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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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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
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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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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;
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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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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
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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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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
