3 Department of Plant Systems Biology, Flanders Interuniversity Institute for Biotechnology, Ghent University, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium
Accepted on March 27, 2002;
Abstract
The onset of nodule development, the result of rhizobialegume symbioses, is determined by the exchange of chemical compounds between microsymbiont and leguminous host plant. Lipo-chitooligosaccharidic nodulation (Nod) factors, secreted by rhizobia, belong to these signal molecules. Nod factors consist of an acylated chitin oligomeric backbone with various substitutions at the (non)reducing-terminal and/or nonterminal residues. They induce the formation and deformation of root hairs, intra- and extracellular alkalinization, membrane potential depolarization, changes in ion fluxes, early nodulin gene expression, and formation of nodule primordia. Nod factors play a key role during nodule initiation and act at nano- to picomolar concentrations. A correct chemical structure is required for induction of a particular plant response, suggesting that Nod factorreceptor interaction(s) precede(s) a Nod factor-induced signal transduction cascade. Current data on Nod factor structures and Nod factor-induced responses are highlighted as well as recent advances in the characterization of proteins, possibly involved in recognition of Nod factors by the host plant.
Key words: legume nodulation/lipo-chitooligosaccharide/Nod factor receptor/signal transduction/three-dimensional Nod factor structure
Introduction
Nodulation (Nod) factors are key signal molecules that play a pivotal role during initiation of nodule development and bacterial invasion (Broughton et al., 2000; Perret et al., 2000
). They are produced by rhizobia, including the genera Allorhizobium, Azorhizobium, Bradyrhizobium, Mesorhizobium, Rhizobium, and Sinorhizobium. The recent discovery of nodulating Methylobacterium sp. (Sy et al., 2001
) and Burkholderia sp. (Moulin et al., 2001
) that belong to the
- and ß-subclass of the Proteobacteria, respectively, calls for expansion of the rhizobia with these latter bacterial genera. Rhizobia nodulate specific leguminous host plants and the nonlegume Parasponia. Such symbioses result in the formation of root nodules, new organs occupied by differentiated bacteria, that fix atmospheric nitrogen and provide it to their respective host plant, thereby promoting plant growth independently of the available soil nitrogen. Mature nodules are either of the determinate or indeterminate type (Crespi and Gálvez, 2000
). Determinate nodules are formed on some tropical and subtropical legumes (e.g., soybean, bean) and are characterized by a round-shaped appearance, initiation of nodule primordia in the outer cortex, and meristematic activity that disappears early after nodule initiation. Oval-shaped, indeterminate nodules usually form on roots of temperate legumes (e.g., pea, alfalfa, vetch), nodule primordia initiate in the inner cortex, the meristematic activity is persistent, and the central tissue consists of a number of distinct zones (Crespi and Gálvez, 2000
). Medicago truncatula (Bell et al., 2001
) and Lotus japonicus (Kawasaki and Murakami, 2000
) are now considered the best model legumes. Their genomes are being sequenced to efficiently determine plant responses occurring during all stages of nodule development.
Nod factors consist of an oligomeric backbone of ß-1,4-linked N-acetyl-D-glucosaminyl residues, N-acylated at the nonreducing-terminal residue (Kamst et al., 1998) and thus are lipo-chitooligosaccharides (LCOs) (Dénarié et al., 1996
). Rhizobia synthesize populations of Nod factors that consist of two (in the case of Rhizobium etli CFN42; Poupot et al., 1995
), to approximately 60 (in the case of R. galegae HAMBI1207; Yang et al., 1999
) different individuals. Qualitative and quantitative aspects of Nod factor populations are strain-specific. The Nod factor structure differs in the number of GlcNAc residues present in the chitooligosaccharide backbone, in the nature of the fatty acyl group, and in the substituents at the nonreducing- and/or reducing-terminal residues. In a few cases, Nod factor substituents are found at nonterminal GlcNAc residues. Nod factor synthesis depends on the expression of nodulation (nod) genes, comprising the nod, nol, and noe genes. Recently, orthologs of the nodA gene, one of the key nod genes encoding an acyl transferase (Kamst et al., 1998
), have been discovered in symbiotic Methylobacterium sp. (Sy et al., 2001
) and Burkholderia sp. (Moulin et al., 2001
).
Major Nod factortriggered responses include the formation and deformation of root hairs, intra- and extracellular alkalinization, membrane potential depolarization, changes in ion fluxes, induction of early nodulin gene expression, and formation of nodule primordia (Broughton et al., 2000; Perret et al., 2000
). Detailed structure analyses of Nod factor populations produced by a variety of rhizobia and phenotypic studies with mutant rhizobia demonstrated the importance of Nod factor structures for causing particular responses. Nod factors act in concentrations as low as 109 to 1012 M, and particular substituents protect against the Nod factor hydrolysis by enzymes of host plant origin. These observations, together with the fact that Nod factors preferentially migrate into root hair cell walls (Goedhart et al., 1999
), suggest that perception by (a) Nod factor receptor(s) may be an initial and essential requirement for Nod factor signaling.
This review summarizes the present data on Nod factor structures and Nod factorinduced responses during nodule initiation, focusing on the importance of the chemical structure for biological activity. Recent advances concerning the characterization of putative Nod factor receptors are highlighted.
Nod factor structures
A landmark in the Rhizobiumlegume symbioses was the report on the structure of the Nod factors of Sinorhizobium meliloti by Lerouge et al. (1990). Currently, Nod factor populations produced by many rhizobial strains from different genera and geographical origins are known in detail (Figure 1; Table I).
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It is interesting that a few Nod factors have exceptional lengths or types of oligosaccharide backbone or substitutions at the terminal residues (Table I). Some Nod factors of M. loti NZP2213 have a dimeric chitooligosaccharidic backbone (Olsthoorn et al., 1998) and some of Rhizobium sp. GRH2 consist of six GlcNAc residues (López-Lara et al., 1995b
), whereas Sinorhizobium fredii USDA191 can synthesize a pentameric Nod factor whose middle GlcNAc residue is replaced by a glucosyl group (Bec-Ferté et al., 1996
). The C-1 position of the reducing-terminal GlcNAc residue of some Nod factors produced by Bradyrhizobium elkanii USDA61 and R. tropici CIAT899 are substituted with a glycerol (Carlson et al., 1993
; Stokkermans et al., 1996
) and a mannosyl (Folch-Mallol et al., 1996
) moiety, respectively. In part of the Nod factor population synthesized by M. huakuii Ra5, the CH3CO group of the reducing-terminal GlcNAc residue is replaced by a CH2OHCO group (Yang et al., 1999
). Also the 3-O-S-2-O-Me-fucosyl and 4-O-Ac-2-O-Me-fucosyl residues found at C-6 of the reducing-terminal residue are unique for Sinorhizobium sp. NGR234 Nod factors (Price et al., 1992
, 1996). Furthermore, the reducing-terminal GlcNAc can be transformed into an open ring structure, such as an acetylated glucosaminitol group in the case of Rhizobium sp. BR816 (Snoeck et al., 2001
). Finally, some Nod factors carry modifications at the GlcNAc residue proximate to the nonreducing-terminal residue, for example, an
-1,3-linked fucosyl group at C-3 in M. loti NZP2213 (Olsthoorn et al., 1998
), an acetyl group at C-3 in R. galegae HAMBI1207 (Yang et al., 1999
), or an acetyl group at C-6 in Rhizobium sp. BR816 (Snoeck et al., 2001
). Diglycosylated Nod factors produced by S. teranga bv. sesbaniae ORS604 (Lorquin et al., 1997a
), S. saheli ORS611 (Lorquin et al., 1997a
), and Azorhizobium caulinodans ORS571 (Mergaert et al., 1997
), carrying both a fucosyl and an arabinosyl group at the reducing-terminal residue, have been observed only in rhizobia that nodulate the tropical legume Sesbania rostrata (Table I).
