Download Symplasmic fields in the tunica of the shoot

1477
Development 125, 1477-1485 (1998)
Printed in Great Britain © The Company of Biologists Limited 1998
DEV0145
Symplasmic fields in the tunica of the shoot apical meristem coordinate
morphogenetic events
Päivi L. H. Rinne1,* and Christiaan van der Schoot2,†
1Department of Plant Physiology, Agricultural University Wageningen, Arboretumlaan 4, NL-6703 BD Wageningen, The
Netherlands
2ATO-DLO, Bornsesteeg 59, P.O. Box 17, NL-6700 AA Wageningen, The Netherlands
*Present address: Department of Biology, University of Oulu, P.O. Box 333, FIN-90571 Oulu, Finland
†Author for correspondence (e-mail: [email protected])
Accepted 5 February; published on WWW 18 March 1998
SUMMARY
In plants, complex cellular interactions, which require the
exchange of morphogenetic signals, underlie morphogenesis
at the shoot apical meristem. Since all apical meristem cells
are interconnected by plasmodesmata, we have investigated
if symplasmic paths are available which may preferentially
channel metabolites and potential morphogens in the apical
meristem, and whether they could support both the
formation of determinate appendages and the sustainment
of an undifferentiated centre. Experiments in which the
permeability of the symplasm was probed with fluorescent
dye revealed that the tunica of the apical meristem of birch
seedlings (Betula pubescence Ehrh.) is symplasmically
compartmentalized into two concentric fields, which restrict
the symplasmic diffusion of small potential morphogens to
the cells inside their boundaries. A transient connection
between the two fields was established early in a
plastochron, potentiating the radial exchange of
symplasmically diffusing signalling molecules. We suggest
that the symplasmic subdivision of the tunica offers a
means to unite cells into communication compartments,
invoke boundary interactions between them, and
shield the distal meristem cells from organogenesis.
Electrophysiological measurements indicate that, in
addition, the cells of these fields constitute metabolic
working units. The relevance of these symplasmic fields for
morphogenesis was established experimentally by
treatment with short photoperiod, which induced
breakdown of the fields into symplasmically isolated cells.
Tannic acid staining and in situ immunolocalisation
revealed that cell isolation was due to the activation of
glucan synthase complexes intrinsic to sphincters. As a
result callose plugs were formed on all plasmodesmata
leading to morphogenetic deactivation.
INTRODUCTION
cooperatively bring about a specific set of structures’ (Müller,
1997). In the ‘positional information model’ (PI model) the
patterning of morphogenetic fields, e.g. in limb formation,
requires the graded presence of diffusing or non-diffusing
morphogens (Wolpert, 1989, 1996). Such gradient fields (e.g.
de Robertis et al., 1991) may provide positional information
which is interpreted by the cells and recorded as positional
values necessary for regeneration and development (Wolpert,
1989, 1996). In plants, morphogenesis at the AM has been
explained in terms of the PI model by assuming the presence
of morphogen gradients in the extracellular space (for review
see Holder, 1979). In the AM two deviations from this situation
may occur, however. Gradients may also be set up in the
symplasmic space, and cells may be engaged in active forms
of conversation. The morphogenetic field hosting such
conversation networks must sustain infinite and repetitive
development of the shoot. Regeneration as well as development
at the AM would then involve self-organizing and selfmaintaining signal networks in addition to gradients, a
In all higher plants the vegetative apical meristem (AM) gives
rise to the shoot. The basic structure of the AM is formed as
an integral part of the embryo (Jürgens, 1995), but the open
growth habit of the shoot and its phyllotactic pattern emerge
from the exclusive activity of the AM (Steeves and Sussex,
1989). The organizing principles are intrinsic to the AM proper
since isolated AMs, with the primordia removed, sustain the
ability for primary morphogenesis (e.g. Steeves and Sussex,
1989; Sachs, 1991). Even small groups of cells, surgically
isolated at different AM loci, can rebuild an AM identical in
size and function to the original (e.g. Sussex, 1989; Steeves
and Sussex, 1989). How the AM is able to regenerate and
sustain its dynamic organization is a key question in plant
morphogenesis, but it has received surprisingly little attention.
In animal systems the ability of fragments to produce copies
of the whole, demonstrated during early embryogenesis, is due
to a morphogenetic field, i.e. ‘an area where the cells
Key words: Dormancy, Apical meristem, Vegetative meristem,
Morphogenesis, Plasmodesmata, Plasmodesmal sphincter, Tunica
1478 P. L. H. Rinne and C. van der Schoot
situation which seems to go beyond the PI model. It is not clear
how these gradients and hypothetical signal networks are
positioned in the cellular matrix of the AM.
