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Development 125, 339-349 (1998)
Printed in Great Britain © The Company of Biologists Limited 1998
DEV0146
Nod factor internalization and microtubular cytoskeleton changes occur
concomitantly during nodule differentiation in alfalfa
Antonius C. J. Timmers, Marie-Christine Auriac, Françoise de Billy and Georges Truchet*
Laboratoire de Biologie Moléculaire des Relations Plantes-Microorganismes, CNRS-INRA, BP 27, 31326 Castanet-Tolosan
Cédex, France
*Author for correspondence (e-mail: [email protected])
Accepted 20 November 1997: published on WWW 13 January 1998
SUMMARY
Reorganization of the plant cytoskeleton is thought to play
an important role during nodule ontogeny. In situ
immunolocalisation of tubulin reveals that important
cytoskeletal changes, implying a transient disorganization
followed by a newly patterned reorganization, occur in
indeterminate and determinate nodules. In alfalfa nodules,
cytoskeletal changes closely parallel the symbiotic
differentiation features related to cell infection, bacterial
release, endopolyploidization, cell enlargement, cell spatial
organization and organelle ultrastructure and positioning.
Moreover, the fact that microtubule disorganization can be
correlated with Nod factor internalization in central
infected cells suggests that Nod factors are possibly
involved in the control of cytoskeletal changes which direct
the differentiation of bacteria-containing cells.
INTRODUCTION
In legumes such as alfalfa and vetch, the nodule meristem
remains active for several weeks, thus leading to the formation
of elongated indeterminate nodules comprising central and
peripheral tissues. Histologically, central tissues are organized
into five well-defined zones: the apical meristematic zone I, the
prefixing (infection) zone II, the starch-rich interzone II-III, the
nitrogen-fixing zone III and the proximal senescent zone IV.
These central zones are surrounded by a parenchyma, vascular
bundles, an endodermis and a cortex (Vasse et al., 1990). The
differentiation of the central cells depends on whether the host
cells are invaded or remain bacteria-free. Fully differentiated
invaded cells are dramatically enlarged in size, highly
polyploid and filled with nitrogen-fixing bacteroids (Truchet,
1978; Vasse et al., 1990). In contrast, non-invaded cells are of
small size and their DNA content remains monosomatic
(Truchet, 1978). Different developmental features characterize
determinate nodules formed, for example, on the roots of
siratro or soybean. In such nodules, meristematic activity is
transient and the central cells differentiate almost
simultaneously (Newcomb, 1981). As a result, round-shaped
mature determinate nodules increase in size by cell
enlargement and all central cells are more or less at similar
stages of differentiation.
The plant cytoskeleton mediates several functions in living
cells (reviewed by Seagull, 1989; Cyr and Palevitz, 1995). The
cytoskeleton is dynamic in the sense that it continually
rearranges by virtue of its two major components,
microfilaments and microtubules which assemble and
disassemble in response to extracellular and intracellular
stimuli. This property can be correlated with fundamental
The symbiotic interactions between soil bacteria of the genera
Rhizobium, Azorhizobium or Bradyrhizobium (here referred
to as rhizobia) and plants of the Leguminosae family result in
the formation of nodules, new organs in which the bacteria
reduce nitrogen into ammonia which can subsequently be
utilized by the plant.
Nodule organogenesis starts with a molecular dialogue
between symbionts and takes place through a series of
developmental stages. Rhizobia produce Nod factors (NFs), the
synthesis of which is under the control of nodulation (nod)
genes which are transcribed in the presence of plant flavonoids.
Chemical studies have shown that NFs from all rhizobial
species are lipochitooligosaccharides consisting of a backbone
of N-acetylglucosamine residues which is decorated on its two
terminal residues (reviewed by Dénarié et al., 1996; Long,
1996; Schultze and Kondorosi, 1996; Spaink, 1996). Each
rhizobial species possesses a characteristic set of nod genes
that specifies the length of the backbone and the nature of the
decorations at both ends of the molecule, thus making the NFs
specific for a given plant host (Roche et al., 1991). NFs are
signal molecules involved in most of the early developmental
responses which are elicited by the corresponding bacteria.
Early rhizobia-dependent responses lead to various cellular
events such as root hair induction and deformations, plant
infection by means of tubular structures called infection
threads and the formation of a nodule meristem whose activity
ensures nodule growth (reviewed by Newcomb, 1981; Brewin,
1991; Roth and Stacey, 1991; Hirsch, 1992; Kijne, 1992).
Key words: Microtubular cytoskeleton, Nod factors, Nodule
differentiation, Rhizobium meliloti, Alfalfa
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A. C. J. Timmers and others
cellular traits that the cytoskeleton is thought to regulate, such
as, for example, cell division, cell shape and polarity, cell
trafficking and the spatial organization of the cytoplasm
(reviewed by Seagull, 1989; Goddard et al., 1994). An
involvement of the plant cytoskeleton during early stages of
nodulation has been suggested by studies showing that
cytoskeletal reorganizations occur at the tip of root hairs treated
with Nod factors (Allen et al., 1994) or in the root cortex of
Vicia hirsuta either infected by its specific symbiont
(Bakhuizen, 1988) or treated with Nod factors (Van Spronsen
et al., 1995). Moreover, if one considers the general roles of
the cytoskeleton in the context of symbiotic responses such as
root hair induction and deformation (Ardourel et al., 1994),
activation of cortical cells (Van Brussel et al., 1992; Ardourel
et al., 1994) which enter the cell cycle but arrest in G2 (Yang
et al., 1994), oriented growth of an infection network in plant
tissues (Bakhuizen, 1988; Ridge, 1992; Van Brussel et al.,
1992), formation of nodulation-related division centers
(Truchet et al., 1991) and the cell enlargement and DNA
endopolyploidization typical of invaded central cells (Truchet,
1978), then it is reasonable to anticipate that variations in the
cytoskeletal structure are likely to be involved in many of the
symbiosis-related steps directing nodule development.