Nod factor responses and their structural requirements
Most commonly, rhizobia enter the host plants root tissue through the intracellular infection thread mode, after they have colonized root hair tips (Kijne, 1992). Based on the developmental stage of root hairs, three zones from the root tip toward the older part of the root are defined: zone I with growing root hairs, located near the root tip; zone II or "susceptible zone," carrying root hairs that terminate growth; and zone III, which consists of mature, fully grown root hairs. Generally, root hairs belonging to the susceptible zone respond to Nod factorproducing rhizobia, with root hair deformation and curling as a result. Small confinements appear that are formed by the curl, called shepherds crooks, in which rhizobia are entrapped. At these sites, rhizobia enter the root hair intracellularly, via an inward-growing infection thread. These infection threads guide the rhizobia to the newly developing nodule primordia, which are localized foci of cortical cell divisions. Subsequently, rhizobia become internalized in the cytoplasm of young plant cells. Many early nodulation events are induced by Nod factors. These plant responses occur at the epidermis, cortex, and pericycle and are summarized in Table II. Some of the early responses in epidermis and cortex are described in more detail.
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Depolarization of the membrane.
Ehrhardt et al. (1992) were the first to demonstrate that 109 M NodSm-IV (C16:2,S) applied to alfalfa root hairs depolarizes the membrane within 10 min. NodSm-IV (C16:2,S) and NodSm-IV (Ac,C16:2,S) are equally active and more active than NodSm-V (C16:2,S) and NodSm-IV (C16:0,S). NodSm-IV (C16:2) and chitotetraose cannot induce an important depolarization of the membrane potential (Felle et al., 1995
; Kurkdjian, 1995
) (Table II). At a concentration of 107 M, chitotetraose does not trigger this response, in contrast to chitohexaose, -heptaose, and -octaose that weakly depolarize the membrane (Felle et al., 2000
) (Table II).
Ion fluxes.
The Nod factorinduced depolarization of the membrane potential and extracellular alkalinization of the alfalfa root surface are inhibited on deactivation of the plasma membrane H+ ATPases, suggesting a role for proton pumps (Felle et al., 1998). A rapid increase in [Cl] is seen at the root surface when 107 M NodSm-IV (C16:2,S) is added to alfalfa roots, whereas, concomitantly, [Cl] decreases within the root hairs (Felle et al., 1998
). This response occurs after [Ca2+] changes but simultaneously with depolarization of the membrane potential and extracellular alkalinization (Felle et al., 1998
). Slightly later than the Cl efflux, a transient increase of [K+] is noticed in the root hair zone (Felle et al., 1998
). Chitooctaose induces similar ion fluxes, although less pronounced (Felle et al., 2000
). By using a single-electrode voltage-clamp technique, the involvement of anion and [K+] channel currents together with those of H+ pumps in Nod factorinduced electrical responses was confirmed (Kurkdjian et al., 2000
).
Changes in [Ca2+
].
Rapid changes in cytosolic [Ca2+], which occur in different patterns, have been documented in a variety of systems (Table II). Approximately 9 min after Nod factors of S. meliloti are added to alfalfa root hairs, regular oscillations in [Ca2+] are induced, referred to as spiking, that last for 2060 min (Ehrhardt et al., 1996). The response is specific because NodSm-IV (Ac,C16:2,S) induces [Ca2+] spiking, in contrast to chitotetraose or NodRlv-V (Ac,C18:4), whereas the latter Nod factor induces [Ca2+] spiking on Vicia hirsuta. No significant response can be monitored when S. meliloti Nod factors are added to Lycopersicon esculentum (tomato) root hairs (Ehrhardt et al., 1996
). [Ca2+] spiking is also observed in M. truncatula (Wais et al., 2000
) and Pisum sativum (pea), in which 109 M NodRlv-V (Ac,C18:4) first induces an increase in [Ca2+], 12 min after Nod factor addition, followed 515 min later by [Ca2+] spiking that lasts for 4060 min (Walker et al., 2000
). Surprisingly, 106 M chitotetraose and -pentaose induce [Ca2+] spiking in pea root hairs, similarly to Nod factors (Walker et al., 2000
). Recently, mutants of M. truncatula (Catoira et al., 2000
) and pea (Walker et al., 2000
) mutants, affected in initial steps of nodulation (e.g., root hair responses and induction of early nodulins), have been analyzed for their ability to exhibit Nod factorinduced [Ca2+] spiking. This analysis allowed particular plant genes to be positioned in the cascade of Nod factor perception[Ca2+] spikingroot hair deformationexpression of early nodulins (Wais et al., 2000
; Walker et al., 2000
).
In other systems, more general increases in [Ca2+] have been observed (Table II). A plateau-like increase in [Ca2+] is induced by 109 M NodNGR [Ac,S] or NodSm-IV,V [Ac,S] on Vigna unguiculata within seconds (Gehring et al., 1997). Such a response is not observed when V. unguiculata and Arabidopsis thaliana roots are treated with chitotetraose and with Nod factors or chitin fragments, respectively (Gehring et al., 1997
). In V. unguiculata root hairs, Ca2+ localization reveals that within 2 min after Nod factor application, [Ca2+] increased at the very tip of the root hair and in some spots throughout the root hair (Gehring et al., 1997
). In alfalfa root hairs, an extracellular decrease in [Ca2+] is measured in the responsive root zone after application of NodSm-IV (C16:2,S) (Felle et al., 1998
). The Nod factorinduced Ca2+ influx precedes the depolarization of the membrane potential and the transient extracellular alkalinization by several s. The Ca2+ influx is required for triggering these events. Ca2+ channel antagonists prevent both membrane depolarization and pH changes (Felle et al., 1998
). On Nod factor treatment of root hairs of Phaseolus vulgaris (common bean), cytosolic [Ca2+] increases in the root hair tip and in spots near the nucleus, often spreading all over the cytoplasm (Cárdenas et al., 1999
). These responses occur within seconds; 1015 min later, fluctuations and changes in [Ca2+] in the nuclear region are observed. Chitin fragments fail to change the cytosolic Ca2+ (Cárdenas et al., 1999
).
