Altered patterns of tubulin polymerization in dividing leaf cells of
Chlorophyton comosum
after a hyperosmotic
treatment
G. Komis, P. Apostolakos and B. Galatis
Faculty of Biology, Department of Botany, University of Athens, Athens 157 84, Greece
Summary
• Microtubule organization and tubulin polymerization in meristematic leaf cellsof
Chlorophyton comosum
treated with an aqueous solution of 1 M mannitol,inducing plasmolysis, were examined with immunofluorescence and transmissionelectron microscopy.• Hyperosmotic treatment induced disintegration of the interphase microtubulesystems. Free tubulin, either liberated from the depolymerized microtubules or pre-existing as a nonassembled pool, was incorporated into a network of paracrystals.In most of the dividing cells, mitotic and cytokinetic microtubule systems werereplaced by atypical spindle-like structures displaying bipolarity and atypical phrag-moplasts, respectively. These atypical mitotic and cytokinetic structures consisted oflarge densely packed bundles of macrotubules (32 nm diameter) or macrotubulesand paracrystals. Tubulin paracrystals also occurred in ectopic positions in plasmolysedmitotic and cytokinetic cells. Dividing cells displaying paracrystals only did not formatypical mitotic and cytokinetic apparatuses.• Short hyperosmotic stress causes disintegration of all microtubule arrays individing cells of
C. comosum
. Free tubulin is incorporated into macrotubules andtubulin paracrystals. The latter exhibit definite periodicity and characteristic finestructure.
Plants continuously encounter different types of environmentalstresses, among which water deficiency is very critical for theirsurvival. They have developed particular mechanisms to toleratethese stresses. Plant cells responding to environmental stresses,the osmotic stress included, display rapid transient elevationsin cytosolic free Ca
2+
concentration (Knight
et al.
, 1991,1998; Bush, 1995; Tazawa
et al.
, 1995; Taylor
et al.
, 1996;Takahashi
et al.
, 1997; Cessna
et al.
, 1998). It has been foundthat these calcium signals finally lead to the increased expressionof stress-responsive genes, including those encoding proteins ofprotective function (Knight
et al.
, 1996, 1997). It is generally
accepted that the cytosolic Ca
2+
controlling calmodulin activityis directly involved in the organization of the microtubule (Mt)cytoskeleton (Cyr, 1991; Fisher & Cyr, 1993; Zielinski, 1998).
Considering the above information it seems reasonable tosuggest that osmotic stress may cause extensive changes ofMt organization in plant cells. Examination of the availableliterature reveals that the existing knowledge on this topicis limited. Roberts
et al
. (1985) reported that epidermalcells of
Pisum sativum
treated with a solution of sucrose at aconcentration of > 0.23 M undergo cortical Mt reorganiza-tion. Bartolo & Carter (1991) found that exposure of leafsections of
Spinacia oleracea
to solutions of sorbitol or poly-ethylene glycol (PEG) caused Mt depolymerization. Blancaflor
NPH033.fm Page 193 Wednesday, December 20, 2000 2:55 PM
to a 350mOsm nonanionic (sorbitol) and KCl osmotic stress, which doesnot induce plasmolysis. The sorbitol treatment disturbedthe organization of the cortical Mts. Slaninova
et al
. (2000)found that transfer of exponentially-growing cells of theyeast
Saccharomyces cerevisiae
to hyperosmotic growth mediumcaused a temporary disassembly of Mts and actin filaments.Hyperosmotic treatment of dividing PtK-1 animal cells intissue culture, in a medium enriched with 0.5 M sucrose, inducesdistinct changes in spindle organization and function. In fewminutes, Mts form highly birefringent bundles consisting oftightly packed Mts (Snyder
et al.
, 1984; Mullins
et al.
, 1985;Snyder, 1988; Mullins & Snyder, 1989).
In the present study we examined the effects of nonionichyperosmotic treatment on the organization of the successiveMt systems appearing in dividing leaf cells of
Chlorophytoncomosum
. This species was selected because the meristem-atic leaf cells are vacuolated and the structural effects of thehyperosmotic treatment are very obvious. In particular, wetreated leaf segments, including the leaf meristem, with anaqueous solution of 1 M mannitol. Under these conditionswater is flowing out of the vacuole, the cell volume is reducedand the protoplast is mechanically removed from the cell wall.Severe changes on Mt organization and tubulin poly-merization, induced by this treatment, were examined withimmunofluorescence and transmission electron microscopy(TEM).
Materials and methods
Treatment
Segments 1-mm-long taken from the leaf base of
Chlorophytoncomosum
Thunb. were immersed in 1 M aqueous solution ofmannitol for 15 –30 min. Some of the samples were broughtback to distilled water to confirm that hyperosmotic shockwas not lethal. Subsequently, leaf segments were processed fortubulin immunolocalization and electron microscopy. Thecourse of plasmolysis as well as the ability of cells to recoverwas monitored with differential interference contrast (DIC)optics on fresh sections.
Preparation of leaf segments for immunofluorescence
Leaf segments were fixed for 90 min in 8% paraformaldehydein microtubule-stabilizing buffer (MSB; 50 mM PIPES, 5 mMMgSO
4
·7H
2
O and 5 mM EGTA, pH 6.8) at room temperature.After thorough washing with MSB, cell walls were digestedwith 1% pectinase (Fluka), 1% cellulase Onozuka R10 (YakultHonsha), 1% cellulysin (Calbiochem) and 1% B-glucuronidase(Sigma) in MSB pH 5.6 for 90 min. Following washing withMSB, cells were separated by gently forcing leaf segmentsthrough a Pasteur pipette.
The resulting cell suspension was layered onto poly L-lysine
coated acid-washed coverslips, excess liquid blotted, and leftto air-dry. Cells were then extracted with 1% Triton X-100 inphosphate-buffered saline (PBS) for 15 min. Afterwards, speci-mens were subsequently incubated with a rat monoclonalanti-
α
tubulin antibody (clone Yol 1/34, Sera-Lab Ltd,Sussex, England), diluted 1 : 80 in PBS containing 1% BSA,followed by a FITC-conjugated anti-rat IgG (Sigma) diluted1 : 80 in the same buffer. Chromatin was counter-stained withHoechst 33258 at 1
µ
g/ml in PBS and the specimens werefinally mounted in 90% glycerol solution in PBS, pH 7.8containing 0.1% p-phenylenediamine. The specimens wereobserved with a Zeiss Axioplan light microscope equippedwith epifluorescence illumination, standard UV, FITC andrhodamine filter set and Neofluar objectives. Photomicro-graphs were captured on Kodak T-MAX 400 pushed at1600 ASA.