Very little is known about the early molecular mechanisms
through which the host plant responds to rhizobial infection or
NF treatment. The current hypothetical model proposes that
NFs bind to plasmalemma-located receptors (Bono et al., 1995;
Niebel et al., 1997), followed by subsequent signal
transduction. Data indicate that Rhizobium meliloti NFs induce
a depolarization of the plasma membrane potential (Ehrhardt
et al., 1992; Felle et al., 1995), cytoskeletal changes (Allen et
al., 1994) and calcium spiking (Ehrhardt et al., 1996) in alfalfa
root hairs. In contrast, it has been shown that Rhizobium
leguminosarum bv. trifolii NFs are internalized specifically into
clover root hairs (Philip-Hollingsworth et al., 1997). In this
study, we show that the microtubular cytosketon (MC)
dramatically reorganizes in differentiating infected cells of
both indeterminate and determinate nodules. In alfalfa nodules,
MC changes initiate in the nodule zone where rhizobial Nod
factors are internalized in infected cells and strongly correlate
with symbiosis-specific cell differentiation traits. Thus, NF
internalization, MC changes and cell differentiation are tightly
coupled in the course of nodule development.
MATERIALS AND METHODS
Bacterial strains and plant assays
The list of bacterial strains and plant species used in this study is given
in Table 1. Respective bacterial and plant growth conditions were as
described by the references listed in Table 1. Spontaneous and NFinduced nodulation in alfalfa were achieved as described by Truchet
et al. (1989) and Truchet et al. (1991), respectively.
Microscopic methods
Nodules harvested at different developmental stages depending on the
nodulation type were processed for histological or ultrastructural
observations as described by Vasse et al. (1990).
Immunolocalization of microtubules
In situ visualization of the microtubular cytoskeleton was as follows.
Nodules were fixed in 4% formaldehyde (prepared from
paraformaldehyde) in microtubule stabilizing buffer (MSB) consisting
of 60 mM Pipes, 10 mM EGTA, 1 mM MgCl2, 0.1% Triton X-100 and
10% DMSO (pH 6.9, 30 minutes at room temperature) followed by a
subsequent 30 minute fixation in 4% formaldehyde in phosphatebuffered saline (PBS) pH 7.4. After rinsing with PBS, the samples were
infiltrated with sucrose up to 1 M in PBS, and cut into 8 and 50 µm
thick sections at −20°C, using a MICROM HM500 M cryostat.
Sections were deposited on poly-L-lysine-coated slides or in small
containers, and tubulin immunolocalized by subsequent incubation
with monoclonal anti-α-tubulin (Sigma T-5168) and an anti-mouse
IgG-FITC antibody (Boehringer Mannheim, 821462). After staining in
Evans blue (0.1% in PBS), to quench autofluorescence, the labeled
sections were mounted in glycerol containing 1% 1,4-diazabicyclo(2.2.2) octane as an anti-fading agent and 4,6-diamidino-2phenylindole as a nuclear stain and viewed by laser confocal
microscopy (Zeiss, LSM 410 Invert). Images were recorded either
from a single focal plane with an average thickness of 0.5 µm or as an
extended focus in which several confocal planes were superimposed.
The gain and offset were chosen in such a way that all the 255 grey
values were used resulting in 0 for the background (i.e. the slide
without section) and 255 for the maximum fluorescence in the
specimen. As a result of the chosen gain and offset, the scaling of the
pseudo-color look up table is comparable for all images. Images were
displayed, after background subtraction, as false colour images
indicating increasing fluorescence intensity on a colour scale ranging
from blue to red.
Immunolocalization of Nod factors
Rabbit anti-NF polyclonal antibody was prepared by Biocytex
(Marseille, France) using R. meliloti purified NodRm-IV(S,C16:2) as
immunogen. Immunserum was tested by ELISA according to standard
procedures (Engvall and Perlmann, 1971). For immunocytochemistry
several protocols were tested. Optimal results regarding labeling
efficiency and preservation of cellular structure were obtained by fixing
the specimen for 1 hour in 2% formaldehyde (prepared from
paraformaldehyde) and 0.5% glutaraldehyde in 0.12 M cacodylate pH
7.2, or 2.5% glutaraldehyde in 0.2 M cacodylate pH 7.2 respectively
for light and electron microscopy. After subsequent dehydration with
ethanol and infiltration with LR white resin, polymerization was
performed at low temperature under UV irradiation. For labeling, 1 µm
semithin sections stuck onto poly-L-lysine-coated slides or ultrathin
sections picked up on nickel grids covered with formvar were first
incubated overnight at 4°C in 10% goat serum, 0.5% Triton X-100,
0.5% Tween 20 in Tris-buffered saline (TBS) to prevent non-specific
binding of antibodies. For immunolocalization at the histological level,
sections were incubated either overnight at 4°C or for 2 hours at 37°C
with anti-NF immunserum diluted 1:50 in TBS. After rinsing in TBS,
incubation was continued with anti-rabbit IgG antibody:1 nm gold
(BioCell) for 1 hour at room temperature. The signal was amplified by
using the BioCell silver enhancing kit. Sections were then stained
briefly in 0.02% toluidine blue, mounted in DPX (BDH Laboratories
Supplies) and observed by bright field or dark field microscopy with an
Olympus Vanox light microscope. For immunolabeling at the
ultrastructural level, sections were incubated as described above with
the immunserum diluted 1:10 and an anti-rabbit IgG:20 nm gold was
used as secondary antibody. Ultrathin sections were stained with uranyl
acetate and lead citrate and observed with a Hitachi H600 transmission
electron microscope.