Striking similarities in [Ca2+] fluctuations between growing root hairs and pollen tubes have been reported. In growing pollen tubes of Lilium, the apical gradient of [Ca2+] is steep and is required for pollen tube growth (Miller et al., 1992). Also, root hair growth has been correlated with an increased cytosolic [Ca2+] in the apex (de Ruijter et al., 1998). Miller et al. (1992)
proposed that the region of elevated [Ca2+] may create conditions favoring vesicle fusion. Within the cytosol of living cells, increased [Ca2+] activates arrays of both rapid and sustained responses. Oscillations and their frequencies may have both quantitative and qualitative influences on gene expression (Meldolesi, 1998
). Consequently, changes in [Ca2+]either rapid plateau-like increases, slightly later occurring [Ca2+] spiking, or bothmay activate a set of genes involved in triggering the onset of nodule development.
Effects on root hair formation or shape.
Depending on the symbiotic system, Nod factors can induce the formation of root hairs (Hai) and/or deform existing ones (Had) (e.g., Roche et al., 1991a; Spaink et al., 1991
; López-Lara et al., 1995a
) (Table II). Only in Macroptilium atropurpureum (ReliM et al., 1993), V. unguiculata (Gehring et al., 1997
), and L. japonicus (Niwa et al., 2001
) Nod factors are sufficient to induce the formation of the typical shepherds crooks (Table II). Root hairs of V. unguiculata treated with NodNGR factors deform rapidly. Within 1 min, a bulge appears that carries an initiation of a root hair branch; at 2 min, branched root hairs are formed; at 4 min, root hairs are deformed and twisted; and at 32 min, root hairs with a shepherds crook are apparent (Gehring et al., 1997
). Nod factorinduced root hair deformation has been followed also in time on Vicia sativa (Heidstra et al., 1994
). Root hairs of the susceptible zone respond to 109 M NodRlv-V (Ac,C18:4), and at 30 min an increased cytoplasmic streaming is observed; at 1 h, root hair tips start to swell; at 2 h, polar growth is initiated; and at 3 h, 80% of the root hairs in zone II is deformed (Heidstra et al., 1994
). Strikingly, Nod factorinduced root hair deformation on V. sativa takes much longer than on V. unguiculata and requires de novo protein synthesis.
Root hair cytoskeleton rearrangements.
Control root hairs or root hairs treated with 107 M chitopentaose show long actin bundles running along the root hair from the tip to the base (Cárdenas et al., 1998). After 108 M Nod factors of R. etli have been applied for 510 min, a breakdown of microfilament bundles is observed. Approximately 1 h later, the structure of the cytoskeleton is partially recovered, but the actin microfilament stain still accumulates in the root hair tip. These cytoskeleton rearrangements may be a prerequisite for root hair deformation (Cárdenas et al., 1998
).
An extensive study of Nod factorinduced cytoskeleton rearrangements has been performed on root hairs of V. sativa (de Ruijter et al., 1998). Zone I root hairs exhibited a polar organization of the cytoplasm. In a clear zone at the tip, the cytoplasm contains almost exclusively Golgi vesicles. The subapical region is rich in organelles (endoplasmic reticulum, Golgi apparatus, mitochondria, plastids) and has small vacuoles. Zone II root hairs also have a cytoplasmic polarity and contain large organelles, including vacuoles, up to the tip. Zone III root hairs lack cytoplasmic polarity, and the large vacuoles, which occupy nearly the complete root hair cell, are surrounded by only a tiny layer of cytoplasm (de Ruijter et al., 1998
). Spectrin, a large multifunctional protein, that has actin- and calmodulin-binding sites and is part of the membrane-associated cytoskeleton, has been used as a molecular marker. Spectrin accumulates strongly in the cytoplasm of the apex of zone I, only very weakly in zone II, and not at all in zone III root hairs. After 1010 M NodRlv-V (Ac,C18:4) is added to V. sativa roots, zone II root hair tips start to swell, a small clear zone at the tip is formed from which a new tip emerges. After Nod factor treatment for 12 h, spectrin accumulates and is found at the plasma membrane of the tip swelling, suggesting that Nod factors reinitiate tip growth with features comparable to those of growing root hairs in zone I (de Ruijter et al., 1998
). Zone I root hairs have an increased cytosolic [Ca2+] in the tip region; 70 min after addition of Nod factors, a high cytoplasmic [Ca2+] is observed at the plasma membrane of the swelling zone II root hair tips and at the reinitiated tip, which originates from the swelling and forms a new root hair on an existing one (de Ruijter et al., 1998
).
During root hair development, one of the first visible events is the formation of a swelling, called a bulge, on an epidermal cell (Miller et al., 1999). These bulges develop into a growing root hair, the apex of which exclusively contains a vesicle-rich zone. Actin filament bundles within growing root hairs are oriented longitudinally, and perpendicularly to those in the epidermal cell. The subapical part exposes fine bundles of actin (Miller et al., 1999
). Actin filaments in Nod factorinduced bulges are positioned close to the plasma membrane in various orientations. At that point, the root hair cell changes its growth from unidirectional, as bulge, to polar growth, thereby initiating root hair deformation (Miller et al., 1999
). Thus, Nod factors reinitiate elongation of fine-bundle actin and vesicle delivery to the apical region that is free of actin filament bundles. Indeed, cytochalasin D (a compound that binds to the distal part of growing actin filaments, thereby blocking filament elongation and tip growth) blocks Nod factorinduced reinitiation of root hair tip growth but not bulge formation (Miller et al., 1999
).
Cortex and pericycle
Induction of early nodulins.
Nod factors induce several plant genes that are expressed during early stages of nodulation, the so-called early nodulin or ENOD genes. PsENOD12, encoding a hydroxyproline-rich protein, is first expressed in root hairs, 24 h after inoculation. Two days after inoculation, transcripts are detected in root hairs, in cortical cells that contain an infection thread, in cells that are ready for infection thread passage, and in inner cortical cells that will form the nodule primordium (Scheres et al., 1990). PsENOD12 expression is induced transiently after 108 M of Nod factors from R. leguminosarum bv. viciae, have been applied for 12, 24, and 48 h (Horvath et al., 1993
) (Table II). For MtENOD12 expression to be induced by S. meliloti Nod factors, final concentrations of 1012 M for NodSm-IV (Ac,C16:2,S) and NodSm-IV (C16:2,S), 1011 M for NodSm-IV (Ac,C16:0,S), and 109 M for NodSm-IV (Ac,C16:2) are required to obtain comparable responses, suggesting that both the presence of a C16:2 fatty acid and a sulfate ester are important. Chitotetraose, even at a concentration of 106 M, does not induce MtENOD12 expression (Journet et al., 1994
). The Rhizobium-induced peroxidase gene (rip1) of M. truncatula, in which the in situ transcript localization coincides with early infection events and initiation of nodule development, is induced by a mixture of NodSm-IV (Ac,C16:2,S) and NodSm-IV (C16:2,S) at a final concentration of approximately 109 M (Cook et al., 1995
) (Table II).
