Download Structure–function relationships during secondary phloem

Tree Physiology 20, 777–786
© 2000 Heron Publishing—Victoria, Canada
Structure–function relationships during secondary phloem
development in an angiosperm tree, Aesculus hippocastanum:
microtubules and cell walls
NIGEL CHAFFEY,1,2 PETER BARLOW2 and JOHN BARNETT1
1
Department of Botany, University of Reading, Reading, U.K.
2
IACR—Long Ashton Research Station, Department of Agricultural Sciences, University of Bristol, Long Ashton, Bristol BS41 9AF, U.K.
Received April 28, 1999
Summary We studied the dynamics of the cortical microtubule (CMT) cytoskeleton during differentiation of axial secondary phloem elements in taproots and epicotyls of Aesculus
hippocastanum L. (horse-chestnut) saplings. Indirect immunofluorescence microscopy of α-tubulin and transmission electron microscopy revealed that fusiform cambial cells possessed a reticulum of CMTs in which individual microtubules
were randomly arranged. During differentiation of these
cambial cell derivatives into secondary phloem cells, the
CMTs were rearranged to become helically oriented, regardless of phloem cell type. Although helical CMTs were a persistent feature of all axial elements of the secondary phloem
(sieve elements, companion cells, phloem parenchyma, and fiber-sclereids), some modifications of this arrangement occurred as cells differentiated. Thus, at late stages of cell differentiation, sieve elements possessed nearly transverse CMTs,
pronounced bundling of CMTs was seen in phloem parenchyma, and the density of CMTs in the helical arrays of fibers
increased markedly. Additionally, phloem parenchyma possessed rings of CMTs in association with developing pit areas.
Aspects of the development and chemistry of cell walls were
also examined during phloem cytodifferentiation.
Keywords: cambium, cytodifferentiation, horse-chestnut, indirect immunofluorescence microscopy, sieve tube members.
Introduction
In view of the importance of trees as CO2 sinks in stabilizing
global climate changes, and as providers of wood, there is considerable interest in studying the process of xylogenesis. Despite the importance of secondary phloem in xylogenesis, e.g.,
as transport conduits for delivery of leaf-derived photosynthates, little is known about the morphogenesis and physiology of secondary phloem.
However, new cell biology techniques are facilitating study
of cambium and secondary vascular differentiation (Chaffey
1999). One of the most promising of these techniques is immunolocalization of the cytoskeleton (e.g., Lloyd 1987). The
cortical microtubule (CMT) component of the plant cytoskele-
ton has been implicated in a wide variety of morphogenetic
phenomena (e.g., Lloyd 1991, Baluška et al. 1998) with most
attention focused on cells or tissues of the primary plant body.
Although primary growth participates in the early development of seedlings, much growth at later stages, particularly in
dicotyledon perennial species, is provided by secondary meristems, such as the vascular cambium, and the tissues that they
produce. Recently, considerable success in understanding the
cell biology of xylogenesis has been achieved by applying immunolocalization techniques to both the CMTs and microfilament components of the cytoskeleton in softwoods (Funada
et al. 2000) and hardwoods (Chaffey 2000). However, this
technique has not yet been applied to a detailed study of
morphogenesis of secondary phloem.
Although ultrastructural studies of secondary phloem have
been undertaken for more than 30 years, microtubules (MTs)
within this tissue have rarely been studied (e.g., Evert and
Deshpande 1969). Attempts to relate changes in distribution,
orientation and abundance of MTs to other changes within a
differentiating phloem element in angiosperm species have
been limited to ultrastructural studies of phloem (Northcote
and Wooding 1966, 1968) and protophloem sieve elements in
cotton roots (Thorsch and Esau 1982) and grasses (Eleftheriou
1995).
Previously, we used indirect immunolocalization of α-tubulin to demonstrate that CMTs within fusiform cambial cells
of the angiosperm tree, Aesculus hippocastanum L. (horsechestnut), are randomly arranged in a reticulum (Chaffey et al.