Quantification of anti-Nod factor labeling intensity
The intensity of NF immunolabeling in different nodule zones was
quantified by silver grain counting on labeled 1 µm thick semithin
sections. The region of interest was identified under bright field optics
at 100× magnification and outlined on the screen. The quantity of light
reflected by each area was then measured under fluorescent epiillumination using a computer-based image analysis system according
to the procedure described by Blanchard et al. (1993) and converted
NF internalization and cytoskeletal changes in nodules
341
Table 1. Plants and bacterial strains used in this study
Plants
Medicago sativa
M. sativa
M. sativa
M. sativa
M. sativa
Trifolium repens
Vicia sativa
Macroptilium atropurpureum
Bacteria
Relevant characteristics*
Rhizobium meliloti RCR 2011
R. meliloti GMI6390, Nod factor overproducing strain
R. meliloti GMI6371, Nod factor non producing strain
R. meliloti fixK− GMI942
R. meliloti exoA−
R. leguminosarum bv trifolii ANU843
R. leguminosarum bv viciae 248
Rhizobium sp. NGR234
WT, IN
WT, IN
Nod−, nodA−
Fix−, IN
Fix−, IN
WT, IN
WT, IN
WT, DN
References
Ardourel et al. 1994
Roche et al. 1991
Roche et al. 1991
Foussard et al. 1997
Leigh et al. 1985
Djordjevic et al. 1985
Van Brussel et al. 1986
Lewin et al. 1990
*DN, determinate nodule; Fix−, non-fixing nodule; IN, indeterminate nodule; WT, wild type.
into the number of grains per square µm. Background grain density
measured on the resin but out of the section was finally subtracted
from the grain density of each region to obtain the density per zone
due to NF immunolabeling.
RESULTS
orientated perpendicular to the cell wall, whilst those in the center
remain randomly distributed within the cytoplasm (Fig. 1C).
The positioning and the ultrastructural differentiation of cell
organelles and of bacteroids were studied in more detail by
electron microscopy. Both plastids (as undifferentiated
proplastids) and rod-shaped mitochondria were found to be
randomly distributed within the plant cytoplasm of bacteria-free
cells observed either in the meristematic zone I or in the most
distal border of zone II where the infection network develops
(Fig. 2A). Progressive changes in ultrastructure and positioning
of organelles occur after the release of bacteria along the
prefixing zone II (Fig. 2B). First proplastids, and then
mitochondria increase in size (Fig. 2B) and in length (Fig. 2C,D)
and move to the periphery of the cytoplasm where they orientate
parallel to, and in close contact with, the plasmalemma,
particularly at intercellular spaces (Fig. 2E). Very short
microtubules are often seen in the limited cytoplasmic space
Variations in shape, cytoplasmic organization and
ultrastructural differentiation in invaded cells of
alfalfa nodules
Speculating that cellular features occurring during nodule
growth might reflect cytoskeletal changes (see Introduction),
we first studied, in detail, the histological and ultrastructural
differentiation and the cytoplasmic organization of central cells
in alfalfa nodules. The terminology used in this study to
describe the histological organization of alfalfa nodules and the
ultrastructural stages in
bacteroid differentiation is
as described by Vasse et al.
(1990).
Invaded cells enlarge
isodiametrically in the distal
part of zone II and become
round shaped in proximal
zone II, interzone II-III and
zone III (Fig. 1A-C).
Simultaneously,
the
clustering of cell organelles
(plastids and mitochondria)
and bacteroids at the cell
periphery, shows that a
modification
in
spatial
organization takes place in
zone II, particularly in the
proximal part of this zone
(Fig. 1B). Interestingly, as
cells mature, the spatial
positioning of bacteroids
Fig. 1. Cell morphology and bacteroid positioning in alfalfa nodules. Bright-field microscopy.
changes from a random
(A) Longitudinal semithin section of a 3 week-old wild-type nodule showing the meristematic zone I (small
orientation in the cells of the
asterisk), the prefixing zone II (small star), the interzone II-III (large asterisk) and the nitogen-fixing zone
prefixing zone II (Fig. 1B,C),
III (large star). Bar, 50 µm. (B) Changes in morphology of the invaded cells in zone II. Cells are small and
to a precise positioning
isodiametric in the distal part (asterisk) and enlarge to become round-shaped in the proximal region (star).
which depends on bacteroid
The clustering of bacteroids (blue dots) at the cell periphery, and vacuolar parceling are seen. Bar, 25 µm.
location in the interzone II-III
(C) Bacteroid positioning in the cytoplasm of invaded cells. Bacteroids are randomly orientated in proximal
and zone III. In the two latter
zone II (small asterisk). In interzone II-III (large asterisk) and zone III (star), the outermost bacteroids are
zones, the bacteroids located
orientated perpendicular to the cell wall, while those in a central location remain randomly distributed. Note
the increase in bacteroid elongation at the interface zone II-interzone II-III. Bar, 100 µm.
at the cell periphery are
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A. C. J. Timmers and others
between the organelles and the plasmalemma (Fig. 2F). In a
similar way, actively dividing bacteroids (type I bacteroids;
Vasse et al., 1990) and bacteroids which have ceased to divide
and have started to differentiate (type II bacteroids; Vasse et al,
1990), also migrate to the cell periphery where they were seen
randomly orientated in the cytoplasm (Fig. 2C,E).
