Abstract

Background

Nitrogen is an essential nutrient in plant growth. The ability of a plant to supply all or part of its requirements from biological nitrogen fixation (BNF) thanks to interactions with endosymbiotic, associative and endophytic symbionts, confers a great competitive advantage over non-nitrogen-fixing plants.

Scope

Because BNF in legumes is well documented, this review focuses on BNF in non-legume plants. Despite the phylogenic and ecological diversity among diazotrophic bacteria and their hosts, tightly regulated communication is always necessary between the microorganisms and the host plant to achieve a successful interaction. Ongoing research efforts to improve knowledge of the molecular mechanisms underlying these original relationships and some common strategies leading to a successful relationship between the nitrogen-fixing microorganisms and their hosts are presented.

Conclusions

Understanding the molecular mechanism of BNF outside the legume–rhizobium symbiosis could have important agronomic implications and enable the use of N-fertilizers to be reduced or even avoided. Indeed, in the short term, improved understanding could lead to more sustainable exploitation of the biodiversity of nitrogen-fixing organisms and, in the longer term, to the transfer of endosymbiotic nitrogen-fixation capacities to major non-legume crops.

INTRODUCTION

Nitrogen is an essential element in plant development and a limiting factor in plant growth. It represents about 2 % of the total plant dry matter that enters the food chain. Nevertheless, plants cannot directly access dinitrogen gas, which makes up about 80 % of the atmosphere. Plants absorb the available nitrogen in the soil through their roots in the form of ammonium and nitrates. The limited bio-availability of nitrogen and the dependence of crop growth on this element have spawned a massive N-based fertilizer industry worldwide (Dobermann, 2007; Westhoff, 2009). About 60 % of synthetic nitrogen fertilizers are presently used for cereals, with irrigated rice production accounting for approx. 10 % of the use. Since >50 % of the fertilizer applied is actually used by plants, the inefficient use of nitrogen contributes to nitrate contamination of soils and ground water, leading to health hazards and compromising agricultural sustainability. Moreover, manufacturing N fertilizer requires six times more energy than that needed to produce either P or K fertilizers (Da Silva et al., 1978).

Only some prokaryotes are able to use atmospheric nitrogen through a process known as biological nitrogen fixation (BNF), which is the conversion of atmospheric N2 to NH3, a form that can be used by plants (Lam et al., 1996; Franche et al., 2009). The bacteria responsible for nitrogen fixation are called diazotrophs; they encode nitrogenase, the enzyme complex that catalyses the conversion of N2 gas to ammonia. The nitrogenase complex is highly conserved in free-living and symbiotic diazotrophs. Various types of associations/interactions occur between diazotrophs and their host plants. The highly specific and most efficient processes for nitrogen fixation involve the formation of root nodules on legumes and non-legumes. The diazotrophic bacteria involved in these endosymbiotic interactions include rhizobia (Gram negative) members of the alpha-subgroup of the phylum proteobacteria that associate with legumes (family Fabaceae) (not included in this review; see Schultze and Kondorosi, 1998; Oldroyd and Downie, 2008; Desbrosses and Stougaard, 2011) and the non-legume Parasponia species (family Cannabaceae), and Frankia sp. (Gram positive) members of the actinomycete family that associate with a broad spectrum of plants belonging to eight families collectively called actinorhizal plants. In addition, nitrogen-fixing cyanobacteria (mainly Nostoc sp.) have also been found to colonize different plant organs, either intracellularly in the family Gunneraceae or extracellularly in liverworts, hornworts, Azolla and Cycadaceae. In contrast with these symbioses, some diazotrophs, such as Azospirillum spp., Azoarcus spp. and Herbaspirillum, form associative and/or endophytic relationships with a wide variety of plant roots including those of cereals. In all these associations and symbioses, for the host plants the expected benefit of the interaction is the fixed nitrogen provided by the symbiotic partner, which, in return, receives reduced carbon and possibly all the other nutrients it requires. In addition, the symbiotic or endophytic plant structure colonized by the nitrogen-fixing microorganisms may provide the appropriate conditions for protecting the nitrogenase complex from oxygen exposure.