Importantly, the proliferating AM restricts pattern
formation to its periphery and maintains an undifferentiated
centre (e.g. Sawhney et al., 1981; Steeves and Sussex, 1989)
which suggests the presence of physiological barriers
(Sawhney et al., 1981), the position of which must be
independent of cell proliferation. The mechanisms by which
these barriers in the morphogenetic field are maintained and
by which cell cycling is coordinated in space and time
(Ferreira et al., 1994) operate at a level above that of the
individual cell (Sachs, 1991; van der Schoot, 1996). The
coordination of cellular events may involve biophysical
processes (Green, 1985; Selker et al., 1992; Hernández and
Green, 1993; Green et al., 1996; Fleming et al., 1997),
morphogen diffusion and protein trafficking (Jackson et al.,
1994; Hantke et al., 1995; Furner et al., 1996; Perbal et al.,
1996; Szymkowiak and Sussex, 1996; van der Schoot, 1996).
The possibility that the organization of the symplasm
underlies the functional subdivision of the AM into zones has
not been previously considered as important in morphogenesis.
Since an elaborate symplasmic network is present within the
AM in the form of numerous cytoplasmic strands,
plasmodesmata, this may provide a channel for distribution of
signals. The way the symplasmic organization of the AM is
built-up depends on the patterns of cell division. Cytoplasmic
unity in cell lineages is created by incomplete cell divisions or
‘chambering’ (Kaplan and Hagemann, 1991) yielding cells
which are exclusively connected by cytokinetic or primary
plasmodesmata (e.g. Gunning, 1978). The multiple branched
lineages in the AM are postcytokinetically connected through
existing walls by secondary plasmodesmata (van der Schoot,
1996; Bergmans et al., 1997) indicating that a threedimensional symplasm is required in AM functioning (van der
Schoot and Rinne, 1996). Primary and secondary
plasmodesmata have been suggested to play partially different
roles in the spatiotemporal control of signal flow (Lucas et al.,
1993), but the specific distribution of plasmodesmata types
within the AM (van der Schoot, 1996; Bergmans et al., 1997)
is not necessarily sufficient to achieve adequate control over
signal flow. In the floral meristem (FM) of Antirrhinum the
difference in the transport of the transcription factor,
GLOBOSA, between and within layers was suggested to be
due to differential regulation of secondary and primary
plasmodesmata, respectively (Perbal et al., 1996). Since
secondary plasmodesmata are also formed within a layer,
between and within branched clonal cell groups (van der
Schoot, 1996; van der Schoot and Rinne, 1996, Bergmans et
al., 1997), GLO would be able to reach all FM cells without
restrictions. Additional mechanisms, supracellular in nature,
must therefore be operational in the spatial control over
morphogen distribution.
The existence of two separate growth zones, tunica and
corpus, is also considered of importance in morphogenesis
(Steeves and Sussex, 1989), but their individual roles and the
way they integrate their activities are unclear. Bidirectional
interactions between tunica and corpus occur at the AM
periphery during the development of leaf primordia, but
spatiotemporal positioning of a primordium and the
development of dorsiventrality rely on mechanisms that act in
a radial fashion (Steeves and Sussex, 1989). Mechanisms that
shield a central population of tunica cells from organogenesis
must also act radially (van der Schoot and Rinne, 1996). When
the FM is formed, the entire tunica, not only its peripheral part,
responds to signals send out from the corpus resulting in the
sequential production of floral organs (Szymkowiak and
Sussex, 1992, 1996). Biophysical models depict primordium
formation as an event initiated in the tunica (e.g. Green, 1985;
Hernández and Green, 1993; Green et al., 1996), but input from
morphogen sources (e.g. Fleming et al., 1997) may be
necessary to sustain the local growth alterations at the future
site of primordia.
Therefore, the organization of the symplasm may underlie
not only the division of the AM into a central and peripheral
area, but also the division into a tunica and corpus. Diffusion
may thus be limited to compartments in the AM that correspond
to this pattern of subdivision. Diffusion of morphogens in
communication compartments is important in developing
animal systems where gap junctions, the ‘equivalents of
plasmodesmata’ (Pitts, 1990; Lucas et al., 1993), are involved
in cell determination (de Laat et al., 1980; Canveney, 1990;
Serras, 1989), but direct experimental investigations of
diffusion paths in the AM are lacking. We here report on the
presence of symplasmic fields in the tunica of proliferating AMs
of birch seedlings, which potentially harbour gradient fields of
diffusing morphogens. These symplasmic fields subdivide the
tunica into two radially organized diffusion compartments,
which keep their position in the proliferating cellular context
and may support the peripheral formation of leaf primordia and
the maintenance of an undifferentiated centre. AMs often
switch to an alternative behaviour in response to environmental
changes. Photoperiod length, for example, influences flower
evocation, tuberization and dormancy (Bernier, 1988; VincePrue, 1994). In general, environmental stimuli may affect
symplasmic communication (e.g. McLean et al., 1997), and in
the case of photoperiodically induced switches this may lead to
altered symplasmic patterns at the AM, serving its new state or
function. We have demonstrated here for birch that exposure to
short photoperiod induced breakdown of the symplasmic fields,
stopped primary morphogenesis, and eventually resulted in
dormancy. We therefore propose that the symplasmic network
in the tunica is important in the coordination of morphogenetic
events at the AM.
