Even in mature trichomes that have completed the cell expansion program, the F-actin appears to be dynamic; CD rapidly fragments the cortical filaments. In stage 6 trichomes, the actin bundles are somewhat evenly spaced, and are oriented longitudinally in the stalk (Fig. 4A). After 30 min of CD treatment, the actin cytoskeleton is completely fragmented and disorganized (Fig. 4B). After 2 h of drug treatment, most stalks contain actin that is assembled into heavy

10 Szymanski

rods (Fig. 4C), and after 24 h, bright punctate spots and heavy rods of F-actin are frequently detected (Fig. 4D). Surprisingly, cells in which the actin cytoskeleton has been completely depolymerized can recover from drug treatment and repolymerize oriented F-actin (Mathur et al., 1999). A direct analysis of actin dynamics in living cells would provide insight into the assembly of ordered F-actin arrays in trichomes.

Figure 4. Time course of cytochalasin D (CD) effects on actin organization in mature trichomes. Each image is a maximum projection of confocal images of anti-actin immunofluorescence. (A) Typical wild-type trichome on a plant treated only with buffer. (B) Trichome treated for 30 min with 50 µM CD. (C) Trichome treated for 2 h with 50 µM CD. (D) Trichome treated for 24 h with 50 µM CD. Bar for (A–D), 10 µm.

The current data suggest that, at a very superficial level, there is a hierarchy of cytoskeletal control during trichome formation, with microtubule-dependent function acting before actin-dependent function. However, the temporal and


functional requirments for microtubules are not entirely clear. Inhibitors of microtubule polymerization inhibit both polarized elongation during stage 2 and branch formation at stage 3 (Figs. 3C, 3D). Microtubule-dependent function is not limited to the onset of polarization. Branched wild-type trichomes treated with oryzalin display severe swelling and branch distortion during stages 4 and 5 (Fig. 3D). These data point to unique, but partially overlapping, roles for the microtubule and microfilament arrays during trichome development.

The relationship between microtubule and F-actin organization is not clear. To begin to address this issue, Mathur et al. (1999) used a GFP–TALIN fusion protein as a probe for actin organization (Kost et al., 1998, 1999; see also Kost et al., this volume). Analysis was performed on living cells in which microtubules were disrupted with drugs and in zwi trichomes. In both cases F-actin was detected, suggesting that actin polymerization does not require native microtubule organization. However, based on the limited actin localization data in drug-treated and zwi trichomes, and the absence of microtubule localization data, it is difficult to conclude that microtubules do not influence the organization of microfilaments (see also Collings & Allen, this volume).

5.2 Actin organization in developing trichomes

A survey of the actin organization in trichomes at each developmental stage has been conducted using both immunolocalization of actin in fixed samples (Szymanski et al., 1999) and the actin-binding domain of mouse TALIN fused to GFP (Mathur et al., 1999). In many cases the description of actin at specific developmental stages differed. For example, antibodies clearly detected F-actin structures in fixed stage 1 and 2 trichomes, however no similar structures were detected with GFP–TALIN. Clearly, both techniques have limitations. For example, the controversial dense actin patch at the tip of growing pollen tubes, that was detected using chemical-fixation methods, appears to be an artifact (Miller et al., 1996). Steady-state actin organization in pollen tubes reported with GFP-TALIN resembles the structures revealed with independent techniques (Kost et al., 1998, 1999). However, the effects of GFP–TALIN on plant cell growth are not known. GFP–actin fusions fail to complement actin mutants in yeast, and cause defects during cytokinesis in Dictyostelium (Doyle & Botstein, 1996; Westphal et al., 1997). GFP fusions to actin-binding proteins such as TALIN may display similar toxicity or may alter actin organization in living cells. Therefore, at this time, it is not clear which method provides the best description of actin organization in developing trichomes.

Nevertheless, both methods for actin localization reveal a similar general relationship between actin organization and morphological transitions in trichomes. In regions of the cell in which growth patterns are being established,

12 Szymanski

such as branch buds and the apical regions of stage 2 stalks, the actin signal is diffuse (Szymanski et al., 1999; see Fig. 2D). The absence of extensive arrays of F-actin in these regions of the cell is similar to what is observed in the tips of growing pollen tubes (Miller et al., 1996; Kost et al., 1998, 1999). Subsequently, in other cellular domains in which the growth pattern is established, such as the central branch domains at stages 4 and 5, or the stage 6 stalk, F-actin dominates the anti-actin signal. A subset of the F-actin is aligned with the growth axis. This is consistent with a role for F-actin during the reinforcement and maintenance of cell growth pattern in trichome development.