Another interesting case is ENOD2 of Glycine soja, which is expressed in nodule parenchyma (Minami et al., 1996a) but not when roots are treated with either LCO-V (C18:1
11, MeFuc), LCO-V (C16:0,MeFuc), LCO-V (C18:1
9,MeFuc), or LCO-IV (C16:0); however, GsENOD2 expression is induced by particular combinations of these Nod factors (Table II), and GsENOD2 transcripts can be detected only in the parenchyma of nodule structures induced by a Nod factor mixture. Thus such a mixture may be required for nodule ontogeny to progress, which may explain why rhizobia produce populations of structurally different Nod factors (Table II) (Minami et al., 1996a
). More examples of ENOD genes that are Nod factorinduced are listed in Table II.
Cytoplasmic bridges.
Cytoplasmic bridges are observed in outer cortical cells of V. sativa (van Brussel et al., 1992) and pea (Bakhuizen, 1988
), are positioned in line with young radial walls in the inner cortex, and form preinfection threads through which infection threads grow. Mitogenic Nod factors, which induce nodule primordia formation in V. sativa at a final concentration of 107 M, are sufficient to form preinfection threads (van Brussel et al., 1992
) (Table II). Interestingly, Niwa et al. (2001)
observed that M. loti JRL501 Nod factors induced preinfection thread formation in outer cortical cells of L. japonicus. Yang et al. (1994)
showed that outer cortical cells of pea entered the cell cycle but were arrested in G2 to form preinfection threads, whereas inner cortical cells went through the complete cell cycle, leading to nodule primordium formation.
Nodule primordium formation.
In many systems, Nod factors induce local foci of cell divisions, forming nodule primordia that develop into nodular structures (Table II). In M. sativa, 107 M NodSm-IV (Ac,C16:2,S) and NodSm-IV (C16:2,S) but not NodSm-IV (Ac,C16:2), NodSm-IV (C16:2), or NodSm-IV (C16:0,S) form discrete foci of cell division, which develop into structures that are microscopically comparable to genuine nodules (Truchet et al., 1991). A mixture of NodSm-IV (Ac,C16:2,S) and NodSm-IV (C16:2,S) (40/60) give a stronger response than each Nod factor separately. Demont-Caulet et al. (1999)
illustrated how important the correct fatty acid is for induction of cortical cell divisions. A chitotetramer carrying a sulfate ester at the reducing-terminal residue does not trigger any response on alfalfa roots. The length as well as the number and positions of the double bonds of the fatty acids are important for efficient formation of nodule primordia (Table II). Maybe one particular fatty acid at the nonreducing-terminal residue of the Nod factor serves as hydrophobic tail that allows insertion in lipid bilayers, such as the plant plasma membrane (Demont-Caulet et al., 1999
). However, foci of cell division can be induced when chitopentaose, substituted with an O-acetyl group at the nonreducing-terminal residue is delivered, together with uridine, to cortical cells of V. sativa by ballistic microtargeting. This response is observed neither when chitopentaose is used as projectile, nor when O-acetylated chitopentaose is applied externally to roots of V. sativa (Schlaman et al., 1997
).
A mixture of 105 M Nod factors, produced by B. elkanii USDA61, induces complete nodule structures on spot-inoculated roots of G. soja (Stokkermans and Peters, 1994). An extensive structurefunction study has been performed on G. soja with natural and synthetic Bradyrhizobium Nod factors and derivatives (Stokkermans et al., 1995
) (Table II). All compounds that cause root hair deformation elicit also nodule primordium formation. The methyl fucosyl group at the reducing-terminal residue is required for biological activity, but the place of the double bond in the fatty acid is not critical. When a pentameric LCO is replaced by a tetramer while retaining all the other substitutions, activity is lost (Table II). However, LCO-IV (C16:0) can induce root hair deformation and nodule primordium formation, suggesting that the MeFuc group at the reducing-terminal residue, required for activity of a pentameric Nod factors, hinders activity when present at the reducing-terminal residue of a tetrameric Nod factor (Stokkermans et al., 1995
).
More general Nod factorinduced responses.
Colonization of G. max roots by the mycorrhizal fungus Glomus mosseae is stimulated by 109 M NodNGR-V [MeFuc,Ac], but not by NodNGR-V [MeFuc,S] (Xie et al., 1995). Mathesius et al. (1998)
constructed transgenic Trifolium repens (white clover) plants that carried an auxin-responsive promoter, GH3, fused to a ß-glucuronidase (gusA) reporter gene, allowing changes to be assessed in auxin balances during early stages of nodule development. Spot inoculation with 108 M Nod factors of R. leguminosarum bv. trifolii results in a local and acropetally transient down-regulation and basipetal up-regulation of auxin transport in the cortex and vascular bundles 24 h after Nod factors have been added. This response is similar to that caused by naphthylphthalamic acid, a common auxin transport inhibitor. O-acetylated chitopentaose and, to a lesser extent, O-acetylated chitotetraose and chitohexaose (but not chitin oligomers) can mimic this response, suggesting that during early stages one of the effects of Nod factors may be the perturbation of auxin transport (Mathesius et al., 1998
).
Suspension-cultured cells of M. sativa and Nicotiana tabacum (tobacco) have been used to demonstrate differences between chitooligomers and Nod factors in induction of extracellular alkalinization and release of hydrogen peroxide (Table II). Chitotrimers, chitotetramers, chitopentamers, and chitohexamers induce an increase of the extracellular pH of M. sativa and N. tabacum suspension-cultured cells, at a concentration of 104, 104, 106, and 104 M, respectively, and within 5 min. Nod factors of S. meliloti, however, induce alkalinization only in N. tabacum suspension-cultured cells, 30 min after Nod factor application. Both 105 M NodSm factors and chitopentaose increase the production of hydrogen peroxide in N. tabacum but not in M. sativa suspension-cultured cells (Baier et al., 1999) (Table II).