MATERIALS AND METHODS
Plant material
Field-grown Northern Scandinavian seedlings of birch (Betula
pubescence Ehrh.), approximately 40 cm in height, were transplanted
in pots at the start of the second growing season, and grown further
in a growth chamber at 20°C and 16 hours photosynthetically active
radiation (PAR) with photon flux density of 100 µmol·m−2·s−1. After
one month, about 10 plastochrons, half of the plants was subjected to
a short photoperiod, 8 hours of PAR. Growth activity, elongation and
production of new leaves were monitored throughout.
Iontophoretic microinjection and electrophysiology
Iontophoresis and electrophysiological measurements were
performed as described previously (van der Schoot and Lucas, 1995).
Apical shoot parts (with 3 mm of stem), with the AM exposed, were
submerged and fixed upright in a bathing chamber. The AM surface
Symplasmic fields in the tunica 1479
was inspected under low-irradiance white light. The extent of green
autofluorescence of cut leaf tissues was assessed by short epiillumination with blue-violet (BV) and blue light (B) in a
fluorescence microscope (Nikon Optiphot II, Nikon, Tokyo, Japan).
The (untouched) AM was free of green autofluorescence but could
show red fluorescence. Excitation and barrier filters were standard
BV and B filter-sets (BV: excitation 400-440/barrier 470/dichroic
mirror 455; B: excitation 470-490/barrier 515/dichroic mirror 510).
Entry of the microelectrode into a tunica cell was signalled by a sharp
voltage drop. After the recording a stable membrane potential (Em)
(a criterion for the ability of the impaled cell to make an adequate
membrane seal around the microelectrode tip) Lucifer Yellow CH
(LYCH; 457 Da) was iontophoresed (I = −1 to −5 nA) into the cell.
In most experiments, however, the small tunica cells (diameter
approx. 10 µm) did not survive the impalement with the
microelectrode (diameter approx. 1 µm), or they had difficulties in
sealing the membrane around the tip. In the latter case LYCH-input
resulted in leakage through membranes, and in restricted dye-transfer
to adjacent cells due to early collapse of the impaled cell. These
experiments were discarded altogether. Vigorously growing plants
were most suited for microinjection, yielding a stable Em and fastforming dye-coupling patterns. In case of deactivated (dormant)
plants we relied on the absence of leakage and the presence of a
stable Em. Dye outflow from the microelectrode into the L1 cell was
examined directly using the microscope. Patterns of LYCH
distribution in actively growing plants (success rate, 9-10%; see
Table 1), and in dormant plants (success rate, 45%) were
photographed on Ektachrome-400 super slide film, and reproduced
in photographs and diagrams.
Light (LM)- and transmission electron microscopy (TEM)
Leaves (but not primordia) were removed from apical shoots, and the
stem trimmed back to 3 mm in fixative, 2% (v/v) glutaraldehyde in
100 mM phosphate buffer (pH 7.2). Samples in fixative with and
without tannic acid (1% w/v) were vacuum-infiltrated for 15 minutes
and incubated in fresh fixative at 20°C for 4 hours. Samples, part of
them postfixed with OsO4 for 1 hour, were embedded in LR White
resin (Agar Scientific, Stansted Essex, UK) according to standard
procedures. For LM 1 µm thick sections were cut, stained with
toluidine blue and studied with the light microscope. For TEM thin
sections (90 nm) were made, double stained with uranyl acetate and
lead citrate and studied in a Phillips CM12 TEM (Eindhoven, The
Netherlands).
In situ immunolocalisation
AMs were fixed in 0.75% (v/v) glutaraldehyde, 1.6% (w/v)
paraformaldehyde and 0.75% (v/v) acrolein. Sections were blocked
for 1 hour with 3% (w/v) bovine serum albumin (BSA) in TBS (10
mM Tris, pH 7.4; 150 mM NaCl). After washing, the sections were
labelled with polyclonal primary (anti-1,3-β-glucan; Genosys
Biotechnologies Inc., UK) and secondary antibodies as described
previously (Northcote et al., 1989). Sections were examined directly
or after fixing in 2% (v/v) glutaraldehyde followed by double
staining.
RESULTS
Symplasmic fields in the tunica layer of the
proliferating AM
The AM of birch is composed of a tunica, of one or
occasionally two layers, superimposed on a corpus (Fig. 1A,C),
and all its cells are interconnected by plasmodesmata (data not
shown) potentially allowing diffusion of morphogens between
all cells of the AM. Diffusing morphogens could be of
significance in the AM, particularly if there were preferential
symplasmic pathways to distribute them.