Preparation of leaf segments for electron microscopy
After treatment with hypertonic mannitol solution, leaf seg-ments were fixed with 3% glutaraldehyde and 1% tannic acidin 100 mM cacodylate buffer pH 7.2 for 3 h at roomtemperature. Following washing with 100 mM cacodylatebuffer, segments were postfixed with 1% OsO
4
in 100 mMcacodylate buffer for 5 h at 4
°
C. Subsequently, segmentswere dehydrated in a graded series of acetone followed bytwo changes in anhydrous propylene oxide. Specimens werefinally infiltrated and embedded in Spurr’s resin and sectionedfor TEM. Semithin sections were stained with 1% (w/v)toluidine blue in 1% (w/v) aqueous borax solution. Thinsections were stained with uranyl acetate and lead citrate andexamined with a Philips 300 EM.
Results
General remarks
The meristematic leaf cells of
Chlorophyton comosum
areconfined to a narrow basal zone. They contain the usualorganelle complement and a rather well developed vacuolarsystem (Fig. 1a). After treatment with 1 M aqueous mannitolsolution for 15–30 min these cells became plasmolysed.Their protoplast became almost completely separated fromthe cell wall, while their volume was drastically reduced(Fig. 1b). This decrease in volume was primarily due to theloss of water from the vacuoles. The hyperosmotic treatmentinduced profound changes in chromatin organization. Chromatinwas rapidly transformed into a more or less compact massand in no case individual chromosomes could be distinguishedin mitotic cells after Hoechst staining of the immunofluorescencespecimens and toluidine blue staining of semithin sections (seealso Snyder
et al.
, 1984).Examination of thin sections with TEM showed clearly
the diminution of the vacuoles and the disruption of the
NPH033.fm Page 194 Wednesday, December 20, 2000 2:55 PM
plasmodesmata. The change in chromatin organization ob-served in plasmolysed cells, after Hoechst staining, waseven more evident in thin sections. The affected nuclei weresmaller and contained much more condensed chromatin(Fig. 1b; cf. Fig. 1a). It seems likely that most, if not all, of
the chromatin had been changed into some form of ‘con-densed’ chromatin.
Apart from these nuclear changes, the mode and the extentof tubulin polymerization were also dramatically altered, asassessed by tubulin immunofluorescence and TEM describedbelow.
Interphase and preprophase cells
In the vast majority of control interphase cells Mts were foundin the cell cortex, while their orientation was predominantlyperpendicular to the main cell axis (Fig. 2a). Mts in the endo-plasm were, mainly in the perinuclear. Control cells, withcondensed chromatin (Fig. 2b
3
), exhibited typical preprophaseMt band (ppb) as well as perinuclear Mts (Fig. 2b
1
, b
2
).Examination of a large number of plasmolysed inter-
phase cells, in more than 12 experiments after tubulin immuno-labelling, showed that nearly all the Mts had disintegrated. Infew cells some cortical Mts persisted, but most of the inter-phase cells exhibited another type of tubulin polymer. Thisappeared in the form of fluorescing spots (Fig. 2c) or as anextensive and, in some cases, continuous network of anastom-osing strands (Fig. 2d
1
, d
2
). The latter fluoresced intenselyand ran through the cytoplasm in various directions. In someplaces these tubulin polymers approached the plasmalemmaand the nucleus (Fig. 2c,d). PPBs were also absent from mostof the preprophase-prophase plasmolysed cells. Very fewsmall cells displayed a ppb (Fig. 2e
1
). In these the protoplastremained attached to the cell wall at the ppb region (Fig. 2e
2
).Since ppbs were common in the control tissues it is clear thatplasmolysis was causing their disassembly, but more slowlythan the interphase cortical Mts. These findings show clearlythat these Mt arrays are extremely sensitive to the hypero-smotic shock.
Examination of the interphase plasmolysed cells with TEMconfirmed the absence of cortical Mts. In rare cases few Mtspersisted in regions where the plasmalemma remained attachedto the cell wall. However, the plasmolysed cells displayed sev-eral profiles of elongated electron-dense structures (Fig. 3a).They were localized at different cytoplasmic sites and showedpreferential associations with lipid bodies and particularlyvacuoles (Fig. 3a,b). The latter association was also obvious inimmunofluorescence specimens examined with DIC optics.In higher magnification these structures seem to consist of finesheets or filaments (Fig. 3a), displayed a 24-nm periodicity.Therefore, they represent sections of elongated anastomosingprotein-paracrystals (Fig. 3a). Fig. 3b shows the filament-ous substructure of the paracrystals. The number, distribu-tion and thickness of the paracrystals indicates that theyare almost certainly the tubulin polymers observed in theimmunolabeled plasmolysed interphase cells. This view isfurther supported by the fact that similar paracrystals appearin contact with the chromatin in plasmolysed mitotic cells(see next section).
Fig. 1 (a) Median paradermal view of a protodermal area taken from a control leaf of Chlorophyton comosum. (b) Median longitudinal section of a plasmolysed interphase mesophyll cell. The nucleus (N) displays obviously more condensed chromatin than those of the control cells (see Fig. 1a). Abbreviations: V, vacuole. Bars, 5 µm.
NPH033.fm Page 195 Wednesday, December 20, 2000 2:55 PM
Control prophase cells display mature ppb and bipolarprophase spindle (Fig. 4a). At late prophase/prometaphase,the ppb disappears and the nuclear envelope breaks down. Themetaphase and anaphase spindles show the usual organization.The metaphase spindle consists mainly of kinetochore-Mtbundles, which converge on broad polar areas (Fig. 4b). Pole-to-pole Mts can also be seen. The anaphase spindles display shortkinetochore-Mt bundles as well as pole to pole Mt bundles(Fig. 4c,d).
In plasmolysed cells, due to the chromatin change, it wasnot always possible to identify unequivocally all the stages inthe cell cycle. For instance, interphase and preprophase/prophasecells could not be distinguished at both light microscope andTEM. Therefore, the results described below are not exactlycell cycle specific, except in cases, where the organization ofthe affected chromatin resembels the control, for examplethe metaphase and anaphase chromosome arrangements.
Telophase and cytokinesis are also obviously discrete. Consider-ing the relatively short time of plasmolysis (15 –30 min), themitotic examined cells must have entered mitosis before theonset of this treatment.
In the plasmolysed prophase-like cells, the chromatin appearedas a compact mass with a smooth outline after Hoechst staining,an observation indicating that nuclear envelope was still intact(Fig. 4e inset, h inset). In these cells polymerized tubulin wasmainly found in the form of thick and intensely fluorescingstrands arranged in a bipolar spindle-like manner around thenucleus (Fig. 4e,g,h). These tubulin strands were straight,more rigid and more prominent than those at interphase(Fig. 4e,g,h; cf. Figure 2d), being visible even under conven-tional DIC optics without contrast enhancement (Fig. 4e
3
).The ‘spindles’ formed by these strands often had focal contactswith the plasmalemma (Fig. 4f ). In some prophase-like cellsindividual tubulin strands traversed the perinuclear cytoplasmin an uninterrupted pole-to-pole fashion, converged on singlepointed foci and were interconnected in a meshwork by fine
Fig. 2 Tubulin immunolabelling in control (a, b) and plasmolysed (c –e) cells of Chlorophyton comosum. (a) Cortical Mts of an interphase cell of Chlorophyton comosum. (b) Preprophase/prophase cell in surface (b1) and median (b2) optical section. Note the dominant presence of a mature ppb and of the perinuclear Mts. (b3) The nucleus after Hoechst staining. (c) Plasmolysed interphase cells. Tubulin polymers, appearing as fluorescing spots, are dispersed in the cytoplasm. Some of them are localized on the surface of the nucleus (N). (d) Surface (d1) and median (d2) optical section of an interphase plasmolysed, probably, protodermal cell. Tubulin strands traverse the cortical cytoplasm as well the endoplasm. (e) (e1) Preprophase plasmolysed cell, the ppb of which has not been disorganized. Inset. The nucleus after Hoechst staining. (e2) The plasmolysed cell viewed by DIC optics. The arrow points to the ppb region, which has not been separated from the cell wall. Bar, 10 µm.
