A quantitative electron microscopic study of changes in microtubule arrays
and wall microfibril orientation during in vitro cotton fiber development
ROBERT W. SEAGULL
United States Department of Agriculture/Agriculture Research Service, Southern Regional Research Center, 1100 Robert E, Lee BLVD., POBox 19687, New Orleans, LA 70179, USA
Summary
A quantitative electron microscopic (E/M) study of thechanges in microtubule arrays and wall microfibrilorientation has been done on in vitro grown cotton fibers.Microtubules change orientation during cotton fiberdevelopment. During fiber initiation and earlyelongation, microtubules have a generally randomorientation. Microtubules re-orient into shallow pitchedhelices as elongation and primary wall depositioncontinue, and into steeply pitched helices during second-ary wall deposition. Accompanying the changes inorientation are increases in microtubule length, number,proximity to the plasmalemma and a decreased varia-bility in orientation of the microtubules. Based on theseobservations, three pivotal stages in microtubule pat-terns were identified during fiber development: (1) thetransition between fiber initiation and elongation, where
microtubules develop a shallow pitched helical orien-tation; (2) the transition between primary and secondarywall synthesis, where microtubules abruptly shift orien-tation to a steeply pitched helical pattern; and (3) earlyhi secondary wall synthesis, where there is a four foldincrease in microtubule number. Microfibrils exhibitchanges hi orientation similar to the microtubules;however significant differences were found when theprecise orientations of microtubules and microfibrilswere compared. During secondary wall synthesis, wallmicrofibrils exhibit some variability in orientation due tointer-fibril bundling, thus indicating that components ofthe wall may also influence final microfibril orientation.
Microtubule arrays in higher land plants are capable ofchanging organization and function in response tovarious stimuli (for review see Seagull, 1989a, 1991).This is apparent not only in cell cycle transitions, butalso within interphase arrays of the cortical micro-tubules. There is growing evidence that construction ofpoly-lamellate cell walls in many cell types is dependanton re-orientations of cortical microtubules (Lloyd,1984; Hogetsu, 1986; Hogetsu and Oshima, 1986; Lloydand Seagull, 1985; Lloyd et al., 1985) and that these re-orientations can be under hormonal control (Ishida andKatsumi, 1991; Akashi and Shibaoka, 1987; Mita andKatsumi, 1986; Roberts et al., 1985).
The developmental regulation of cortical micro-tubule arrays and their involvement in microfibrildeposition remains unclear. We know that arrays ofcortical microtubules are not static since there is anincrease in their numbers during cell extension (Hard-ham and Gunning, 1979; Seagull and Heath, 1980;Seagull, 1983, 1989c). Modifications to microtubulealignment are also possible, with microtubulesexhibiting changes in order (Seagull and Heath, 1980),
clustering during cell differentiation (Seagull andFalconer, 1991) and shifts in orientation (Seagull,1989c). Deposition of wall microfibrils is also not static.The orientation of the most recently deposited layer ofwall microfibrils is under developmental control inmany systems, and this results in the construction ofpoly-lamellate cell walls (Roland and Vian, 1979). Ifthere is a direct relationship between the cytoskeletonand microfibril patterns, then one would predict thatmodifications in microtubule patterns throughout de-velopment would be reflected in changes to theorganization of the inner-most layers of wall microfi-brils. General qualitative studies on the co-alignment ofmicrotubules and wall microfibrils have been done(Sawhney and Srivastava, 1975; Seagull and Heath,1980; Gunning, 1981; Lang et al., 1982; Wada et al.,1990), as have detailed quantitative analyses of micro-tubule arrays (Hardham et al., 1980; Gunning, 1981;Bergfeld et al., 1988; Wasteneys and Williamson, 1987)and wall microfibrils (Hogetsu and Shibaoka, 1978;Takeda and Shibaoka, 1981a,b; Folsom and Brown,1987). However, detailed quantitative studies, comparingmicrotubule and microfibril arrangements during poly-lamellate wall production have not been performed.
562 R. W. Seagull
Quantitative analysis can be used to help define thefunction of cortical microtubule arrays. While generallybelieved to function in controlling wall microfibrilorientation, there is no conclusive data to support aspecific model or mode of action for cortical micro-tubules (Heath and Seagull, 1982; Seagull, 1991).Descriptions of the quantitative relationship betweenmicrotubule and microfibril arrays, with respect toorganization and orientation, may help elucidate themode of action of microtubules and the mechanism(s)for controlling cell wall organization.
Developing cotton fibers appear to have many of thecharacteristics needed for detailed quantitative analysisof microtubule dynamics and their affects on wallmicrofibril deposition. These exceedingly long, single-cell trichomes contain extensive arrays of cytoskeletalelements (both microtubules and microfilaments) thatappear to be involved in controlling the organization ofmicrofibrils in both primary and secondary cell walls.Fibers contain highly organized arrays of microtubuleswhich appear to shift orientation in a developmentallycontrolled manner (Seagull, 1986, 1989c; Yatsu andJacks, 1981; Westafer and Brown, 1976; Itoh, 1974).Pharmacological studies using microtubule-disruptingagents have provided circumstantial evidence for adirect role of microtubules in regulating microfibrilorientation (Seagull, 1989b, 1990a). Preliminary quanti-tative observations of the cytoskeleton during fiberdevelopment, comparing young (3 and 10 days postan thesis (DPA)) and old (30 DPA) fibers, indicate thatbesides changes in orientation, microtubules appear tochange in length, number and organization (Seagull,1989c). These changes are consistent with an observedincrease in tubulin content (Kloth, 1989). Increases intubulin content and microtubule arrays appear tocorrelate temporally with secondary wall synthesis;however detailed descriptions of microtubule arrays atstages leading up to, and immediately after, the onset ofsecondary wall synthesis have not been done. Cottonfibers are a fairly synchronous population of single cellsthat develop in a highly predictable manner (Seagull,1990b), thus facilitating any future biochemical analysisof cytoskeletal elements.
