Cytoskeletal involvement in cotton fiber growth and development

Micron,Vol. 24, No. 6, pp. 643~660,1993 Copyright © 1994ElsevierScienceLtd Printed in Great Britain.All rights reserved 0968~1328/93$24.00

Pergamon

REVIEW PAPERS

Cytoskeletal Involvement in Cotton Fiber Growth and Development R O B E R T W. S E A G U L L

Department of Biolo#y, Hofstra University, Hempsted, NY 11550, U.S.A. Abstract--The organization of cellulose microfibrils in plant cell walls influences physical properties of the wall and thus cell expansion characteristics. Developingcotton fiber represents an excellentmodel systemfor the analysis of the biologicalregulation of cell wall patterns. Current research indicates that the cytoskeleton has a major role in directing the deposition and organization of cellulosemicrofibrilsin the cellwallsof many plant systems,includingdevelopingcotton fibers. Both microtubules and microfilaments appear to be involvedin regulating changes observed in microfibrilpatterns during fiber development.The polylamellatearchitecture of the fiber wall can be attributed to changes in the orientation of cytoplasmicmicrotubules which appear to direct the orientation of microfibrildeposition in each successivelayer of the fiber wall. In the drug-induced absence of microtubules, celluloseis deposited in the fiber wall in a swirled pattern of bundled microfibrils. Interaction between adjacent microfibrils may influence cell wall organization on a localizedlevel.In contrast to the direct involvementofmicrotubules on wall organization, microfilamentsappear to be indirectly involved in the deposition of cellulose microfibrils. Current evidence indicates that microfilaments influence wall organization by controlling changes in microtubules patterns. Although a greater understanding of the relationship between the cytoskeleton and the fiber wall is needed, there is sufficientevidenceto indicate that geneticmanipulation of cytoskeletalcomponents is one path toward future direct manipulation of cell expansion characteristicsin many plant systemsand may lead to improvementsin the textile qualities of cotton fibers. Key words: Cellulose, cotton fiber, cytoskeleton, plant cell wall, microtubule, microfibril, microfllament. CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Why study cotton fibers? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Cell wall organization during development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Primary wall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Secondary wall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Co-alignment of microfibrils with microtubules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Involvement of microfilaments in microfibril patterning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Microfibril order irrespective of the cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Future . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Regulation of the cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I. I N T R O D U C T I O N

A. Why study cotton fibers? 1. Cotton fiber as a model system for cell morphooenesis and cytoskeletal function P l a n t morphogenesis is defined by the e x p a n s i o n characteristics of i n d i v i d u a l cells in the p l a n t body. T o control p l a n t m o r p h o g e n e s i s we m u s t u n d e r s t a n d the mechanism(s) which regulate cell e x p a n s i o n a n d shape. Wall e x p a n s i o n characteristics a n d thus cell shape, are controlled by the o r g a n i z a t i o n of the i n n e r - m o s t layer of wall microfibrils ( R o l a n d a n d Vian, 1979). Cellulose microfibrils are synthesized at the p l a s m a l e m m a by a n array of m e m b r a n e b o u n d proteins f o u n d at the growing ends of microfibrils (Brown, 1985; E m o n s , 1991). As in other systems, c o t t o n fibers c o n t a i n m e m b r a n e - b o u n d

643 643 645 645 648 650 654 655 656 656 657 659

cellulose synthetic complexes (Willison a n d Brown, 1977). Directing the path of these enzyme complexes in the plane of the p l a s m a l e m m a is one m e c h a n i s m for regulating the p a t t e r n i n g of wall microfibrils. There are volumes of evidence illustrating c o - a l i g n m e n t between the i n n e r - m o s t microfibril layer a n d cortical microtubules, thus indicating that m i c r o t u b u l e s m a y be involved in directing the path of cellulose synthetic complexes. Also, if m i c r o t u b u l e s are disrupted or modified, then s u b s e q u e n t microfibril a n d cell e x p a n s i o n patterns are modified (Hepler a n d Palevitz, 1974; H e a t h a n d Seagull, 1982; G u n n i n g a n d H a r d h a m , 1982; Seagull, 1991). This correlation between m i c r o t u b u l e s a n d wall microfibrils has resulted in the l o n g - s t a n d i n g belief that a cause a n d effect relationship with respect to l o c a t i o n a n d o r i e n t a t i o n exists between the two (Heath a n d Seagull, 1982; R o b i n s o n a n d Q u a d e r , 1982; G i d d i n g s a n d Staehelin, 643

644

R.W. Seagull

1991 ; Seagull, 1991 ). The precise involvement and control of cytoskeletal elements (both microtubules and microfilaments) in regulating cellulose microfibril deposition and orientation remains a central unresolved question in plant cell morphogenesis. Cytoskeletal arrays (microtubules and microfilaments) of plants are dynamic, changing organization and function in response to developmental signals (Seagull, 1989a; Giddings and Staehelin, 1991). Transitions in cytoskeletal organization occur not only during the cell cycle but also at interphase, during polylamellate cell wall construction (Lloyd, 1984; Lloyd and Seagull, 1985). In some systems interphase transitions are mediated by hormones (Akashi and Shibaoka, 1987; Ishida and Katsumi, 1991; Sauter et al., 1992; Seagull 1992b), however the mechanism for this is not known. Cotton fibers offer some unique properties that facilitate the examination of cytoskeletal elements during polylamellate wall production. Growing as trichomes on the surface of developing seeds, fiber development is highly synchronous with most, if not all fibers on a seed exhibiting identical developmental stages (Schubert et al., 1976). Cotton fibers can be grown in culture, thus facilitating experimental manipulation and observation (Beasley, 1984). As a result of its economic importance (see Section I.A.2) the sequence and patterns of cotton fiber development are well characterized.

dramatic impact on fiber mechanical properties (Rebenfeld, 1965; Zeronian, 1991). The orientation of the cellulose microfibrils in the fiber wall has a direct impact on final fiber strength (Egle and Grant, 1970), thus changing microfibril patterns during fiber development may be a way of modifying (improving) fiber strength. The orientation of these microfibrils can also affect subsequent expansion characteristics of the fiber (Houwink and Roelofsen, 1954; Seagull, 1990a). Thus modification of microfibril patterns during fiber growth could directly impact final fiber length. For these reasons it is critical to understand how microfibril patterns are