Rhizosphere associations between nitrogen-fixing microorganisms and plants have been a major driving force in allowing organisms to spread across the biosphere, occupy new niches, and adapt to a variety of environmental stresses. This review presents an overview and recent advances in the understanding of the associations between a wide range of diazotrophs and non-legumes. Discoveries and breakthroughs in legume and non-legume nitrogen fixation provide new insight into ways of manipulating key steps in this process. Finally, new perspectives to engineer nitrogen-fixing ability in non-legume crops based on knowledge of endosymbiotic processes in non-legumes are discussed.

ACTINORHIZAL SYMBIOSES

Actinorhizal plants and their major ecological role

Actinorhizal plants have the ability to develop an endosymbiosis with the nitrogen-fixing soil actinomycete Frankia. The establishment of the symbiotic process results in the formation of root nodules in which Frankia provides fixed nitrogen to the host plant in exchange for reduced carbon. Actinorhizal plants represent a diverse group of about 220 species belonging to eight plant families distributed in the three orders, Fagales (Betulaceae, Casuarinaceae and Myricaceae), Rosales (Rosaceae, Eleagnaceae and Rhamnaceae) and Cucurbitales (Datiscaceae and Coriariaceae) (Wall, 2000; Pawlowski, 2009; Franche and Bogusz, 2011). All actinorhizal species belong to the Rosid I clade, thus sharing a common ancestor with legumes (Fabaceae), but differing from them in their wide distribution in numerous botanical families. It has been suggested that 100 million years ago (Mya), the common ancestor of Rosid I acquired a unique feature upon which a root nodule symbiosis (RNS) could evolve, and that this evolution occurred several times 50–60 Mya (Doyle, 2011). Three to four independent evolutionary origins have been postulated for actinorhizal symbiosis (Swensen, 1996).

Actinorhizal plants are woody shrubs and trees, except for the genus Datisca, which is herbaceous. They are distributed worldwide from cold regions (except Antartica) with, for example, Alnus (alder), to warm latitudes with, for example, Casuarina (beef wood). Many actinorhizal plants are also capable of forming mycorrhizal associations, and this tripartite symbiosis (host plant–Frankia–mycorrhiza) gives them a propensity to grow in marginal and poor soils (Dawson, 2008). Some species are very well adapted to flooded land, arid regions, contaminated soils, extreme pH and high salinity. Due to these properties, some actinorhizal trees are pioneer species that colonize disturbed areas; they play extremely important ecological roles and are intensively used in the re-vegetation of different landscapes or to prevent desertification. For example, in Africa, Casuarinaceae are planted to stabilize coastal and desert dunes, and for reclamation of salt-affected soils as well as in inter-cropping systems (Diem and Dommergues, 1990). In these arid soils, the species Casuarina equisetifolia fixes an average of 15 kg N ha−1 year−1. But in temperate climates, nitrogen-fixation activity in actinorhizal plants could be similar to the rate of 300 kg ha−1 year−1 measured in legumes (Wheeler and Miller, 1990). In addition, these perennial plants contribute to the N cycle through litter fall and soil decomposition.