Observations using immunocytochemistry and lightmicroscopy reveal a correlation between microtubuleand microfibril orientation during fiber development(Seagull, 1986). Changes in microtubule orientationappear to predict the subsequent changes in microfibrilorientation that occur with the initiation of secondarywall synthesis (Seagull, 1989b). However, the limitedresolution of the techniques employed in this previousstudy may not have detected subtle differences betweencytoskeleton and wall microfibril patterns. Thesedifferences may shed light on the mechanisms respon-sible for the dynamic interaction between microtubulesand microfibrils. This study provides detailed quantitat-ive and qualitative observations at the electron micro-scope (E/M) level, describing shifts in orientation, thedegree of co-alignment within the microtubule popu-lations and between microtubules and wall microfibrilsthat went undetected in previous studies.
Materials and methods
CulturingSeeds of cotton, Gossypium hirsutum L., variety SJ-2 (CottonIncorporated, Raleigh, NC), were sown in soil and grownunder glass house conditions (day/night temperatures of25/20°C, light/dark cycle of 16/8 h). Flowers were harvested 2days post anthesis (DPA). Ovules were removed asepticallyand floated on growth medium in 100x25 mm Petri dishes.Ovules were grown in vitro as per the techniques of Beasleyand Ting (1973) using a NAA:GA3 concentration of 5 mM:0.2mM. Cultures were grown at 34CC in the dark.
Electron microscopyCotton fibers attached to ovules of various ages were fixedusing standard chemical fixation (Seagull, 1986). Briefly,fibers of various ages were fixed in 0.1 M PIPES buffer (pH6.8), containing 1.0% glutaraldehyde and 2 drops of Tween80/10 ml, at room temperature for 1-2 h. After 3 washes inPIPES buffer, fibers were post-fixed for 1 h in 1.0% OsO4,buffered as above. Rapid dehydration of specimens wasaccomplished using two 15 min treatments with 2,2-dimeth-oxypropane, followed by two 30 min treatments with 100%ethanol. Fibers were embedded in Spurr's resin, betweensilicone-coated microscope slides.
Cotton fibers to be sectioned were selected based onstructural integrity at the light microscope level. Serial sectionreconstructions of microtubule arrays were performed asdescribed previously (Seagull and Heath, 1980). Reconstruc-tions were made from a minimum of 10 consecutive sections.Observations were made using a Philips 300 electronmicroscope and recorded on Kodak E/M film SO-281.
Microfibril patterns were examined using platinum-coatedwall fragments prepared as follows: ovules with attachedfibers were frozen in liquid nitrogen. Samples were gentlyground with a chilled mortar and pestle to separate fibers fromthe ovule. Ovules were removed and the remaining fibersvigorously ground at liquid nitrogen temperature. Fiberfragments were wanned to room temperature and washedrepeatedly with water to remove cytoplasmic components.Wall fragments were washed until minimal cytoplasmiccontamination was evident by examination with DIC lightmicroscopy. Clean wall fragments were layered onto Form-var-coated mesh grids, air dried and coated with platinum at ashadow angle of approximately 20 degrees.
Calculations of microtubule and microfibril parametersChanges in number and orientation of microtubules weredetermined from serial section reconstructions. A minimumof three reconstructions, each from different fibers, were usedfor the analysis. Microtubule number was calculated bydetermining the number of microtubules per unit length ofcell within the area of the reconstruction. A minimum of fivereadings were taken from each reconstruction. Analysis ofcross sectioned fibers produced values for microtubulenumbers which were not significantly different from recon-struction analysis (data not shown). However, cross sectionscould not be used for accurately determining angulardeviation of microtubules relative to the fiber long axis.
Angular deviation of microtubules was calculated fromreconstructions, by measuring the orientation of individualmicrotubules relative to the axis of fiber elongation (i.e.0°=parallel to long axis of fiber; 9O°=perpendicular to longaxis of fiber). Fifty microtubules from each reconstructionwere measured.
Microtubule proximity to the plasmalemma was measured
Microtubules and microfibrils in cotton fiber development 563
using sections in which microtubules appeared in cross section(in fibers less than 24 DPA these sections were approximatelymedian longitudinal; in fibers 24 DPA and older, sectionswere more oblique). A minimum of 800 microtubules wereobserved from at least three different fibers at each age.
Angular deviation of wall microfibrils was determined fromwall fragments of three different fibers from each age. Theorientation of fifty individual microfibrils was measuredrelative to the axis of fiber elongation. Only wall fragmentswhich were long enough to reveal a detectable fiber axis wereused for these measurements. Due to the preservationtechniques required to visualize individual wall microfibrils,the same fiber could not be used to analyze the orientation ofboth microfibrils and microtubules.