SW

2. The fiber wall and textile properties

Recent advances in textile processing technology and competition from synthetic fibers and foreign markets are two of the driving economic forces for the improvement of domestic cotton fiber. Length, strength and fineness are predicted to be fiber properties that need improvement to meet these new economic pressures (Deussen, 1989). Because these characteristics are under genetic control (Basra and Malik, 1984), modification of fiber development through genetic manipulation (recombinant DNA) can potentially result in well-defined, specific changes to the textile properties of mature cotton. For example, changing the timing of pivotal steps or modifying key biological processes in fiber development could result in desirable improvements in the textile properties of cotton. However, while it is clear that direct manipulation of genes controlling fiber quality can have a tremendous impact on the textile properties of cotton fibers, there is very little information on cotton fibers as living cells and the genetic mechanisms which control fiber quality. Therefore, before cotton fibers can be genetically engineered in a well-defined, predictable manner, we need a greater understanding of the genes and gene products regulating fiber growth and development. Specific cell wall characteristics influence a number of textile properties in cotton. Of obvious importance are chemical composition of the wall, degree of cellulose polymerization and amount of cellulose deposited (Timpa and Ramey, 1989; Gupta et al., 1979; Rowland et al., 1976). On a larger scale, the organization and orientation of cellulose microfibrils within the wall have a

t

wl

law

(3

Q Fig. 1. A montage composite illustrating microfibril orientation in various layers of the cotton fiber wall. The fiber is coated by a waxy cuticle (c) throughout development. The primary wall (pw) contains two distinguishable microfibril orientations, axial in the outer layer and transverse (to the fiber long axis) in the innermost layer. The winding layer (wl) is the first helically oriented wall microfibril layer. The secondary wall (sw) is comprised of layers of helically orientated microfibrils, with the pitch of the helix increasing with subsequent layers of microfibrils. Cytoskeletal components in the cytoplasm (cyt) predict all microfibril patterns during fiber development. This figure does not illustrate reversals.

Cotton Fiber Growth and Development

645

Fig. 2. A light micrograph of a 1 day post-anthesis (DPA) ovule epidermal surfacewith developingfiber initials (arrows). Initials appear as swellingsabove the epidermal surface. Scalebar = 100 lam. Fig. 3. An electronmicrographofa thin sectionthat grazesthrough the primarywall (pw)and corticalcytoplasm(c).Wallmicrofibrils (arrows) exhibit a random orientation relative the axis of fiber elongation (doublearrow). Scale bar = 0.5 rtm biologically regulated. This review will concentrate on this higher order of wall organization. For details on wall composition and developmental changes in wall content see Huwyler et al. (1979), Meinert and Delmer (1977).

II. C E L L WALL O R G A N I Z A T I O N D U R I N G DEVELOPMENT During growth, the fiber deposits successive layers of microfibrils in a highly regulated, predictable pattern (Waterkeyn, 1985). The end result is the construction of a thick, polylamellate structure, composed of layers of microfibrils with different orientations relative to the fiber long axis (Fig. 1). Fiber cells initiate from epidermal cells

and development can be divided into four phases: (1) initiation, (2) elongation, (3) secondary wall thickening and (4) maturation (Jasdanwala et al., 1977). Except for fiber maturation (i.e. boll opening, death of the living cytoplasm and desiccation of the wall), all stages of wall development exhibit specific microfibril patterns that can be followed using calcofluor staining with light microscopy (Seagull, 1986) or electron microscopy [EM] (Seagull, 1992a).

A. Primary wall

The primary cell wall is deposited during fiber initiation and throughout most of the elongation phase of the developing fiber, except for the final stages of elongation

646

R.W. Seagull

Fig. 4. An electron micrograph of a thin section that grazes through the primary wall of a 2 DPA fiber. The inner-most layer of microfibrils (arrow) is organized into parallel arrays oriented transverse to the axis of cell elongation (double arrow). Scale bar =0.5/am. Fig. 5. Light micrograph of a 10 DPA fiber viewed with polarized light. Little organization of the microfibrils is detected, as evidenced by the slight negative birefringence (faint glow, arrow). Scale bar = 30.0/am. Fig. 6. An electron micrograph of a thin section that grazes through the primary wall of a 15 DPA fiber. The inner-most layer of wall microfibrils (arrows) is oriented transverse to the axis of fiber elongation (double arrow). The outer layers ofmicrofibrils (arrowheads) have re-oriented due to intercalary fiber expansion and parallel the axis of fiber elongation. Scale bar =0.5/am. Fig. 7. Light micrograph of 15 DPA fibers viewed with polarized light. Primary walls exhibit greater degree of negative birefringence (arrow) than in Fig. 5, indicating a buildup of parallel wall microfibrils. Scale bar = 50.0/am

when secondary wall synthesis is initiated (Meinert and Delmer, 1977). Fiber initiation, 0-1 day post anthesis (DPA), is the emergence of the fiber primordium from the epidermal cell (Berlin, 1986; DeLanghe et al., 1979). At

this stage the fiber appears as a bulge on the surface of the epidermis (Fig. 2). The cell wall contains microfibrils that are organized in a random pattern (Fig. 3). These randomly organized microfibrils do not provide sufficient

Cotton Fiber Growth and Development

647

Fig. 8. Carbon-platinumreplica of windinglayer.Non-uniformbands of wall microfibrils(arrows)are interspersedwith a finelacy network of microfibrils(arrowheads).The alteration of these two arrays is irregular. Scalebar = 0.1 lam.

reinforcement strength for the wall to resist lateral cell expansion. This results in the swollen appearance of the fiber initials (Fig. 2). The elongation phase of fiber growth proceeds for several weeks. During this time the fiber increases its length by several orders of magnitude depending on genotype and growth conditions. Throughout elongation, wall microfibrils are deposited in a shallow pitched helical pattern, generally oriented transverse to the axis of fiber elongation (Fig. 4). No accumulation of oriented microfibrils is apparent during this stage since the walls exhibit little birefringence during most of this phase (Fig. 5). Microfibrils within the primary wall re-orient during

fiber elongation, thus indicating intercalary growth of the wall (Roelofsen, 1951; see Seagull, 1990a for review). In very young fibers this re-orientation is evidenced by the primary walls containing arrays of spirally organized microfibrils (Balls, 1923; Hock et al., 1941; Osborne, 1935). Thus only the most recently deposited layer of microfibrils exhibits the transverse orientation that they had at deposition. By 10 D PA the outer-most layers of the primary wall contain microfibrils with an axial orientation (Fig. 6). As fiber elongation rate declines there is an increase in wall birefringence (Fig. 7), due to a build-up of transversely oriented wall microfibrils. It is not known if the build-up of ordered wall material is the result of

648

R.W. Seagull

Fig. 9. Scanningelectronmicrograph of an'S' reversal in a mature cotton fiber. At the reversal point (arrow) the helicalarrays of microfibrilsreversesits gyre (arrowheads).In the scanningelectron microscope,reversals are detectedby the change in directionof wrinkles in the primarywall. (From deGruy et al., 1974.) Scalebar = 10 ~tm. decreased re-orientation of wall microfibrils as elongation declines or an increase in the rates of microfibril synthesis and/or deposition. B. Secondary wall