NPH033.fm Page 196 Wednesday, December 20, 2000 2:55 PM
). In other prophase-like cells, tubulinstrands were arranged in two subsets, forming a spindle-likestructure, which also displayed a clear bipolarity (Fig. 4g).The strands of each subset converged on the pole region onone side, while on the other they terminated on the nuclearsurface. The ‘half spindles’ sometimes appeared intercon-
nected by thinner tubulin strands (Fig. 4h). The nucleus of thecell shown in Fig. 4(i), is probably at a prometaphase-likestage and exhibits associations with short tubulin strands.Some of these are localized on the surface of the nucleus, whileothers enter or travers the chromatin (Fig. 4i
1
, i
2
). Thesetubulin strands do not form a bipolar system (Fig. 4i; cf.Figure 4e,g,h).
In cells in which the nuclear envelope had broken down,the chromosomes appeared as a lobed mass of compactchromatin (Fig. 5 insets). In many of these metaphase-likecells, tubulin strands, similar to those observed in prophase-like cells, appeared to interact with chromatin at variouspoints. These tubulin strands formed ‘spindle’ structureswith pointed poles (Fig. 5a,b,c). The metaphase-like ‘spindles’appeared elongated compared to the controls and consisted oftwo half-spindles (Fig. 5a,b; cf. Fig. 4b). In many cases theyappeared anchored on the plasmalemma (Fig. 5a
2
). Tubulinstrands diverging from the ‘spindle’ (Fig. 5c) or embeddedwithin the chromatin mass were also visible in metaphase-like cells.
An early anaphase-like ‘spindle’ in a plasmolysed cell isillustrated in Fig. 5d. This shows chromosome-attached tubulinstrands converging on the poles and some tubulin strands inthe interzonal region. The advanced anaphase-like ‘spindles’in plasmolysed cells exhibited tubulin strands at the poles aswell in the interzonal area, which appeared in contact with thedaughter chromosome groups (Fig. 5e).
An exceptional feature in many plasmolysed mitotic cells wasthe presence of tubulin polymers in cytoplasmic sites distantfrom the ‘spindle’. The ectopic tubulin polymers of prophase(Fig. 5f ), metaphase (Fig. 5g,h) and anaphase plasmolysedcells appeared similar to the tubulin strands found in theplasmolysed interphase cells. Although these were most abund-ant in the cytoplasm away from the spindle (Fig. 5f,g,h), theywere also seen approaching or entering it (Fig. 5f,g, h
;cf. Fig. 5f,g). A few plasmolysed mitotic cells display onlyectopic tubulin polymers. Some of these appeared to con-tact the chromatin mass but did not form a spindle-like struc-ture (Fig. 5i
1
, i
2
).Longitudinal and transverse sections of more than 10
metaphase-like plasmolysed cells were examined with TEM.They were characterized by the presence of a compact chro-matin mass (Fig. 6a,b). In some cases remnants of nuclearenvelope remained attached to the chromatin. This examina-tion revealed that the metaphase-like ‘spindles’, observed inthe immunofluorescence specimens, consisted of bundles ofmacrotubules or of macrotubules and tubulin paracrystals(Fig. 6c–f ). Both the macrotubule bundles and the paracrys-tals entered and/or traversed the chromatin mass (Fig. 6c,d).The macrotubules had a diameter varying between 24 and40 nm with a mean value 32 nm. These values were derivedfrom measurement of 230 macrotubules. In this population,
Fig. 3 Electron micrographs of tubulin paracrystals found in plasmolysed interphase cells of Chlorophyton comosum. (a) Anastomozing paracrystals. Periodicity is clearly evident. Abbreviations: L, lipid body; V, vacuole. (b) Transverse section of paracrystals. They display locally a filamentous substructure (arrows). V, vacuole. Bars, 200 nm.
NPH033.fm Page 197 Wednesday, December 20, 2000 2:55 PM
14% of macrotubules displayed a diameter 24 –28 nm, 70%28 –36 nm and the remaining 16% 36 – 40 nm. The meanvalue of Mt diameter in control cells was 23 nm. The macro-tubule and microtubule diameter was measured by meansof a grid covered by a grating replica (Polysciences, Ltd).The macrotubules usually form well-organized bundles(Fig. 6e,f ). Cross bridges between the macrotubules withinthe bundles were observed (Fig. 6f ). A few of the macrotubuleswere decorated by tubulin hooks (Fig. 6f ). In addition, singleor small groups of macrotubules were seen embedded in thechromatin mass (Fig. 6d inset).
The strands of paracrystalline tubulin in the metaphase-
like ‘spindle’ had a structure similar to that of the interphaseparacrystals (Fig. 6d; cf. Figure 3a,b). They displayed a 24-nmperiodicity and seemed to consist of tubulin sheets or fila-ments. In anaphase-like plasmolysed cells the daughter chro-matin masses were distinct in thin sections (Fig. 7a). TypicalMts were not observed in these cells. The interzonal regionwas traversed by bundles of macrotubules and/or tubulinparacrystals (Fig. 7b,c). In thin sections it was also confirmedthat the metaphase-and anaphase-like plasmolysed cells pos-sessed an extensive network of ectopic tubulin paracrystalsrunning through the cytoplasm in many directions. Theywere most frequently in the polar regions.
Fig. 4 Tubulin immunolabelling in dividing control (a–d) and plasmolysed (e–i) cells of Chlorophyton comosum. (a) Prophase spindle. (b) Metaphase spindle. Inset. The chromosomes after Hoechst staining. (c) The spindle at the beginning of anaphase. (d) Advanced anaphase spindle. Inset. The daughter groups of chromosomes after Hoechst staining. (e) (e1, e2) Successive optical sections of a prophase-like plasmolysed cell. Intensely fluorescing perinuclear tubulin strands form a spindle-like structure displaying pointed poles. Inset. The nucleus after Hoechst staining. (e3) The perinulear tubulin strands (arrows) viewed by DIC optics. (f) Prophase-like plasmolysed cell viewed by DIC optics. One of the poles of the ‘mitotic spindle’ appears to be in contact with the plasmalemma (arrow). (g, h) Prophase-like plasmolysed cells, displaying a ‘mitotic spindle’ consisting of two half-spindles. Inset in h. The nucleus after Hoechst staining. (i) Optical sections through surface (i1) and median (i2) planes of a nucleus in a prometaphase plasmolysed cell. Tubulin strands line the nuclear surface and intervene among the chromosomes. Inset. The prometaphase nucleus after Hoechst staining. Bar, 10 µm.