Fig. 2. Ultrastructure and
positioning of cell organelles
and bacteroids in invaded cells
of zone II. Transmission
electron microscopy. (A) Cells
at the distal border of zone II
in which cell organelles are
randomly distributed. The
arrowhead points to an
infection thread. Bar, 5 µm.
(B) Bacterial release of
rhizobia at the unwalled
extremity of an infection
thread. Remnants of the thread
cell wall (arrows), released
bacteroids (arrowheads) and
enlarging plastids (white star)
are seen. Bar, 2.5 µm.
(C) Distal zone II. Progressive
clustering of plastids (large
arrows), mitochondria (small
arrows) and type 2 bacteroids
(lower cell) at the cell
periphery. Note the random
positioning of type 1
bacteroids in the upper cell.
Bar, 5 µm. (D) Detail of a cell
in distal zone II, showing cell
organelle and bacteroid
clustering at the periphery of
the cell and abundant
endoplasmic reticulum profiles
(arrowheads) in the cytoplasm.
Bar, 1 µm. (E) Cell organelles
in proximal zone II. Note the
elongation of plastids (arrows)
parallel to and in close contact
with the cell wall and
mitochondria (arrowheads)
applied against plastids. Bar, 1
µm. (F) Microtubules
(arrowheads) at the interface
between a plastid (white
asterik) and the cell wall (star).
Bar, 1 µm.
Changes continued in interzone II-III. Here, we noted the
exceptional elongation of both mitochondria and plastids, which
accumulate very large starch granules in their stroma (Fig. 3A;
see Vasse et al., 1990) and we confirmed our observation, made
by light microscopy, that the spatial positioning of type III and
type IV bacteroids (in interzone II-III and zone III, respectively;
NF internalization and cytoskeletal changes in nodules
343
Major microtubular cytoskeleton changes are
observed during cell differentiation in alfalfa
nodules
Immunolocalization of MC in alfalfa enabled the
differentiation features described above to be correlated with
variations in cytoskeletal structure. As expected, meristematic
cells fluoresced strongly, while a loss in staining due to the
progressive disorganization of both endoplasmic and cortical
microtubules was observed in cell layers of the prefixing zone
II (Fig. 4A-C). Such a MC disorganization continues
throughout the entire zone II to the point where only very low
level staining can be detected at the periphery of the most
proximal cells of this zone (Fig. 4D).
A signal is suddenly recovered at the interzone II-III, a zone
which differentiates from cell to cell in indeterminate nodules
(Vasse et al, 1990). In this zone, a fluorescent band underlining
the cell periphery without interruption (Fig. 4D), indicates that
a cortical cytoskeleton has reformed. Moreover, short radial
microtubular appendages are seen, which then develop
centripetally from the cell wall into the cytoplasm of each
invaded cell. As cells mature, the number and the length of
these appendages increase significantly. In the cells of zone III,
radial appendages are orientated in the cell cytoplasm like the
spokes of a bicycle wheel (Fig. 4E), i.e. parallel to the
individual outermost bacteroids (compare Figs 1C, 3A and 4E).
These appendages which most likely correspond to those
observed in close contact with symbiosomes (see Fig. 3B), do
not extend to the central region of the invaded cells where
bacteroids are randomly oriented in the cytoplasm (compare
Figs 1C and 4E). Such an organization of the endoplasmic MC
is progressively lost in the most proximal part of the nodule,
where a new decrease in fluorescence is observed (Fig. 4F). It
is worth mentioning that (i) MC changes described above are
not related to the nitrogen-fixing capacity of nodules since
similar variations were observed in equivalent zones of Fix−
alfalfa nodules elicited by a R. meliloti fixK mutant (Vasse et
al., 1990; not shown), and (ii) no labeling was seen in controls
where the primary antibody was omitted, thus confirming the
specificity of the labeling (not shown).
Fig. 3. Ultrastructure and positioning of cell organelles and
bacteroids in invaded cells of zone III. Transmission electron
microscopy. (A) Part of a nitrogen-fixing cell in zone III. Positioning
of type IV bacteroids perpendicular to the cell wall. Bar, 5 µm.
(B) Microtubules (arrowheads) in contact with a type IV bacteroid
sectioned tangentially at one pole. Bar, 1 µm. (C) Proximal
inefficient zone III. Small-sized plastids (arrows) and mitochondria
(arrowheads) at the cell periphery. The star indicates the section of an
infection thread. Bar, 1 µm.
Vasse et al., 1990), indeed depends on their location (Fig. 3A,
compare with Fig. 1C). Interestingly, the positioning of the
outermost bacteroids often correlated with the presence of
microtubules orientated parallel to, and often in close contact
with, symbiosomes containing nitrogen-fixing type IV
bacteroids (Fig. 3B). Such a distribution is progressively lost in
proximal zone III (not shown), a nodule region where both
plastids and mitochondria gradually lose their elongated form
and return to being rod-shaped (Fig. 3C).