Statistical analysisThe angle means of microtubules and microfibrils werecompared among the different fiber ages using a t-test tocompare all possible pairs of fiber ages. If the variancediffered between the angles observed at the two ages beingcompared, then a Satterthwaite approximation was used tocompare the mean angles (Steel and Torrie, 1980). Thisprocedure was also used to compare data on microtubulelength and proximity to the plasmalemma. Comparison of themeans for microfibril and microtubule angles at each age wasalso conducted using Mests with approximations when thevariances were unequal.
Variability within populations of microtubules and microfi-brils were monitored using a comparison of angle variancesamong the fiber ages and the Bartlett-Kendall log s (squared)homogeneity of variance test (Anderson and McLean, 1974).Mests were used to compare microtubule and microfibrilangle variances for each age (Steel and Torrie, 1980).
A trend analysis was done for both the populations ofmicrotubules and microfibrils. Regressions were comparedusing ANOVA analysis of covariance.
The relationship between microtubule number and fiberage was analyzed using regression analysis.
Results
Qualitative analysisDeveloping cotton fibers contain extensive arrays ofcortical microtubules throughout development (Figs 1-7). Both single sections that graze through the corticalcytoplasm and cell wall (Figs 1A-7) and serial sectionreconstructions (Figs 1B-7B) clearly illustrate thepreviously documented (Seagull, 1986) reorientation ofmicrotubule arrays. Reconstruction plots however alsoclearly illustrate the degree of variability in microtubuleorientation, a characteristic of the microtubule arraysthat was not evident in previous studies. Grazingsections also reveal the general order of the innermostlayer of wall microfibrils (Figs 3A-7A), but do notreveal the degree of variability in microfibril orientationthat is observed in shadowed wall preparations (Figs2C-7C). A general comparison of microtubule (Figs 1B-6B) and microfibril (Figs 2C-7C) orientations indicatethat arrays show more variability in younger (2, 3 DPA)cells than in older ones (30, 36 DPA).
Quantitative analysisUsing reconstruction analysis of microtubule popu-lations and shadowed wall preparations, quantitative
descriptions of the changes in microtubule and microfi-bril arrays during fiber development can be obtained(Figs 8-12).
Microtubule analysisRegression analysis of microtubule numbers duringcotton fiber development indicates that numbers ofmicrotubules slightly decline over the first 19 DPA,then begin to increase between 19 and 24 DPA (Fig. 8).The incorporation of the 30 and 36 DPA data results ina steeper positive slope in the regression line, thusindicating a continued increase in microtubule numberto 36 DPA. Paralleling this increase in number is anincrease in the relative length of microtubules (Fig. 9).With increasing fiber age there is a decreasing pro-portion of microtubule ends compared to the totalnumber of microtubules observed in the reconstructions(Figs 1B-7B), indicating that microtubules becomerelatively longer during fiber development. Statisticalanalysis indicates that there is no significant change inthe length of microtubules between 2 and 3 DPA butthen microtubules become progressively longer at eachsampling date (Fig. 9).
Potential for interaction between microtubules andthe plasmalemma was assessed by calculating theproportion of microtubules within 25 nm of theplasmalemma (Fig. 10). At all ages, less than 20% ofthe microtubules are located more than 25 nm from themembrane. There is, however, a steady increase inmicrotubule proximity to the plasmalemma as fibersdevelop. By 36 DPA, only about 5.0% of themicrotubules are located more than 25 nm from theplasmalemma. The data form three groups: 2 and 3DPA are not significantly different; 7, 19 and 24 DPAnot significantly different; 30 and 36 DPA not signifi-cantly different. However, the three groups are signifi-cantly different from one another. Microtubules thatare not associated with the plasmalemma are oftenfound in small clusters and have other microtubuleslocated between them and the membrane (data notshown).
Changes in microtubule angle (relative to the axis ofcell elongation) can be plotted (Fig. 11) to illustratetrends in orientation during development. 1 DPA fibers(Fig. 1) have a somewhat random orientation (42.4°from axial) and a large degree of variability (plus/minus24.2°). Microtubule orientation gradually changes to ashallow pitch helical pattern (80° from axis of cellelongation) by 7 DPA and remains constant through 19DPA. Although quantitative data were not collected,the microtubule arrays appear to maintain this orderthrough 22 DPA (data not shown). The comparison ofmeans for microtubule angle indicates that from fiberinitiation to primary wall synthesis there is a transitionfrom random to shallow pitched helical arrays. Duringsecondary wall synthesis (beginning at 24 DPA) themicrotubules shift to steeply pitched helical arrays.Steeply pitched helical arrays of microtubules are firstdetected at 24 DPA (Fig. 11). The pitch of the helixvaries with subsequent development but maintains ageneral steep pitch (Fig. 11, Table 1). The variance for
564 R. W. Seagull
B
Fig. 1. One day post anthesis (DPA) cotton fibers. Axis of fiber elongation is parallel to the page width. (A) A thinsection that grazes through the cell wall and cortical cytoplasm reveals cortical microtubules (arrowheads) which areoriented both parallel and oblique to the axis of fiber elongation. Bar, 0.5 ism. (B) Serial section reconstruction of themicrotubule population reveals a random organization of both long (large arrowheads) and short (small arrowheads)microtubules. Bar, 1.0 /an.