Nod factor processing
The chitin backbone of Nod factors can be hydrolyzed by chitinases. Table III gives an overview of leguminous and nonleguminous chitinases and other enzymes that degrade Nod factors. Plant chitinases can be components of plant defense reactions, have an antifungal activity, play a role in carrot somatic embryo development, and be involved in plant development (see Collinge et al., 1993; Brunner et al., 1998
). A growing body of evidence suggests that chitinases may also play a role during nodule development. Chitinases are present in the cortex of soybean nodules, induced by B. japonicum 61-A-101, supposedly to protect central tissues against pathogen invasion (Staehelin et al., 1992
); Chinese G. max cultivars produce several chitinase isoforms during nodule development (Xie et al., 1999
); during the S. melilotiM. sativa symbiosis, chitinases have been detected in necrotic cells in the cortex, involved in infection thread abortion (Vasse et al., 1993
); and, during stem nodule development on S. rostrata, Srchi13, an early nodulin gene, encoding class III chitinase is expressed around infection pockets, around the developing nodule, and in uninfected cells of the central tissue (Goormachtig et al., 1998
). Recently, eight different chitinase genes from M. truncatula have been isolated and characterized, some of which are expressed during S. melilotiinduced nodule development. The expression patterns during nodule development differ from those generated on mycorrhiza or pathogen infection (Salzer et al., 2000
).
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Remarkably, on M. sativa roots, Nod factors induce their own breakdown by enhancing the production of a "dimer-forming hydrolase" (Staehelin et al., 1995). The presence of an O-acetyl group at the nonreducing-terminal residue, but not a sulfate ester at the reducing-terminal residue, protects Nod factors against degradation by the dimer-forming hydrolase. This Nod factor hydrolase is strongly induced by NodSm-IV (Ac,C16:2,S), whereas NodSm-IV (C16:2), LCO-III (C16:2), chitotetraose, or chitopentaose have no effect. Possibly, a rapid Nod factorinduced degradation of Nod factors may be part of Nod factor signaling, required for induction of plant genes involved in nodule development, or necessary to finely regulate Nod factor amounts and to avoid a continuous stimulation of the Nod factor perception system(s) (Staehelin et al., 1995
), eventually eliciting defense-like reactions (Savouré et al., 1997
). Also P. sativum roots respond to NodRlv factors and their fucosylated derivatives by an enhanced production of a dimer-forming Nod factor hydrolase (Ovtsyna et al., 2000
) (Tables II and III). As illustrated in Table III, a fucosyl group at the reducing-terminal residue of NodRlv factors renders the neighboring ß-1,4-glycosidic linkage inaccessible for cleavage by the N. tabacum class I chitinase Chi32 and class III chitinase lys28a (Ovtsyna et al., 2000
). The importance of the O-acetyl group at the nonreducing-terminal residue for protection against degradation has been demonstrated by Chi32, which degrades NodSm-IV (C16:2), but not NodSm-IV (Ac,C16:2) (Schultze et al., 1998
) (Table III). Finally, Staehelin et al. (2000)
showed that a partial de-N-acetylation of NodSm-IV (C16:2,S) rendered the compounds more resistant against degradation by the dimer-forming hydrolase. A mixture of de-N-acetylated compounds (Table III) was approximately 10,000-fold less active in inducing root hair deformation on M. sativa roots and does not increase production of the dimer-forming hydrolase. However, Nod factorinduced responses are not inhibited in the presence of de-N-acetylated compounds, indicating that the latter products are probably not perceived (Staehelin et al., 2000
).
Nod factor localization
Little is known about the in situ localization of Nod factors and their eventual degradation products. Philip-Hollingsworth et al. (1997) synthesized R. leguminosarum bv. trifolii Nod factor analogs that carry the 7-nitrobenzo-2-oxa-1,3-diazole (NBD) group at the C1 position of the reducing-terminal residue. During nodule initiation on T. repens, fluorescence has been observed in the cell wall and the plasma membrane of root hairs and other epidermal cells of the young part of the root. Part of the label is seen in the vicinity of the nucleus, another fraction at the root hair base, and later in underlying cortical cells. NBD-related fluorescence has been detected within four to five underlying cortical cell layers, but NBD-labeled chitotriose is not taken up by plant cells (Philip-Hollingsworth et al., 1997
).
In an independent study, fluorescent R. leguminosarum bv. trifolii Nod factor analogs have been synthesized by deacetylating chitotetraose using NodB, thus creating a free amino group subsequently acylated with a commercially available fluorophore-carrying Bodipy-C16:0 fatty acid (Gadella et al., 1997). By using LCO-IV (Bodipy-C16:0), fluorescence has been noticed predominantly at the level of the root hairs of V. sativa, which confirms earlier observations using [3H]-NodRlv-V (Ac,C18:0) (Heidstra et al., 1994
). The binding is not restricted to root hairs and epidermal cells of the susceptible zone but encompasses also mature and young actively growing root hairs (Heidstra et al., 1994
). In addition, an increased labeling has been observed in root hair tips (Gadella et al., 1997
). By using a variety of Bodipy-fatty acidlabeled Nod factor analogs and advanced microscopy techniques, it has been shown that Nod factors have a high tendency to be inserted into micelles or vesicles and to be transferred from vesicles to root hair cell walls (Goedhart et al., 1999
). Already 3 h after application of labeled Nod factors to V. sativa roots at a final concentration of 108 M, most Nod factors accumulate in the cell wall of root hairs. Occasionally, a very low level of fluorescence has been detected in the plasma membrane. The Nod factorinduced outgrowth, however, is not fluorescent in its cell wall, suggesting that Nod factors are immobilized or that Nod factor migration is slow. Only at high Nod factor concentrations (106 M) do Nod factors also accumulate in the plasma membrane (Goedhart et al., 2000
).
Antibodies raised against NodSm-IV (C16:2,S) have been used for in situ visualization of Nod factors during nodule development on M. sativa (Timmers et al., 1998). Signals are predominantly present in infection threads, but are also in the cytoplasm of invaded cells and in bacteroids throughout all stages of development. Immunolocalization on nodule sections that exhibit infection threads, internalization events, and infected cells suggests that Nod factors may be internalized in the cytoplasm while bacteria are internalized in plant cells. The spatial-temporal characteristics of this proposed internalization of Nod factors are identical to those of microtubular cytoskeleton disorganizations, which occur when bacteria are taken up by plant cells after release from infection threads. Moreover, the latter microtubular rearrangements have not been observed in bacteria-free nodules. Thus, the concomitant occurrence of internalization of Nod factors and microtubular rearrangements suggests that Nod factors possibly control cytoskeleton changes, which direct the differentiation of bacteria-containing cells (Timmers et al., 1998
).
Nod factor perception by putative receptors
Many indications support the hypothesis that Nod factors are perceived by plant receptors. Final concentrations of Nod factors as low as 109 to 1012 M provoke particular responses on roots, often requiring a defined Nod factor structure (Table II). Valuable information concerning putative Nod factor receptors has been obtained by a study of invasion phenotypes of S. meliloti nodL and nodFE mutants, and a nodL/nodF double mutant on M. sativa (Ardourel et al., 1994). Dependent on the mutantand thus on the type of Nod factorsinitiation of infection thread formation and subsequent penetration can be uncoupled from induction of nodule primordia formation. Based on these observations, Ardourel et al. (1994)
proposed the presence of a signaling receptor that recognizes Nod factors even when the nonreducing-terminal residue is altered and that would pave the way for infection (controlling infection thread growth), and an entry receptor that demands more stringent structural requirements for recognition, allowing rhizobial ingestion and control of the door opening.