1996, 1997a, 1997c). The CMTs undergo cell-type-specific
changes during subsequent xylogenesis, and are intimately associated with all forms of wall-elaboration in this tissue (e.g.,
Chaffey 1999). We have now applied the same technique to
determine the role of the CMT cytoskeleton in the development of the axial elements of the secondary phloem of A. hippocastanum. We found evidence of a degree of dynamism in
the CMT cytoskeleton of the phloem that has not previously
been reported. We also attempted to correlate differentiationrelated alterations in the CMTs with changes in cell wall
chemistry.
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CHAFFEY, BARLOW AND BARNETT
Materials and methods
Plant material, growth conditions, and sampling
Seedlings of A. hippocastanum (Hippocastanaceae) were raised
as described previously (Chaffey et al. 1996). Samples were
taken during the first 3 years of seedling growth, during periods when the vascular cambium was active in cell production.
Indirect immunofluorescence microscopy of α-tubulin
As described more fully elsewhere (Chaffey et al. 1996), radial
longitudinal slivers of cambial tissue and derivatives were excised from taproots and epicotyls and fixed for about 4.5 h in a
3.7% (w/v) solution of paraformaldehyde in microtubule-stabilizing buffer (MTSB; 12.5 mM piperazine-N,N′-bis-(2ethylsulfonic acid) (PIPES), 5 mM ethylene glycol-bis-(βaminoethyl ether) N,N,N′,N′-tetraacetic acid (EGTA) and
5 mM MgSO4, pH 6.9) in 10% (v/v) dimethyl sulfoxide. The
slivers were then dehydrated and embedded in Steedman’s low
melting point wax, and sectioned at 6 µm. After dewaxing and
dehydration, sections were incubated with the primary antibody, monoclonal mouse anti-α-tubulin (Amersham, Little
Chalfont, U.K.), diluted 1:200 in phosphate-buffered saline
(PBS), for 1 h at 37 °C in the dark. Following an MTSB rinse,
sections were incubated with the secondary antibody, fluorescein isothiocyanate (FITC)-conjugated goat-anti-mouse IgG
(Sigma Chemical Co., Poole, U.K.) diluted 1:80 or 1:160 in
PBS, and incubated as for the primary antibody.
Some sections were stained with Calcofluor White M2R
New to allow study of cell walls (e.g., Chaffey 1994). Autofluorescence of tissues was reduced by lightly staining sections with Toluidine Blue (Chaffey et al. 1996) before mounting in anti-fade mountant (p-phenylenediamine in glycerol).
Sections were examined with a Zeiss Standard 18 microscope
with epifluorescence illumination and standard filter combinations for FITC (e.g., Staiger 1994) and Calcofluor (e.g.,
Chaffey and Pearson 1985) fluorescence. Appropriate controls
were employed as previously described (Chaffey et al. 1996).
Tissue was also embedded in butyl-methylmethacrylate.
Tissue was processed as described above for wax-embedment,
except that butyl-methylmethacrylate resin mixture containing
5 mM dithiothreitol was substituted for wax, and blocks were
cured by UV illumination at –20 °C (Chaffey et al. 1997d).
Sections were incubated in a blocking solution (bovine serum
albumin, fish-skin gelatin, normal goat serum and glycine in
PBS) for about 45 min before application of the primary antibody, monoclonal mouse-anti-α-tubulin, diluted 1:10 in PBS.
Sections were incubated for about 2 h at room temperature before a thorough wash in PBS and application of the secondary
antibody, FITC-conjugated goat-anti-mouse monoclonal, diluted 1:30 in a solution containing bovine serum albumin,
fish-skin gelatin, and NaN3 in PBS. Sections were incubated
for about 1 h at room temperature, washed in PBS, lightly
stained with Toluidine Blue and mounted in Vectashield®
anti-fade mountant (Vector Laboratories Ltd., Peterborough,
U.K.), and examined in a Zeiss Axiophot light microscope. A
Zeiss LSM410 inverted laser scanning microscope was used to
produce confocal images of some of the butyl-methylmeth-
acrylate-embedded material immunostained for localization
of α-tubulin. A detailed description of the experimental design
is given in Chaffey et al. (1999).