In search of such pathways we have microinjected the nontoxic and membrane-impermeable fluorescent probe LYCH,
which visualizes diffusion pathways in the symplasm (Stewart,
1981). By direct observation we established that injection of
LYCH, into cells at the peripheral zone of the AM (Fig. 2A),
resulted in consistent dye-coupling patterns. In some cases
(Table 1) LYCH injected into a single L1 cell, moved full circle
in both directions along the base of the AM dome. In
approximately 20 seconds LYCH occupied the entire
peripheral zone of the tunica layer, excluding the leaf
primordia, rising up along the outer edge of the peripheral zone
(Fig. 2B). A single radial diffusion boundary inside the
peripheral zone was typically present, where plasmodesmata
were not able to pass on LYCH to neighbouring cells (Fig. 2C).
In agreement with the literature this is interpreted as being due
to a decrease in the size exclusion limit of the plasmodesmata
(references in Lucas et al., 1993), and which may even lead to
complete plasmodesmata closure (Duckett et al., 1994).
Hereafter we will refer to these permeability changes as
‘plasmodesmata narrowing or closure’, which probably occurs
at the plasmodesmata neck region and/or at the level of
subchannels (Lucas et al., 1993). In some cases the injection
into the AM periphery resulted only in the formation of
segments of the symplasmic ring (Table 1). In all cases,
however, LYCH never entered subjacent tissue, nor the central
AM area, which remained sharply delineated from the
peripheral zone of the AM.
Microinjection of LYCH into a single cell of the central zone
of AMs revealed the presence of a central zone symplasmic
field in L1, complementary in shape to that at the peripheral
zone (Fig. 3). The boundaries became visible within seconds
and were usually irregular due to the protrusion of small rows
of cells, probably part of a lineage, which still retained
symplasmic continuity with the central zone field (Fig. 3).
When the AM was relatively small, i.e. at the beginning of
a plastochron, microinjection of LYCH into a cell of the central
zone revealed the presence of a central zone field which
connected to a minor segment of the peripheral zone field (Fig.
4A-C). This fusion event was encountered a few times (Table
1) and probably existed only briefly.
Distinct Em-values in cells of the central zone and
peripheral zone symplasmic fields
The symplasmic fields at the periphery and the centre of the
AM (Figs 2, 3) matched the central and peripheral zones of the
Table 1. Observations on symplasmic permeability in
actively proliferating (under long day) and deactivated
(under short day) AMs
Plant type
LD
LD
LD
SD
Type of symplasmic field
Number of AM
czf, distinct border
pzf, distinct border to czf
closed ring
closed ring, 2-3 segments
czf fused with pzf segment
cell isolation, in pzf and czf
23
12
5
7
3
12
LD, long day grown plants; SD, short day grown plants; czf, central zone
field; pzf, peripheral zone field.
1480 P. L. H. Rinne and C. van der Schoot
Fig. 1. Birch seedlings exposed to long photoperiod and short
photoperiod, and micrographs of their AM in median-longitudinal
sections, stained with toluidine blue. (A) AM of long-day plant
showing histological zonation in four superimposed cell layers.
Central histological zone, between solid arrowheads; peripheral
histological zone between solid and open arrowheads; p,
primordium. Scale bar, 50 µm. (B) Actively growing seedling with
new leaves continuously unfolding from the apex. (C) AM of shortday plant, exposed for 2 weeks to short photoperiod, showing
absence of histological zonation. Scale bar, 50 µm. (D) Deactivated
seedling, exposed for 2 weeks to short photoperiod, showing
formation of a terminal bud with brown scales (arrow).
zonation model (Fig. 1A; Steeves and Sussex, 1989; Sachs,
1991) both in position and activity level. Electrophysiological
recording of the cell Em in the symplasmic field at the AM
periphery showed strongly negative values (Table 2) due to the
presence of an electrogenic component (∆Ψ), which may
locally drive active protonated uptake of substrates (van der
Schoot and van Bel, 1990, and references therein).
In contrast, the Em values of tunica cells in the central zone
fields were considerably smaller than those in peripheral
zone fields of the AM (Table 1), indicating a low proton
pumping activity and carrier-mediated uptake of substrates.
The low variation of Em values between cells of the same
Table 2. Membrane potentials of cells in symplasmic fields
at the peripheral zone and central zone loci of actively
proliferating and deactivated AMs
AM status
Active
Deactivated
AM locus
Em (mV)*
±s.e.
n
cz
pz
cz+pz
−52
−98
−39
2
4
6
40
21
26
*The mean Em values of the cells at various loci were statistically different
from each other (P<0.001, Student’s t-test, paired comparisons). cz, central
zone; p2, peripheral zone; Em, membrane potential.
Fig. 2. Dye-coupling pattern emerging after iontophoresis of LYCH
into a single L1 cell at the AM periphery. Diagrams indicate position
of dye-coupled cells: top, longitudinal section; bottom, surface view.