The arrangement and location of F-actin and G-actin appear to be under strict spatial control within a given cell. This is most evident during the transition from stage 3 to stage 4. In stage 3 branch buds, actin signal is very strong and diffuse. Expanding stage 4 branches, which are located just 20 µm away, display a fine meshwork of polarized F-actin with a reduced diffuse actin signal at the tip. Similar spatial compartmentation of F-actin has been observed in tip-growing pollen tubes (Miller et al., 1996; Kost et al., 1998; see also Vidali & Hepler, this volume). In contrast to growing pollen tubes, stage 3 trichomes contain three separate elongation domains that appear to undergo similar rearrangements of F-actin. Each domain sequentially executes a highly regulated branch initiation and maintenance program. It is not known how the spatial control of actin polymerization and organization is achieved during branch growth, but almost certainly actin-binding proteins are involved.

Anti-actin antibodies detect actin filaments or fine bundles at each stage of trichome development. After stage 1, many of these bundles are aligned with the local axis of elongation. Similar aligned arrays of fine actin bundles have been observed in a variety of elongating cell types (Parthasarathy, 1985; Heslop-Harrison et al., 1986; Jackson & Heath, 1993). Even after cell expansion has ceased, stage 6 trichomes display fine, parallel arrays of evenly spaced F-actin bundles. Several groups have proposed that the presence of fine actin bundles, as opposed to heavy bundles of closely associated filaments, is associated with the ability to transport vesicles (Thimann et al., 1992; Foissner et al., 1996; Waller & Nick, 1997; Miller et al., 1999). The detection of actin-associated vesicle-like structures at all developmental stages and the observation of rapid longitudinal vesicle transport along the cortex of living stage 6 trichomes (Szymanski, unpublished) are consistent with this idea.

Figure 5. Actin organization in wild-type and distorted mutant trichomes. Each image is a maximum projection of confocal images of anti-actin immunofluorescence. (A) Three-branched stage 5 trichome. (B) High magnification of cortical actin signal derived from the boxed region in (A). (C) Stage 6 wrm trichome. (D) High magnification of cortical actin signal derived from the boxed region in (C). (E) Stage 6 crk trichome. (F) High magnification of cortical actin signal derived from the boxed region in (E). Arrows mark branch tips. Arrowheads mark the stalk region. Bar for (A–F), 10 µm.


Regulated vesicle transport to the cell cortex is probably essential for the spatial control of cell expansion during stage 5. The length and diameter of stage 5 stalks and branches increase in a highly-ordered fashion. The actin

14 Szymanski

cytoskeleton during stage 5 consists of an intricate network of actin filaments or bundles, some of which display clear longitudinal orientation (Fig. 5A). High magnification images of these structures reveal an intricate branched network of filaments or bundles (Fig. 5B). This organization resembles the reticulating lattice of cortical F-actin in elongating cotton fibers (Seagull, 1990). Stage 5 trichomes are sensitive to F-actin disrupting agents, and display localized cell swelling and branch shape defects after 12 h of treatment. It is possible that disruption of actin organization in stage 5 cells leads to misregulated accumulation and fusion of vesicles at random positions within the cell. This could explain the variable shape defects observed in growing trichomes treated with CD.

5.3 The distorted trichome shape mutants

Precise actin-dependent function is required to coordinate and maintain cell expansion during later stages of trichome morphogenesis. The striking similarity of the distorted class of trichome shape mutants to CD-treated trichomes suggests that constrained cytoskeletal function during trichome morphogenesis may provide a useful genetic system to study actin function (Mathur et al., 1999; Szymanski et al., 1999). The distorted1 (dis1) and distorted2 trichome mutants have been used as visual markers for classical genetic mapping experiments for more than 20 years (Feenstra, 1978). Six additional mutants of this class were identified based on the aborted branch and swollen phenoytype, which reflect defects in maintaining a normal growth pattern. (Hulskamp et al. 1994).