Fig. 2. Two days post anthesis (DPA) cotton fibers. Axisof fiber elongation is parallel to the page width in (A,Band C). (A) A thin section that grazes through the cellwall (C) and cortical cytoplasm. Numerous microtubules(arrowheads) are oriented generally transverse to the axisof cell elongation, while others have very differentorientations (arrows). Bar, 0.5 /zm. (B) Serial sectionreconstruction of the microtubule population showsvariability in microtubule orientation. Microtubulediameters are not drawn to scale. Both long (large
arrowhead) and short (small arrowheads) microtubules areevident. Bar, 1.0 /.cm. (C) Platinum-shadowed wallpreparation shows both the inner (I) and outer (O) walllayer. Microfibrils (arrowheads) with various orientationsare observed on the inner-most wall layer. Bar, 0.5 /im.(D) Low magnification of a typical wall fragment used todetermine fiber axis of elongation. The outer (O) and inner(I) wall layers are evident. The cylindrical nature of thefragment readily reveals the axis of fiber elongation(double headed arrow). Bar, 2.0 /tm.
Microtubules and microfibrils in cotton fiber development 565
566 R. W. Seagull
Fig. 3. For legend see p. 568.
Microtubules and microfibrils in cotton fiber development 567
Fig. 4. For legend see p. 568.
568 R. W. Seagull
Fig. 3. Three DPA cotton fibers with axis of fiberelongation paralleling the page width. (A) Grazing sectionthrough the cell wall and cortical cytoplasm of a chemicallyfixed fiber. Microtubules (arrowheads) with variousorientations are evident in the cortical cytoplasm. Themicrofibrils of the innermost wall layer appear (arrows) tohave an orientation that mimics the microtubules. Bar, 0.5fan. (B) Serial section reconstruction of microtubule arrayshows both long (large arrowheads) and short (smallarrowheads) elements. The most predominant orientationto the tubules is a shallow pitched helix. Bar, 1.0 fan.(C) Fiber wall fragment shadowed with platinum, showingboth inner (I) and outer (O) wall layers. Innermost layerof wall microfibrils is oriented transversely to the axis ofelongation; however there are significant numbers ofmicrofibrils that deviate from the predominant orientation(arrows). Bar, 0.5 fim.Fig. 4. Nineteen DPA cotton fibers with the axis of fiberelongation paralleling the page width. (A) Grazing sectionthrough the cell wall and cortical cytoplasm of a chemicallyfixed fiber. Microtubules are oriented in a shallow pitchedhelix and are parallelled by the innermost layer of wallmicrofibrils (arrows). Bundles of microfilaments(arrowheads) lie just beneath the microtubules and parallelthe length of the fiber. Bar, 0.5 fan. (B) Serial sectionreconstruction of the microtubule array shows fewermicrotubule ends and less variability in microtubuleorientation. Bar, 1.0 fan. (C) Platinum-shadowed wallfragment shows both the inner (I) and outer (O) walllayers. The microfibrils of the innermost wall layer(arrowheads) are predominantly oriented transverse to thefiber long axis. Bar, 0.5 fan.
the angular deviation of microtubule arrays shows asteady decrease during fiber development, indicative ofincreasing order within the microtubule array. Meanscomparison tests of the variability in microtubule angleindicate that there is a constant degree of variabilityearly in development (1-3 DPA), followed by a gradualincrease in order within the microtubule arrays withincreasing fiber age (Table 1). Superimposed onchanges in the angle of microtubule populations arecontinued increases in order (Table 1), length (Fig. 9)and proximity to the plasmalemma (Fig. 10).
Table 1. Analysis of means test
Fiberage
Microtubule angle Microfibril angle
Means Variance Means Variance
1237
1619243036
42.4A
64.5B"68.8B"74.7C "79.9C
79.6C a
17.5D '21 .5 E "1 7 . 1 D a
475.2A
644.7A a
345.3AB a
263.0AB °89.3C
94.6C >
26 .5 D a
6.5E»9 . 1 E >
6 5 . 1 A a
78.8B b
75.3C '
72.4C b
15.7D b
2 3 . 1 E a
17.8F '
312.0* "107.7BC b
144.5A "
115.1BD a
10.3E b
37.0CF b
25 .2 F b
A comparison of means and variance for angular deviationvalues, between microtubules and microfibrils at various stages offiber development. Upper case letters designate comparisons incolumns only. Lower case letters designate row comparisons ofmeans or variances of microtubules and microfibrils.
Fig. 5. Twenty-four DPA fibers oriented with the long axisparalleling the width of the page. (A) Grazing sectionthrough the cell wall (W) and cortical cytoplasm of achemically fixed fiber. Numerous microtubules(arrowheads) are oriented in a steeply pitched helix.Microfibrils of the secondary cell wall (arrows) appear tohave an orientation that parallels the microtubules. Bar,0.5 fim. (B) Serial section reconstruction shows a steeplypitched array of microtubules. (C) Platinum-coated wallfragment shows both the outer (O) and inner (I) layer ofwall microfibrils. Microfibrils exhibit some bundling(arrowheads) with bundles of parallel microfibrils orientedin slightly different directions. Bar, 0.5 fan.