Microtubular cytoskeleton changes are common
traits of nodule differentiation and are rhizobiarelated
To examine whether MC changes are general traits of nodule
differentiation, we studied MC in various indeterminate and
determinate nodules. Changes identical to those described
above in alfalfa, i.e. transient microtubule depolymerization
followed by a newly patterned reorganization, were observed
in clover and vetch indeterminate nodules (not shown). A
progressive MC disorganization was also seen in developing
determinate siratro nodules (6-10 days after inoculation) from
the outer meristematic cells, which fluoresced strongly, to the
cells of the central region where labeling could hardly be
detected (Fig. 4G). In this legume, MC reformed in the
nitrogen-fixing cells of fully differentiated nodules. However
the signal was restricted to the cell periphery (Fig. 4H),
indicating that the cortical cytoskeleton, but not the
endoplasmic cytoskeleton, reorganizes in determinate
nodules.
To determine whether cytoskeletal changes are rhizobiarelated, we studied the cytoskeleton in bacteria-free nodules
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A. C. J. Timmers and others
Fig. 4. Microtubular cytoskeletal rearrangements in nodules. Laser confocal microscopy. Fig. 4E, F and G are extended focus compositions of
10 sections each separated by 1 µm. The others are single focal plane images. (A-F) Three-week old alfalfa nodules. (A) Longitudinal section
showing a significant decrease in signal in zone II (asterisk). Bar, 25 µm. (B) Decrease in labeling in the distal part of a nodule, from
meristematic zone I (star) to the proximal part of zone II (asterisk). Arrows indicate the middle region of zone II. Bar, 50 µm. (C) Faint staining
at the periphery of the cells in proximal zone II. Bar, 25 µm. (D) Signal at the transition between proximal zone II (asterisk) and interzone II-III
(star). Bar, 25 µm. (E) Newly formed microtubules which radiate from the cell cortex to the cytoplasm in nitrogen-fixing cells of zone III. Bar,
20 µm. (F) Loss in fluorescence in the proximal cells of zone III. Bar, 25 µm. (G) and (H) Siratro nodules. (G) Eight-day old nodule. Decrease
in fluorescence with a centripetal gradient. Bar, 100 µm. (H) Three-week old nodule. Labeling restricted to the cell periphery. Bar, 25 µm.
(I) Two-week old alfalfa nodule elicited by a R. meliloti exoA mutant. A fluorescent signal is observed both in the meristematic region (star) and
in the central uniform tissue (asterisk). Bar, 50 µm.
induced either by a R. meliloti exoA mutant (Leigh et al., 1985)
or by NFs purified from R. meliloti (Truchet et al., 1991) or
following spontaneous development (NAR nodules; Truchet et
al., 1989). All these nodule types have a uniform central tissue
made of bacteria-free cells. In contrast to wild-type nodules, it
was found that a predominantly cortical MC remained present
NF internalization and cytoskeletal changes in nodules
345
Fig. 5. Nod factor immunolocalization in alfalfa nodules. (A,E) Dark-field microscopy; (B-D,F) bright-field microscopy. (A,B) Same
longitudinal section of a 2-week-old alfalfa nodule. NF internalization (bright area in A; dark area in B) in zone II. Note the decrease in signal
from prefixing zone II (asterisks) to nitrogen-fixing zone III (stars). The arrows point to infection threads. Aspecific labeling is seen on the
nodule endodermis (large arrow) and amyloplasts (arrowheads). Bars, 10 µm. (C) Labeling in the invaded cells of zone II. The arrows point to
infection threads. Bar, 25 µm. (D) NF immunolocalization on bacteroids (arrowheads) and in the vacuole (star) of a cell in proximal zone III.
Bar, 100 µm. (E) Serial section to A and B. Control assay using preimmune serum. Aspecific labeling on nodule endodermis (arrow) and
amyloplasts (arrowheads). Bar, 10 µm. (F) Serial section to (A and B). Control assay using NF-adsorbed immunserum. Note the absence of
labeling on infection threads (arrows) and in zone II (asterisk). Bar, 10 µm.
throughout the differentiation of these three nodule types,
indicating that MC changes require the presence of bacteria in
nodules (Fig. 4I).
Nod factors are immunolocalized in alfalfa nodules
Our observation that MC changes are rhizobia-dependent (see
above) together with results showing that nod genes are
transcribed in rhizobia that are still enclosed in infection
threads (Schlaman et al., 1991), prompted us to examine
whether MC changes could be correlated with the presence of
NFs in central nodule cells. Therefore, rabbit polyclonal
antibodies directed against purified NFs of R. meliloti were
prepared and their reactivity verified by ELISA. Briefly, we
found that coated NFs reacted with antiserum in a time- and
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A. C. J. Timmers and others
0.09
0.08
Average grain density per µm2
0.07
0.06
0.05
0.04
0.03
0.02
0.01
0
Zone I
Zone II
Zone III
Nodule zones
Fig. 6. Nod factor quantification. Cytoplasmic silver grain density in
different central zones of alfalfa nodules after NF
immunolocalization. Average values with standard errors obtained
from five nodules (3-weeks old). The total measured areas were 1.57
mm2, 2.48 mm2 and 1.55 mm2 for zones I, II and III, respectively.
concentration-dependent manner (data not shown). Out of four
sera tested, the one which gave the highest positive signal with
a concentration of NFs as low as 10−9 M, was used for the
experiments. We then controlled the specificity of the immune
serum by inoculating axenic plants with R. meliloti strains
known either to overproduce NFs (strain GMI6390, Roche et
al., 1991), or to be unable to synthesize NFs (strain GMI6371,
Roche et al., 1991). Immunostaining of plants 2 days postinoculation showed that the NF-producing strain fluoresced
strongly while no signal was detected on alfalfa inoculated
with rhizobia which do not synthesize NFs (not shown). This
result indicated clearly that the antibodies were specific for
NFs and did not immunoreact with plant and bacterial surface
components.