In both drug-treated and gnarled (grl) distorted trichomes, abnormal stalk swelling and expansion along the apical face of the cell often occur during the transition to stage 4 (Figs. 6A, 6B; see also Szymanski et al., 1999). Similar stage-specific defects are observed in crooked (crk) trichomes (Figs. 6C, 6D). The cellular reorganization during the transition to diffuse branch and stalk expansion is complex and likely requires several components. For example, reinforcement of an established cell growth pattern during stage 4 might require the regulated delivery or recycling of a specific class of vesicles. Any mutation that alters the timing, position, or specificity of vesicle transport could cause a breakdown in coordinated cell growth. The existence of at least eight trichome mutants with a similar distorted phenotype is consistent with the idea that multiple components are required to coordinate F-actin-dependent branch and stalk elongation after stage 4.


Figure 6. Cell shape defects of grl and crk trichomes. (A) Adaxial leaf epidermis of grl containing stage 1 and stage 2 trichomes. Arrow indicates normally-shaped stage 2 trichome. (B) Epidermis of grl leaves containing stage 4 and stage 5 trichomes. Swollen stage 4 trichomes are labeled with arrows. (C) Stage 4 crk trichome. (D) Stage 6 crk trichome.

The actin organization of several distorted trichome mutants has been examined (Mathur et al., 1999; Szymanski et al., 1999). A detailed examination of actin organization in the crk and grl mutants detected altered F-actin organization in developing trichomes that was coincident with cell shape defects. These results do not prove that CRK and GRL directly affect actin organization, but demonstrate a close temporal link between misregulated F-actin organization and growth in the mutants. The actin organization in alien, klunker, worm (wrm), and dis1 trichomes also were examined using GFP–TALIN (Mathur et al., 1999). In each case, the actin organization of stage 6 mutant trichomes differed from that of the wild type. An example of the actin immunolocalization in mature wrm trichome is shown in Figure 5. The steady state actin organization in stage 6 wrm (Figs. 5C, 5D) trichomes is similar to that of grl (Szymanski et al., 1999). Both display extensive branching of actin filaments in the stage 6 stalk, the extent of branching correlating with the severity of cell shape defects in almost all cases. The existence of heavily branched F-actin structures in the stalks of stage 6 trichomes is abnormal,

16 Szymanski

however, similar structures are observed in wild-type stage 5 trichomes (Fig. 5B). The grl and wrm mutations may affect the spatial or temporal control of actin organization. Mathur et al. (1999) reported that the actin organization in crk trichomes appeared distinct from other distorted trichome mutants. Antibody signal from fixed specimens also consistently revealed an extensively cross-linked actin cytoskeleton in stage 6 crk trichomes (Fig. 5E). High magnification images of the cortical actin cytoskeleton in the base of an elongated crk stalk detected dense populations of extremely fine actin filaments or bundles, some of which display clear alignment with the long axis of the cell (Fig. 5F). It is possible that CRK affects F-actin bundling. The gene encoding the F-actin binding protein villin is transcribed at elevated levels in several elongating cell types in Arabidopsis, including trichomes (Klahre et al., 2000). Perhaps mutations in genes like villin will be identified in trichome morphology screens.

Although the shape of distorted mutants and CD-treated cells are indistinguishable, it is interesting that none of the distorted mutants have the actin organization of CD-treated cells. Perhaps F-actin-dependent function is highly constrained during the transition to the elongation and expansion phase of trichome growth (stages 4 and 5). Subtle defects in actin organization during this transition may have dramatic effects on trichome growth. Mutations that cause F-actin fragmentation or depolymerization in diverse cell types would most likely be lethal; however, additional screens for distorted trichome mutants may identify a trichome-specific gene with more dramatic effect on actin organization.

It is not known at present if mutations in distorted group genes affect actin organization directly or indirectly. It may be that F-actin, membrane recycling, and vacuole biogenesis are inter-dependent processes, and that distorted mutants may affect any one of them. Regardless, these cellular processes are fundamental to plant cell growth and are not well understood. A genetic analysis of distorted trichome mutants may identify both weak alleles of essential genes and mutations in genes that have a more specialized role in polarized cell elongation. Molecular and biochemical analysis of the distorted group gene products will lead to a more mechanistic understanding of plant cell growth control.

ACKNOWLEDGEMENTS

This work was made possible by support from David Marks and Sue Wick at the University of Minnesota, and the National Science Foundation Cytoskeleton Training Grant DBI 96002237. This project was also supported


by NRI Competitive Grants Program/USDA Grant No. 99-35304-8525 to DBS.