Transmission electron microscopy
Tissue was fixed in a mixture of glutaraldehyde and paraformaldehyde in PIPES buffer, post-fixed in OsO4 and embedded in
Spurr’s resin (Chaffey et al. 1997a, 1997c). Ultrathin sections
were contrasted with uranyl acetate and lead citrate and
viewed in a Hitachi H-7000 electron microscope operating at
75 kV. Additionally, semi-thin sections were stained with
Toluidine Blue (O’Brien and McCully 1981) or Methylene
Blue/Azur A (Clark 1981).
Thiéry reaction (PATAg test) for polysaccharides
For the PATAg test, which was based on a modification of the
method of Thiéry (1967) (Chaffey 1985b), we used ultrathin
sections of material that had been processed for transmission
electron microscopy, either with or without an osmicationstep. Sections were collected on gold grids and examined by
transmission electron microscopy without further contrasting
at 50 or 75kV.
Immunolocalization of pectins in cell walls
Material was processed as described previously by Chaffey et
al. (1997c). Briefly, tissue was fixed for about 4.5 h in a mixture of 3.7% paraformaldehyde and 0.2% glutaraldehyde in
25 mM PIPES buffer, pH 6.9, then dehydrated in a graded water–ethanol series and embedded in LR White resin. Sections
were cut either at 2 µm and mounted on poly-lysine-coated
glass microscope slides for immunofluorescence, or at about
100 nm and collected on uncoated gold grids for immunogold
localization in the transmission electron microscope. Sections
were then incubated in blocking solution (as for immunofluorescence above) for about 45 min before application of the
undiluted primary rat-monoclonal antibody, JIM5 or JIM7
(gifts from Dr J.P. Knox, University of Leeds, Leeds, U.K.).
Typically, JIM5 recognizes unesterified epitopes of pectin and
JIM7 recognizes methyl-esterified epitopes of pectin (Knox et
al. 1990). Sections were incubated for about 2 h at room temperature and then washed with PBS before application of the
secondary antibody. We used FITC-conjugated anti-rat monoclonal antibody (Sigma Chemical Co., Poole, U.K.) for indirect immunofluorescence microscopy, and anti-rat monoclonal antibody linked to 10 nm gold particles (British BioCell
International, Cardiff, U.K.) for transmission electron microscopy. Secondary antibodies were prepared in a solution containing bovine serum albumin, fish-skin gelatin and NaN3 in
PBS, and sections were incubated for about 1 h at room temperature. After washing in PBS, slides were further processed
for immunofluorescence microscopy and examined as above.
Grids were allowed to air dry for transmission electron microscopy and examined at 75 kV either with or without contrasting with lead citrate.
Cytochemistry of cell walls
Hand-cut sections of fresh root tissue were stained with Phlo-
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roglucinol/HCl for identification of lignin (O’Brien and
McCully 1981) and examined in transmitted light. Other sections were stained with Aniline Blue for localization of callose
(e.g., Clark 1981) and Calcofluor White M2R New for identification of β-linked glucans (e.g., Chaffey 1985a, 1996) and
examined with epifluorescence illumination.
Results
General anatomy
Changes in the CMT cytoskeleton described for axial elements of the secondary phloem of the taproot of Aesculus were
similar to those in the epicotyl of this species. In transverse
section, the secondary phloem was distinguished from the
cambial zone by the presence of the products of anticlinal radial division of cambial derivatives, and consisted of companion cells, phloem parenchyma, sieve tube members, and
lignified fibers (which first appeared during the second year of
seedling growth). Differentiation of cambial derivatives to
secondary phloem elements was accompanied by an increase
in diameter and cell wall thickening (Figure 1). With reference
to the fusiform cambial cells, phloem parenchyma cells were
shorter, fibers were considerably longer, and sieve tube members were approximately the same length, or sometimes
shorter; companion cells were usually shorter than their associated sieve tube member (Figure 2).