Nitrogen-fixing actinobacteria Frankia

Frankia is a genus of soil actinomycetes in the family Frankiaceae that fix nitrogen, both under symbiotic and free-living aerobic conditions, while most rhizobia do not (Benson and Silvester, 1993). Phylogenetically, the filamentous gram-positive Frankia sp. and the unicellular gram-negative paraphyletic rhizobia are quite distant, suggesting that these two major groups of nitrogen-fixing symbionts have acquired mechanisms for nitrogen fixation from different evolutionary origins (Normand et al., 1996). The first successful isolation of Frankia was reported relatively recently from Comptonia peregrina root nodules (Callaham et al., 1978). At present, over 200 strains of Frankia have been isolated from many, although not all, actinorhizal plant species. Phylogenetic analyses revealed that Frankiae form a coherent clade within actinobacteria, and that strains generally fall into three major groups or clusters (Normand et al., 1996; Benson and Clawson, 2000; Hahn, 2008). Cluster I consists of strains isolated from plants belonging to the order Fagales; they have the most specific host range and are only able to interact with plants belonging to this clade. Strains belonging to cluster III have a wider host range and can interact with plants belonging to five families within the two distant plant orders Rosales and Fagales. No strain belonging to cluster II has yet been cultured; actinobacteria are thus hypothetical obligate symbionts, even though no genome reduction has been observed in Frankia Dg1, the endosymbiotic strain of Datisca glomerata (Persson et al., 2011). A fourth group of non-infective or non-effective Frankia strains forms a more deeply branching cluster (Kucho et al., 2010). Each group has its own set of characteristics regarding symbiotic properties such as host-plant specificity, physiology and symbiotic relationships (Benson and Clawson, 2000).

Although Frankia strains are highly diverse in terms of ecological niches in the soil, current knowledge is focused on its life as an endophyte in root nodules. The difficulties sometimes encountered in growing Frankia in culture or in isolating Frankia from some plant species such as Datisca, reflect this limited knowledge. In pure culture, cultivable Frankia form vegetative hyphae that grow slowly, requiring >3–7 d before the colony appears on solid medium (Benson et al., 2011). One striking feature of Frankia is its ability to differentiate into two unique structures besides hyphae: (1) spores, which occur in sporangia and contribute to natural dissemination of the actinobacteria (Huss-Danell et al., 1997); and (2) vesicles, which are the sites of nitrogen fixation and differentiate at the tips of hyphae under nitrogen limitation. The vesicles are surrounded by a laminated envelope composed of hopanoid lipids that provide the necessary O2 protection to prevent nitrogenase inactivation (Harriott et al., 1991; Berry et al., 1993; Dobritsa et al., 2001).

Molecular biology of Frankia is limited by the absence of tools for genetic transformation and transposon mutagenesis (Kucho et al., 2009). However, progress in genome sequencing, transcriptome and proteome analyses has provided some important insights into Frankia. Genomes of three Frankia strains belonging to different host-compatibility groups were sequenced in 2007, revealing the absence of the canonical nodABC genes that code for the lipochito-oligosaccharidic Nod factors in rhizobia (Normand et al., 2007). Only low similarity nodB and nodC homologues have been detected; these data are in agreement with the absence of functional complementation of rhizobia nod mutants with Frankia DNA (Cérémonie et al., 1998). Moreover, genes known to be involved in symbiosis such as nif (nitrogenase), hup (hydogenase uptake) and suf (sulfur-iron cofactor synthesis) are scattered throughout the genomes. The absence of symbiotic islands in Frankia genomes, in contrast to rhizobia, complicates the identification of symbiotic genes. Nevertheless, with the sequencing of several other Frankia genomes in progress, valuable information is expected to become available from further genome comparisons. Preliminary analysis of the uncultured Dg1 symbiont of Datisca glomerata reveals that it contains nodABC-like genes, thus suggesting the use of Nod factors-like compounds during the infection process (Persson et al., 2011).

Several transcriptome data analyses of free-living and symbiotic Frankia have been carried out. Alloisio et al. (2010) showed that nodule-induced genes of Frankia alni were mostly distributed over several regions with high synteny between three Frankia genomes and that, as expected, several genes linked with nitrogen fixation (nif, suf, hup2, ispg) were up-regulated. In addition, comparison with rhizobia transcriptome revealed that Frankia is metabolically more active in nodules, due to vesicle biosynthesis. Overall, Frankia is less dependent on the host plant than rhizobia for symbiotic requirements and, as a result, metabolic activity is higher in symbioses (Alloisio et al., 2010). Interestingly, expression of genes necessary for ammonium assimilation was reduced in symbiosis, suggesting that symbiotic Frankia may not perceive N starvation and that nif genes are regulated by signals other than those involved in N metabolism (e.g. O2 level) (Benson et al., 2011). In addition, using high throughput RNA deep sequencing, Bickhart and Benson (2011) demonstrated significant heterogeneity in cell populations of Frankia sp. CcI3 growing in different conditions (NH4+ added vs. N2 fixing; young and ageing bacterial culture). Interestingly, the high expression of transposase ORF (open reading frame) observed in 5-d-old and nitrogen-fixing cultures suggests that the stationary phase of growth and nitrogen starvation trigger mechanisms that favour modification of the genome.