Curved arrows show direction of spread of the dye and the straight
arrow indicates field boundary. (A) Symplasmic movement of LYCH
from injected L1 cell in both directions along the AM periphery,
visible in individual cells. (B) After approx. 20 seconds LYCH
moved full circle through the L1 at the side of the pronounced AM
dome. A symplasmic barrier is present close to the microelectrode
(arrowhead). (C) Unchanged dye-coupling pattern after 30 minutes.
Some L1 cells do not contain dye. Insert shows detail of the
overexposed area. Scale bar, 50 µm. AM, apical meristem; cz, central
zone; L, leaf; me, microelectrode; pz, peripheral zone.
field showed that the cells making up those fields may have
a strong electric coupling and may also represent metabolic
coupling groups.
Short day induced breakdown of tunica fields
An important effect of short photoperiod, exploited here, is
the deactivation of the AMs of birch seedlings (Rinne et al.,
1994). To test the significance of the observed symplasmic
fields for primary morphogenesis, we deactivated birch
seedlings by treatment with short photoperiod. A 2-week
exposure to short photoperiod induced growth cessation and
disappearance of histological zonation (Fig. 1). In plants
exposed to 4 subsequent short days, cells in the peripheral
zone and central zone of the AM tunica, microinjected with
LYCH, did not pass on the dye to other cells anymore (Fig.
4D) indicating a breakdown of both types of symplasmic field.
Cell isolation was accompanied by a decrease of the Em values
Symplasmic fields in the tunica 1481
in both fields (Table 2). Apparently, some days after short
photoperiod exposure the symplasmically isolated cells
retained only the basic activity level characteristic for cells in
a resting state.
Reversibility, up to a point, is typical for photoperiodic
responses in general (e.g. Bernier, 1986). In birch, growth
reversibility is already severely reduced after 10 days of
exposure to short photoperiod (Welling et al., 1997). We have
used this property to test in a separate experiment whether this
was reflected in the symplasmic structure of the AM. The
described breakdown of symplasmic fields by short
photoperiod appeared to be reversible after 1 but not 3 weeks
under short photoperiodic conditions. After 3 weeks,
impalement of a cell with a microelectrode resulted in a strong
osmotic withdrawal of LYCH solution from the electrode tip
(Fig. 4E). The cells retained the LYCH and their swollen state
at least until the next day, indicating firm plasmodesmata
closure and water-proofing of cell walls.
Since all main temperate tree species have photoperiodic
ecotypes differing in their response to day length (references
in Olsen et al., 1997), we have confirmed in a separate
experiment that a more southern ecotype (from Southern
Scandinavia) showed the same response although the
dissipation of the symplasmic fields was delayed in
correspondence with their morphogenetic activity by
approximately 1 week.
Different plasmodesmata mechanisms are involved
in the formation and dissipation of fields
In active AMs versatile mechanisms must be operational for the
S3
cz
pz
Fig. 3. Dye-coupling pattern emerging from a single injected L1 cell
at the AM centre. Diagrams indicate position of dye-coupled cells in
surface views and longitudinal section; curved arrows show direction
of spread. (A) LYCH moved symplasmically in approx. 5 seconds
through the AM centre. Irregularity of the boundary is due to cell
proliferation. (B) Final dye-coupling pattern reached after approx. 15
seconds. Photograph was taken after 30 minutes. Scale bar, 50 µm.
AM, apical meristem; cz, central zone; L, leaf; me, microelectrode;
pz, peripheral zone; s, section plane for cross section diagram.
pz
Fig. 4. Dye-coupling pattern representing a ‘fusion field’ (A-C) and
lack of dye-coupling under short photoperiod conditions (D,E).
Diagrams indicate the positions of dye-coupled L1 cells and
symplasmically isolated cells in surface views and longitudinal
sections. The top four diagrams refer to A-C and the bottom two to
E. (A) LYCH moves symplasmically in approx. 5 seconds from the
injected L1 cell at the AM centre towards the AM periphery (curved
arrow in diagram). (B) After 15 seconds symplasmic boundaries are
distinct. (C) Dye-coupling pattern in the L1 after 30 minutes.
(D) AM L1 cell injected after 4 days exposure to short photoperiod
shows absence of dye-coupling. (E) AM L1 cells in former
peripheral zone and central zone, injected after 4 weeks of short
photoperiod. Absence of dye-coupling and swelling of dehydrated
cells due to water absorption from the impaling microelectrode.
Scale bar, 50 µm. AM, apical meristem; cz, central zone; L, leaf;
me, microelectrode; pz, peripheral zone; s1-3, section planes 1-3 for
cross section diagrams.
1482 P. L. H. Rinne and C. van der Schoot
transient narrowing or closure of the plasmodesmata at the
boundaries of a symplasmic field. Local activation of 1,3-βglucan synthase resulting in controlled callose deposition
around the plasmodesmata neck has been considered as an
important mechanism for the regulation of the plasmodesmal
permeability (e.g. Lucas et al., 1993). The lack of labelling with
anti-callose antibody shows, however, that this mechanism is
not involved in the creation of symplasmic field boundaries (e.g.