Ultrastructure
At the ultrastructural level, the axial phloem cell types were
readily distinguishable at maturity. Sieve tube members were
essentially empty-looking cells (Figure 2) with P-protein filaments and starch-bearing plastids within the lumen, and tubular membranous structures closely appressed to the radial and
tangential walls. Their oblique end walls were perforated by
callose-lined pores within the sieve plates. At maturity, a nucleus appeared to be absent, as did most other organelles.
Companion cells had a considerably elongated nucleus and a
full complement of organelles, although the plastids generally
appeared devoid of starch (Figure 2). The cytoplasm was
densely stained because of the large number of free ribosomes.
Complex plasmodesmata were present in the walls between
the companion cell and sieve tube member and were branched
on the side adjacent to the companion cell. The cytoplasm of
parenchyma cells was less electron-dense than that of companion cells, but contained abundant starch-bearing plastids;
no complex plasmodesmata were observed (Figure 3). In sectioned material, vacuoles occasionally contained circular profiles of material that was assumed to be tannin, based on its
green coloration with Toluidine Blue (O’Brien and McCully
1981) and its pronounced osmiophilia (Esau 1969). Presence
of starch within the plastids was inferred on the basis of staining with PATAg (Figure 4), confirming that the plastids of
sieve tube members are S-type (cf. Behnke 1981). Fibers were
generally devoid of cell contents and had thick, lignified cell
walls (Table 1).
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Cell walls
Information obtained from studies of the cell wall chemistry of
fusiform cambial cells and their axial phloem derivatives is
presented in Figures 4 and 5 and Table 1. Although phloem
cell walls were thicker than those of their cambial precursors,
they showed the same staining reactions, except for lignification of fiber walls and the presence of callose in the sieve plate
of sieve tube members. In appropriately oriented sections, layering was seen within the secondary phloem cell wall (e.g.,
Figure 12). Such layering was not observed within the walls of
active cambial cells. In agreement with the observations of
Esau and Cheadle (1958) on sieve tube members of Aesculus
shoot tissue, there was no sign of a nacreous wall in taproot
material of this species.
Microtubule cytoskeleton
Fusiform cambial cells possessed a reticulate arrangement of
randomly oriented CMTs, but there was no obviously preferred angle of orientation to the component microtubules
(Figures 6 and 7).
In elements of the secondary phloem at their earliest stages
of development, i.e., in cells within the cambial zone, the
CMTs as seen by immunofluorescence were arranged helically (Figure 7), apparently extending throughout the length of
the cell. Transmission electron microscopy showed parallel
CMTs at both tangential walls of the cell, also arranged in a
manner consistent with a helical arrangement (Figure 8). At
this early stage of differentiation it was not possible to identify
which cell type the element would become.
In cells close to the cambial zone, which could be identified
as phloem cell types, parallel CMTs were observed by
epifluorescence microscopy (fiber-sclereids; Figure 9) and
confocal laser scanning microscopy (parenchyma cells; Figure 12). The CMTs within the helical arrays of fibers were
much more dense than in other axial phloem cell types (cf.
Figure 9 with Figures 10 and 15). The only deviation from the
helical arrangement of CMTs in axial phloem cells was found
in some parenchyma cells, where rings of MTs were observed
in association with the periphery of primary pit fields (Figure 11), but co-existing with the helical array. Study of parenchyma cells by TEM corroborated the presence of helical
arrays of CMTs (Figure 12), and demonstrated their bundled
nature (Figure 13). The CMT bundles were closely associated
with particulate structures (Figure 13), which may be cytoplasmic ribosomes or MT-associated proteins (MAPs). Coorientation of CMTs and microfibrillar features within the
walls of these cells was also evident (Figure 13).
Microfibrillar structures observed by TEM in the walls of
secondary phloem parenchyma cells close to the boundary
with the primary phloem, which are among the oldest phloic
derivatives of the cambium, were observed to swirl around the
pit fields (Figure 14). In similar cells immunostained for localization of α-tubulin, helices of CMTs were evident (Figure 15). Staining of such cells with Calcofluor showed helical
structures apparently within the cell walls (Figure 16).