NPH033.fm Page 198 Wednesday, December 20, 2000 2:55 PM
In control late anaphase-early telophase cells, an interzonalMt system was localized between the daughter chromosomegroups. It consisted of two sets of interdigitating Mts (Fig. 8a).Progressively, Mt length was reduced and a barrel-shapedphragmoplast was organized between the daughter nuclei(Fig. 8b). The cell plate developed along the plane where theMts were overlapping. The phragmoplast-cell plate system
gradually expanded towards the cell periphery, where the cellplate fused with the parental wall.
In late anaphase-early telophase plasmolysed cells intenselyfluorescing tubulin strands traversed the daughter chromo-some ‘masses’. They were directed from the latter towards theinterzonal region, forming two independent systems (Fig. 8c
1
).In late telophase cells, a dominant system of tubulin strandswas found in the interzonal region. It consisted of continu-ous and densely arranged strands traversing the whole
Fig. 5 Tubulin immunolabelled plasmolysed mitotic cells of Chlorophyton comosum. (a1, b, c) ‘Mitotic spindles’ in metaphase-like cells. They exhibit pointed poles. In the cell shown in c some tubulin strands seem to diverge from the‘spindle’ area towards cell periphery. Insets. The chromosomes after Hoechst staining. (a2) The cell shown in a1 viewed by DIC optics. (d,e) Early (d) and advanced (e) anaphase-like plasmolysed cells. Insets. The chromosomes after Hoechst staining. (f, g) Prophase-like (f) and metaphase-like (g) plasmolysed cells displaying tubulin polymers far from the ‘spindle’. Inset in g. The chromosomes as they appear after Hoechst staining. (h) Surface (h1) and median (h2) optical sections of a metaphase-like cell. Numerous tubulin polymers are localized outside the ‘spindle’ region. As it can be observed in h2 the latter is not well organized. Inset in h2. The chromosomes after Hoechst staining. (i) Successive optical sections of a metaphase-like plasmolysed cell. Intensely fluorescing tubulin strands not forming a ’spindle’ can be observed. Inset in i2. The chromosomes after Hoechst staining. Bar, 10 µm.
NPH033.fm Page 199 Wednesday, December 20, 2000 2:55 PM
Fig. 6 Electron micrographs of plasmolysed mitotic cells of Chlorophyton comosum. (a, b) Mitotic cells in which the ‘spindle’ has been sectioned longitudinally (a) and transversely (b). Note the compactness of the chromatin. Chromosomes cannot be distinguished. (a) Bar, 5 µm (b) Bar, 3 µm. (c) Region of the chromatin of the cell shown in (a) in higher magnification. The arrows point to macrotubules, while the arrowhead to a tubulin paracrystal. Bar, 200 nm. (d) Region of the chromatin of the cell shown in (b) in higher magnification. The arrows indicate tubulin paracrystals running through the chromatin mass. Bar, 200 nm. Inset. A pair of macrotubules embedded in the chromatin mass. Bar, 200 nm. (e) Transverse section of macrotubule bundles taken from a mitotic plasmolysed cell. Bar, 200 nm. (f) Higher magnification of macrotubules. The arrow marks a macrotubule decorated by a tubulin hook, the arrowhead indicates macrotubules interconnected by electron-dense bridges. Bar, 100 nm.
NPH033.fm Page 200 Wednesday, December 20, 2000 2:55 PM
distance between the daughter nuclei (Fig. 8d, h2). In somecells a few tubulin strands were found around the nuclei(Fig. 8d, h2).
TEM examination of telophase cells revealed that the tubulinstrands were made of macrotubules arranged into thick bundles(see Fig. 7c). Paracrystals traversing the space between thedaughter nuclei were also observed. Macrotubule bundles ortubulin paracrystals were often localized on the polar surfaceof the daughter nuclei.
In cytokinetic cells, displaying cell plates (Fig. 8e2), elongatedtubulin strands were found running through the cytoplasmbetween the daughter nuclei, but they did not show the struc-ture of a phragmoplast (Fig. 8e1; see also Fig. 8f1, g). Theywere continuous and traversed the whole distance between thedaughter nuclei (Fig. 8e,g) or were confined to the marginsof the cell plate (Fig. 8f ). Examination of cytokinetic cellswith TEM (Fig. 9a) revealed that the tubulin strands repres-ented well-organized bundles of macrotubules and tubulinparacrystals (Fig. 9b). In contrast to phragmoplast, theyusually formed a continuous system of tubulin polymers(Fig. 8e,g; cf. Figure 8b). Ectopic tubulin polymers were alsofound in telophase and cytokinetic plasmolysed cells, most
abundantly in the cell cortex (Fig. 8g, h1). In these cells, thoseshowing numerous ectopic tubulin polymers displayedpoorly organized atypical cytokinetic apparatuses. There alsooccurred plasmolysed telophase/cytokinetic cells, withoutan atypical cytokinetic apparatus, displaying only ectopictubulin polymers (Fig. 8i1–i3). In these cells a cell plate wasnot observed between the daughter nuclei with a DIC opticalsystem (Fig. 8i4).
Finally, it should be noted that the study of the plasmolysedinterphase and dividing cells, after tubulin immunolabelingas well as with TEM gave the impression that much moretubulin was in polymeric form than in control cells.
Discussion
General remarks
The results presented in this study revealed for the first timethat a short hyperosmotic nonionic treatment disintegrates allMt arrays in dividing cells of Chlorophyton comosum. Duringthis treatment the protoplast, decreasing in volume, becomesseverely separated from the cell wall. In the non-detached
Fig. 7 (a) Late-anaphase plasmolysed cell of Chlorophyton comosum. Note the organization of the daughter chromosome groups. Bar, 3 µm. (b, c) Tubulin paracrystals (b) and macrotubule bundles (c) taken from the interzonal region of the cell shown in a. (b) Bar, 200 nm (c) Bar, 200 nm.
NPH033.fm Page 201 Wednesday, December 20, 2000 2:55 PM
regions of the plasmalemma, cortical Mts sometimes persist.This phenomenon is more obvious in some preprophase cells,in which the ppb persists. In the ppb cortical region theplasmalemma is not separated from the cell wall. A similar
phenomenon has been observed in plasmolysed fern protonemalcells (Kagawa et al., 1992).