REFERENCES

Alessa L and Kropf DL (1999) F-actin marks the rhizoid pole in living Pelvetia compressa zygotes. Development 126: 201-209

Ayscough KR (1998) In vivo functions of actin-binding proteins. Curr Opin Cell Biol 10: 102-111

Ayscough KR, Stryker J, Pokala N, Sanders M, Crews P and Drubin DG (1997) High rates of actin filament turnover in budding yeast and roles for actin in establishment and maintenance of cell polarity revealed using the actin inhibitor latrunculin-A. J Cell Biol 137: 399-416

Bender H (1960) Studies on the expression of various singed alleles in Drosophila melanogaster. Genetics 45: 867-883

Brawley SH and Robinson KR (1985) Cytochalasin treatment disrupts the endogenous currents associated wth cell polarization in fucoid zygotes: Studies of the role of F-actin in embryogenesis. J Cell Biol 100: 1173-1184

Cant K, Knowles BA, Mooseker MS and Cooley L (1994) Drosophila singed, a fascin homolog, is required for actin bundle formation during oogenesis and bristle extension. J Cell Biol 125: 369-380

Doyle T and Botstein D (1996) Movement of yeast cortical actin cytoskeleton visualized in vivo. Proc Natl Acad Sci USA 93: 3886-3891

Drubin, DG and Nelson, WJ (1996) Origins of cell polarity. Cell 84: 335-344 Duke SO (1994) Glandular trichomes: a focal point of chemical and structural interactions. Int J

Plant Sci 155: 617-620 Eaton S, Wepf R and Simons K (1996) Roles for Rac1 and Cdc42 in planar polarization and hair

outgrowth in the wing of Drosophila. J Cell Biol 135: 1277-1289 Eisner T, Eisner M and Hoebeke ER (1998) When defense backfires: detrimental effect of a

plant's protective trichomes on an insect beneficial to the plant. Proc Natl Acad Sci USA 95: 4410-4414

Feenstra WJ (1978) Contiguity of linkage groups I and IV as revealed by linkage relationship of two newly isolated markers dis-1 and dis-2. Arab Inf Serv 15: 35-38

Foissner I, Lichtscheidl IK and Wasteneys GO (1996) Actin-based vesicle dynamics and exocytosis during wound wall formation in characean internodal cells. Cell Motil Cytoskeleton 35: 35-48

Folkers U, Berger J and Hülskamp M (1997) Cell morphogenesis of trichomes in Arabidopsis: Differential control of primary and secondary branching by branch initiation regulators and cell growth. Development 124: 3779-3786

Gibbon, BC, Kovar, DR and Staiger, CJ (1999) Latrunculin B has different effects on pollen germination and tube growth. Plant Cell 11: 2349-2363.

Heslop-Harrison J, Heslop-Harrison Y, Cresti M, Tiezzi A and Ciampolini F (1986) Actin during pollen tube germination. J Cell Sci 86: 1-8

Hoover KK, Chien AJ and Corces VG (1993) Effects of transposable elements on the expression of the forked gene of Drosophila melanogaster. Genetics 135: 507-526

Hopmann R, Cooper JA and Miller KG (1996) Actin organization, bristle morphology, and viability are affected by actin capping protein mutations in Drosophila. J Cell Biol 133: 1293-1305

18 Szymanski

Hülskamp M, Folkers U and Grini PE (1998) Cell morphogenesis in Arabidopsis. Bioessays 20: 20-29

Hülskamp M, Misra S and Jürgens G (1994) Genetic dissection of trichome cell development in Arabidopsis. Cell 76: 555-566

Jackson SL and Heath IB (1993) The dynamic behavior of cytoplasmic F-actin in growing hyphae. Protoplasma 173: 23-34

Johnson HB (1975) Plant pubescence: an ecological perspective. Bot Rev 41: 233-258 Klahre U, Friederich E, Kost B, Louvard D and Chua N-H (2000) Villin-like actin-binding

proteins are expressed ubiquitously in Arabidopsis. Plant Physiol 122: 35-48 Kost B, Spielhofer P and Chua N-H (1998) A GFP-mouse talin fusion protein labels plant actin

filaments in vivo and visualizes the actin cytoskeleton in growing pollen tubes. Plant J 16: 393-401

Kost B, Lemichez E , Spielhofer P, Hong Y, Tolias K, Carpenter C and Chua N-H (1999) Rac homologues and compartmentalized phosphatidylinositol 4,5-bisphosphate act in a common pathway to regulate polar pollen tube growth. J Cell Biol 145: 317-330