At a late stage of sieve tube member development, as indi-
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CHAFFEY, BARLOW AND BARNETT
Figures 1–5. Transmission electron micrographs showing the
structure and cell wall chemistry
of axial elements of the secondary
phloem in taproots of Aesculus
hippocastanum. Bars: 5 µm (Figure 1), 4 µm (Figures 2 and 3),
2 µm (Figure 4), and 0.2 µm (Figure 5). Abbreviations: C, cambial
zone; CC, companion cell; F, fusiform cambial cell; I, intercellular
space; N, nucleus; P, phloem parenchyma cell; SP, secondary
phloem tissue; S, sieve tube member; and W, cell wall. Figure 1.
Radial longitudinal section showing the cambial zone and adjacent
secondary phloem tissue. Note the
phragmoplast (䊐) within the fusiform cambial cell that has been
fixed during periclinal division,
and the increase in cell diameter
and wall thickness as cambial derivatives differentiate as secondary
phloem elements (direction of vascular differentiation indicated by
arrow). Figure 2. Radial longitudinal section showing a cytoplasmic
companion cell dominated by an
elongate nucleus, and portions of
two adjacent sieve tube members
that are nearly devoid of cell contents. Figure 3. Radial longitudinal
section showing portions of two
adjacent phloem parenchyma
cells. Figure 4. Transverse section
showing portions of two sieve
tube members, companion cells,
and surrounding cells stained with
PATAg to reveal polysaccharides.
Note the intense staining of the
cell walls and starch (䉳) within
the plastids of the sieve tube member, and the weak staining of the
callose region of the sieve pores
(夽). Figure 5. Transverse section
showing immunogold JIM 5-staining of pectins within cell walls
surrounding an intercellular space.
Note the pronounced gold labeling
of the middle lamella region of the
cell wall (夽).
cated by their cell diameter and distance from the cambial
zone, helices of CMTs were still evident in sieve tube members. However, the helices were much shallower—almost
transverse in some instances—than in other axial phloem cells
(cf. Figures 15 and 17).
Discussion
Early stages of differentiation
The marked change in CMT orientation, from random in
cambial cells to helical in secondary phloem vascular derivatives, is regarded as evidence that determination, i.e., the establishment of a pathway of vascular differentiation, has taken
place (Chaffey et al. 1997a). All derivative cell types, whether
sieve tube members, companion cells, parenchyma cells or fibers, appeared to have similar helical CMT orientations that
differed from those in the cambium. This observation confirms the difficulty of distinguishing among the different
phloem cell types at an early stage of differentiation (e.g.,
Parthasarathy 1975, Evert 1990, Romberger et al. 1993, Iqbal
1995, Iqbal and Zahur 1995).
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Table 1. Comparison of cell wall chemistry of fusiform cambial cells and their secondary phloem derivatives in taproots of Aesculus hippocastanum.
Procedure
Cambium
Phloem
Comments
MeB/AzA
Toluidine Blue
Phloroglucinol/HCl
Aniline Blue
Calcofluor
CTEM
Purple
Purple
—2
+ 4,5
+++
++
Purple/blue-green1
Purple/blue-green1
—/red 3
+++ 6
+++ 7
++
PATAg
++
++
JIM5
++ 8
++ 9
JIM7
++ 10
++ 10
Methylene Blue/Azur A (Clark 1981)
O’Brien and McCully 1981
Stain for lignin (O’Brien and McCully 1981)
Fluorescent stain for callose (Clark 1981)
Fluorescent stain for cell walls (e.g., Chaffey 1994)
Conventional transmission electron microscopy using uranium and
lead-contrasting
Periodic acid-thiocarbohydrazide-silver proteinate stain
for polysaccharides (Thiéry 1967)
Monoclonal antibody recognizing unesterified epitopes of pectin
(Knox et al. 1990)
Monoclonal antibody recognizing methyl-esterified epitopes of pectin
(Knox et al. 1990)
1
2
3
4
5
6
7
8
9
10
Blue-green-staining reaction only in fibers.