An unexpected finding of this work is that in interphasecells, free tubulin either liberated from the depolymerized
Fig. 8 Tubulin immunolabelled control (a, b) and plasmolysed (c–i) telophase/cytokinetic cells of Chlorophyton comosum. (a) Interzonal Mt system in early telophase control cell. (b) Phragmoplast. Inset. The daughter nuclei after Hoechst staining. (c, d) Early (c) and advanced (d) telophase plasmolysed cells. Intensely fluorescing tubulin strands traverse the interzonal region. They form two independent systems (c1) or a united system (d). (c2) The daughter chromosome groups after Hoechst staining. (e) Cytokinetic plasmolysed cell. The space between the daughter nuclei is traversed by tubulin strands, which do not form a phragmoplast (e1). Examination of the cell by DIC optics (e2) reveals the existence of a cell plate (arrow in e2). (f) Plasmolysed cytokinetic cells (f1) Atypical tubulin strands are localized at the margins of the cell plate. (f2) The daughter nuclei after Hoechst staining. (g) Telophase/cytokinetic cell. Tubulin polymers are localized in the interzonal region as well as in other cell sites. (h) Optical sections through a surface (h1) and a median plane (h2) of a plasmolysed telophase cell. Tubulin fluorescence is observed in the interzonal region (h2), on the surface of the daughter nuclei (h2) as well as in many other cortical cytoplasmic sites (h1). (i) (i1–i3) Successive optical sections of a cytokinetic plasmolysed cell. The entire cytoplasm is characterized by the presence of tubulin polymers,which do not form a phragmoplast. (i4) The cell viewed by DIC optics. A cell plate can not observed between the daughter nuclei (compare with e2). Bar, 10 µm.
NPH033.fm Page 202 Wednesday, December 20, 2000 2:55 PM
Mts or preexisting as a nonassembled pool, is incorporatedinto a network of paracrystals. In most of the dividing cells,mitotic and cytokinetic Mt systems are replaced by atypicalspindle-like structures displaying intense bipolarity and atypicalphragmoplasts, respectively. They consist of macrotubule
bundles or macrotubules and tubulin paracrystals. Moreover,in dividing cells tubulin paracrystals were found in ectopicpositions far from the atypical mitotic spindles and atypicalphragmoplasts. Some plasmolysed dividing cells possessedonly tubulin paracrystals, which did not form atypical mitotic orcytokinetic systems.
The view that the paracrystals in the plasmolysed cellsconsist of, or at least contain tubulin, is supported by thefollowing observations: (a) The paracrystals are localized influorescence microscope after tubulin immunolabelling. (b) Inmitotic cells they appear connected with as well as embeddedin the chromatin mass. (c) They display structural similaritiesto tubulin paracrystals formed in cells treated with anti-Mtdrugs (Figs 3a and 6d; cf. Figures 4 and 5 in Bensch &Malawista, 1969, Figs 2F-2K in Starling, 1976 and Figs 11and 13 in Apostolakos et al., 1990). As far as we know thisis the first time in which tubulin paracrystal formation hasbeen induced in plant cells by a non-chemical treatment. Suchtubulin polymers have been observed in plant cells after treat-ment with anti-Mt drugs (Apostolakos et al., 1990; Karagian-nidou et al., 1995; and literature therein).
Microtubule disorganization
The sensitivity of Mts to osmotic stress seems to be a generalphenomenon. Apart from Chlorophyton comosum (presentstudy) changes in Mt organization have been observed: (a) Inspinach mesophyll cells that underwent a sorbitol or PEGtreatment (Bartolo & Carter, 1991). (b) In root cells of Zeamays treated with sorbitol, PEG or sucrose (Blancaflor &Hasenstein, 1995). (c) In cells of Saccharomyces cerevisiae grow-ing in a medium containing 0.7–1 M KCl, 1 M mannitoland/or 1 M glycerol (Slaninova et al. 2000). (d) In withdrawingreticulopodia of the protozoan Allogromia induced by thewithdrawal stimulus (seawater substitute made hypertonicwith MgCl2) (Rupp et al., 1986; Welnhofer & Travis, 1996).
Concerning the mechanism(s) of Mt depolymerization inplasmolysed cells examined two possible hypotheses cometo mind, although definite conclusions cannot be drawn.According to the first hypothesis, mechanical tensions exertedon the plasmalemma during plasmolysis may induce dir-ectly cortical Mt cytoskeleton disassembly. The persistenceof cortical Mts in the nondetached regions of the plas-malemma in plasmolysed cells, and their disappearancefrom the separated plasmalemma regions supports thishypothesis.
The sensitivity of the Mt cytoskeleton to mechanical stressis fairly well documented. The application of external pressureon plant cells lead to depolymerization (Cleary & Hardham,1990 and literature therein) or disorganization (Hush &Overall, 1991; Cleary & Hardham, 1993; Zandomeni &Schopfer, 1994; Wymer et al., 1996) of the Mt cytoskeleton.The imposition of mechanical stresses also disturbs theorganization of cytoskeleton in animal cells (Ingber, 1997).
Fig. 9 (a) Electron micrograph of a median section of a cytokinetic plasmolysed cell of Chlorophyton comosum. CP: cell plate. Bar, 5 µm. (b) The cytoplasmic region marked by the arrow in a in higher magnification. Many macrotubules and a paracrystal (arrow) can be observed. Bar, 200 nm.
NPH033.fm Page 203 Wednesday, December 20, 2000 2:55 PM
It has been suggested that the plant cytoskeleton particip-ates in the functional continuum with the plasmalemmaand the cell wall intimately involved in cell morphogenesis.Plasmalemmal proteins probably connect the cytoskeletalelements with particular cell wall elements (Williamson,1991; Cyr, 1994; Kropf, 1994; Shibaoka, 1994; Fowler &Quatrano, 1997). The disturbance of the cell wall-plasmalemmafunctional continuum in plasmolysed cells may induce Mtdepolymerization.
According to the second hypothesis, which in our opin-ion is the more plausible, Mt disintegration in plasmolysedcells may be induced by rapid and profound changes in cellmetabolism triggered by the hyperosmotic shock. It is welldocumented that the plant cells responding to osmotic stressdisplay rapid transient elevations in cytosolic Ca2+ concentra-tion (Knight et al., 1991, 1997, 1998; Brownlee et al., 1999).Mt polymerization/depolymerization is controlled, amongothers, via changes of Ca2+ concentration (Cyr, 1991; Fisher& Cyr, 1993; Fisher et al., 1996; O’Brien et al., 1997). Theincrease of cytosolic Ca2+ in plasmolysed cells (Cessna et al.,1998; Knight et al., 1998) may induce Mt disorganization byseveral different mechanisms, e.g. by MAP activation throughCa2+/calmodulin complex, which induces Mt disassembly(Fisher et al., 1996) or by increase of the rate of GTP hydrol-ysis within the GTP cap end of the Mts (O’Brien et al., 1997).The persistence of some cortical Mts in nonplasmolysedregions of the cells in our material suggest that interconnec-tions between Mts-plasmalemma-cell wall protect Mts fromthe destabilizing effect of the elevated cytosolic Ca2+ (see alsoShibaoka, 1994).