Secondary wall synthesis refers to the deposition of large quantities of cellulose with a high degree of polymerization (DP). Chemically the secondary wall is relatively simple, being composed almost exclusively of cellulose. Several changes in wall organization patterns occur at this time. Microfibrils are deposited in layers with a steep helical pattern. The pitch of the helix increases as subsequent layers of microfibrils are deposited (i.e. microfibrils closest to the cell lumen have a more axial orientation than previously deposited microfibrils). 1. Winding layer

Early researchers made a distinction between the first layers of the secondary wall and those subsequently deposited. The first layer of the secondary wall has been termed the 'winding layer' (Anderson and Kerr, 1938) or the 'inner sheath' (Rollins, 1945). This layer consists of microfibrils oriented in a steep pitched spiral on the inside of the primary wall. Deposition of the winding layer coincides with decreasing cell elongation (Maltby et al.,

1979; Schubert et al., 1973) and may be involved in limiting subsequent fiber elongation (Basra and Malik, 1984). The winding layer was distinguished from other secondary wall layers by its microfibrils which have a greater diameter and have an opposite helical gyre to the rest of the secondary wall layers (Anderson and Kerr, 1938; Flint, 1950; Rollins, 1945). The winding layer is not uniform in the distribution of cellulose microfibrils (deGruy et al., 1974). Densely packed arrays of helical microfibrils occur in bands that are separated by an open 'lacy' fibrillar network (Fig. 8). While a distinction has been made between the winding layer and both the primary and secondary wall (Rollins, 1945), the evidence thus far gathered is contradictory (Flint, 1950). The use of microfibril orientation to distinguish 'winding layer' from secondary wall is not valid since the winding layer will parallel secondary wall microfibrils in certain regions, depending on wall reversals (see below). When the wall is fragmented the winding layer separates from the secondary wall, often remaining attached to the primary wall. This observation is consistent with the winding layer being part of the primary wall. However, the simultaneous deposition of the winding layer with the chemical onset of secondary wall synthesis is consistent with the winding layer being chemically similar to the secondary wall. The bulk of the secondary wall is deposited as layers of

Cotton Fiber Growth and Development

649

Fig. 10. Electron micrograph of a thin section that grazes through the primary wall (PW) and cortical cytoplasm (C) ofa 2 DPA fiber. The inner-most layer of wall microfibrils(arrows) parallels microtubules (arrowhead) that are oriented transverselyto the axis of fiber elongation (double arrow). Scale bar = 1.0 gum. Fig. 11. Electron micrograph of a thin section that grazes through the primary wall (PW) and cortical cytoplasm (C) of a 10 DPA fiber. The inner layer of mierofibrils (arrows) parallels mierotubules (arrowheads). Both are transversely oriented to the axis of fiber elongation (double arrow). Scale bar = 1.0 gm. Fig. 12. Electron micrograph of a thin section that grazes through the secondary wall (SW) and cortical cytoplasm (C) of a 36 DPA fiber. Wall microfibrils (arrows) parallel cortical microtubules (arrowhead) in a steep helical pattern, almost paralleling the axis of fiber elongation (double arrow). Scale bar = 1.0 lain. parallel microfibrils, organized in helical arrays with increasing pitch (Fig. 1). I n cross section, the walls exhibit a lameUar p a t t e r n when treated with caustic soda (for review see Flint, 1950). This p a t t e r n results from differential rates of cellulose a c c u m u l a t i o n d u r i n g daily tempera-

ture cycling ( G i p s o n a n d Ray, 1970; Haigler et al., 1991). U n d e r c o n s t a n t t e m p e r a t u r e these rings are n o t evident. It is n o t k n o w n if the lameUae seen after swelling also c o r r e s p o n d to layers of microfibrils with different helical orientations.

650

R.W. Seagull

Fig. 13. Light micrograph illustrating immunofluorescentstaining of cortical microtubule arrays in a 10 DPA fiber. Microtubules are organized into shallow-pitched helical arrays. This technique facilitatesthe cell-wideanalysis of microtubuledistributions. The white rectangle illustrates the area of the microtubule array visualized using electron microscopy,serial section reconstruction analysis, as illustrated in Fig. 14. Scale bar = 10 ~tm. Fig. 14. Reconstruction drawing from serial section electron microscopic analysis of microtubule arrays of a 10 DPA fiber. Microtubules (arrowheads) exhibit variability in orientation and length that is not evident from immunofluorescent images of comparable cells (compare with Fig. 13). Microtubule ends are easily identified (arrowheads). The area of the cell depicted in this figure is represented by the rectangle in Fig. 13. Scale bar= 1.0 ~tm.

2. Reversals Arrays of secondary wall microfibrils exhibit 'reversals' in their helical gyre (Fig. 9). Reversals first appear in the winding layer and continue to be evident through fiber maturation. Two forms of reversals are evident. 'Z' reversals are characterized by an abrupt change in microfibril orientation and are most prevalent early in secondary wall synthesis. 'S' reversals exhibit a more gradual change in microfibril orientation (Fig. 9) and are found in later stages of secondary wall deposition (Seagull, 1986). Reversals appear randomly along the length of the fiber and their frequency varies with genotype (Wakeham and Spicer, 1955). Although reversals may be found along the entire length of an individual fiber, in general, the frequency of reversals is greater in the tip one-third of the fiber than in the basal two-thirds (Wakeham and Spicer, 1955; Seagull and Timpa, 1990).

The frequency of reversals increases significantly during fiber growth (Hebert and Boylston, 1984), thus indicating developmental regulation of this process. The biological control of reversal frequency and location remains unknown. III. C O - A L I G N M E N T O F M I C R O F I B R I L S W I T H MICROTUBULES The plant cytoskeleton is involved in the deposition of organized arrays of cellulose microfibrils in most, if not all higher plant systems (Seagull, 1989a, 1991; Williamson, 1991). There is strong circumstantial evidence that microtubules in the cortical cytoplasm somehow determine location and orientation of microfibrils deposited into the developing cell wall (Heath and Seagull, 1982; Seagull, 1991).