— indicates no staining noted.
Red-staining reaction only in fibers.
Abbreviations: +, ++, +++ denote relative intensity of staining reaction.
Only in association with nascent tangential cell walls.
Only at sieve plates, and displacing Calcofluor-staining from the sieve pores.
Particularly at sieve plates.
Throughout cell wall, but concentrated at middle lamella.
Throughout cell wall, but particularly at the middle lamella and cell–cell junctions bordering intercellular spaces.
Throughout cell wall.
Cortical MTs have been implicated in cell wall formation
(e.g., Lloyd 1991). Reorientation of the random CMTs to a helical arrangment coincided with increased rates of synthesis
and deposition of wall material, suggesting that the CMT
reorientation is associated with the onset of cell wall thickening, a feature that is prominent in the development of phloem
derivatives.
Although it is uncertain whether true secondary thickening
takes place within phloic derivatives (e.g., Esau 1969, 1979,
Iqbal and Zahur 1995), considerable deposition of wall material occurs as differentiation proceeds (e.g., Evert 1990).
Catesson (1992) stated that cell walls of tree secondary phloem are characterized by the deposition of a “more ordered”
cellulose microfibril framework early in differentiation. This
view is supported by our finding of wall layering in phloem
cell walls, but not in the precursor cambial cell walls. The observed co-orientation of putative cell wall microfibrils with
helical CMTs supports the widely held view that MTs are involved in aligning cellulose microfibrils (e.g., Giddings and
Staehelin 1991, Hable et al. 1998) in secondary phloem cells.
These observations suggest that there are structural differences in those regions of the phloem cell wall formed against a
background of helical CMTs, and that phloem cell walls in
Aesculus are not simply thicker versions of their cambial progenitors. An association between parallel MTs and wall thickening is also found in cambial cells during dormancy (Chaffey
et al. 1998), during late stages of secondary xylem fiber differentiation (Chaffey et al. 1999), and in secondary xylem vessel
element development (Chaffey et al. 1997b, 1999).
We were unable to detect significant differences between
the thickened walls of the phloem and primary walls of the
cambium (cf. xylem and cambium cell walls in Chaffey et al.
1997b) based on the use of monoclonal antibodies against different polysaccharide epitopes, JIM5 and JIM7, suggesting
that the walls of phloem elements are similar in composition to
primary walls. This conclusion accords with other work on
secondary phloem cell walls of shoots of different tree species
(e.g., Catesson 1973, Freundlich and Robards 1974, Evert
1990).
Differentiating cells
Although helical CMTs are a consistent feature of all secondary phloem cells in Aesculus, in companion and parenchyma
cells the CMTs also show bundling. Such bundling is likely to
be promoted by MT-associated proteins (MAPs) (e.g., Cyr
1991). Whether the electron-dense particles seen in association with the CMTs in Figure 13 are MAPs or ribosomes is not
known. The significance of CMT bundling is also not known.
In parenchyma cells, it may be related to the formation of the
helical wall thickenings that are revealed by Calcofluor staining (Figure 16). The thickenings possibly act to strengthen the
wall, and may be analogous to the wall elaborations and modifications in the secondary xylem. The present observation may
also have some relevance to the crossed helical striations previously recorded within the sieve cell walls of Pinus strobus
by Chafe and Doohan (1972).
The swirled nature of the cell wall microfibrils around the
pit fields of phloem parenchyma cells is similar to the micro-
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CHAFFEY, BARLOW AND BARNETT
Figures 6–11. Radial longitudinal sections of the microtubule
cytoskeleton of axial secondary
phloem cells and their cambial
precursors in taproots of
Aesculus hippocastanum. Figures 6 and 7 are transmission
electron micrographs; Figures
8–11 are fluorescence micrographs of sections immunolocalized for α-tubulin. Bars:
0.5 µm (Figures 6 and 8), and
20 µm (Figures 7 and 9–11).