NFs were immunolocalized in wild-type alfalfa nodules
elicited by the wild type R. meliloti RCR 2011 strain. By light
microscopy, the strongest signal was observed in prefixing zone
II, particularly associated with the infection threads, while a
lower, but still significant labeling, was detected in the
cytoplasm of the invaded cells (Fig. 5A-C). In contrast, low
levels of staining were found in interzone II-III and zone III
(Fig. 5A and 5B). This variation was confirmed by quantifying
the density of gold particles per surface unit (µm2), in
conditions where the highly reactive infection threads and large
plastids were excluded from the scanned regions. We found in
prefixing zone II, a higher density than in meristematic zone I
and distal nitrogen-fixing zone III, respectively (Fig. 6).
Detailed observations showed that NFs are internalized in the
cytoplasm of the infected cells in zone II (Fig. 5C). There is
also a constant, albeit discrete, level of immunolabeling
associated with bacteroids at all stages of their differentiation
(Fig. 5D). The labeling of bacteroids might account for the
slight, but significant (t=2.8, P<0.025), increase in gold particle
density detected in the cells of zone III as compared to the
meristematic cells (Fig. 6). Moreover, a signal was also
observed in the large vacuoles of the most proximal cells of
zone III (Fig. 5D). This could be due to an accumulation in the
vacuole of NFs released from bacteroids which undergo a
degeneration process in the vacuole of infected cells in this
region of the nodule (Truchet and Coulomb, 1973). Finally, and
unexpectedly, labeling was seen on starch granules
accumulating in amyloplasts and on the cell wall of endodermal
cells (Fig. 5A and 5B). This latter labeling, which varied
strongly between assays, could be due either to the recognition
by the immune serum of epitopes localised at these sites and/or
to non-specific labeling. Comparative studies between assays
and controls omitting the primary antibodies (not shown), using
the preimmune serum (Fig. 5E, compare with 5A), or where
immune serum was exhaustively adsorbed with purified NFs
prior to the staining procedure (Fig. 5F, compare with 5B) left
open the two possibilites. However, in contrast to the labeling
in infection threads and cell cytoplasm in zone II, it seems very
unlikely that labeling on the endodermis accounts for the
presence of NFs, since this tissue is totally bacteria-free (Vasse
et al., 1990). NF immunolocalization was confirmed at the
ultrastructural level (not shown).
DISCUSSION
In this paper, we provide experimental evidence of NF
internalization and MC architectural rearrangements during
nodule differentiation. That similar changes are observed in
different nodule types and depend on bacterial release indicate
that cytoskeleton changes are symbiosis-specific and rhizobiadependent. Our results lead to several conclusions and
hypotheses concerning the general role that the cytoskeleton
plays in mediating plant cell differentiation and suggest that
NF internalization and MC changes are correlated during
nodule maturation.
The cytoskeleton in relation to nodule infection and
bacterial release in host cells
In indeterminate nodules, MC changes occur in nodule central
zones in which at least six major cellular changes are observed
microscopically. The first cellular event which determines the
individualization of the prefixing zone II is the penetration of
meristem-derived cells by the growing part of the infection
network. A number of studies either provide direct evidence or
imply that the cytoskeleton is involved in infection thread
growth: (i) microtubules are present at the original site of
penetration (Ridge and Rolfe, 1985); (ii) microtubules
interconnect the host cell nucleus to the growing infection
thread tip (Bakhuizen, 1988); (iii) cytoskeleton-determined
cytoplasmic bridges guide the growth of infection threads on
their way down to the nodule primordium (Van Brussel et al.,
1992; Bakhuizen, 1988); (iv) the cytoskeleton plays a major
role in cell wall formation (reviewed by Seagull, 1989; Cyr,
1994), and infection threads are cell wall limited structures
(Callaham and Torrey, 1981) and (v) an interesting model has
been proposed in which Rhizobium infection through root
hairs may result from the mobilization of normal root hair tip
growth machinery, including the cytoskeleton, at the infection
site (Ridge, 1992). Taken together, these and our data indicate
that cytoskeletal integrity and functions are sufficiently
preserved to sustain infection thread growth at the meristemprefixing zone II interface of developing nodules.
The second cellular event is the release of bacteria in the cell
NF internalization and cytoskeletal changes in nodules
cytoplasm which is restricted to the distal part of prefixing zone
II (Vasse et al., 1990). The signals that trigger bacterial release
are not known. However, the fact that release occurs exclusively
at unwalled parts of infection threads (Roth and Stacey, 1989),
has suggested the possibility of an enzymatic degradation of the
thread cell wall either by enclosed rhizobia (Truchet and
Coulomb, 1973) or by a mechanism which would involve the
mobilization of the endoplasmic reticulum (Roth and Stacey,
1991) which is particularly abundant in the region of bacterial
release (Truchet and Coulomb, 1973; this study). Our results
open up another possibility. We hypothesize that the release
might also be a consequence of discrete and local variations of
the endoplasmic cytoskeleton in the direct vicinity of the
growing tip of infection threads, i.e. at sites where the developing
cell wall is not yet completely organized. As a consequence,
some of the cytoskeleton-controlled functions, such as cell wall
building (for example), could be locally modified, or slackened,
to an extent and for a period of time sufficient to allow bacterial
release through an opened door at infection extremities.