Microfibril analysisWall microfibrils exhibit shifts in orientation compar-able to those seen in the microtubule populations,becoming more ordered within the array during fiberdevelopment (compare Figs 2C-7C). The statisticalanalysis of microfibril order (Fig. 12, Table 1) indicatessignificant changes in orientation early in development(2-7 DPA), followed by a period of constant orientation(7-19 DPA). Wall microfibril orientation exhibits a shiftto shallow pitch helical during primary wall synthesis,followed by orientation in steeply pitched helical duringsecondary wall synthesis. Microfibril orientation doesnot remain constant after the transition to steeplypitched arrays, as there are significant differences inorientation between 24 and 30 DPA (Table 1). A meanscomparison test indicates shifts in the degree of orderwithin the wall microfibrils (Table 1). Trend analysisindicates that populations of microfibrils exhibit de-creasing variance, thus becoming more ordered duringfiber development.
Microtubule/microfibril co-alignmentA comparison of microtubule and microfibril angles atvarious stages of development shows similar, but notprecise co-alignment (Table 1). The same developmen-tal pattern (i.e. increases in order and decreases invariability) is seen when comparing the variabilitywithin the populations of microtubules and microfibrils(Table 1). Trend analysis showed that both exhibitdecreasing variance with increasing age. The variancewithin the microtubule population is greater than thatof the microfibril population early in development, butdecreases significantly (P=0.0002) faster than themicrofibril variance. This results in the microtubulepopulations having a lower variance than the microfibrilpopulations during secondary wall synthesis. Specificage comparisons (i.e. 3, 19, 24 DPA) exhibit small butsignificant differences in mean angle and variancebetween microtubules and microfibrils.
Discussion
Changes in microtubule arrays during developmentDuring early fiber development, at or near thetransition between fiber initiation and elongation (i.e. 1DPA), cortical microtubules have a random orientation
Microtubules and microfibrils in cotton fiber development 569
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570 R. W. Seagull
Fig. 6. For legend see p. 572,
Microtubules and microfibrils in cotton fiber development 571
c
Fig. 7. For legend see p. 572.
572 R. W. Seagull
Fig. 6. Thirty DPA fibers oriented with the long axisparalleling the width of the page. (A) Grazing sectionthrough the secondary cell wall (W) and cortical cytoplasmof a chemically fixed fiber. Plasmalemma appears as a darkgrey area (PM) upon which the closely packedmicrotubules (arrowheads) are found. (B) Serial sectionreconstruction shows a steeply pitched helical array oftightly packed microtubules. While many of themicrotubules are long enough to traverse the entirereconstruction (large arrowhead), relatively shortmicrotubules are evident (small arrowhead). (C) Platinum-coated wall fragment reveals both the outer (O) and inner(I) layers of microfibrils. The innermost layer ofmicrofibrils is oriented in a steeply pitched helix thatmimics the orientation of the microtubules. Microfibrilbundling is evident at the fragmented ends of the wall(arrows). Bar, 0.5 fan.Fig. 7. Thirty-six DPA fibers oriented with the long axisparalleling the width of the page. (A) A grazing sectionthrough the secondary cell wall and cortical cytoplasm of achemically fixed fiber. Arrays of microtubules (arrows) arehighly organized and tightly packed. Bar, 0.5 fan. (B)Serial section reconstruction shows the tightly packed arrayof microtubules. Most microtubules (large arrowheads)traverse the entire reconstruction, while others (smallarrowhead) are relatively short. Bar, 1.0 fim. (C) Platinum-coated wall fragment showing both the outer (O) and inner(I) wall layers. The innermost layer of microfibrils isorganized into parallel arrays, oriented in a steeply pitchedhelix. Undulation of bundles of microfibrils is evident(arrows). Bar, 0.5 fffn.
0.90 r
m
0.75 -
0.60 -
g 0.45
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0.30 |-
0.15 I-
0.00
aI
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1 1
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1 CI
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0 5 20 25 30 35 40
FIBER AGE (DPA)
Figs 8-12. Histograms represent mean values for eachanalysis, with vertical bars representing standard errorvalues.Fig. 8. Changes in microtubule concentration (number ofmicrotubules • fan"1) during fiber development. Linearregression analysis illustrates a slight decrease inmicrotubule concentration through 19 DPA. Microtubuledensity increases gradually by 24 DPA and sharply by 30DPA, resulting in a four-fold increase in microtubulenumber between primary (2-19 DPA) and secondary (30-36) wall synthesis.