Abbreviations: F, fusiform
cambial cells; FS, fibersclereids; P, phloem; R, ray
cells; SP, secondary phloem elements; and W, cell wall. Figure 6. Randomly oriented
cortical microtubules (arrows)
in a fusiform cambial cell. Figure 7. Immunofluorescence micrograph showing the change
in cortical microtubule orientation from random in fusiform
cambial cells to helical in axial
phloem elements. Figure 8.
Parallel-oriented cortical
microtubules (arrow) within
differentiating axial secondary
phloem elements. Figure 9.
Dense helical arrays of cortical
microtubules in developing fiber-sclereids. Figure 10. Confocal laser scanning image of
helical microtubules in axial
cells of the secondary phloem.
Figure 11. Rings of cortical
microtubules (䉰) in a differentiating phloem parenchyma
cell.
fibril patterns around xylem bordered pits (Preston 1939) and
illustrated for a parenchyma cell in the coleoptile of wheat
(Preston 1974). The pit-associated rings of CMTs found in
some phloem parenchyma cells represent the only deviation
from the helical orientation in secondary phloem cells that we
found. It is noteworthy that they co-exist with helical CMTs.
The rings of CMTs thus appear to represent a distinct subset of
CMTs, rather than helical CMTs deviating around the pit field.
Their significance remains unknown. However, by analogy
with the similar MT rings associated with formation of bordered pits (Chaffey et al. 1997b, 1999), contact pits (Chaffey et
al. 1999), and perforations (Chaffey et al. 1997d, 1999) in secondary xylem vessel elements of this species, we suggest that
the ring of MTs defines a domain in the cell wall within which
thickening will not take place, i.e., at the primary pit field.
We observed the presence of CMTs within sieve tube members of the secondary phloem. Although we do not know
whether such cells were functional, their distance from the
cambial zone and their diameter suggested that they were at
late stages in their development. In the same sections, it was
also possible to observe similarly oriented CMTs within sieve
tube members of primary phloem (N.J. Chaffey, unpublished
observations), which must be considered functional. The generally prevailing view for this phloem element is that “the
microtubules typically disappear entirely from the cell” (Evert
1990); however, the relevant observations have been derived
solely from ultrastructural studies. The observation presented
here—derived from a different fixation and visualization pro-
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783
Figures 12 and 13. Radial longitudinal sections showing the
ultrastructure of the cell wall–
plasma membrane–cytoplasm
interface of secondary phloem
parenchyma cells in taproots of
Aesculus hippocastanum. Bars:
1 µm (Figure 12), and 0.25 µm
(Figure 13). Abbreviations: P,
phloem parenchyma cell; W,
cell wall. Figure 12. Bundling
of cortical microtubules (arrows) near the cell wall; note
also the pronounced wall layering. Figure 13. Particles (in encircled region) associated with
the bundles of microtubules
(black arrows). Note also the
co-orientation of microtubules
and cell-wall microfibrillar features (white arrows) (䊐).
cedure—raises the possibility that MTs may not necessarily be
absent from sieve tube members; however, further work will
be necessary to establish the longevity of these cell features
within the sieve tube members.
The decrease in pitch of the CMT helix—from high-angled
to near transverse—in sieve tube members during differentiation and the concomitant increase in diameter is consistent
with the view that these CMTs are intimately associated with
the plasma membrane, possibly through cross-linking element(s) (e.g., Traas 1990). However, this reorientation provides no information on whether an increase in cell circumference is, in part, driven by a change in CMT arrangement, or
whether CMTs are dragged to a more shallow-pitched helix as
a consequence of such an increase. We note that secondary xylem vessel elements also possess transversely-oriented CMTs
at a late stage of their development (Chaffey et al. 1997b,
1999).