Furthermore, recent studies have revealed that a transmem-brane hybrid-type histidine kinase in Arabidopsis functionsas an osmosensor and transmits a stress signal to a down-stream mitogen-activated protein kinase cascade (Urao et al.,1999), as happens in yeast (for literature see Brownlee et al.,1999). The involvement of a mitogen-activated protein kinasecascade in osmotic stress response in higher plants has alsobeen suggested previously (Popping et al., 1996; Mizoguchiet al., 1997; Hirt, 2000). In addition, the hyperosmotic stressinduces rapid synthesis of a polyphospoinositide-like lipidin several plant species (Meijer et al., 1999), which is part ofthe inositol signalling system. These plant responses toosmotic stress, associated with changes in cytosolic Ca2+
concentration, may disturb phosphorylation/dephosphoryla-tion, processes which control of the assembly/disassemblyof Mts (for literature see Desai & Mitchison, 1997; Quader,1998; Vaughn & Harper, 1998; Bögre et al., 2000). Thiscould easily result in Mt disorganization in plasmolysedcells.
Macrotubule and tubulin paracrystal assembly
The induction of macrotubule formation in cells after ahyperosmotic stress is described for the first time here. In
addition, measurements, from published micrographs, of thediameters of the Mts in the elongated spindles in Pt K-1 cells,induced by a hyperosmotic treatment (Figs 4 in Snyder et al.,1984, Figs 5 and 7 in Snyder, 1988), reveal that these also maybe macrotubules. If this is true, the animal cells may respondto a hyperosmotic shock in a similar manner to plants.
Macrotubules have been repeatedy observed in specializedanimal cells or are assembled in vitro under specific condi-tions (for literature see Unger et al., 1990; Vater et al., 1997).Macrotubules display more than 13 protofilaments. Forinstance those in wing epidermal cells of Drosophila have adiameter about 30 nm, and are made of 15 protofilaments(Tucker et al., 1986; Mogensen & Tucker, 1987). In plantcells macrotubules have been observed in oil-body cells ofthe liverwort Marchantia paleacea (Galatis & Apostolakos,1976; Apostolakos & Galatis, 1998) and in root-tip cellsof Triticum turgidum treated with aluminium (Frantzioset al., 2000).
The formation of tubulin paracrystals in plasmolysed cellsof Chlorophyton comosum was unexpected. It is noteworthythat Mt cytoskeleton in withdrawing reticulopodia of the pro-tozoan Allogromia is transformed into a tubulin-containingparacrystal made of helical elements. Although paracrystalformation in Allogromia is induced by hyperosmotic stress(Rupp et al., 1986; Welnhofer & Travis, 1996), and by variousphysical stimuli, it may also be seen sometimes in normal con-ditions (Rupp et al., 1986 and literature therein). Moreover,in Ptk-1 cells, cultured in presence of 0.5 M sucrose, straight,short fluorescing strands have been observed after tubulinimmunolabelling (see Figs 4, 5 and 8 in Mullins & Snyder,1989). These are comparable to the tubulin strands observed inthe plasmolysed cells of Chlorophyton comosum and thereforethey may represent tubulin paracrystals. These data suggestthat tubulin paracrystal formation may be a rather generalresponse of animal and plant cells to hyperosmotic stress.
What then are the conditions favouring macrotubule andtubulin paracrystal assembly in plasmolysed cells of Chloro-phyton comosum? In vitro studies show that macrotubule ortubulin paracrystal formation is a phenomenon determinedby the mode of interaction between tubulin protofilaments,which consequently is controlled by Ca2+ concentration. Thekind of tubulin assemblies formed, essentially seems to dependon the action of Ca2+ during tubulin polymerization (Vateret al., 1997).
Based on the above information it may be suggested thatin Chlorophyton the concentration of Ca2+ formed duringthe initial stages of the hyperosmotic shock favours Mt dis-assembly. Later the Ca2+ concentration changes and the newconditions favour macrotubule and tubulin paracrystal forma-tion. Recent data have shown that the Ca2+ concentration changesduring the osmotic shock (Takahashi et al., 1997; Cessnaet al., 1998), a fact supporting the above hypothesis. It hasbeen also reported that Mt disorganization induces the act-ivation of molecular Ca2+ pumps in the plasmalemma, the
NPH033.fm Page 204 Wednesday, December 20, 2000 2:55 PM
tonoplast and other biomembranes which results in the sig-nificant increase of cytosolic Ca2+ in cells lacking cytoskeletalelements (Mazars et al., 1997).
Therefore, it may be suggested that, in plasmolysed cells ofChlorophyton comosum, cytosolic Ca2+ concentration changesfurther after Mt disassembly. In some cells these changes mayresult in the establishment of conditions favouring macro-tubule and in others tubulin paracrystal assembly. Moreover,the establishment of different Ca2+ concentrations in differentcell regions may explain the formation of both paracrystalsand macrotubules in the same cell. The consistent relation-ship between tubulin paracrystals and vacuoles may supportthe above hypothesis. The vacuoles are Ca2+-stores while inthe tonoplast there are voltage and ligand-activated calciumchannels (for literature see Mazars et al., 1997). These chan-nels may be activated by the osmotic stress and create onthe tonoplast surface Ca2+ concentration favouring tubulinparacrystal assembly.
Organization of the atypical tubulin polymers in plasmolysed cells
The pattern of organization of the atypical tubulin polymersassembled in the plasmolysed cells of Chlorophyton comosumdepends on the stage in the cell cycle when they are exposedto the hyperosmotic solution. For example in most of themitotic and cytokinetic plasmolysed cells atypical tubulin poly-mers form mitotic-and cytokinetic-like apparatuses, respect-ively. Tubulin polymers also appear in ectopic positions inthe form of paracrystals. The presence of tubulin polymersin regions far from the mitotic spindle has been describedin drug-treated metaphase Ptk-1 cells following hyperosmotictreatment (Mullins & Snyder, 1989). Therefore, in dividingplant and animal cells under particular conditions, tubulinpolymerization may take place in positions far from thoseforming the mitotic and cytokinetic apparatuses, a fact suggest-ing the existence of a large tubulin pool apart from the spindle.It is also important that in plasmolysed mitotic cells macro-tubules and tubulin paracrystals are found in immediatecontact with the chromatin, an observation denoting thattubulin polymerization may take place on the chromatin. Thissupports the hypothesis that in angiosperms the Mts of themetaphase spindle are formed in the immediate vicinity ofthe chromosomes and not at the pole regions (Binarova et al.,2000; Bögre et al., 2000; Zachariadis et al., 2000; and literaturetherein).
A consistent feature of plasmolysed cells is the extremebipolarity attained by mitotic-like spindles expressed by theformation of sharply pointed poles. In normal mitotic animaland plant cells, Mt convergence at the poles is thought tobe an intrinsic property of mitotic Mt arrays, driven by theaction of Mt-associated motor and structural proteins (forliterature see Bögre et al., 2000; Zachariadis et al., 2000).However, in many plant cells, the mechanical hindrance
imposed by chromosome arms arranged between kinetochoreMt bundles, seems to prevent Mt convergence (Zachariadiset al., 2000). This obstacle appears to be eliminated during thehyperosmotic stress where the chromosomes coalesce in asingle mass and in no case were chromosome arms seen toextend between the macrotubule bundles. Macrotubules arethen free to converge, a finding implying that the mechanismunderlying the establishment of the spindle shape is still func-tional. This mechanism does not seem to apply to the case oftubulin paracrystals, which apparently do not display anydistinct organizational pattern.