(Figs 1,10, Table 1). The rather bulbous appearance ofthe fibers at this stage of development (Berlin, 1986)indicates some lateral cell expansion. This is consistent
24
E 20
oi 16
3
o 8cco^ 4
0 5 10 15 20 25 30 35 40
FIBER AGE (DPA)
Fig. 9. Relative changes in microtubule length during fiberdevelopment, determined from reconstruction plots. Theratio of the number of microtubule ends (terminations) pertotal number of microtubules observed in a reconstructionis indicative of relative length. The closer the ratioapproaches the value 2 (i.e. that both ends of themicrotubule are present in the area of analysis), the shorterthe microtubules in the population. Letters assigned toeach histogram represent statistical comparisons.Histograms with the same letter are not significantlydifferent from one another using f-test analysis.Microtubule lengths do not change significantly between 2and 3 DPA; however by 7 DPA the microtubules haveincreased in length significantly. The length appears toremain constant during primary wall synthesis.Microtubules exhibit increasing lengths during secondarywall synthesis (24-36 DPA).
with the observed pattern of microtubule orientation.Analysis of microfibril orientation at this stage ofdevelopment was inconclusive because of the extremedifficulty in assessing the axis of fiber elongation in wallfragments (data not shown). As a result, no comparisonof orientations between microtubules and microfibrilswas done at this early stage of elongation.
During the next two days of development, micro-tubules organize into shallow pitched helices (Figs2B,3B,11, Table 1), with the pattern stabilizing by 7DPA at an orientation of about 75-80° from axial. Thispattern is maintained throughout primary wall syn-thesis.
The arrays of microtubules do not remain staticduring primary wall synthesis. Variability within themicrotubule population steadily declines during fiberdevelopment (Table 1), indicating some type of align-ment mechanism. Accompanying the decrease invariability in orientation, arrays appear to be modifiedthrough changes in microtubule length (Fig. 9) andproximity to the plasmalemma (Fig. 10). These changescould account for the observed increase in microtubuleorder (Table 1). Increasing the length of microtubulesand restricting the location of these linear structures tonear the plasmalemma would result in the formation ofmore parallel arrays.
To function in directing wall microfibril orientation,cortical microtubules are believed to interact with the
LJ
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8 -
4 -
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Microtubules and microfibrils in cotton fiber development 573
100
0 5 20 25 30 35 40
FIBER AGE (DPA)
Fig. 10. Relative changes in the proximity of microtubulesto the plasmalemma as measured by the percentage ofmicrotubules found at distances greater than 25 /xm fromthe cell membrane. Letters assigned to each histogramrepresent statistical comparisons. Histograms with the sameletter are not significantly different, based on Mestcomparisons. Throughout development, the majority ofmicrotubules (> 80%) are within 25 /m\ of theplasmalemma; however there is a significant trend towardscloser association with the plasmalemma as fibers age.Microtubules are significantly closer to the plasmalemmaduring secondary wall synthesis (30, 36 DPA), but at theinitiation of secondary wall (24 DPA) microtubuleproximity to the plasmalemma is the same as duringprimary wall deposition (7, 19 DPA).
O
3003
Oono
80
60
40
20
• 1
•
•
-
1
i i
0 5 10 15 20 25 30 35 40
FIBER AGE (DPA)
Fig. 11. Changes in orientation of the microtubule arrayrelative to the axis of fiber elongation. During fiberinitiation and early elongation (1-3 DPA) microtubulesbecome progressively more organized into shallow pitchedhelices (relative to the fiber long axis). Once established,the orientation of the array remains constant throughprimary wall synthesis. At the onset of secondary wallsynthesis there is an abrupt shift of orientation to a steeplypitched helix.
LJ_JO
80
_, 60
CDLI 40o
y 20
0 5 10 15 20 25 30 35 40
FIBER AGE (DPA)
Fig. 12. Changes in orientation of the microfibril arrayrelative to the axis of fiber elongation. Microfibrils becomeprogressively more organized into shallow pitched helicesduring early fiber elongation (2-3 DPA) and maintain thatorientation during primary wall synthesis (7-19 DPA).Onset of secondary wall synthesis is signaled by an abruptshift in orientation to a steeply pitched helix.
plasmalemma (Heath and Seagull, 1982; Seagull,1989a). Interaction between microtubules and theplasmalemma could occur via an array of associatedproteins or by direct tubulin membrane interaction (fordiscussion see Hepler, 1985). While projections can bevisualized between the membrane and microtubules incotton fiber (Westafer and Brown, 1976; Ryser, 1979),there is often a very precise separation without anydetectable structures between the two. For this reasonthe potential for interaction between microtubules andthe plasmalemma has been assessed by describing theproximity of the two structures, with 25 nm being wellwithin the length range for microtubule associatedproteins (MAPs) extending from the surface of themicrotubule (Mandelkow and Mandelkow, 1989).While cortical microtubules can tightly interact with theplasmalemma (Lloyd et al., 1980) and this interactionmay be ATP-sensitive (Marchant, 1978), a specificprotein or set of proteins thus involved has yet to beidentified in higher plants. The specific associationbetween cortical microtubules and the plasmalemma(Fig. 10) is consistent with the postulated presence ofsome type of linking agent(s) or protein(s). Suchbinding agents must be present throughout develop-ment since over 80% of the microtubules associate withthe plasmalemma at any one time. However, thegradual yet significant increase in association of theseelements indicates some type of developmental regu-lation of the interaction.