It also remains unclear whether an increase in diameter of
the CMT helix, as would occur with cell widening, relates to
overlapping CMTs (e.g., Hardham and Gunning 1977) sliding
further apart, so that they overlap less, or to production and insertion of additional CMTs, or a combination of both. In accordance with the ideas of Marchant (1982), it may be that,
under certain conditions, CMTs stabilize the changing shape
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CHAFFEY, BARLOW AND BARNETT
Figures 14–17. Radial longitudinal sections of cell walls and
the cortical microtubule cytoskeleton in axial elements of
the secondary phloem in taproots of Aesculus hippocastanum. Bars: 1 µm (Figure 14),
and 20 µm (Figures 15–17).
Abbreviations: P, phloem parenchyma cell; PF, pit field;
and S, sieve tube member. Figure 14. Electron micrograph of
a glancing section through the
wall of a parenchyma cell
showing the swirled nature of
microfibril features around the
pit fields, which are perforated
by numerous plasmodesmata.
Figure 15. Immunofluorescence micrograph of helical
cortical microtubules in a
phloem parenchyma cell. Figure 16. Fluorescence micrograph of phloem parenchyma
cells stained with Calcofluor,
revealing prominent helical
structures within the cell wall.
Figure 17. Immunofluorescence micrograph of nearly
transversely oriented cortical
microtubules within a sieve
tube member at a late stage of
differentiation.
of a cell that, before wall thickening and wall hardening, is
bounded only by a thin, pliant primary cell wall. Thus, any
physical association between CMTs and plasma membrane
could promote a close appression of the latter against the cell
wall (Barnett 1981), facilitating a more ready transfer of
polysaccharide precursors from cytoplasm to sites of cell wall
biosynthesis. This suggestion implies that interactions between CMTs and plasma membrane are important in cell
morphogenesis (e.g., Traas 1990), and highlights the role of
the cell wall–plasma membrane–cytoskeleton continuum
(e.g., Wyatt and Carpita 1993) in the process of phloem
cytodifferentiation.
We obtained no evidence for the involvement of CMTs in
sieve plate formation in Aesculus sieve tube members. It might
be that CMTs have no role to play in sieve plate formation, the
endoplasmic reticulum perhaps being of greater importance
(e.g., Northcote 1968, Iqbal 1995). The apparent absence of a
role for CMTs in sieve plate formation may be because
callose, rather than cellulose, is deposited at the sieve pore. A
detailed ultrastructural study by Eleftheriou (1990) also failed
to demonstrate an involvement of microtubules in sieve plate
formation in wheat root protophloem.
On the basis that fibers were absent close to the cambial
zone and were present only during the second and subsequent
years of growth, they are considered to be fiber-sclereids (e.g.,
Fahn 1990, Romberger et al. 1993) as opposed to bast fibers.
TREE PHYSIOLOGY VOLUME 20, 2000
PHLOEM DEVELOPMENT IN TREES
Lack of detection of early stages of differentiation of these
cells suggests that they develop from already differentiated
parenchyma cells that subsequently undergo “renascent
growth and sclerification” (Romberger et al. 1993). Despite
their apparent development from axial parenchyma cells, the
fiber-sclereids of the secondary phloem share similarities with
the secondary xylem fibers that develop directly from fusiform cambial cells. For instance, both possess a much greater
density of helically arranged CMTs than is seen in other secondary phloem cells (this study) or secondary xylem cells
(Chaffey et al. 1999), and develop lignified, thickened cell
walls that bear simple pits oriented at a steep angle to the long
axis of the cell (N.J. Chaffey, unpublished observations). Further investigation of the controls over lignification and morphogenesis of these phloem cells is also likely to increase our
understanding of cytodifferentiation of xylem fibers, a wood
cell type of considerable economic importance.
Acknowledgments
N.J.C. thanks Dr. J.P. Knox for the gift of the JIM antibodies, and
Dr. C. Hawes for use of his confocal laser scanning microscope. This
project was funded by the BBSRC under a LINK award to Reading
University and IACR—Long Ashton Research Station. IACR receives grant-aided support from the Biotechnology and Biological
Sciences Research Council of the United Kingdom.
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