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J. Cell Sci. Suppl. 2, 143-155 (1985)
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THE CYTOSKELETON UNDERLYING SIDE WALLS
AND CROSS WALLS IN PLANTS: MOLECULES AND
MACROMOLECULAR ASSEMBLIES
C. W. LLO Y D 1, L. C LA Y TO N 1, P. J. D AW SON1 J. H. D O O N A N 1,
S. H U LM E2, I. N. R O B ER T S1 a n d B. W ELLS1
lJohn Innes Institute, Colney Lane, Norwich NR4 7UH, U.K.
2Unilever Research Colworth Laboratory, Shambrook, Bedfordshire. U.K.
J.
SUMMARY
Plant cells organize their growth by reinforcing side walls during interphase (causing them to
elongate) and by positioning and orienting the cross wall at cytokinesis. In the first part of this
presentation we review progress made in identifying different cytoskeletal components that underlie
side walls and that are involved in the deposition of the cross wall.
During interphase, the cortical microtubule arrays co-distribute with an antigen recognized by
a ‘universal’ monoclonal antibody to intermediate filaments.
Using rhodaminyl-lysine-phalloidin no F-actin could be detected at the cortex but endoplasmic,
axial cables were found. The cytokinetic apparatus - the phragmoplast - contains microtubules and
we find that F-actin and the intermediate filament antigen also co-distribute with this array.
We describe the three-dimensional arrangement of microtubules forming the interphase array in
cells enlarging by both tip-growth and intercalary growth. In root hairs of higher plants and in apical
cells of the filamentous stage of moss Physcomitrella patens, microtubules (MT) are detected at the
apices and it is suggested from this that fragmentation of microtubules and absence of MTs from
the tip are preparation artefacts.
Using human serum from a scleroderma patient, possible microtubule nucleating sites are detected
in meristematic cells; these segregate with the broad spindle poles and they surround the nucleus
during early interphase - implying a peri-nuclear origin for the cortical MT array.
The interphase microtubule array is described in terms of a dynamic helical model, which
proposes: that the MT array is an integral complex; that microtubules form helices; that helices can
change their pitch - the array converting to the various conformations.
IN TRO DU CTIO N
Development involves the acquisition of polarity and, especially since plant cells
are not free to move, the ability to shift the polar axis in response to external cues.
Development of an axis involves the reinforcement of side walls against the lateral
expansion that would otherwise be uncontrolled as the turgid cell expands. A possible
macromolecular basis for cytoplasmic control over directional enlargement was
reported by Ledbetter & Porter (1963) after they saw for the first time transverse
hoops of cortical microtubules parallel to the innermost fibrils of the cell wall, which
were presumably resisting transverse strain. Since that time, microtubules (MT) have
been considered to be templates for the oriented deposition of wall fibrils but it has
become clear, as more evidence accumulates, that microtubules (and inner wall
fibrils) can adopt non-transverse orientations; that is, oblique as well as longitudinal.
Accordingly, the concept of cortical MTs as transverse hoops has to be revised and
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C. W. Lloyd and others
any new model will also have to take into account the ability of MTs to shift orienta­
tion. In this paper we present a three-dimensional model of microtubules capable of
undergoing re-alignments in time.
The particular way in which the new cell plate partitions the mother cell also has
a clear and direct effect upon the shaping of tissues, determining the symmetry of the
division products and the way they are related in space (i.e. separated by a transverse
wall or by a periclinal or anticlinal longitudinal wall). Microtubules help form the
cytokinetic phragmoplast and the orientation of this device is often predicted, before
mitosis, by the preprophase band of microtubules. Both before and after mitosis,
different microtubule arrays therefore define the division plane, but they are
separated in time by another transient apparatus, the mitotic spindle, and it is not
known how the ‘memory’ of preprophase band orientation persists until cytokinesis,
nor is it known how the division plane is determined in the first place. It is likely that
intracellular positional information is conveyed, not by one cytoskeletal element alone
but by a combination of elements, and in the first part of this paper progress in
characterizing the cytoskeleton and in studying the co-distribution of different com­
ponents around the cell cycle is reported.