Fig. 5A,B).
In contrast, callose was evidently involved in the breakdown
of symplasmic fields in the AM during morphogenetic
deactivation by short photoperiod (Fig. 5C-E). This was shown,
firstly, by fixation in the presence of tannic acid, which
visualizes putative glucan synthase complexes at the
plasmodesmata neck (Olesen, 1980; Olesen and Robards, 1990)
in patterns typical for sphincters (Fig. 5C). These sphincters
could not be detected in the morphogenetically active AM (Fig.
5A,B). Secondly, anti-callose antibodies clustered at the
plasmodesmata neck (Fig. 5D,E) in patterns identical to those
found with tannic acid (Fig. 5C), demonstrating that short
photoperiod induced formation and subsequent activation of
glucan synthase complexes at the plasmodesmata entrances.
Physical closure of the plasmodesmata (Fig. 5D,E),
categorically preventing symplasmic signalling, effectively
blocked further formation of leaf primordia (not shown).
Fig. 5. Ultrastructure of plasmodesmata in active (A,B) and deactivated
(C-E) AMs. (A) Standard fixation with tannic acid. Absence of
sphincters on plasmodesmata between L1 and L2 (as elsewhere).
(B) Immuno-fixation and anti-callose labelling. Scant labelling of
plasmodesmata between L1 and L2 (as elsewhere). (C) Standard
fixation with tannic acid. Sphincters at the cytoplasmic ends of
plasmodesmata between cell membrane and wall. (D) Immuno-fixation
and anti-callose labelling. Sphincters, slightly visible due to naturally
occurring tannins. Antibodies cluster at sphincters. (E) As in D, but
without staining. c, plasmodesmata channel; cw, cell wall; s, section
through sphincter ring. Scale bar (A-E), 100 nm.
DISCUSSION
Control over AM function resides at the supracellular level
(e.g. Poethig, 1990; Sachs, 1991; Steeves and Sussex, 1989;
van der Schoot, 1996). We therefore addressed the question
of whether the organization of the symplasm could provide
a setting for such control. We investigated the tunica, which
in birch is normally one layer thick (Fig. 1). Whether the
degree of AM stratification is significant for primary
morphogenesis has remained unclear (Szymkowiak and
Sussex, 1996). In sunflower, for example, the two layers
show a similar zonation (Sawhney et al., 1981), and in
Arabidopsis similar growth dynamics occur (Schnittger et
al., 1996). It is feasible that when more tunica layers are
present they form distinct physiological compartments with
very similar growth dynamics. Below we discuss the
functional structure of the symplasmic network in the tunica
of birch and its relevance for the coordination of
morphogenetic events at the AM.
Symplasmic fields subdivide the tunica
We have probed the symplasm of the AM by microinjecting
LYCH into single tunica cells at different locations. Since
LYCH does not pass through the cell membrane (Stewart,
1981), its spread from injected cells visualizes all
symplasmic pathways available to small (up to 1 kDa)
diffusing morphogens. The groups of symplasmically united
tunica cells in which gradients of diffusing morphogens can
be formed have been defined here as ‘symplasmic fields’. In
mature tissues of plants, e.g. stem and leaves, groups of
symplasmically coupled cells have been characterised as
‘symplastic domains’ (Erwee and Goodwin, 1985), which
stabilize the tissue and serve as storage and for internal
redistribution of metabolites (van der Schoot and Van Bel,
1990; Lucas et al., 1993). Our earlier use of the term
‘symplasmic domain’ for coupling groups in the AM (van
der Schoot and Rinne, 1996, and references therein) seems
now inappropriate; since their cellular composition changes
continuously due to cell proliferation and they collectively
constitute a morphogenetic field, these cell-coupling groups
should be conceptualized as ‘fields‘ rather than ‘domains’.
In the AM of birch we found two different symplasmic
fields. A concentric, ring-like field was present in the
primordium forming part of the tunica (Fig. 2) which
enclosed a second field, situated in the central
undifferentiated part of the tunica (Fig. 3). The partial fields
observed at the AM periphery may in some cases represent
the complementary field of a fusion-field. Since the
peripheral zone symplasmic field could be locally only a few
cells wide, particularly where central cells were protuding
into the periphery of the AM (Fig. 3C), the symplasmic
movement may be slowed down or stopped at these locations.