The cytoskeleton in relation to cell morphology and
endopolyploidy
The third cellular event observed in prefixing zone II is that,
once invaded, host cells enlarge isodiametrically in distal zone
II, leading to a round-shaped morphology in the proximal cells
of this zone. The cytoskeleton plays an active role in cell
elongation and cell shape control by directing the ordered
synthesis of cellulose microfibrils via a microtubule/plasma
membrane/cell wall functional continuum (reviewed by
Seagull, 1989; Giddings and Staehelin, 1991; Williamson,
1991; Cyr, 1994). Thus the isodiametric enlargement of the
invaded cells in the distal part of zone II, should indicate that,
in this region, the cortical array maintains its function in cell
wall building necessary for cell enlargement. In contrast, the
changes in both cell shape and morphology in zone II and the
arrest in cell enlargement occurring in the central nodule
tissues (this study) might result from the progressive lost in
cytoskeletal integrity observed in the infected nodule cells. It
might seem contradictory to hypothesize that in distal cells of
zone II the cytoskeleton still functions efficiently in plant cell
wall organization whilst infection thread cell wall formation is
impaired. However, as stated above, we propose that local
changes in the cytoskeleleton centered around the extremity of
an infection branch and which are not discernable by
microscopy, locally modify cytoskeleton functioning.
The fourth cellular event is that simultaneously to cell
enlargement, endopolyploidy takes place gradually and
exclusively in the invaded cells of prefixing zone II in
indeterminate pea and alfalfa nodules (Truchet, 1978). The
cytoskeleton is subjected to a series of specific rearrangements
that are essential for karyokinesis and cytokinesis to proceed
successively (reviewed by Seagull, 1989; Goddard et al.,
1994). The fact that, in nodules, the impairment of these two
cellular events occurs simultaneously to cytoskeletal
disorganization raises the question of the mechanisms that are
responsible for DNA amplification. Particularly relevant to our
study are the results showing that drug-induced microtubule
depolymerization is sufficient to initiate both DNA synthesis
and the entry of human and animal fibroblast-like cells into the
proliferative cycle (Crossin and Carney, 1981; Bershadsky et
al., 1996) and that microtubule depolymerization is directly
347
responsible for DNA synthesis (Crossin and Carney, 1981). It
would be interesting to determine if a similar mechanism is
involved in the establishment of the polyploid gradient in the
prefixing zone II of indeterminate nodules.
The cytoskeleton in relation to cytoplasmic spatial
organization and ultrastructural differentiation
The fifth cellular event concerns the major variations in the
spatial distribution and the positioning of both organelles and
bacteroids in the infected cells. The capacity of the cytoskeleton
to mediate the movement and distribution of organelles in a cell
is supported by several correlative studies (reviewed by Cole and
Lippincott-Schwartz, 1995). For example, the distribution of
organelles frequently parallel that of the cytoskeleton and the use
of drugs which depolymerise microtubules or microfilaments
results in modified spatial distribution of organelles (Yaffe et al.,
1996). Our data are in agreement with these studies: the random
distribution of cell organelles and bacteroids in the most distal
cells of zone II is associated with an organized cytoskeleton; the
progressive migration of organelles and bacteroids to the
periphery of the cells in proximal zone II occurs simultaneously
to endoplasmic MC disorganization. Moreover, our study
suggests that the positioning of both cell organelles and
bacteroids is microtubule-mediated. Thus, in zone III, the
peculiar positioning of the outermost bacteroids, perpendicular
to the cell wall, parallels that of radial microtubules originating
from the cortical cytoskeleton and radiating centripetally to inner
regions of the cytoplasm. In plants, a radial cytoskeleton pattern
has been described in meristematic and elongating cells
(Bakhuizen et al., 1985; Baluska et al., 1992), and in the
syncytial stage during endosperm development (Brown et al.,
1994). Generally, radial patterns are thought to originate from
the nuclear periphery (Lambert, 1993) and to provide an anchor
for the nucleus at its appropriate position during cell
differentiation. Similar functions for the radial cytoskeleton in
nitrogen-fixing cells are unlikely. In nodules, radial arrays
obviously originate in the cell cortex, then elongate into the cell
cytoplasm as cells mature and are not connected to other
structures in the cytoplasm. Thus, we provide further evidence
for the presence of microtubule organizing centers (MTOCs) in
the cortex of plant cells (Liu et al., 1994; Panteris et al., 1995).
The situation is different in mature determinate nodules in which
the random distribution of large symbiosomes containing many
bacteroids in the cell cytoplasm (Newcomb, 1981) is in
agreement with our data showing that a cytoplasmic
cytoskeleton does not reform in the invaded cells of nitrogenfixing siratro nodules.
The sixth cellular event concerns the ultrastructural changes
which characterize cell organelles and bacteroids in invaded cells.