MICROTUBULES
It is difficult to extract from plants the high concentrations of tubulin required to
initiate self-assembly into microtubules. However, taxol lowers the critical concentra­
tion for assembly and Morejohn & Fosket (1982) have begun the characterization of
taxol-MTs from rose cells. We have modified this method (Dawson & Lloyd,
unpublished) according to Dawson, Gutteridge & Gull (1983), in order to purify
carrot tubulin. Negative staining of taxol-MTs from carrot suspension cells shows
they are 24-30 nm in diameter and staining with tannic acid reveals a range of
protofilament numbers. By gel electrophoresis the subunits co-migrate in the vicinity
of brain tubulin except, as with Physarum tubulin (Clayton et al. 1980), the alpha
subunit migrates ahead of the beta (Fig. 1). It has been demonstrated by peptide
Œ1
Œ3
1
Fig. 1. 2-D gel electrophoresis of taxol-MTs from carrot suspension cells. Peptide map­
ping and immunoblotting confirm that the ^-tubulins migrate more slowly in. the
SD S-PA G E dimension (i.e. their positions are reversed relative to brain tubulins). In
addition, 2-D analysis reveals the multiplicity of subunits.
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145
mapping and by Western (immuno) blotting using subunit specific monoclonal
antibodies that this faster migrating species is alpha tubulin, heterologous to brain
alpha tubulin. On two-dimensional gels, it is apparent that there are multiple
subunits: two major and three minor beta, and four alpha tubulins. Heterology
of plant alpha tubulins and multiplicity of subunits has also been demonstrated
for homogenates of French bean (Hussey & Gull, 1985). Taxol-MTs are coldstable and hence do not cycle, but in pilot studies we have found that short MTs,
though few in number, can be prepared using 20 % dimethylsulphoxide to initiate
assembly.
F-ACTIN
There are few reports of the visualization of actin filaments in higher plant cells.
However, phallotoxins are now established as specific probes for F-actin (Pesacreta,
Carley, Webb & Parthasarathy, 1982; Wieland, Miura & Seelinger, 1983) and we have
used rhodaminyl-lysine phallotoxin (kind gift from Professor Wieland) to identify
actin filaments in formaldehyde-fixed onion root meristematic cells (Clayton & Lloyd,
1985). In elongated interphase cells, actin cables are seen in the endoplasm. These run
Fig. 2. Co-distribution of F-actin and microtubules in the phragmoplast. Formaldehydefixed onion root tip cells double-stained with: a , rhodaminyl-lysine phalloidin; b ,
monoclonal anti-tubulin (Yol/34). Bar, 10/im.
C. W. Lloyd and others
longitudinally, close to the nucleus. In cells double-stained for F-actin and for
microtubules it is clear that these endoplasmic cables do not co-distribute with
microtubules for the latter mostly occur in the cell cortex and at right angles to the
actin cables (i.e. transversely). No specific actin fluorescence is observed at pre­
prophase when microtubules form the pre-prophase band, nor are actin filaments seen
during the early stages of mitosis. Towards telophase, however, when the
phragmoplast microtubules help deposit the new cell plate between the sister nuclei,
specific patterns of fluorescence are restored, and in double-stained cells it is apparent
that F-actin and microtubules co-distribute throughout all phases of cytokinesis (Fig.
2). The phragmoplast consists of short, oppositely directed (Euteneuer & McIntosh,
1980), overlapping microtubules forming a palisade that grows centrifugally as the
cell plate expands within it. We interpret this co-distribution to mean that actin
filaments help move wall-containing vesicles to the mid-line marked by the zone of
overlap amongst the interdigitating phragmoplast microtubules. Antibody to yeast
actin (Kilmartin & Adams, 1984) also stains the phragmoplast of onion meristematic
cells (Clayton & Lloyd, 1985).
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OTHER FILA M EN TS
Cross-reactivity, often across kingdoms, of probes directed against the cytoskeleton, illustrates the ‘primitive’ nature of these scaffolding structures. Proteins of
the actomyosin and microtubule systems are well established as existing in plants but
although various other filaments are sometimes encountered (e.g. see Powell, Peace,
Slabas & Lloyd, 1982) there is as yet no evidence that intermediate filaments occur
in the plant kingdom. The amino acid sequences of various intermediate filaments can
be quite variable (Geisler et al. 1983) but a monoclonal antibody raised by Pruss et
al. (1981) immuno-blots with all five classes of filament and recognizes proteins from
annelids as well as from vertebrates. The consensus amino acids sequence recognized
by this antibody was recently described by Geisler et al. (1983). Immunoblots of
homogenates from onion roots and from carrot suspension cells (Dawson, Hulme &
Lloyd, 1985) show that this antibody cross reacts with plant proteins of approximately
68 and 50 (X 103)Mr. By indirect immunofluorescence it stains all four microtubulecontaining arrays in fixed onion meristematic cells (interphase, pre-prophase band,
mitotic spindle and phragmoplast). Staining of the phragmoplast is illustrated in
Fig. 3. One explanation would be that this ‘universal’ anti-intermediate filament is
cross-reacting with a microtubule subunit but this does not appear to be the case, for
the following reasons: taxol-MTs from carrot cells do not react positively on
immunoblots with anti-IFA, whereas proteins from isolated fibrillar bundles do; taxol
MTs can be immunoblotted with anti-tubulins but fibrillar bundle proteins cannot.