Changes in microfibril orientations duringdevelopmentWall organization appears to be developmentallyregulated, with microfibril deposition changing in anage-dependent manner (Fig. 12, Table 1). The organiz-ation of the innermost layer of microfibrils is ofparticular interest since this region is proposed to be the
574 R. W. Seagull
load-bearing wall layer, responsible for defining cellexpansion characteristics (Richmond et al., 1980). Thepatterns of microfibril orientation at the cell wall-plasmalemma interface dramatically impact cell expan-sion characteristics and cell shape. The more parallelthe organization within the microfibril arrays, thestronger the cell wall and thus the more capable ofrestricting turgor pressure-driven cell expansion intospecific directions. In addition, being the most recentlydeposited, this layer should reflect the orientation ofthe proposed control system governing microfibrilorientation, i.e. the microtubules (Robinson andQuader, 1982). If microtubules are directly responsiblefor controlling the orientation of microfibrils duringdeposition, then characteristics of the microfibril pat-tern should mimic precisely the microtubules.
There is a significant shift in microfibril depositiontowards a direction transverse to the cell's long axisearly in the elongation phase (2,3 DPA). The shiftappears to stabilize by 7 DPA and remain constantthrough 19 DPA (Fig. 12, Table 1). Accompanying thisshift in orientation is a decreased variability within thepopulation of microfibrils (Table 1). Variability withinthe microfibril population is less than that of themicrotubules, indicating that other components may beinvolved in producing final wall organization.
The organization of microfibrils can be modifiedduring wall development through interactions with avariety of wall components (Preston, 1979). There arechanges in the chemical composition of the fiber wallduring primary wall synthesis (Meinert and Delmer,1977) that may account for the observed decrease invariability of microfibril orientation. Variability inmicrofibril orientation may be impacted by the degreeof interaction between adjacent microfibrils. A numberof non-cellulosic components, as well as cellulose itself,have been shown to cross-link wall microfibrils(McCann et al., 1990). Factors such as the changingrates of cellulose deposition or changes in proteincontent and non-cellulosic components (Meinert andDelmer, 1977) may be involved in controlling andmodifying variability within the cell wall.
Microtubule/microfibril co-alignmentBoth the microtubules and microfibrils appear to beunder developmental regulation as both exhibit specificshifts in orientation and changes in variability withintheir populations (Figs 11, 12, Table 1). If microfibrilorientation is controlled by the orientation of corticalmicrotubules, then one would predict co-alignment ofthe two. The degree of co-alignment may be defined bythe mechanism of the interaction (Heath and Seagull,1982; Seagull, 1991). The correlation between micro-tubules and microfibrils with respect to orientation andvariability within the population will provide clues as tothe factors responsible for microfibril patterns.
Microtubules and microfibrils (1) exhibit similarorientations throughout development; (2) exhibit anabrupt shift in orientation at the same developmental
stage and (3) become more ordered during develop-ment (compare Figs 11 and 12, Table 1). These resultsgenerally agree with previous observations made usingimmunofluorescence microscopy (Seagull, 1986).
Detailed comparisons between microtubules andmicrofibrils indicate small but significant differences inorientation at several stages of development (i.e. 3, 19and 24 DPA). Microtubule arrays exhibit more varia-bility than microfibrils at 3 and 24 DPA and lessvariability at 30 and 36 DPA. These observationsindicate several possibilities: (1) that the interactionbetween the two is not direct, as suggested by Heplerand Palevitz (1974) and Staehlin and Giddings (1982);(2) that other components (in the cytoplasm or the cellwall) are involved in defining the orientation ofmicrofibrils; (3) that not all the microtubules function inwall deposition (i.e. the more randomly orientedmicrotubules do not function in orienting wall microfi-brils); (4) that the observed differences, while math-ematically significant, are not biologically significant.
There is no compelling reason to assume that all ofthe microtubules in the cortical cytoplasm function indirecting wall microfibril deposition. It is feasible thatsome of these microtubules may have other, undes-cribed functions in the cell. Cortical microtubule arraysin developing cotton fiber are not homogeneous withrespect to stability against microtubule-disruptingagents (Seagull, 1990). The final microfibril orientationin the cell wall may be the accumulated result ofinteractions with microtubules, and possibly with othercytoplasmic or membrane components and with otherwall components.
It is generally accepted that microtubule orientationcontrols microfibril patterns in many if not all higherplant systems (Seagull, 1989a; Lloyd, 1987). In cottonfiber development this would imply that microtubules(followed by microfibrils) should undergo re-orien-tations from random (fiber initiation) to shallow pitchhelical (primary wall synthesis) to steeply pitchedhelical (secondary wall synthesis). However, in bothprimary (3 and 19 DPA) and secondary (24 DPA) walldeposition of cotton fibers, the orientations of microfi-brils are significantly different from microtubules. In 3DPA fibers, the helically organized microfibrils have asteeper pitch than the microtubule arrays, very muchlike the microfibrils in the primary walls of older fibers(i.e. 7-19 DPA). In 24 DPA fibers, the helicallyorganized microfibrils have a shallower pitch than theairays of microtubules, very much like the microfibrilsin older fibers (i.e. 30 and 36 DPA). The differences inorientation between the arrays of microtubules andmicrofibrils are not large, but they are statisticallysignificant (Table 1). A simple explanation for thisapparent discrepancy is not readily apparent.