Many attempts have been undertaken to identify putative Nod factor receptors. Some excellent reviews dealing with Nod factor perception have been published recently (Niebel et al., 1999; Cullimore et al., 2001
; Oldroyd, 2001
).
Nod factorbinding sites
Tritiated NodSm-IV (Ac,C16:2,S) has been used to identify Nod factorbinding sites (NFBSs) in particular fractions of M. truncatula roots. Binding is saturable and reversible but independent of the presence of an O-acetyl group at the nonreducing-terminal residue, a sulfate ester at the reducing-terminal residue, or an unsaturated fatty acid. Nevertheless, chitotetraose is a poor competitor. This NFBS has been designated NFBS1 (Bono et al., 1995). Two NFBSs for [35S]NodSm-IV (Ac,C16:2,S) have been found in microsomal fractions of M. varia suspension-cultured cells, one corresponding most probably to NFBS1, the other designated NFBS2. Initial characterization of NFBS2 showed a higher affinity for NodSm-IV (Ac,C16:2,S) and chitotetraose was a poor competitor (Niebel et al., 1997
). An O-acetyl group and a C4-hydroxyl group at the nonreducing-terminal residue, but not a sulfate ester at the reducing-terminal residue, are determinants for high-affinity binding to NFBS2. The length of the fatty acid, but not the number or position of double bonds, is important for efficient binding. Also the oligomerization degree of the Nod factor chitooligosaccharide backbone influences the affinity for binding to NFBS2 (Gressent et al., 1999
). Recently, a novel protein P60 has been purified from M. sativa roots with a high affinity for GlcNAc but also for NodSm factors because the latter are strong competitors for binding of GlcNAc monomers (Minic et al., 2000
).
G proteincoupled receptors
The cleavage of phosphatidylinositol (4,5)-biphosphate into inositol (1,4,5)-triphosphate and diacylglycerol (DAG) is a common feature of signal transduction pathways in animals and is often catalyzed by heterotrimeric GTP-binding regulatory protein (G protein)mediated activation of specific phospholipase C (PLC) isoenzymes. Inositol (1,4,5)-triphosphate can release Ca2+ from internal calcium stores, whereas DAG is converted into phosphatidic acid (PA) by DAG kinase. Alternatively, PA is formed by G protein-mediated activation of phospholipase D (PLD), by which structural lipids, including phosphatidyl choline, are hydrolyzed to produce PA. PA kinase converts PA into diacylglycerol pyrophosphate (see den Hartog et al., 2001).
The rapid activation of the Nod factor-induced MtENOD12 promoter (Table II) has been used to study components of Nod factor signal transduction in alfalfa transgenic lines (Pingret et al., 1998). Nod factorinduced MtENOD12 expression can be mimicked by mastoparan, an amphipathic tetradecapeptide that activates animal G proteins. Pertussis toxin, an antagonist of G protein activation, and neomycin, a PLC antagonist, block both Nod factor and mastoparan-induced MtENOD12 expression. Interestingly, exposure of roots to Nod factors for 15 min is sufficient to activate downstream processes. Furthermore, the release of Ca2+ from both internal stores and the external environment is required for MtENOD12 expression. These indications strongly suggest a role for heterotrimeric G proteins during Nod factorinduced MtENOD12 expression. Because in animal cells, G proteins are almost invariably coupled to a family of seven transmembrane span receptors, Nod factors may be perceived by such putative plant receptors (Pingret et al., 1998
).
Recently, den Hartog et al. (2001) further illustrated the importance of G proteinmediated signal transduction pathways in Nod factor or mastoparan-induced root hair deformation on V. sativa. Both Nod factors and mastoparan increase the level of PA, which was caused by increased DAG kinase and PLD activity. In addition, Nod factor and mastoparan-induced PA production was inhibited by neomycin, a PLC inhibitor, or primary butyl alcohols, which block PLD activity, confirming the involvement of PLC and PLD activities, respectively.
In conclusion, at least one of the Nod factorinduced downstream signaling pathways may involve G proteinmediated activation of both PLC and PLD, leading to the release of Ca2+ from internal stores, required for Nod factorinduced MtENOD12 expression in M. sativa (Pingret et al., 1998) and root hair deformation in V. sativa (den Hartog et al., 2001
).
Lectins and apyrases
Carbohydrate-binding lectin proteins are good candidates to function as putative Nod factor receptors. Transgenic T. repens hairy roots expressing a pea lectin gene (PSL) are nodulated by the pea symbiont R. leguminosarum bv. viciae (Díaz et al., 1989); when they carry a PSL analog mutated in its sugar-binding site, they can form only pseudonodules after inoculation with R. leguminosarum bv. viciae, illustrating the importance of the lectin sugar-binding domain (van Eijsden et al., 1995
). The presence of the nodABCIJ, nodD, and nodFEL genes in R. leguminosarum bv. viciae is necessary for nodule development on transgenic white clover roots that express PSL, suggesting that PSL may be involved in the recognition of R. leguminosarum bv. viciae Nod factors (Díaz et al., 1995
). Recently, PSL has been introduced into hairy roots of T. pratensis to test several purified Nod factors, including NodRlt-IV/V (Ac,C18:2/C18:4), NodRlv-V (Ac,C18:4), NodRlv-IV (Ac,C18:4), NodRlv-V (Ac,C18:1), NodSm-IV (Ac,C16:2,S), and NodMl-V (Me,Cb,C18:0/C18:1, AcFuc), for their capacity to trigger cortical responses. All of these Nod factors induce cortical cell division with the formation of structures resembling nodule primordia as a result, but a similar response is triggered by chitobiose, chitotriose, chitotetraose, and chitopentaose (Díaz et al., 2000
).
On transgenic Lotus corniculatus true nodules are formed that express the soybean lectin gene (SBL) upon inoculation with B. japonicum, a symbiont of soybean, but not of L. corniculatus. SBL has been localized in L. corniculatus root hairs. Mutation of the lectin sugar-binding site abolishes infection thread formation and nodulation by B. japonicum. By inoculating a variety of purified Nod factors on both wild-type and transgenic plants, Nod factorinduced nodule primordium formation is found to be influenced not directly by the presence of SBL. Seemingly, a component of extracellular polysaccharides of B. japonicum, rather than Nod factors, extends the host range to transgenic L. corniculatus plants (van Rhijn et al., 1998). Inoculation of SBL and PSL transgenic M. sativa lines with B. japonicum and R. leguminosarum bv. viciae, respectively, leads to the formation of nodule-like structures only when S. meliloti Nod factors are produced (van Rhijn et al., 2001
). In the first case, only empty nodule-like structures are observed devoid of infection threads, whereas in the latter infection thread formation appears and some nodule-like structures are infected, but no features indicative for nitrogen fixation are seen. Interestingly, the production of extracellular polysaccharides is a prerequisite for both induction of nodule development and infection thread formation (van Rhijn et al., 2001
).