To rule out the possibility that anti-IFA may be non-reactive to sodium dodecyl
sulphate (SDS)-denatured proteins but reactive towards aldehyde-fixed proteins,
we fixed plant taxol-MTs in the standard manner used for cells. These MTs could
be stained, using indirect immunofluorescence, with anti-tubulins but not with
anti-IFA.
The plant cytoskeleton
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Fig. 3. Monoclonal antibody to intermediate filament antigen (Pruss et al. 1981) stains
the phragmoplast in fixed, onion root tip cells. Bar, 10 /im .
It is concluded that a protein containing an amino acid sequence common to all
classes of intermediate filament in animal cells, is present in plant cells and codistributes with microtubules. If this is correct, then the cytokinetic phragmoplast is
not only composed of microtubules and F-actin but of another cytoskeletal protein
also. In the same way, the interphase microtubule array may eventually prove to be
associated with components of this other system, but if actin microfilaments co-align
with cortical microtubules then this is beyond the resolution of fluorescence micro­
scopy.
T H E I N T E R P H A S E M I C R O T U B U L E ARRAY
Initiation of the array
Involving four different arrays, the microtubule cycle raises questions about the
timing and location of tubulin assembly, which relate to our understanding of cellular
morphogenesis. Where, for instance, are new interphase arrays re-established follow­
ing cytokinesis, at the nucleus or at the cell cortex? Are the nucleation sites different
from those involved in mitosis and cytokinesis; if not, do the sites migrate from the
peri-nuclear region to the cortex in order to initiate the new array? In files of cells, new
arrays can be regenerated transversely, as they were in the mother cell and there are
various explanations (see Lloyd & Barlow, 1982) for this apparent continuity between
generations: each new array may orientate itself by taking independent spatial readings
C. W. Lloyd and others
148
or else spatial information may be inherited by way of the pre-positioning of
microtubule-organizing centres. For all these questions it is clearly important to deter­
mine the location of microtubule-nucleating sites, but a persistent problem in higher
plant cell biology has been the lack of recognizably structured organizing material. This
problem is only rarely encountered with animal cells where the amorphous nucleating
components may surround the unmistakable centriole. However, the meiotic spindle of
the unfertilized mouse egg is also barrel-shaped and lacks a centriole, but CalarcoGillam et al. (1983) have identified the amorphous nucleating sites using serum from a
human scleroderma patient. On fixed onion cells this serum (Clayton, Black & Lloyd
1985) stains material that segregates with the spindle poles at prophase; at early inter­
phase, immediately following cytokinesis, fluorescence is found around the sister nuclei
and not at the cell edges (Fig. 4b ). If microtubule-nucleating sites are indeed being
stained then this would imply that microtubules grow out radially from the nucleus
towards the cortex and only later establish arrays that circumnavigate the cell, perhaps
as a result of MT—plasma membrane and/or M T—MT interaction.
Fig. 4. a . Following cytokinesis, as the phragmoplast disappears, the new interphase
array of M Ts forms radially between nucleus and cortex in fixed onion root tip cells. At
this stage the mature, transverse, cortical array is not yet established. B. At this postcytokinetic stage, human scleroderma serum, known to stain amorphous microtubule
nucleating sites in animal cells (Calarco-Gillam et al. 1983), stains the perinuclear region
rather than the cell cortex, suggesting that microtubules are initiated from the nuclear
region. Bar, 10 /im .
The plant cytoskeleton
149
Three-dimensional arrangement of microtubules in interphase arrays
Cells may expand in different ways: tip-growing cells, like root hairs, are usually
exposed to the environment and are thought to expand only at the apical dome,
whereas cells within tissues undergo a more uniform intercalary growth. The cyto­
skeleton may be specialized in different ways in support of these different modes of
growth.
Tip-growing cells. Colonies of the moss, Physcomitrella patens, can be ‘blotted’
onto glutaraldehyde-derivatized coverslips such that they do not disintegrate when
subjected to subsequent treatments making the cells permeable to antibodies
(Doonan, Cove & Lloyd, 1985). In this way, developmental lineages are preserved.
In tip cells of caulonemal filaments, the microtubules run more or less axially from
basal cross wall to the cell’s apex. Towards the tip, anti-tubulin fluorescence is more
abundant and this corresponds with the asymmetric distribution of organelles known
to occur in the closely related moss, Funaria hygrometrica (Schmiedel & Schnepf,
1980). The nucleus maintains an approximately constant distance from the tip and
endoplasmic microtubules are associated with this organelle. There is surprisingly
little to suggest from the literature that microtubules are associated with the tip but
anti-tubulin immunofluorescence clearly establishes their presence (Fig. 5a ).