Cotton Fiber Growthand Development

651

Fig. 15. Electronmicrographofgrazingsectionsthroughthe cellwalland corticalcytoplasmof l0 DPA control (a) and 24 hr taxol treated (b) fibers.Microtubules(arrowheads)are evidentand are more numerousis taxol treatedfibers (comparea and b). Scale bar=0.5 ~tm.FigureadaptedfromSeagull,1990b. In cotton fiber, wall microfibril organization determines cell growth characteristics and physical properties of the mature fiber (DeLanghe, 1986). A relationship between microtubule function, microfibril patterns and fiber quality was demonstrated by Yatsu (1983). Modifications in wall organization due to chemical disruption of the cytoskeleton result in significant decreases in fiber textile properties (Yatsu, 1983). Understanding the involvement and function of cytoskeletal elements in controlling microfibril patterns during cotton fiber development is essential to studies directed towards genetic modification (improvement) of specific fiber wall and textile properties and will provide valuable information on the fundamental principles involved in controlling cell morphogenesis. Specific components of the cytoskeleton may play regulatory roles in defining fiber development and may provide convenient 'handles' or 'markers' for the genetic manipulation of not only fiber characteristics but perhaps general plant cell morphogenesis. Early ultrastructural studies of fiber indicate that microtubules are present in the cortical cytoplasm of developing cotton fibers and generally parallel the innermost layer of microfibrils (Itoh, 1974; Ryser, 1979; Westafer and Brown, 1976; Willison and Brown, 1977; Yatsu and Jacks, 1981). During both primary and secondary wall synthesis, the innermost layer of microfibrils parallels the orientation of cortical microtubules (Figs 10-12). Young fibers (2-3 DPA), whose ordered microfibril arrays are only visualized at the EM level, also exhibit co-alignment of microfibrils with microtubules (Fig. 10). Two microscopic approaches have been used to assess cytoskeletal function during fiber development. Indirect

immunofluorescence techniques were used to obtain a better understanding of cell wide control of wall patterns (Fig. 13). Electron microscopic examination of cytoskeletal elements reveals high resolution details of cytoskeletal organization and interaction with various cytoplasmic components. Quantitative information on microtubule patterns can be collected using serial section reconstruction analysis (Seagull and Heath, 1980; Hardham and Gunning, 1978). Resulting plots (Fig. 14) provide details on specific characteristics of the microtubule array, such as length, number, orientation and population variability. While immunofluorescence light microscopy facilitates the examination of large areas of numerous fibers, the resolution of the cytoskeletal arrays is much reduced (compare Figs 13, 14). At all stages of fiber development, changes in microfibril orientation mimic changes that occur in the orientation of cytoplasmic microtubules located adjacent to the plasma membrane (Ryser, 1985; Seagull, 1986; Quader et al., 1987). The mechanisms involved in controlling microtubule patterns must function very quickly and over the total expanse of the fiber as no intermediate arrays (i.e. orientations between transverse and helical) of microtubules were found and all microtubules within a fiber had the same general orientation (i.e. no instances of transversely and helically oriented microtubules occurring simultaneously in the same fiber). The dependence of microfibril patterns on microtubule arrays is substantiated by experiments where cytoskeletal elements are disrupted with various chemical agents (Seagull, 1989b, 1990b). Microtubules within a fiber exhibit differential sensitivity to these chemical agents,

652

R.W. Seagull

Y

Figs 16--19:Microtubule plots (A) using reconstruction techniques from at least 10 consecutive sections (Scale bars=0.5 lam). Carbon-platinum coated wall fragments (B) showing orientation of the inner-most layer of microfibrils relative to the axis of fiber elongation (double arrow). Figs a and b are from different fibers of the same sample. I--inner microfibril layer, O--outer wall. Scale bar = 1.0 l.tm. Figs adapted from Seagull, 1992. Fig. 16. Microtubule reconstruction plot and carbon platinum shadowed inner cell wall layer for a 2 DPA fiber. (A) Long (large arrow) and short (smallarrow) microtubulesare organizedinto shallowpitch helicalarrays, generallyoriented transverse to the axis of fiber elongation (double arrow). (B) Wall microfibrils (arrow) exhibit a pattern similar to the microtubules (compareA and B) with significant variability in orientation within the population.

indicating a chemical (and perhaps functional) heterogeneity within the microtubule population. Treated fibers often contain widely spaced arrays of microtubules that maintain orientation within the fiber, as well as function. Moreover, the co-localization of newly deposited microfibrils in association with these fragmented arrays of microtubules supports the proposed function of microtubules in directing wall microfibril orientation (Seagull, 1989b, 1990b). Interference with microtubule dynamics prevents the production of a normal poly-lamellate cell wall in the cotton fiber. Taxol, a plant alkaloid, stabilizes microtubule arrays. When applied to growing fibers, taxol induces a rapid increase in the number of microtubules in the cortical cytoplasm (Fig. 15) and prevents the normal

shifts in microtubule orientation, thus directly impacting cell wall patterns (Seagull, 1990b). The mechanisms that regulate microtubule dynamics are important in controlling wall development and affect physical and thus expansion characteristics of the cell wall. Chemical treatments that either disrupt or stabilize cytoskeletal patterns often result in a non-uniform distribution of microtubules (Seagull, 1989b, 1990b). Subsequently synthesized wall microfibrils appear deposited in specific association with microtubules. In fact, other cell systems which deposit microfibrils in localized arrays (i.e. developing stomatal and conductive cells) do so via a non-uniform distribution of microtubules (Seagull, 1989a, 1991). These observations are consistent with the existence of as yet undescribed elements (proteins)

Cotton Fiber Growth and Development

653

! i

Fig. 17. Microtubule(A)and microfibril(B)patternsfrom3 DPAfibers.Microtubulearraysexhibitlessvariabilityin orientationthan at 2 DPA (comparewithFig. 16A)but stillexhibitvariabilityin length.The mostrecentlydepositedwallmicrofibrils(arrow)havean orientationthat mimicsthe microtubules. linking the cytoskeleton, plasmalemma and microfibril synthesizing complexes. Characterization of these elements may provide information useful for future regulation of cytoskeletal function and fiber development. Low-resolution immunofluorescence light microscopy indicates that a direct relationship between microtubule and wall microfibril patterns may exist (Seagull, 1990b). To further probe this possibility, quantitative analyses of the relationship between microtubules and wall microfibrils were conducted using serial section reconstruction analysis coupled with quantification of microfibril order (Seagull, 1992a). From these reconstructions, qualitative information about microtubules, such as length, proximity to the plasmalemma, number and organization can be obtained. Each stage of fiber development was examined for both microtubule and microfibril organization (Figs 16-19) and statistical comparisons done to determine not only orientation but also variability within each of the populations (Table 1). These analyses indicate

that during the transition between fiber initiation and elongation, cortical microtubules change organizational pattern from random to shallow-pitch helical (compare Figs 16 and 17). Microtubule length increases during fiber development, with significant changes occurring at the transition between initiation and elongation and between primary and secondary wall synthesis (Fig. 20). To function in controlling cell wall events, microtubules must be near the plasmalemma. Median sections through the fiber illustrate microtubules in close proximity to the plasmalemma, often appearing bridged to it (Fig. 21). While the majority of microtubules maintain a close association with the plasmalemma, proximity does appear to be developmentally regulated (Fig. 22). Comparisons of microtubule and microfibril angles at various stages of fiber development (Table 1) indicate a similar, but not precise co-alignment. Both microtubules and wall microfibrils appear to be under developmental regulation as both exhibit stage specific shifts in often-