Possible local damage to the symplasmic connections in these
corridors, due to primordia removal, cannot be ruled out. The
presence of these fields demonstrates that plasmodesmata
connections, although produced continuously between
dividing cells, and within and between cell layers (van der
Schoot, 1996; Bergmans et al., 1997), are narrowed or closed
at certain boundaries within the AM. The resulting
symplasmic fields, in analogy to those created by gap
junctions in animal systems (de Laat et al., 1980; Serras,
Symplasmic fields in the tunica 1483
1989), are suited to harbour gradient fields of diffusing
morphogens, which provide positional information to the
cells within the fields (Wolpert, 1978, 1989). Such gradient
fields may be represented by simple diffusion patterns or by
gradients of interacting morphogens (Turing, 1952;
Meinhardt, 1989; Wolpert, 1989). The small size of
morphogenetic fields in animals supports the hypothesis that
such gradients are important as ‘prepatterning’ agents (Crick,
1970; Müller, 1997). The size of the central zone symplasmic
field in birch, 80-125 µm in diameter (Fig. 3), is well within
this range (Holder, 1979; Müller, 1997). Since at the
proliferating AM cells are displaced towards the periphery,
plasmodesmata must be narrowed or closed by a positiondependent mechanism which keeps the size of the central
zone field in proportion to the size of the AM. The fact that
initial cells, occasionally drifting from their distal position,
can be replaced by derivatives (Steeves and Sussex, 1989),
indicates that this particular position in the overall signal
network of the AM (but not the cell as such) provides an
anchoring point for the mechanism which controls the size of
the central zone field. The mechanism which controls
plasmodesmata narrowing or closure at the field boundaries
could, therefore, be based on a gradient set up by the initial
cells. The central zone field in birch corresponds to the
histological central zone (Fig. 1A) and, as schematically
shown in Fig. 6B, the steepness of the hypothetical gradient
inside the central zone field may determine the size of the
field by influencing where the size exclusion limit of the
plasmodesmata is lowered.
The low Em values in the central zone field (Table 2) match
the low metabolism and cell cycling rate described for this
distal area (Esau, 1977; Steeves and Sussex, 1989). This
histological zone often extends beyond the tunica into the
corpus (Steeves and Sussex, 1989) but the tunica, which lies
within the central zone (Fig. 6), can be very distinctive and
physiologically different from the subjacent central zone cells
(Sawhney et al., 1981). This fact was, rightly in view of our
findings, considered to be due to physiological isolation of
these tunica cells, but the presence of numerous
plasmodesmata connections did not seem to support such
isolation (Sawhney et al., 1981). As it appears now it is not
the absence of plasmodesmata which physiologically
separates the distinct zones but their temporary narrowing or
closure, as demonstrated by the present microinjection
experiments. The large Em values of cells in the peripheral
zone field reflect the active proton pumping which drives
competitive nutrient uptake, needed to maintain a high
metabolic activity during cell proliferation. The large
difference in cell Em values between the two fields (Table 2)
indicates that the symplasmically united tunica cells are
functioning as metabolic working units with distinct
morphogenetic tasks. Subjacent cell layers may also be
subdivided into metabolic working units since they also show
a histological zonation (see e.g. Fig. 1A).
The internal diffusion boundaries in the AM of birch
coincide with patterns of gene expression in other species.
Certain genes, some of which have a ‘household’ function, are
expressed in the entire vegetative AM, but not in the upper
layer, or vice versa (Hake and Freeling, 1986; Pri-Hadash et
al., 1992; Fleming et al., 1993). These expression patterns can
change during the floral transition (Pri-Hadash et al., 1992) and
may be influenced by spatial alterations in symplasmic fields.
In Iris, plasmodesmata frequencies between L2 cells and
surrounding cells diminish in early floral development
(Bergmans et al., 1997), which implies a specific alteration in
the density of connections in the overall symplasmic network.
This probably affects diffusion patterns in the AM
quantitatively, but it is unknown if the pattern of symplasmic
fields have also changed during the transition to flowering. This
is feasible, however, since the physiological zonation of the
AM is also altered.
The diffusion boundaries described here may not present a
barrier to diffusing molecules and ions that are smaller than
LYCH (457 Da). In the case of root hair cells of Arabidopsis,
which exhibit a high degree of electrical isolation (Meharg et
al., 1994), the absence of dye-coupling has been argued to be
due to occlusion or developmentally regulated closure
(Duckett et al., 1994). The distinct difference in Em’s of the
central and peripheral field in the birch tunica similarly
suggests that the plasmodesmata size exclusion limit at the
field boundaries is close to zero. Certain larger factors may
pass the field boundaries by plasmodesmata gating, e.g. those
involved in the regulation of non-autonomous gene
expression or those involved in the coordination of cellular
events across layers. For example, GLO in Antirrhinum is
suggested to move between layers through plasmodesmata
(Perbal et al., 1996), and may pass the diffusion boundary by
transient opening and widening of plasmodesmata. Such
trafficking would also provide a possibility for transient
cotransport of diffusing morphogens. In addition, a lipid
signalling pathway in the endoplasmic reticulum (Grabski et
al., 1993) may be operational across symplasmic field
boundaries.
Symplasmic fields may temporarily fuse or dissipate
A fusion between the two symplasmic fields in the tunica was
observed a few times when the AM was relatively small (Fig.