Furthermore, the microtubules are organized at the apex where the more-or-less axial
elements become concentrated at one or a few foci. Unlike meristematic cells of higher
Fig. 5. Caulonemal tip cell of the moss P. patens stained by indirect immunofluorescence
with monoclonal anti-tubulin. In a , the more or less axial M Ts can be seen to focus
towards a point at the cell’s tip. This apical concentration of microtubules does not
depolymerize, as interphase microtubules do in higher plant cells, but persists through
mitosis ( b ) and cytokinesis (c). Bar, 20/Llm.
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C. W. Lloyd and others
Fig. 6. By fixing radish root hairs in a buffer composed of lOOmM-PIPES, 5 mM-Mgz+,
5 mM-EGTA, microtubules do not fragment as they do in phosphate buffer, nor are they
absent from the tip. The microtubules are net axial. Bar, lO^um.
plants, these interphase microtubules do not depolymerize during mitosis but are
present throughout the cell cycle and appear to interconnect with both the spindle and
the phragmoplast in turn (Fig. 5b , c ) .
Again, from electron microscopic (EM) studies, microtubules have been concluded
to be absent or poorly organized in the apical portion of root hairs of higher plants.
This may, however, be due to the conditions under which the cells are fixed. Using
PIPES, Mg2+, EGTA as a microtubule stabilizing buffer for immunofluorescence,
microtubules are seen to be arranged net axially in radish root hairs and to invade the
apical dome (Fig. 6). In long, highly vacuolate root hairs (Lloyd & Wells, 1985) it
becomes important to adjust the osmoticum by adding mannitol, otherwise - and this
is especially so when phosphate buffer (routinely used for EM) is substituted for
PIPES - the microtubules appear to be fragmented and no longer penetrate the apical
dome. It is concluded that the interphase MT array is a unitary assembly that only
appears fragmented when poorly fixed. Shadow casts of root hairs from which wall
matrix materials have been removed confirm that the innermost wall fibrils exist in
steeply pitched off-axial configurations that match the patterns formed by
microtubules. This observation is contrary to the multi-net growth hypothesis
The plant cytoskeleton
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(Roelofsen & Houwink, 1953), which holds that axial fibrils should only occur in the
outer wall layers as a result of originally transverse inner layers being re-aligned by cell
extension. One alternative explanation is that some of the primary wall texture is
generated by the advancing tip. It is thought that in such cells cellulose microfibrils
extend at the cell tip (Willison, 1982), which agrees with findings that presumptive
cellulose-synthesizing particles are concentrated towards the apex of tip-growing cells
(Wada & Staehelin 1981; Reiss, Schnepf & Herth, 1984). Microtubules, too, are
present at the tip and in onion root hairs, where they form 45 0 helices (Lloyd, 1983),
and have been seen to spiral into the apex. As the tip extends, microtubules would also
extend by following a spiral course. The rate of this rotation at the tip, balanced
against the rate of cell elongation, should influence the helical angle of fibrils deposited
in the wake of the apical synthesizing complexes.
In onion root hairs, microtubules form 450helices as do inner wall fibrils. However,
successive wall layers alternate their helical sign and, if microtubules are involved in
this, in order to remain parallel to the innermost fibrils they must then either undergo
rapid dis-assembly and re-assembly into arrays of opposite helical sign or else re­
orientate by twisting the entire array without depolymerization (cf. Robinson &
Quader, 1982). The possibility of dynamic behaviour cannot be ignored if it is accepted
both that microtubules parallel inner wall fibrils and that the stratigraphic record of
wall sections shows lamellae to be deposited in differing orientations. Confirmatory
evidence for such behaviour cannot be easily obtained from fixed systems, but in Fig.
7 whole cell immunofluorescence does demonstrate that microtubules are capable of
Fig. 7. In onion root hairs, microtubules generally form 45 ° multi-start helices. Occasion­
ally, other arrangements are encountered. Here, from the base of the cell, the microtubules
successively tend oblique, transverse, oblique and longitudinal angles, indicating both the
integral nature of the assembly and the fact that it can vary its helical pitch. Compare the
three orientations simultaneously occurring within this single microtubule array with the
separate orientations successively formed by epidermal cells treated with ethylene (see
Fig. 8). Bar, 20/im.