654

R.W. Seagull

Fig. 18. Microtubule(top) and microfibril(bottom) patterns from 24 DPA fibers. Microtubulesexhibit a steep pitch helical pattern relativeto the axis of fiberelongation.The most recentlydepositedmicrofibrilsparallelmicrotubulearrays. Microfibrilsexhibitsome bundling (arrowheads)with bundles exhibitingsomewhat differentorientations. tation and changes in population variability. Significant differences in orientation and organization exist between microtubules and microfibrils but the reasons for this are unknown. It is possible that differences may be mathematically significant but not biologically significant with respect to the mechanisms by which microfibril patterns are controlled. During secondary wall deposition, microfibrils exhibit a significantly different degree of variability. Interactions between fibrils may result in subtle changes in microfibril order and the degree of variability within the population. Without the influence of microtubules, microfibrils tend to form swirled clusters in the fiber wall (Seagull, 1990b), indicating that inter-microfibril associations are possible.

IV. I N V O L V E M E N T O F M I C R O F I L A M E N T S IN MICROFIBRIL PATTERNING Improvements in preservation techniques for both light and electron microscopy have expanded our understand-

ing of microfilament distribution and function in plants (Seagull, 1989a). In developing cotton fibers the first specific observation of microfilaments was reported by Seagull (Seagull, 1990b). Two populations of microfilaments are described, large cables of filaments running parallel to the axis of elongation and a fine three dimensional network of filaments in the cortical cytoplasm. Using improved EM preservation techniques, microfilaments are observed in close association with some microtubules (Fig. 23). Not all microtubules exhibit associated microfilaments and the precise relationship between them remains unclear. Involvement of microfilaments in controlling wall patterns is indicated by rapid changes in microtubule and microfibril patterns upon chemical disruption of the microfilaments (Seagull, 1990b). The involvement of microfilaments appears to be indirect, through a regulation of microtubule patterns. Rapid re-organization of microtubules in response to disruption of microfilament systems indicates a functional relationship between microtubules and microfilaments with microfilaments

Cotton Fiber Growth and Development

655

Fig. 19. Microtubule (A) and microfibril (B) patterns in 30 DPA fibers. Microtubules are organized in a tightly packed, steep helical array. Fewer short microtubules are evident (compare with Figs 16A, 17B). Bundling of wall micro fibrils is evident at the fragmented ends of the wall (arrowheads).

controlling overall, cell-wide microtubule and microfibril patterns. Table 1. Analysis of means test Fiber age

1 2 3 7 16 19 24 30 36

Microtubule angle Means

Variance

42.4 A 64.5 B~ 68.8 B~ 74.7 c~ 80.3 c 78.2 ca 17.5 D~ 21.5 E~ 17.10~

475.2 A 644,7 xa 345.3 AB" 263.0 ABa 89.3 c 81.0 c~ 26.5 c~ 6.5 ~ 9.1E~

Microfibril angle Means

Variance

65.1 A* 78.8 Bb 75.3 c~

312.0 Aa 107.7 acb 144.5 Aa

72.4 cb 15.7cb 23.1E~ 17.8 v~

115.1 aD~ 10.3 Eb 37.0 CFb

25.2 Fb

A comparison of means and variance for angular deviation values, between microtubules and microfibrils at various stages of fiber development. Upper case letters designate comparisons between columns only. Lower case letters designate row comparison (within a column) of means or variances of microtubules and micro fibrils. (From Seagull, 1992a.)

V. M I C R O F I B R I L O R D E R IRRESPECTIVE O F THE CYTOSKELETON

Wall microfibrils are not deposited in a completely random pattern in the absence of cytoskeletal elements. While it is clear that cytoskeletal elements play a crucial role in the development of wall patterns in cotton fiber (see previous sections), certain characteristics of order are controlled by other factors, irrespective of the cytoskeleton. This level of control is documented (i) in cells where the cytoskeleton has been removed by chemical treatments (Seagull, 1990b) or (ii) by high-resolution, quantitative analysis of microtubule/microfibril co-alignment (Seagull, 1992a). In the complete absence ofmicrotubules, wall microfibrils exhibit a swirled pattern in the cell wall (Fig. 24). These walls exhibit discrete areas of birefringence, indicating that the swirls are composed of

656

R.W. Seagull

0.90'u.I 0.75 m 0.60 -

bundles of microfibrils) involves other, perhaps wallassociated, factors.

aa

b b

c d

___

0.30 0.45 O3 za 0.15 [tLl 0.00 : 0

VI. CONCLUSIONS e

1

1

' ' , t 5 20 25 30 35 40 FIBER AGE (DPA)

Fig. 20. Relative changes in microtubule length during fiber development, determined from reconstruction plots. The ratio of the number of microtubule ends (terminations) per total number of microtubules observed in a reconstruction is indicative of relative length. The closer the ratio approaches the value 2 (i.e. that both ends of the microtubules are present in the area of analysis), the shorter the microtubules in the population. Letters assigned to each histogram represent statistical comparison. Histograms (means and standard error bars) with the same letter are not significantly different (p < 0.001) from one another using t-test, analysis (measurements of at least 800 microtubules from at least 3 different fibers at each age). Microtubule lengths do not change significantly between 2 and 3 DPA; however by 7 DPA the microtubules have increased in length. The length of microtubules remains constant during the remainder of primary wall synthesis but increases during secondary wall synthesis from 24 to 36 DPA. (From Seagull, 1992.)

bundles of parallel cellulose microfibrils (Seagull, 1990b). This swirled pattern is more evident with the disruption of cytoskeletal elements during secondary wall synthesis and may be attributed to increased rates of cellulose synthesis. Detailed quantitative comparisons of microtubule and microfibrils arrays show that, at various stages of development, the two have small but statistically significant differences in orientation (Table 1). During secondary wall synthesis, wall microfibrils have a greater degree of order than the microtubule population. In this case, interactions between neighboring microfibrils may result in subtle changes in alignment, thus decreasing the degree of variability in the microfibril population. From both types of analysis one concludes that cell-wide control of wall patterns directly involves the cytoskeleton, whereas localized order (i.e. between adjacent microfibrils or