A lectin-nucleotide phosphohydrolase (LNP) has been isolated from roots of Dolichos biflorus, a legume that can be nodulated by B. japonicum and Sinorhizobium sp. NGR234. DbLNP-chitin binding is inhibited by high concentrations of GlcNAc monomers. D-N-acetylated chitooligosaccharides are not, but chitobiose, chitopentaose, and chitohexaose are strong competitors as well as Nod factors purified from B. japonicum and Sinorhizobium sp. NGR234, illustrating that DbLNP can bind Nod factors. DbLNP is unique because it is not significantly similar with other known lectins and is an apyrase that can hydrolyze a phosphate group from ATP and ADP residues. In the presence of Nod factors, the enzymatic activity of DbLNP increases. Strikingly, DbLNP is localized in the epidermal cell surface of root hairs and DbLNP antiserum blocks root hair deformation and nodule formation, an inhibitory effect that happens only in the antiserum-treated part of the root (Etzler et al., 1999). These observations suggest a role for DbLNP in Nod factor perception by D. biflorus roots. Closely related orthologs of DbLNP have been found in M. sativa and P. sativum, whereas a second apyrase-encoding gene isolated from D. biflorus, designated apyrase-2, corresponds to sequences in L. japonicus, M. sativa, and A. thaliana (Roberts et al., 1999
). The differential distribution of DbLNP along the surface of the root axis coincides with the nodulation zone on D. biflorus roots. DbLNP is present on the surface of young and emerging root hairs and is redistributed in response to inoculation with a rhizobial symbiont or to application of Nod factors. The redistribution correlates with the localization of rhizobia on the root hair surface (Kalsi and Etzler, 2000
). Two apyrase-encoding cDNAs of G. soja, GS50 and GS52, have been characterized. The level of GS50 expression is not influenced by B. japonicum, and GS50 is localized in the Golgi apparatus. In contrast, expression of GS52, a DbLNP ortholog, is enhanced rapidly by B. japonicum. Anti-GS52 but not anti-GS50 antibodies block nodulation by B. japonicum, suggesting a role for GS52 in soybean nodulation (Day et al., 2000
).
Chitinase-like proteins
A chitinase homolog, Srchi24, has been isolated from S. rostrata (Goormachtig et al., 2001) and its transcript levels increase 4 h after inoculation with Nod factorproducing azorhizobia. Both Srchi24 transcripts and proteins are located in the outer cortical cell layers of developing nodules. An important catalytic glutamic acid residue is replaced in Srchi24, which possibly explains the lack of chitinase activity of the fusion protein between the maltose-binding protein and Srchi24. Currently, research is being performed on the Srchi24 and Nod factor binding (Van de Velde and Holsters, personal communication). Interestingly, a chitinase-related receptor-like kinase (CHRK1) has been isolated from tobacco (Kim et al., 2000
) that consists of a C-terminal kinase domain and a putative extracellular domain, which is closely related to class V chitinases of tobacco. The chitinase domain lacks the glutamic acid residue required for chitinase activity, and chitooligosaccharides of chitin are not degraded. CHRK1 is located in plasma membrane fractions, in agreement with a putative role in perception (Kim et al., 2000
).
Others
Perhaps the best Nod factor receptor candidate is sym10. A sym10 mutant of pea is Nod, but infection by mycorrhizal fungi is not affected. Moreover, the sym10 mutation affects early Nod factor responses, such as [Ca2+] spiking and root hair deformation (Walker et al., 2000).
Finally, nn1 of alfalfa, associated with a Nod phenotype, has been map-based cloned. nn1 encodes a receptor kinase with a leucine-rich repeat region in the putative external domain and may also serve as a candidate Nod factor receptor (Endre et al., 2001).
Nod factorlike molecules in eukaryotes
Because Nod factors can switch on a complex developmental program leading to nodule formation, it is interesting to investigate whether Nod factorlike compounds are also present in (non)legume plants or other organisms and to decipher eventual Nod factorrelated responses.
Transgenic tobacco plants that express nodA, nodB, or both are severely affected in development, as demonstrated by reduced growth and internode distance, rounded and wrinkled leaves, and compact inflorescence. These observations suggest that tobacco contains substrates that can be modified by NodA or NodB and that are involved in plant growth and organ development (Schmidt et al., 1993).
Arrested embryo development in Daucus carota (carrot) cell lines can be rescued by applying 109 M NodRlv-V (Ac,C18:4) or 108 M NodRlv-V (Ac,C18:1), whereas addition of chitopentaose was ineffective, suggesting a role of Nod factorlike molecules in carrot development (De Jong et al., 1993). More recently, a mixture of Sinorhizobium sp. NGR234 Nod factors has been shown to stimulate Picea abies (Norway spruce) protoplast division and regeneration of proembryonic masses from the protoplasts (Dyachok et al., 2000
).
In transgenic Oryza sativa (rice) plants, harboring an MtENOD12gus fusion, the MtENOD12 promoter can be activated by either sulfated or nonsulfated Nod factor mixtures purified from Sinorhizobium sp. NGR234 but not by chitotetraose. MtENOD12 promoter activity is observed in cortical parenchyma, endodermis, and pericycle. NodNGR factors, however, do not induce rice root hair deformations, and MtENOD12 expression is not detected in epidermal cells. Thus, part of the Nod factor signal transduction pathway, required for inducing MtENOD12 expression, exists in rice plants (Reddy et al., 1998).
Xenopus laevis carries a DG42 gene that is similar to nodC of rhizobia and that is expressed during a short period in embryo development. In vitro DG42-dependent synthesis of chitooligomers with an oligomerization degree of 26 has been demonstrated, and longer chitooligosaccharides have been produced as well. Possibly, these molecules are important in vertebrate embryogenesis (Semino and Robbins, 1995). Overexpression of the DG42 gene in Saccharomyces cerevisiae demonstrated that DG42 functions as a hyaluronan synthase that utilizes UDP-GlcA and UDP-GlcNAc to form a hyaluronan polysaccharide of approximately 106107 Da with a repeating unit equal to (
4)-ß-D-GlcA(1
3)-ß-D-GlcNAc(1
) (Pummill et al., 1998
). X. laevis DG42 homologs are found in Brachydanio rerio (zebrafish), human, and mouse; the in vitro synthesis of chitooligosaccharides by zebrafish or mouse extracts from appropriate developmental stages depends on DG42 (Semino et al., 1996
). That chitooligosaccharides are synthesized during particular developmental stages of zebrafish and Cyprinus carpio (carp) was confirmed by Bakkers et al. (1997)
, who purified extracts and labeled chitooligosaccharides by an in vitro transfucosylation reaction, using GDP-[U-14C]fucose and B. japonicum NodZ. Carp produces predominantly chitotetraose and zebrafish chitotetraose and chitopentaose. Strikingly, when fertilized zebrafish eggs are injected with anti-DG42 serum or NodZ protein, but not with rabbit preimmune serum, severe malformations in trunk and tail have been observed (Bakkers et al., 1997
). A similar phenotype is apparent when the B. japonicum nodZ gene is expressed in one-cell zebrafish embryos (Semino et al., 1998
), and when a Streptomyces plicatus chitinase 63 is injected (Semino and Allende, 2000
). These observations suggest that chitooligosaccharides may play a role in the development of these vertebrates.