C. W. Lloyd and others
adopting various helical angles along the length of a root hair. Our results, which show
micro tubules to be present at the tip, therefore support the idea that apical growth
need not automatically be considered random because of a supposed lack of cytoskeletal organization; they also suggest that the component of wall texture generated
by tip advancement may be a reflection of the ability of microtubules to change the
pitch of their helices.
Intercalary-growing cells. Ledbetter & Porter (1963) described transverse cortical
microtubules as forming hoops around the cell. The concept of transverse hoops does
not appear to be sufficiently flexible to account for oblique and axial microtubules but
it should not be forgotten that these authors recognized, although it has been largely
ignored subsequently, that their observations could be explained by the tubules form­
ing low-pitched helices. This explanation was also suggested (Lloyd, 1984) on the
grounds that microtubules, which were long relative to the cell’s circumference,
would be more likely to form helices than hoops. Accepting that microtubules form
helices composed of unfragmented, relatively long individual elements, it only
requires that the entire array be capable of altering its helical pitch, to account for the
various orientations in which microtubules have been encountered in thin-sectioning
studies. That the interphase microtubule array is not a series of unconnected hoop­
like elements but is an entire assembly composed of helically wound microtubules has
been demonstrated for onion root hairs using immunofluorescence (Lloyd, 1983), but
a corollary of the dynamic helical model is that the entire assembly is able to alter its
helical angle. In the epidermal cells of mung bean and pea, the fibrils of each newly
deposited wall layer regularly alter their angle of deposition; the rhythmic shift of
alignment building up a rotating ply or helicoidal texture (Roland, Vian & Reis,
1977). Microtubules also shift their orientation in such cells: they are found in trans­
verse, oblique or longitudinal orientations, which are either parallel to the innermost
wall fibrils or anticipate the next orientation (Lang, Eisinger & Green, 1982).
Ethylene over-rides this programme by switching the alignment, of both microtubules
and inner wall fibrils, towards the longitudinal, and this encourages lateral expansion.
We (Roberts, Lloyd & Roberts, 1985) have used these well-documented effects
of ethylene to test the idea that the interphase microtubule array is based upon
helices capable of changing their pitch. According to this, the 90° switch in polarity
would be caused by the unwinding of helices, like the stretching of an initially com­
pressed spring. Sheets of epidermal cells of mung bean or pea stained with anti152
Fig. 8 . Pea or mung bean epidermal cells stained with anti-tubulin. In a , a sheet of
epidermal cells is shown to contain more-or-less transverse microtubule arrays. All three
orientations (a , b , c,) shown here occur in controls but the axial arrays (c) are scarce. After
treatment with ethylene, which is known to change cell polarity by 90°, inhibiting cell
elongation and encouraging lateral swelling, this balance is shifted as oblique arrays ( b )
predominate and axial microtubules (c) are more commonly encountered. Cells without
microtubules are not found, which suggests that the three conformations are interconvert­
ible forms of the basically helical structure seen to best advantage in the intermediate stage
(b ). Transverse arrays would therefore be based upon fully wound, flat helices and axial
arrays would be unwound, steeply pitched helices. The unwinding of helices shifts orienta­
tion through 90°. Bars, lOjUm.
The plant cytoskeleton
Fig. 8.
153
C.W. Lloyd and others
tubulin (Fig. 8) confirm that transversely wound microtubules occur more frequently
than other orientations, but within 30 min of treatment with ethylene, oblique micro­
tubules (resembling the multi-start 45 ° helices observed in onion root hairs) are more
readily encountered and the incidence of axial microtubules (rare in controls) is
increased. Since no cells are devoid of microtubules, then the three arrays are con­
strued as representing a progressive re-orientation rather than a series of depolymeriz­
ations and repolymerizations as occurs, for instance, between the preprophase band,
the mitotic spindle and the phragmoplast microtubules. However, unwinding a helix
from its fully wound transverse form to an unwound longitudinal form involves axial
extension of the elements (consider stretching a spring), but since the cells are not
elongating it seems likely that some degree of microtubule depolymerization is
occurring.
The significance of the oblique microtubules is that they can clearly be seen by
whole-cell immunofluorescence to be unambiguously helical and intermediate be­
tween the ‘transverse’and the ‘longitudinal’ arrays. Because of this, and as others have
argued from studies on wall fibrils (see Lloyd (1984) for review), the terms ‘flatpitched’ and ‘steeply-pitched’ are preferable since they acknowledge the helical basis;
they are figures at different points in a series, separated by 90 °. Microtubules are often
regarded as templates for the oriented deposition of wall fibrils and their ability to
undergo such re-arrangements offers an explanation for the rotating helical texture of
helicoidal walls and, in various permutations of orientation, an explanation for other
wall textures.
154
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