A number of conclusions can be made based on the above information regarding the functions of the cytoskeleton in controlling wall patterns in cotton fiber: (1) Microtubule-microfibril co-alignment is maintained in the cotton fiber throughout development and in situations where the cytoskeleton has been modified or disrupted. (2) Microtubule orientation must change for the production of a polylamellate wall. (3) The cytoskeleton appears to be involved in controlling the cell-wide organization of the microfibrils, however local order can be maintained even in the absence of microtubules. (4) Microtubules and microfilaments are not static or homogeneous since populations of microtubules change in a developmentally regulated manner and individual elements may exhibit differential sensitivity to disrupting agents. (5) Microfilaments play an active role in establishing and/or maintaining microtubule organization. Observations made on developing cotton fibers have direct application to fundamental questions regarding mechanisms of plant cell growth and wall organization. While being a highly specialized cell depositing large amount of highly organized wall material, there is no reason to believe that the fundamental mechanisms that control wall development in cotton fiber are any different than in many other, less specialized cell types. Many higher plant cells deposit polylamellate walls composed of layers of highly organized wall microfibrils (Seagull, 1991). However, the diversity of developmental stages and low numbers of these cells in most plant tissues makes detailed quantitative characterization very difficult. A major advantage of the cotton fiber system as a model is the large number of developmentally synchronized cells that can be characterized at one time. A. Future

It is now clear that the cytoskeleton of cotton fiber (as well as many other plant cells types) consists of a highly

Fig. 21. Median section through a 12 DPA fiber showing cross sectioned microtubules (arrow) close to the plasmalemma. Small connections (arrowhead) are observed between some of the microtubules and the plasmalemma. Scale bar=0.5 ~tm.

Cotton Fiber Growth and Development 24a

16

~

12 b

0

C

4 0

@

b

I

I

o

5

20

25 30 35

40

FIBER AGE (DPA)

Fig. 22. Relativechangesin the proximity of microtubules to the plasmalemma as measured by the percentage of microtubules found at distances greater than 25 Inn from the plasmalemma. Letters assignedto each histogram represent statistical comparisons. Histograms (meansand standard error bars) with the same letter are not significantlydifferent,based on t-test, comparisons (same as Fig. 20). Throughout development, the majority of microtubules (>80%) are within 25 ~tm of the plasmalemma; however there is a significant trend towards closer association with the plasmalemma as fibers develop. From Seagull, 1992a.

dynamic, three dimensional array of filaments, capable of changing organization and function in a developmentally regulated manner. Descriptions of these arrays and the transitions they go through are valuable in providing evidence for cytoskeletal function and regulation. However, central unresolved questions remain. H o w are cytoskeletal arrays established, maintained and/or reor-

657

ganized during fiber development? What is the mechanism that regulates the observed differential stability of elements within and between cytoskeletal arrays? What are the interactions (proteins involved) between microtubules, microfilaments, the plasmalemma and wall micro fibrils? Are cytoskeletal arrays developmentally regulated and how does this occur? Can the manipulation of cytoskeletal function be used to improve fiber quality parameters or manipulate general plant cell morphogenesis? Study of developing cotton fiber will provide a unique insight into some questions regarding cytoskeletal involvement in wall patterning. For example, at pivotal stages in fiber development, microtubule arrays undergo dramatic reorganizations (i.e. at the transition between fiber initiation and elongation, as well as at the transition between primary and secondary wall synthesis). The predictability of the fiber system and the large amounts of synchronous tissue that can be collected will facilitate an examination of proteins responsible for inducing or regulating these changes. Cotton fiber is one of only a few systems where microfilaments appear to be involved in controlling microtubule patterns. Study of fiber development may reveal the mechanism by which this occurs.

B. Regulation of the cytoskeleton In animal systems, m a n y of the dynamic and functional changes that occur in cytoskeletal elements are mediated

Fig. 23. Grazing section through the cortical cytoplasmand cell wallofa 7 DPA fiberpreservedusing rapid freezing-freezesubstitution techniques. Cortical microtubules (large arrow) are paralleled by wall microfibrils (arrowheads). In many instances, microfilaments are observed in close association with microtubules (small arrow). Scale bar---0.5 pm.

658

R.W. Seagull

Fig. 24. Grazingthin sectionthrough control (A)and colchicine(B) treated cellwallsof 25 DPA fibers.Axisoffiberelongation(double arrow). Control walls exhibit a steep helical pattern of wall microfibrils(arrow). Fibers treated for 7 days with 10-4 Mcolchicine exhibita swirledappearancein the wall (arrow).The cytoplasm(C)in the colchicinetreatedfibershowsno microtubulespresent.Scale bar = 1.0 p.m.

through a variety of chemical (protein) changes in these elements. These include changes in the structural proteins (tubulin or actin) via the production of different protein isotypes (Hightower and Meagher, 1986; Sullivan, 1988; Kabsch and Vanderkerckhove, 1992), post-translational modifications of the structural proteins (Joshi and Cleveland, 1990) and through modulation ofcytoskeletalassociated proteins (Olmsted, 1986; Schliwa, 1981). In plants, information on isotypes and developmental regulation has accumulated with regard to the structural proteins of cytoskeletal elements (Silflow et al., 1987; Fosket, 1989), however, very little is known about

cytoskeletal regulatory proteins (Cyr, 1991; McCurdy and Williamson, 1991). In order to understand the transitions in cytoskeletal organization and function during cell morphogenesis, we must discover how these elements are regulated at the protein and gene levels. This approach is currently limited by the lack of knowledge concerning the regulatory processes of cytoskeletal function, i.e. modifications of structural and/or regulatory proteins. Developing cotton fibers need to be studied with respect to cytoskeletal function and the role regulatory proteins play during fiber development. The observed

Cotton Fiber Growth and Development

dynamics of cytoskeletal arrays in cotton fiber are consistent with the involvement of regulatory mechanisms such as the generation of protein isotypes, posttranslational modification of tubulins and actins, or the generation of different regulatory proteins. The synchrony and amount of tissue that can be collected from various developmental stages will facilitate biochemical investigations into the protein composition of cytoskeletal arrays at various stages of development. In summary, pivotal changes in wall patterns coincide with changes in cytoskeletal organization. From the vast literature on cytoskeletal function, it seems likely that genetic regulation of microtubule and microfilament structural and/or regulatory proteins play a key role in the dynamics of cytoskeletal function during fiber development. Identification of these cytoskeletal proteins and genes involved in the development of specific wall patterns is needed. Once identified and their regulation understood, these proteins and genes may be useful as markers in assessing potential fiber properties and as keys for regulating fiber quality through genetic engineering.