Nod factor modeling
Information about the 3D structure of Nod factors is needed to understand the role of substitutions in biological activity. LCO-IV (C18:1,MeFuc) and LCO-IV (C16:1,S) have been analyzed by nuclear magnetic resonance spectroscopy (Gonzalez et al., 1999), and a study of the crystal structure of NodNGR factors is currently under investigation (Broughton, personal communication). Because these techniques are laborious and their application on many samples is practically impossible, 3D modeling may be a useful alternative. Molecular modeling is a fast and reliable method to obtain preliminary structural information. It has recently been applied to cell wall polysaccharides, such as pectins (Pérez et al., 2000
). However, the data should be interpreted with care; predictions obtained by modeling may not correspond completely with the real structures, particularly for compounds present in defined microenvironments, such as plant cell walls or cytoplasmic membranes.
A case study has been performed on Nod factors produced by A. caulinodans ORS571. As mentioned in Table I, A. caulinodans produces mainly pentameric Nod factors with either a C16:0, C18:1, or C18:0 fatty acid at the nonreducing-terminal residue. All Nod factors contain an N-methyl and a 6-O-carbamoyl group at the nonreducing-terminal residue, and no glycosylations, an L-fucosyl group, a D-arabinosyl group, or both at the reducing-terminal residue (Table I). D'Haeze et al. (2000) suggested that carbamoyl and glycosyl groups of azorhizobial Nod factors may play a role in the recognition of Nod factors by putative receptors. A 3D model of a fully substituted NodARc-V (Me,Cb,C18:0,Fuc,Ara) (Figure 2) illustrates that the fatty acid chain is positioned approximately perpendicularly to the chitooligosaccharide backbone. Neither this feature nor the global structure changes significantly when the C18:0 fatty acid is replaced by a C18:1 or C16:0 fatty acid; when the carbamoyl, fucosyl, and/or arabinosyl groups are removed; or when the carbamoyl or fucosyl groups are replaced by an acetyl group or a sulfate ester, respectively. De-N-acetylation of the second, third, fourth, or fifth GlcNAc residue of NodARc-V(Me,Cb,C18:0,Fuc,Ara) does not influence the global structure (data not shown). However, when the N-methyl group of NodARc-V(Me,Cb,C18:0,Fuc,Ara) is removed, the Nod factor conformation is severely altered in that the fatty acyl chain is oriented almost in parallel with the chitooligosaccharide backbone (Figure 2). The modifications at the reducing- or nonreducing-terminal residue have no effect, also when a carbamoyl group is substituted at the C3 and/or C4 position (data not shown).
|
Some conclusions, leading to many questions
Since the identification of Nod factors, major progress has been made in our knowledge on the molecular mechanisms of controlled bacterial invasion and induction of organ development. Data on nodulation-related gene expression have accumulated steadily over the past decade and are now expected to grow exponentially with the advent of genomics/genetics programs. The next challenges will be data mining and fitting all the data into a functional, biological context. Knowledge from other developmental processes and from studies of pathogen infections provides a solid background for setting up models and hypotheses in the nodulation field.
Some players of the game are known, but the rules of symbiotic development need to be deciphered. An important question, the answer of which will definitely lead to another landmark in symbioses history, is how Nod factors are perceived. Various protein candidates may serve as putative plant Nod factor receptors, and it seems plausible that different Nod factor receptors exist in a particular legume plant, all of them contributing to the final picture. If different Nod factor receptors exist, how is their expression regulated? Where are they expressed? Do they have a specificity for particular Nod factors? Do they govern specific downstream responses? Are the receptors monomers or dimers, or are they part of a more complex perception construction? Based on Nod factor localization and binding studies, the picture emerges that the primary Nod factor perceiving proteins or protein complexes are localized at the cell wall, whereas true Nod factor receptors for downstream signaling may be expected at the plasma membrane. If so, how do Nod factors traverse the cell wall to reach their receptor(s)? Is this a simple diffusion process or do apoplastic adaptor proteins play a role, comparable to the chaperone concept, in transport, protection, or docking with a receptor in the membrane?
The picture of initial Nod factor recognition gets even more complicated, because Nod factors may be degraded by plant-derived Nod factor-hydrolyzing enzymes. What is the turnover of Nod factors in the host environment, and what is the fate of the degradation products? Are these molecules also recognized by the host, and what may be their role during nodule initiation?
What is the sequence of events directly downstream of Nod factor perception? An impressing set of Nod factorinduced early responses have been recorded in different systems, involving Ca2+, membrane potential, and pH. However, a clear picture of causes and consequences is missing, and the order of events and which responses are controlled by which changes are unknown.
Finally, how do Nod factors fit into the more general scheme of oligosaccharide signaling and/or lipid-derived signals in eukaryotes? Rhizobia may mimic fungal chitin signals or endogenous oligosaccharins derived from cell walls or from glycoproteins. For researchers in the symbiosis field, the challenges are numerous and the most tempting ones have yet to come.
Acknowledgments
We are grateful to Russell W. Carlson (University of Georgia) for critical reading of the manuscript and stimulating discussions, to Martine De Cock for help preparing it, and to Rebecca Verbanck and Karel Spruyt for art work. W.D. is indebted to the Belgian-American Educational Foundation and Fulbright for a postdoctoral and a postdoctoral honorary fellowship, respectively.
Abbreviations
DAG, diacylglycerol; LCO, lipo-chitooligosaccharides; LNP, lectin-nucleotide phosphohydrolase; NBD, 7-nitrobenzo-2-oxa-1,3-diazole; NFBS, Nod factorbinding site; Nod, nodulation; PA, phosphatidic acid; PLC, phospholipase C; PLD, phospholipase D.
Footnotes
1 Present address: Complex Carbohydrate Research Center, University of Georgia, Athens, GA 30602, USA
2 To whom correspondence should be addressed; E-mail: mahol{at}gengenp.rug.ac.be
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