During secondary wall deposition the microtubulesco-align with microfibrils; however, the two populationsexhibit differing degrees of variability (Table 1). In thiscase, interactions between fibrils may result in subtlechanges in microfibril order and the degree of varia-bility. Without the influence of microtubules, microfi-
Microtubules and microfibrils in cotton fiber development 575
brils tend to form swirled clusters in the cotton fiberwall (Seagull, 1989b, 1990a), thus indicating that inter-microfibril associations are possible and result in thecurving patterns of microfibrils. This possibility isconsistent with the observed undulating pattern ofmicrofibrils evident as fibril bundles appear to interactwith each other in the wall (Figs 6C,7C). As thesecondary cell wall of the cotton fiber is exclusivelycellulose (Meinert and Delmer, 1977), these interac-tions are most likely via hydrogen bonding betweenglucose chains in adjacent microfibrils. Inter-fibrilhydrogen bonding may modify the microfibril orien-tation imparted via the microtubules and result in theobserved undulations (Figs 6C,7C).
It is also possible that the variability observedbetween the organizations of microtubules and microfi-brils may be the result of differences in the rigidity ofthe two populations. Wall microfibrils are closelypacked into the cell wall, whereas the microtubules arewidely spaced (relative to the packing of microfibrils) inthe cortical cytoplasm. Therefore, microtubules may bemore prone than the microfibrils to slight reorientationsdue to fixation or physical affects on the fibers. Artifact-induced changes in the orientation of microtubules isconsistent with the observed greater variability inmicrotubule arrays. This could account for the slightdifferences in orientation observed between micro-tubules and microfibrils.
It is equally possible that the observed differencesbetween the orientations of microtubules and microfi-brils may be mathematically significant (Table 1) butbiologically insignificant. The observed fluctuations inmicrotubule orientation and organization may be theresult of normal biological variations that do not impactoverall growth and development within fibers. Thispossibility is consistent with the general lack of a trendto alter the variability in angle of either the micro-tubules or microfibrils (i.e. variance in the microfibrilsangle decreases, then increases, then decreases, etc.). Itmay be proposed that the observed variability inorganization is within the "biological limits of toler-ance" for microtubules to influence microfibril orderand for microfibrils to control the expansion character-istics of the developing fiber.
The observations concerning microtubule and micro-fibril co-alignment are consistent with several of thecurrent models explaining the role of the cytoskeletonin regulating wall organization. Indirect models (asdefined by Heath and Seagull, 1982) propose no directinteraction between microtubules and wall microfibrils,thus variability in organization would not negate aninteraction between the two. Staehelin and Giddings(1982) propose another indirect model, with channell-ing of cellulose synthetic complexes between tracks ofmembrane rigidity that are induced by the interactionof microtubules with the fluid plasmalemma. All of theindirect models allow for a certain amount of variabilityin the system and are thus consistent with the observeddifferences in alignment between microtubules andmicrofibrils.
Conclusions
The fluctuations in microtubule and microfibril organiz-ation during cotton fiber development clearly illustratethe complexity of the mechanisms that control cell wallorganization. The control of cell wall organization is offundamental importance to plant morphogenesis andthus the data presented here relate not only to cottonfiber development but also to plant morphogenesis ingeneral.
Arrays of microtubules undergo modification duringfiber development. Three pivotal points in developmenthave been identified: (1) the transition from fiberinitiation to elongation, when microtubules undergo are-orientation and ordering; (2) the transition betweenprimary and secondary wall synthesis, when micro-tubules change orientation from shallow pitch helices tosteeply pitched ones; and (3) early in secondary wallsynthesis when the number of microtubules increasesfour-fold. Other characteristics of the microtubulearrays also appear developmentally controlled (i.e.length and proximity to the plasmalemma); howeverthese transitions occur more gradually over time.
Changes in microtubule arrays relate to in vivodynamics. Clues to possible chemical modifications tothe micTOtubule arrays can be obtained by the analysisof the types of changes observed in microtubules.Increases in microtubule order and proximity to theplasmalemma may be indicative of the presence ofmicrotubule associated proteins (Olmsted, 1986). Thesame type of argument could be made for the observedincrease in microtubule lengths and numbers. Pharma-cological disruption or protein characterization of thecytoskeleton at these points of development mayprovide clues as to the mechanisms of (i.e. proteinsinvolved in) processes such as microtubule organizationand re-orientation, the role of dynamic instability(Mitchison and Kirschner, 1984) in plants for control-ling microtubule turnover and the factors affectingmicrotubule polymerization and number.
In the cotton fiber it appears that both microtubulesand microfibrils themselves may influence the finalorganization of the innermost wall layer. The concomi-tant shifts in orientation of microtubules and microfi-brils strongly indicate a cause and effect relationship.Microfibril arrays exhibit less variability than themicrotubules, thus indicating that another factor,perhaps interaction of wall components, may bemodifying the order imparted by the microtubules.Interaction among wall components has been suggestedpreviously (Seagull, 1989a, 1990b) to explain microfibrilorder in the absence of microtubule influence.
Thanks to Dr. Bryan Vinard and Dr. David F. Millie forassistance with the statistical analysis and preparation of thegraphs and table, as well as to Drs. D. Dixon, A. Lax, T.Lonergan and K. Vaughn for critical reviews and valuablesuggestions for this manuscript.
Mention of a trademark, proprietary product or vendordoes not constitute a guarantee or endorsement of this item bythe USDA and does not imply its approval to the exclusion ofvendors that may also be suitable.
576 R. W. Seagull
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{Received 14 October 1991 - Accepted 16 December 1991)