REFERENCES Akashi, T. and Shibaoka, H., 1987. Effects of gibbereUin on the arrangement and the cold stability of cortical microtubules in epidermal cells of pea internodes. Plant Cell Physiol., 28, 339-348. Anderson, D. B. and Kerr, T., 1938. Growth and structure of cotton fiber. Industr. Engn#. Chem., 30, 48-55. Balls, W. L., 1923. The determiners of cellulose structure as seen in the cell wall of cotton hairs. Proc. R. Soc., B 95, 72-80. Basra, A. S. and Malik, C. P., 1984. Development of the cotton fiber. Int. Rev. Cytol., 89, 65-113. Beasley, C. A., 1984. Culture of cotton ovules. In: Cell Culture and Somatic Cell Genetics of Plants, Vol. l, Vasil I. K., (ed.). Academic, Orlando, pp. 232-240. Berlin, J. D., 1986. The outer epidermis of the cottonseed. In: Cotton Physiology, Number 1, Mauney, J. R. and McD. Stewart, J., (eds.). The Cotton Foundation, Memphis TN, pp. 375-414. Brown, R. M. Jr, 1985. Cellulose microfibril assembly and orientation: recent developments. J. Cell Sci., Suppl., 2, 13-32. Cyr, R. J., 1991. Microtubule-associated proteins in higher plants. In: The Cytoskeletal Basis of Plant Growth and Form, Lloyd, C. W., (ed.). Academic, London, pp. 57~8. deGruy, I. Z., Carra, J. H. and Goynes, W. R., 1974. The Fine Structure of Cotton: An Atlas of Cotton Microscopy, O'Connor, R. T., (ed.). Dekker Press, New York. DeLanghe, E. A. L., 1986. Lint development. In: Cotton Physioloffy, Number 1, Mauney, J. R. and McD. Stewart, J., (eds.). The Cotton Foundation, Memphis TN, pp. 325-349. DeLanghe, E. A. L., Demol, J., Marechal, R., Paes, G., Fransen, J., Verschraeghe, L. and Waterkeyn, L., 1979. Genetical, physiological and ecological influences on the structure and technological properties of cotton fibers. In: Cotton in a Competitive WorM, Harrison, P. W., (ed.). The Textile Institute, Manchester, U.K. Deussen, H., 1989. Cotton Fiber Properties for High-speed Spinninff, Belt-wide Cotton Conferences, pp. 104-106. Egle, C. J. and Grant, J. N., 1970. Interrelations of structural and physical properties of untreated cottons. Text. Res. J., 40, 158-168. Emons, A. M. C., 1991. Role of particle rosettes and terminal globules in cellulose synthesis. In: Biosynthesis and Biodegradation of Cellulose, Haigler, C. H. and Weimer, P. J., (eds.). Dekker Press, New York, pp. 71-98. Flint, E. A., 1950. The structure and development of the cotton fiber. Biol. Rev., 25, 414-434. Fosket, D. E., 1989. Cytoskeletal proteins and their genes in higher plants. In: The Biochemistry of Plants,. Vol. 15, Molecular Biology, Marcus, A., (ed.). Academic, San Diego, pp. 393-454. Giddings, T. H. Jr and Staehelin, L. A., 1991. Microtubule-mediated control of microfibril deposition: a re-examination of the hypothesis.

659

In: The Cytoskeletal Basis of Plant Growth and Form, Lloyd, C. W., (ed.). Academic, London, pp. 85-99. Gipson, J. R. and Ray, L. L., 1970. Temperature variety interrelationships in cotton. 1 Boll and fiber development: 2 Seed development and composition. Cotton Grow Rev., 47, 257-271. Gunning, B. E. S. and Hardham, A. R., 1982. Microtubules. A. Rev. Plant Physiol., 33, 651~i91. Gupta, V. B., Moharir, A. V. and Panda, B. C., 1979. The characterization of some indian cotton varieties with the help of electron microscopy and X-ray diffraction and the correlation of their structure with their mechanical properties. In: Cotton in a Competitive World, Harrison, P. W., (ed.). The Textile Institute, Manchester, U.K., pp. 83-105. Haigler, C. H., Rao, N. R., Roberts, E. M., Huang, J.-Y., Upchurch, D. R. and Trolinder, N. L., 1991. Cultured ovules as models for cotton fiber development under low temperature. Plant Physiol., 95, 88-96. Hardham, A. R. and Gunning, B. E. S., 1978. Structure of cortical microtubule arrays in plant cells. J. Cell Biol., 78, 14-34. Heath, I. B. and Seagull, R. W., 1982. Oriented cellulose fibrils and the cytoskeleton: A critical comparison of models. In: The Cytoskeleton in Plant Growth and Development, Lloyd, C. W., (ed.). Academic, London, pp. 163-182. Hebert, J. J. and Boylston, E. K., 1984. Reversal frequencies in developing cotton fibers. Text. Res. J., 54, 2356. Hepler, P. K. and Palevitz, B. A., 1974. Microtubules and microfilaments. A. Rev. Plant Physiol., 25, 309-362. Hightower, R. C. and Meagher, R. B., 1986. The molecular evolution of actin. Genetics, 11, 315-332. Hock, C. W., Ramsay, R. C. and Harris, M., 1941. Microscopic structure of the cotton fiber. Text. Res. J., 11, 200-217. Houwink. A,L. and Roelofsen, P. A., 1954. Fibrillar architecture of growing plant cell walls. Acta Bot. Neerl., 3, 385-395. Huwyler, H. R., Franz, G. and Meier, H., 1979. Changes in the composition of cotton fiber cell walls during development. Planta, 146, 635-642. Ishida, K. and Katsumi, M., 1991. Immunofluorescence microscopic observation of cortical microtubule arrangement as affected by gibberellin in d 5 mutant of Zea mays L. Plant Cell Physiol., 32, 409-4 17. Itoh, T., 1974. Fine structure and formation of cell wall of developing cotton fiber. Wood Res,. 56, 494il. Jasdanwala, R. T., Singh, Y. D. and Chinoy, J. J., 1977. Auxin metabolism in developing cotton hairs. J. exp Bot., 28, 1111-1116. Joshi. H. C. and Cleveland, D. W., 1990. Diversity among tubulin subunits: toward what functional end? Cell Motil. Cytoskel., 16, 159-163. Kabsch, W. and Vandekerckhove, J., 1992. Structure and function of actin. A. Rev. biophys, biomol. Struct., 21, 49-76. Lloyd, C. W., 1984. Towards a dynamic helical model for the influence of microtubules on wall patterns in plants. Int. Rev. Cytol., 86, 1-51. Lloyd, C. W. and Seagull, R. W., 1985. A new spring for plant cell biology: microtubules as dynamic helices. Trends Biochem. Sci., 10, 476M78. Maltby, D., Carpita, N. C., Montezinos, D., Kulow, C. and Delmer, D. P., 1979. B-(1,3)-glucan in developing cotton fibers. Structure, localization and relationship of synthesis to that of secondary wall cellulose. Plant Physiol., 63, 1158-1164. McCurdy, D. W. and Williamson, R. E., 1991. Actin and actinassociated proteins. In: The Cytoskeletal Basis of Plant Growth and Form, Lloyd, C. W., (ed.). Academic, London, pp. 3-14. Meinert, M. C. and Delmer, D. P., 1977. Changes in biochemical composition of the cell wall of the cotton fiber during development. Plant Physiol., 59, 1088 1097. Olmsted, J. B., 1986. Microtubule-associated proteins. A. Rev. Cell Biol., 2, 421-458. Osborne, G. G., 1935. Microanalysis of textile fibers. Text. Res., 2, 274-307. Quader, H., Herth, W., Ryser, U. and Schnepf, E., 1987. Cytoskeletal elements in cotton seed hair development in vitro: their possible regulatory role in cell wall organization. Protoplasma, 137, 56-62. Rebenfeld, L., 1965. Morphological foundations of fiber properties. J. Polymer Sci.: Pt C, No. 9, 91-112. Robinson, D. G. and Quader, H., 1982. The microtubule microfibril syndrome. In: The Cytoskeleton in Plant Growth and Development. Lloyd, C. W., (ed.). Academic, London, pp. 109-126. Roelofsen, P. A., 1951. Orientation of cellulose fibrils in the cell wall of growing cotton hairs and its bearing on the physiology of cell wall growth. Biochem. biophys. Acta, 7, 43-53. Roland, J. C. and Vian, B., 1979. The wall of the growing plant cell: its three dimensional organization. Int. Rev. Cytol., 61, 129-166.