Krishnakumar S and Oppenheimer DG (1999) Extragenic suppressors of the Arabidopsis zwi-3 mutation identify new genes that function in trichome branch formation and pollen tube growth. Development 126: 3079-3088

Lancelle SA, Cresti M and Hepler PK (1987) Ultrastructure of the cytoskeleton in freeze-substituted pollen tubes of Nicotiana alata. Protoplasma 140: 141-150

Larkin JC, Marks MD, Nadeau J and Sack F (1997) Epidermal cell fate and patterning in leaves. Plant Cell 9: 1109-1120

Larkin JC, Young N, Prigge M and Marks MD (1996) The control of trichome spacing and number in Arabidopsis. Development 122: 997-1005

Lazzaro MD and Thomson WW (1992) Ultrastructural localization of calcium in the organic acid secreting trichomes of chickpea (Cicer arietinum). Can J Bot 70: 2319-2325

Lazzaro MD and Thomson WW (1996) The vacuolar-tubular continuum in living trichomes of chickpea (Cicer arietinum) provides a rapid means of solute delivery from base to tip. Protoplasma 193: 181-190

Lloyd AM, Schena M, Walbot V and Davis RW (1994) Epidermal cell fate determination in Arabidopsis: Patterns defined by a steroid-inducible regulator. Science 266: 436-439

Luo D and Oppenheimer DG (1999) Genetic control of trichome branch number in Arabidopsis: the roles of the FURCA loci. Development 126 5547-5557

Marks MD (1997) Molecular genetic analysis of trichome development in Arabidopsis. Annu Rev Plant Physiol Plant Mol Biol 48: 137-163

Mascarenhas JP and LaFountain J (1972) Protoplasmic streaming, cytochalasin B, and growth of the pollen tube. Tiss Cell 4: 11-14

Mathur J, Spielhofer P, Kost B and Chua N-H (1999) The actin cytoskeleton is required to elaborate and maintain spatial patterning during trichome cell morphogenesis in Arabidopsis thaliana. Development 126: 5559-5568

McKinney EC, Ali N, Traut A, Feldmann KA, Belostotsky DA, McDowell JM and Meagher RB (1995) Sequence-based identification of T-DNA insertion mutations in Arabidopsis: actin mutants act2-1 and act4-1. Plant J 8: 613-622

Miller DD, de Ruijter NCA, Bisseling T and Emons AMC (1999) The role of actin in root hair morphogenesis: studies with lipochito-oligosaccharide as a growth stimulator and cytochalasin as an actin perturbing drug. Plant J 17: 141-154

Miller DD, Lancelle SA and Hepler PK (1996) Actin microfilaments do not form a dense meshwork in Lilium longiflorum pollen tubes. Protoplasma 195: 123-132


Oppenheimer DG, Pollock MA, Vacik J, Szymanski DB, Ericson B, Feldmann K and Marks MD (1997) Essential role of a kinesin-like protein in Arabidopsis trichome morphogenesis. Proc Natl Acad Sci USA 94: 6261-6266

Parthasarathy MV (1985) F-actin architecture in coleoptile epidermal cells. Eur J Cell Biol 39: 1-12

Perazza, D, Herzog M, Hulskamp M, Brown S, Dorne AM and Bonneville JM (1999) Trichome cell growth in Arabidopsis thaliana can be derepressed by mutations in at least five genes. Genetics 152: 461-476

Picton JM and Steer MW (1981) Determination of secretory vesicle production rates by dictysomes in pollen tubes of Tradescantia using cytochalasin D. J Cell Sci 49: 261-272

Pringle JR, Harkins HA, Zahner JE, De Virgilio C, Chant J, Corrado K and Fares H (1995) Establishment of cell polarity in yeast. Cold Spring Harb Symp Quant Biol 60: 729-744

Pyke KA, Marrison JL and Leech RM (1991) Temporal and spatial development of the cells of the expanding first leaf of Arabidopsis thaliana (L.) Heynh. J Exp Bot 42: 1407-1416

Quatrano RS (1973) Separation of processes associated with differentiation of two-celled Fucus embryos. Devel Biol 30: 209-213

Rodriguiez E, Healey PL and Mehta I (1984) Biology and Chemistry of Plant Trichomes, New York, Plenum Press

Schnittger A, Jürgens G and Hülskamp M (1998) Tissue layer and organ specificity of trichome formation are regulated by GLABRA1 and TRIPTYCHON in Arabidopsis. Development 125: 2283-2289