4A-C), probably at the beginning of a plastochron. This event,
possibly coinciding with the initiation of the leaf primordium
that will emerge at the end of the plastochron, may serve the
symplasmic exchange of certain morphogenetic signals
between the two fields. Such signals may be chemical or
electrical in nature (Lucas et al., 1993). Fusion events are also
of interest for photoperiodic processes, such as flowering and
dormancy, that typically take place in the leaf and are expressed
at the AM (Bernier, 1988). In Anagallis (pimpernel), for
example, the AM is only responsive (competent) to the
inductive photoperiod during a short period in its plastochronic
trajectory (Bernier, 1988). As the ‘florigenic signal’ probably
travels via the symplasm from the leaf to the AM (Gunning
and Robards, 1976; Bernier, 1988), it may reach the central
meristem during a fusion event.
The uncoupling of AM cells in birch, induced by short
photoperiod (Fig. 4D-E), was due to plasmodesmata being
plugged with callose (Fig. 5C-E) and is obviously an effective
means of preventing symplasmic diffusion, macromolecular
trafficking or lipid transfer. Symplasmic uncoupling of cells in
fern prothallia releases individual cells from global tissue
constraints and results in redifferentiation and subsequent
development of new prothallia from each individual cell
(Tucker, 1990). In higher plants, some cellular constraints must
remain, since redifferentiation is held in check and tissue
1484 P. L. H. Rinne and C. van der Schoot
Fig. 6. Schematic representation of models defining morphogenesis
at the shoot apical meristem (AM), and their relationship to the
observed symplasmic fields. (A) Structural organization of the AM
reflects the direction of cell divisions (arrows in left diagram), i.e.
anticlinal in the tunica (t) and in all directions in the corpus (c).
Physiological organization is reflected in the histological zonation.
Metabolically active cells with high cell cycling are at the AM
periphery (a) and metabolically less active cells with low cell cycling
occupy the centre (b). (B) The present work shows that the tunica (t)
of the AM is subdivided into symplasmic fields (sf): central field (1),
peripheral field (2), primordium (3), which host hypothetical gradient
fields (gf), created by source/sink mechanisms and by boundary
interaction between fields. According to the PI model (e.g. Wolpert,
1989) cells measure their position in a gradient field which can be
linear due to diffusion from a central source to a peripheral sink, or
exponentially decreasing due to the presence of sinks along the
diffusion paths; beyond a certain morphogen concentration, cells
may exit the morphogenetic field. In addition, ‘reaction-diffusion
systems’ (Turing, 1952) inside the central symplasmic field may
provide positional information of some kind.
functioning can be restored by recoupling the cells (Erwee and
Goodwin, 1984). In birch the symplasmic uncoupling of AM
cells, and the resulting arrest of development, was reversible at
the early stage by transfer to long photoperiod. This indicates
that the inert state of the AM becomes fixed – serving the
survival through winter – by biochemical processes which
develop in symplasmically uncoupled cells. Biochemical
adjustments involve changes in cell-autonomous gene
expression (Welling et al., 1997), but it is unknown if they
relate to the fixing of the AM.
Plasmodesmata closure due to short photoperiod is
controlled by a different mechanism than plasmodesmata
closure/narrowing at field boundaries, since sphincters were
present only in the former (Fig. 5). Sphincters, found in a
limited number of plants, have been suggested to regulate cellcell transfer through plasmodesmata by the local deposition of
callose (Olesen and Robards, 1990; Lucas et al., 1993).
Sphincters are often absent, however, even in species for which
they have been described. Explanations for this include the
possibility that sphincters develop only late at certain stages of
cell differentiation (Olesen and Robards, 1990), that they
reflect a particular functional stage of plasmodesmata, or that
they are stress-induced (Turner et al., 1994). Our observations
show that in the birch AM sphincter formation occurs during
developmental transitions induced by the environment. The
sensing of day length and the subsequent formation of
sphincters are likely part of the phylogenetically acquired
mechanisms by which the plant anticipates the arrival of harsh
conditions, and enters a dormant state.
In the above we have characterized regular and dynamically
sustained symplasmic patterns of cell coupling in the tunica.
The spatiotemporal correlation of these symplasmic fields with
metabolic and organogenetic activities, and field breakdown
during AM deactivation, provide evidence for a role of the
symplasm in coordinating cellular activities. Symplasmic fields
control the symplasmic diffusion of potential morphogens and
metabolites inside the AM and permit field interactions at their
boundaries, thereby mediating various types of cellular
collaborations. Given the generality of AM geometry and
function, symplasmic fields may canalize morphogenesis in all
higher plants.
We thank J. Lodders (Tree Nursery Lodders, Wernhout/Zundert,
The Netherlands) for gifts of birch seedlings; L. van der Plas
(Wageningen Agricultural University) for providing opportunities for
P. R. to carry out this work; R. W. Goldbach, J. van Lent and J.
Groenewegen (Wageningen Agricultural University) for use of TEM
facilities; J. Donkers (ATO-DLO, Wageningen) for assistance; the
Academy of Finland for financial support to P.R.
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