660

R.W. Seagull

Rollins, M. L., 1945. Applications of nitrogen dioxide treatment to the microscopy of fiber cell wall structure. Text. Res. J., 15, 65 77. Rowland, S. P., Nelson, M. L., Welch, C. M. and Hebert, J. J., 1976. Cotton fiber morphology and textile performance properties. Text. Res. J., 46, 194-214. Ryser, U., 1985. Cell wall biosynthesis in differentiating cotton fibers. Eur. J. Cell Biol., 39, 236-256. Ryser, U., 1979. Cotton fiber differentiation: Occurrence and distribution of coated vesicles and smooth vesicles during primary and secondary wall formation. Protoplasma, 98, 223 239. Sauter, M., Seagull, R. W. and Kende, H., 1992. Internodal elongation and orientation of cellulose microfibrils and microtubules in deepwater rice. Planta, in press. Schliwa, M., (1981 ) Proteins associated with cytoplasmic actin. Cell, 25, 587-590. Schubert, A. M., Benedict, C. R., Berlin, J. D. and Kohel, R. J., 1973. Cotton fiber development--kinetics of cell elongation and secondary wall synthesis. Crop Sci., 13, 704-709. Schubert, A. M., Benedict, C. R., Gates, C. E. and Kohel, R. J., 1976. Growth and development of the lint fibers of Pima S-4 cotton. Crop Sci., 16, 539-543. Seagull, R. W., 1986. Changes in microtubule organization and wall microfibril orientation during in vitro cotton fiber development: an immunofluorescent study. Can. J. Bot., 64, 1373 1381. Seagull, R. W., 1989a. The plant cytoskeleton. CRC Rev. Plant Sci., 8, 131-167. Seagull, R. W., 1989b. The role of the cytoskeleton during oriented microfibril deposition. II Microfibril disposition in cells with disrupted cytoskeletons. In: Cellulose and Wood: Chemistry and Technology, Schuerch, C., (ed.). Wiley, NY, pp. 811-825. Seagull, R. W., (1990a) Tip growth and transition to secondary wall synthesis in developing cotton hairs. In: Tip Growth in Plant and Funoal Cells, Heath, I. B., (ed.). Academic, San Diego, CA, pp. 261-284. Seagull, R. W., 1990b. The effects of microtubule and microfilament disrupting agents on cytoskeletal arrays and wall deposition in developing cotton fibers. Protoplasma, 159, 44-59. Seagull, R. W., 1991. Role of the cytoskeletal elements in organized wall microfibril deposition. In: Biosynthesis and Biodegradation of Cellulose, Haigler, C. H. and Weimer, P. J., (eds.). Marcel Dekker, New York, pp. 143-163.

Seagull, R. W., 1992a. A 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. Seagull, R. W., 1992b. Commentary: The hormonal regulation of cell elongation through changes in microtubule orientation. Int. J. Plant Sci., 153, iii-iv. Seagull, R. W. and Heath, I. B., 1980. The organization of cortical microtubule arrays in the radish root hair. Protoplasma, 103, 205-229. Seagull, R. W. and Timpa, J. D., 1990. The relationship between reversal frequency and fiber strength. Belt-wide Cotton conferences, pp. 626. Silflow, C. D., Oppenheimer, D. G., Kopczak, S. D , Ploense, S. E., Ludwig, S. R., Haas, H. and Snustad, D. P., 1987. Plant tubulin genes: structure and differential expression during development. Devl Genet., 8, 435-460. Sullivan, K. F., 1988. Structure and utilization of tubulin isotypes. A. Rev. Cell Biol., 4, 687-716. Timpa, J. D. and Ramey, H. H., 1989. Molecular characterization of three cotton varieties. Text. Res. J., 59, 661-664. Wakeham, H., Spicer, H., 1955. The strength and weakness of cotton fibers. Part II: Reversal distribution and breaking properties. Text. Res. J., 25, 585-591. Waterkeyn, L., 1985. Light microscopy of the cotton fiber. In: Cotton Fibers: Their Development and Properties. A technical monograph from the Belgian Cotton Research Group. International Institute for Cotton, Manchester, U.K., pp. 17-22 Westafer, J. M. and Brown, R. M., 1976. Electron microscopy of the cotton fiber: New observations on cell wall formation. Cytobios, 15, 111-138. Williamson, R. E., 1991. Orientation of cortical microtubules in interphase plant cells. Int. Rev. Cytol., 129, 135-206. Willison, J. H. M. and Brown, R. M., 1977. An examination of the developing cotton fiber: Wall and plasmalemma. Protoplasma, 92, 2141. Yatsu, L. Y., 1983. Morphologieal and physical effects of colchicine treatment on cotton (Gossypium hirsutum L.) fibers. Text. Res. J., 53, 515-519. Yatsu, L. Y. and Jacks, T. J., 1981. An ultrastructural study of the relationship between microtubules and microfibrils in cotton (Gossypium hirsutum L. ) cell wall reversals. Am. J. Bot., 68, 771-777. Zeronian, S. H., 1991. The mechanical properties of cotton fibers. J. appl. Pol. Sci., 47, 445-46l.