Seagull RW (1986) Changes in microtubule organization an wall microfibril orientation during in vitro cotton fiber development: an immunofluorescent study. Can J Bot 64: 1373-1381

Seagull RW (1990) The effects of microtubule and microfilament disrupting agents on cytoskeletal arrays and wall deposition in developing cotton fibers. Protoplasma 159: 44-59

Seagull RW (1992) Quantitative electron microscopic study of changes in microtubule arrays and wall microfibril orientation during in vitro cotton fiber development. J Cell Sci 101: 561-577

Shade RE, Thompson TE and Campbell WR (1975) An alfalfa weevil larval resistance mechanism detected in Medicago. J Econ Entom 68: 399-404

Song H, Golovkin M, Reddy AS and Endow SA (1997) In vitro motility of AtKCBP, a calmodulin-binding kinesin protein of Arabidopsis. Proc Natl Acad Sci USA 94: 322-327

Staiger CJ (2000) Signaling to the actin cytoskeleton in plants. Annu Rev Plant Physiol Plant Mol Biol 51: In press

Staiger CJ, Gibbon BC, Kovar DR and Zonia LE (1997) Profilin and actin-depolymerizing factor: modulators of actin organzation in plants. Trends Plant Sci 2: 275-281

Szymanski DB, Lloyd AM and Marks DM (2000) Progress in the molecular genetic analysis of trichome initiation and morphogenesis in Arabidopsis. Trends Plant Sci, In press

Szymanski DB, Marks MD and Wick, SM (1999) Organized F-actin is essential for normal trichome morphogenesis in Arabidopsis. Plant Cell 11: 2331-2347

Szymanski DB, Jilk RA, Pollock SM and Marks MD (1998) Control of GL2 expression in Arabidopsis leaves and trichomes. Development 125: 1161-1171

Szymanski, DB and Marks MD (1998) GLABROUS1 overexpression and TRIPTYCHON alter the cell cycle and trichome cell fate in Arabidopsis. Plant Cell 10: 2047-2062

Taylor, LP and Hepler PK (1997) Pollen germination and tube growth. Annu Rev Plant Physiol Plant Mol Biol 48: 461-491

Thimann KV, Reese K and Nachmias VT (1992) Actin and the elongation of plant cells. Protoplasma 171: 153-166

Tilney LG, Connelly P, Smith S and Guild GM (1996) F-actin bundles in Drosophila bristles are assembled from modules composed of short filaments. J Cell Biol 135: 1291-1308

20 Szymanski

Tilney LG, Tilney MS and Guild GM (1995) F-actin bundles in Drosophila bristles. I. Two filament cross-links are involved in bundling. J Cell Biol 130: 629-638

Tiwari SC and Polito VS (1990) An analysis of the role of actin during pollen activation leading to germination in pear (Pyrus communis L.): treatment with cytochalasin D. Sex Plant Reprod 3: 121-129

Tiwari SC and Wilkins TA (1995) Cotton (Gossypium hirsutum) seed trichomes expand via diffuse growing mechanism. Can J Bot 73: 746-757

Turner CM and Adler PN (1998) Distinct roles for the actin and microtubule cytoskeletons in the morphogenesis of epidermal hairs during wing development in Drosophila. Mech Dev 70: 181-192

Verheyen EM and Cooley L (1994) Profilin mutations disrupt multiple actin-dependent processes during Drosophila development. Development 120: 717-728

Waller F and Nick P (1997) Response of actin microfilaments during phytochrome-controlled growth of maize seedlings. Protoplasma 200: 154-162

Wang Q-Y and Nick P (1998) The auxin response of actin is altered in the rice mutant Yin-Yang. Protoplasma 204: 22-33

Wasteneys GO, Willingale-Theune J and Menzel D (1997) Freeze shattering: A simple and effective method for permeabilizing higher plant cell walls. J Microsc 188: 51-61

Westphal M, Jungbluth A, Heidecker M, Muhlbauer B, Heizer C, Schwartz JM, Marriott G and Gerisch G (1997) Microfilament dynamics during cell movement and chemotaxis monitored using a GFP-actin fusion protein. Curr Biol 7: 176-83

Wulfkuhle JD, Petersen NS and Otto JJ (1998) Changes in the F-actin cytoskeleton during neurosensory bristle development in Drosophila: the role of singed and forked proteins. Cell Motil Cytoskeleton 40: 119-132

the role of actin during arabidopsis trichome morphogenesis · the role of actin during arabidopsis...

Documents