Regarding the function of the atypical tubulin polymersformed in the plasmolysed cells of Chlorophyton comosum onlyhypotheses can be made. These polymers as well as thoseformed in animal cells after a hyperosmotic stress (Snyderet al., 1984; Mullins et al., 1985; Snyder, 1988) are elongated,straight and rigid structures. They probably offer mechanicalsupport to the protoplast to resist forces exerted on it dur-ing plasmolysis. According to a current view in animal cellsmechanical tensions generated through molecular interactionswithin the cytoskeleton are balanced by external adhesion tothe extracellular matrix. Thus, the cell is in a condition stateof isometric tension. The imposition of external stresses onthese systems, i.e. mechanical stresses, disturbs the organizationof cytoskeleton, in order to restore the equilibrium betweenexternal and internal forces (Ingber, 1997, 1998; Chicurelet al., 1998). It is probable that the changes of the Mtcytoskeleton in plasmolysed cells of Chlorophyton comosum isthe result of the above mechanism.
Note added in proof: Recently Lang–Pauluzzi and Gunning(Protoplasma 212: 174–185, 2000) studied the effects ofplasmolysis on cytoskeletal elements (microtubules and actinmicrofilaments) in onion inner epidermal cells. They showedthat plasmolysis had minor effects on the organization ofcytoskeletal elements. They also demonstrated their occurencewithin Hechtian strands.
Acknowledgements
G. Komis was awarded a scholarship by the State ScholarshipFoundation.
References
Apostolakos P, Galatis B. 1998. Microtubules and gametophyte morphogenesis in the liverwort Marchantia paleacea Bert. In: Bates JW, Ashton SW, Duckett JG, eds. Bryology for the Twenty-First Century. Leeds, UK: Maney & Son Ltd, 205 –221.
Apostolakos P, Galatis B, Katsaros C, Schnepf E. 1990. Tubulin conformation in microtubule-free cells of Vigna sinensis. An immunofluorescent and electron microscopy study. Protoplasma 154: 132 –143.
Bartolo ME, Carter JV. 1991. Microtubules in mesophyll cells of nonacclimated and cold-acclimated spinach: visualization and responses to freezing, low temperature and dehydration. Plant Physiology 97: 175 –181.
NPH033.fm Page 205 Wednesday, December 20, 2000 2:55 PM
Bensch KG, Malawista SE. 1969. Microtubular crystals in mammalian cells. Journal of Cell Biology 40: 95 –107.
Binarova P, Cenklova V, Hause B, Kubatova E, Lysak M, Dolezel J, Bogre L, Draber P. 2000. Nuclear ã-tubulin during acentriolar plant mitosis. Plant Cell 12: 433 – 442.
Blancaflor EB, Hasenstein KH. 1995. Growth and microtubule orientation of Zea mays roots subjected to osmotic stress. International Journal of Plant Science 156: 774 –783.
Bögre L, Calderini O, Merskiene I, Binarova P. 2000. Regulation of cell division and the cytoskeleton by mitogen-activated protein kinases in higher plants. In: Hirt H, ed. MAP Kinases in Plant Signal Transduction. Berlin, Germany: Springer-Verlag, 95 –117.
Brownlee C, Goddard H, Hetherington AM, Peake L-A. 1999. Specificity and integration of responses: Ca2+ as a signal in polarity and osmotic regulation. Journal of Experimental Botany 50: 1001–1011.
Bush DS. 1995. Calcium regulation in plant cells and its role in signalling. Annual Review of Plant Physiology and Plant Molecular Biology 46: 95 –122.
Cessna SG, Chandra S, Low PS. 1998. Hypo-osmotic shock of tobacco cells stimulates Ca2+ fluxes deriving first from external and then internal Ca2+ stores. Journal of Biological Chemistry 273: 27286 –27291.
Chicurel ME, Chen CS, Ingber DE. 1998. Cellular control lies in the balance of forces. Current Opinion in Cell Biology 10: 232 –239.
Cleary AL, Hardham AR. 1990. Reinstatement of microtubule arrays from cortical nucleating sites in stomatal complexes of Lolium rigidum following depolymerization by oryzalin and high pressure. Plant Cell Physiology 31: 903 –915.
Cleary AL, Hardham AR. 1993. Pressure induced reorientation of cortical microtubules in epidermal cells of Lolium rigidum leaves. Plant Cell Physiology 34: 1003 –1008.
Cyr RJ. 1991. Calcium /calmodulin affects microtubule stability in lysed protoplasts. Journal of Cell Science 100: 311–317.
Cyr RJ. 1994. Microtubules in plant morphogenesis: role of the cortical array. Annual Review of Cell Biology 10: 153 –180.
Desai A, Mitchison TJ. 1997. Microtubule polymerization dynamics. Annual Review of Cell and Developmental Biology 13: 83 –117.
Fisher DD, Cyr RJ. 1993. Calcium levels affect the ability to immunolocalize calmodulin to cortical microtubules. Plant Physiology 103: 543 –551.
Fisher DD, Gilroy S, Cyr RJ. 1996. Evidence for opposing effects of calmodulin on cortical microtubules. Plant Physiology 112: 1979 –1087.
Fowler JE, Quatrano RS. 1997. Plant cell morphogenesis: plasma membrane interactions with the cytoskeleton and cell wall. Annual Review of Cell and Developmental Biology 13: 697 –743.
Frantzios G, Galatis B, Apostolakos P. 2000. Aluminium effects on microtubule organization in dividing root-tip cells of Triticum turgidum. I. Mitotic cells. New Phytologist 145: 211– 224.
Galatis B, Apostolakos P. 1976. Associations between microbodies and a system of cytoplasmic tubules in oil-body cells of Marchantia. Planta 131: 217–222.
Hirt H. 2000. MAP kinases in plant signal transduction. Results and Problems in Cell Differentiation 27: 1–9.
Hush JM, Overall RL. 1991. Electrical and mechanical fields orient cortical microtubules in higher plant tissues. Cell Biology International Reports 15: 551–560.
Ingber DE. 1997. Tensegrity: the architectural basis of cellular mechan-otransduction. Annual Review of Physiology 59: 575 –599.
Ingber DE. 1998. Cellular basis of mechanotransduction. Biological Bulletin 194: 323 –327.
Kagawa T, Kadota A, Wada M. 1992. The junction between the plasma membrane and the cell wall in fern protonemal cells, as visualized after plasmolysis, and its dependence on arrays of cortical microtubules. Protoplasma 170: 186 –190.
Karagiannidou T, Eleptheriou EP, Tsekos I, Galatis B, Apostolakos P. 1995. Colchicine induced paracrystals in root cells of wheat (Triticum aestivum L.). Annals of Botany 76: 23 –30.
Knight H, Brandt S, Knight MR. 1998. A history of stress alters drought calcium signalling pathways in Arabidopsis. Plant Journal 16: 681– 687.