By correlating the present data with our previous results on
bacteroid differentiation in alfalfa nodules (Vasse et al., 1990), it
clearly appears that bacteroids (i) divide exclusively in the cells
of distal zone II, where the MC is microscopically observed, (ii)
stop dividing and elongate moderately in the second half of the
proximal zone II where the MC disorganizes and (iii) increase
dramatically in size in a cell to cell fashion (Vasse et al., 1990),
correlating with the differentiation of the interzone II-III where
cortical MC reforms. Interestingly, we found that cell organelles
undergo similar variations in the same regions of the nodule,
being small-sized in distal zone II, increasing moderately in size
in proximal zone II and reaching their maximal size in interzone
348
A. C. J. Timmers and others
II-III, before returning to a smaller size in zone III. The
mechanisms which control the division of cell organelles are
poorly understood, although it is generally admitted that there is
no link between the mechanism that divides the mother cell and
the dividing organelle (reviewed by Warren and Wickner, 1996).
Despite the difficulty in deducing a dynamic relationship simply
from morphological features, our results strongly suggest that the
MC plays a role in bacteroid and cell organelle division and
elongation. This hypothesis implies that bacteroids are perceived
as endogenous organelles. Microtubules mediate the distribution
of cell organelles via membrane-binding microtubule-based
motor proteins such as kinesin and cytoplasmic dynein (Yaffe et
al., 1996). As bacteroids are enclosed by a plasmalemma-derived
peribacteroid membrane (Roth and Stacey, 1989), we anticipate
that the host cell senses intracellular bacteroids as normal cell
organelles via the connection of cytoskeleton microtubular
elements and associated proteins with the peribacteroid
membrane.
Nod factor internalization and cytoskeletal changes
In this paper, we show that NFs are immunolocalized throughout
the central tissues of alfalfa nodules from the prefixing zone II
to the proximal part of zone III. The fact that the strongest signal
is observed in infection threads is in agreement with previous
studies indicating that nod genes are expressed in bacteria
enclosed in infection threads (Schlaman et al., 1991; F. Maillet,
G. Truchet and J. Dénarié, unpublished data). Moreover, in zone
II, a specific signal is also detected in the cytoplasm of the
infected plant cells indicating that NFs are internalized. Finally,
a weak but reproducible labeling is associated with bacteroids,
independent of their differentiation stage and histological
location. This result is in contradiction with previous data
indicating that in vetch and alfalfa nodules, nod genes are not
expressed in intracellular bacteroids (Schlaman et al., 1991; F.
Maillet, G. Truchet and J. Dénarié, unpublished data) and
suggests that NFs are synthesized at a low level by bacteroids.
Our data and the recent results showing that R.
leguminosarum bv. trifolii NFs are internalized in clover root
hairs (Philip-Hollingsworth et al., 1997) indicate that NF
internalization can take place in different tissues amenable to
infection. At the moment, we do not know how NF
internalization occurs. It might involve a simple diffusion
process, or be receptor-mediated. Putative high-affinity NF
receptors could be located in the plasma membrane (Bono et al.,
1995; Niebel et al., 1997), a membrane which also surrounds
infection threads and bacteroids (Hirsch, 1992; Kijne, 1992; see
Fig. 2B). The possible coupling of NF reception and
internalization is also supported by the recent report showing
that purified R. leguminosarum bv. trifolii NFs are internalized
into the root hairs of clover, the homologous host, but not of
alfalfa, an heterologous host for this species (PhilipHollingsworth et al., 1997). Finally, our observation that NFs are
only internalized in the distal region of alfalfa nodules whilst
they are synthesized by bacteroids throughout their entire
differentiation process, suggests that the mechanism(s) directing
NF internalization change(s) during nodule maturation.
Identifying the nature of the causal mechanism(s) resulting
in cytoskeletal disorganization is a major challenge raised by
this study. Our data suggest that NF internalization and MC
disorganization are coupled events. Firstly, NFs are
immunolocalized in the cytoplasm of invaded cells of the
prefixing zone II, where the MC disorganizes, but not in the
cytoplasm of cells in nitrogen-fixing zone III, where the
cytoskeleton reforms. This result highlights the spatiotemporal
correlation between the two processes. Secondly, the MC is not
disorganized in bacteria-free alfalfa nodules indicating that MC
disorganization requires the presence of bacteria. Thirdly,
identical MC disorganization occurs in nodules of different
types induced by rhizobial strains which are taxonomically
distant (Martinez-Romero and Caballero-Mellado, 1996), but
share the capacity to produce NFs belonging to the same
chemical family and displaying similar biological activities
(reviewed by Dénarié et al., 1996; Long, 1996; Schultze and
Kondorosi, 1996; Spaink, 1996). From these considerations, it
can reasonably be hypothesized that the causal mechanism(s)
responsible for cytoskeletal disorganization in nodules is
common to all rhizobia and that NFs, or a structural component
of NFs, are (is) this common link.
In summary, this paper reports several correlations between
NF internalization, MC changes and the particular
differentiation of the invaded central cells in alfalfa nodules.
We propose that, once internalized, rhizobial NFs are involved
in cytoskeletal changes accompanying cell differentiation. A
validation of this model requires a detailed study of the direct
effects of purified NFs on cytoskeletal architecture.
We are very grateful to Fabienne Maillet and Jean Dénarié for
providing purified R. meliloti Nod factors, Philippe Cochard for advice
in using laser confocal microscopy and Michel Petitprez for help in gold
label quantification. We thank very much D. Barker, P. Boistard and J.
Dénarié for their constructive comments regarding the manuscript and
C. Gough and D. Barker for English reviewing. A. C. J. T. was supported
by BIOTECH and TMR postdoctoral fellowships and this work was
funded by grants from the European Community BIOTECH Programme
(PTP CT93-0400) and TMR Programme (FMRX-CT96-0039).
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