Knight MR, Campbell AK, Smith SM, Trewavas AJ. 1991. Transgenic plant aequorin reports the effects of touch and cold-shock and elicitors on cytoplasmic calcium. Nature 352: 524 –526.
Knight H, Trewavas AJ, Knight MR. 1996. Cold calcium signaling in Arabidopsis involves two cellular pools and a change in calcium signature after acclimation. Plant Cell 8: 489 –503.
Knight H, Trewavas AJ, Knight MR. 1997. Calcium signalling in Arabidopsis thaliana responding to drought and salinity. Plant Journal 12: 1067–1078.
Kropf DL. 1994. Cytoskeletal control of cell polarity in a plant zygote. Developmental Biology 165: 361–371.
Mazars C, Thion L, Thuleau P, Graziana A, Knight MR, Moreau M, Ranjeva R. 1997. Organization of cytoskeleton controls the changes in cytosolic calcium of cold shocked Nicotiana plumbaginifolia protoplasts. Cell Calcium 22: 413 – 420.
Meijer HJG, Divencha N, van den Ende H, Musgrave A, Munnik T. 1999. Hyperosmotic stress induces rapid synthesis of phosphatidyl- D-inositol 3,5-biphosphate in plant cells. Planta 208: 294 –298.
Mizoguchi T, Ichimura K, Shinozaki K. 1997. Environmental stress response in plants: the role of mitogen-activated protein kinases. Trends in Biotechnology 15: 15 –19.
Mogensen MM, Tucker JB. 1987. Evidence for microtubule nucleation at plasma membrane-associated sites in Drosophila. Journal of Cell Science 88: 95 –107.
Mullins JM, Snyder JA. 1989. Induction of random microtubule polymerization in cold and drug-treated PtK1 cells following hyperosmotic shock treatment. European Journal of Cell Biology 49: 149 –153.
Mullins JM, Wolf KM, Snyder JA. 1985. Immunofluorescence analysis of sucrose-induced changes in spindle morphology. European Journal of Cell Biology 39: 333 –340.
O’Brien ET, Salmon ED, Erickson HP. 1997. How calcium causes microtubule depolymerization. Cell Motility and the Cytoskeleton 36: 125 –135.
Popping B, Gibbons T, Watson MD. 1996. The Pisum sativum MAP kinase homologue (PSMAPK) rescues the Saccharomyces cerevisiae HOG1 deletion mutant under conditions of high osmotic stress. Plant Molecular Biology 31: 355 –363.
Quader H. 1998. Cytoskeleton: microtubules. Progress in Botany 59: 374 – 395.
Roberts IN, Lloyd CW, Roberts K. 1985. Ethylene-induced microtubule reorientations: mediation by helical arrays. Planta 164: 439 – 447.
Rupp G, Bowser SS, Mannella CA, Rieder CL. 1986. Naturally occuring tubulin-containing paracrystals in Allogromia: immunocytochemical identification and functional significance. Cell Motility and the Cytoskeleton 6: 363 –375.
Shibaoka H. 1994. Plant hormone-induced changes in the orientation of cortical microtubules: alterations in the cross-linking between microtubules and the plasma membrane. Annual Review of Plant Physiology and Plant Molecular Biology 45: 527–544.
Slaninova I, Sestak S, Svoboda A, Farkas V. 2000. Cell wall and cytoskeleton reorganization as the response to hyperosmotic shock in Saccharomyces cerevisiae. Archives of Microbiology 173: 245 –252.
Snyder JA. 1988. Effect of metabolic inhibitors on sucrose-induced metaphase spindle elongation and spindle recovery. Cell Motility and the Cytoskeleton 11: 291–302.
Snyder JA, Golub RJ, Berg SP. 1984. Sucrose-induced spindle elongation in mitotic PtK-1 cells. European Journal of Cell Biology 35: 62 – 69.
Starling D. 1976. The effects of mitotic inhibitors on the structure of vinblastine-induced tubulin paracrystals from sea-urchin eggs. Journal of Cell Science 20: 91–100.
Takahashi K, Isobe M, Knight MR, Trewavas AJ, Muto S. 1997. Hypoosmotic shock induces increases in cytosolic Ca2+ in tobacco suspension-culture cells. Plant Physiology 113: 587–594.
NPH033.fm Page 206 Wednesday, December 20, 2000 2:55 PM
Taylor AR, Manison NFH, Fernandez C, Wood JW, Brownlee C. 1996. Spatial organization of calcium signalling involced in cell Volume control in the Fucus rhizoid. Plant Cell 8: 2015 –2031.
Tazawa M, Shimada K, Kikuyama M. 1995. Cytoplasmic hydration triggers a transient increase in cytoplasmic Ca2+ concentration in Nitella flexilis. Plant Cell Physiology 36: 335–340.
Tucker JB, Milner MJ, Currie DA, Muir JW, Forrest DA, Spencer MJ. 1986. Centrosomal microtubule-organizing centers and a switch in the control of protofilament number for cell-surface associated microtubules during Drosophila wing morphogenesis. European Journal of Cell Biology 41: 279 –289.
Unger E, Bohm KJ, Vater W. 1990. Structural diversity and dynamics of microtubules and polymorphic tubulin assemblies. Electron Microscopy Reviews 3: 355 –395.
Urao T, Yabukov B, Satoh R, Yamaguchi-Shinozaki K, Seki M, Hirayama T, Shinozaki K. 1999. A transmembrane hybrid-type histidine kinase in Arabidopsis functions as an osmosensor. Plant Cell 11: 1743 –1754.
Vater W, Bohm KJ, Unger E. 1997. Tubulin assembly in the presence of calcium ions and taxol: Microtubule bundling and formation of macrotubule-ring complexes. Cell Motility and the Cytoskeleton 36: 76–83.
Vaughn KC, Harper JD. 1998. Microtubule-organizing centers and nucleating sites in land plants. International Review of Cytology 181: 75 –149.
Welnhofer EA, Travis JL. 1996. In vivo microtubule dynamics during experimentally induced conversions between tubulin assembly states in Allogromia laticollaris. Cell Motility and the Cytoskeleton 34: 81–94.
Williamson RE. 1991. Orientation of cortical microtubules in interphase plant cells. International Review of Cytology 129: 135 –206.
Wymer CL, Wymer SA, Cosgrove DJ, Cyr RJ. 1996. Plant cell growth responds to external forces and the response requires intact microtubules. Plant Physiology 110: 425 – 430.
Zachariadis M, Galatis B, Apostolakos P. 2000. Study of mitosis in root-tip cells of Triticum turgidum treated with the DNA-intercalating agent ethidium bromide. Protoplasma 211: 151–164.
Zandomeni K, Schopfer P. 1994. Mechanosensory microtubule reorientation in the epidermis of maize coleoptiles subjected to bending stress. Protoplasma 182: 96 –101.
Zielinski RE. 1998. Calmodulin and calmodulin-binding proteins in plants. Annual Review of Plant Physiology and Plant Molecular Biology 49: 697 –725.
NPH033.fm Page 207 Wednesday, December 20, 2000 2:55 PM