Signals involved in plant–Frankia recognition and in the plant signalling pathway

It is assumed that, as observed in rhizobium–legume symbioses, the compatible interaction between Frankia and actinorhizal plants that leads to the development of nitrogen-fixing nodules is the result of a fine-tuned exchange of signals between the two partners (Franche and Bogusz, 2011).

On the plant side, although the involvement of flavonoids in symbiosis is poorly understood, several studies indicate that they may play a significant role in the early stage of the interaction. These studies also suggest a role for Frankia in chemo-attraction and proliferation (Smolander and Sarsa, 1990), and in the enhancement of nodulation following the addition of seed washes from red alder Alnus rubra (Benoit and Berry, 1997). These results were reinforced by Hughes et al. (1999), who observed that flavonols (quercetin and kaempferol) contained in A. glutinosa (black alder) root exudates enhanced the level of nodulation. In addition, root hair curling, which is the primary event in the symbiotic process, was enhanced by exposure of Frankia to A. glutinosa root filtrate (Prin and Rougier, 1987; Van Ghelue et al., 1997). More recently, Popovici et al. (2010) reported that Myricaceae plants adapt their secondary metabolism in accordance with the compatibility status of Frankia bacterial strains, thus suggesting that flavonoids determine the specificity of the microsymbionts. The main plant compounds differentially affected by inoculation with Frankia are phenols, flavonoids and hydroxycinnamic acids. Interestingly, Beauchemin et al. (2012) demonstrated that Casuarina root extracts containing flavonoids altered the physiology, surface properties and plant infectivity of the compatible Frankia strain CcI3. In addition, several genes of the isoflavonoid biosynthesis pathway were shown to be up-regulated during early steps of interactions between Casuarina glauca and Frankia CcI3 (Auguy et al., 2011). A study is underway to understand the role of flavonoids in the Casuarina/Frankia symbiosis in more detail by using an RNAi approach to down-regulate the level of chalcone synthase transcripts in C. glauca (Rhizogenesis group, Montpellier, France, unpubl. res.).

In legume–rhizobium symbiosis, specific signal molecules secreted by Rhizobium, called Nod factors, play a pivotal role in host-symbiont specificity and the induction of all early plant responses including symbiotic gene activation leading to mitotic reactivation of the cortical cells, and formation of pre-infection threads (Oldroyd et al., 2010). As mentioned above, some specificity has been observed in the interaction between Frankia strains and their host plants. Studies aimed at purifying and characterizing symbiotic molecules from Frankia first relied on a bioassay based on root hair deformation with culture supernatants (Van Ghelue et al., 1997; Bhuvaneswari and Solheim, 2000). Preliminary characterization of Frankia root hair deforming factor(s) indicated that their chemical properties differed from those of rhizobium Nod factors (Cérémonie et al., 1999). However, since N-acetyl-glucosamine, the backbone of rhizobium Nod factors, has been detected in the Frankia root hair-deforming active fraction, it is possible that the two compounds are structurally related (Cérémonie et al., 1999). Recently, a sensitive and reproducible bioassay based on early expression of symbiotic C. glauca genes was developed (Rhizogenesis group, Montpellier, France, unpubl. res.). This should help future work dealing with purification and complete characterization of Frankia signalling molecules involved in the early dialogue with the root system.

Sugars and phytohormones may also be involved in the molecular dialogue. Interestingly, a study of whole-cell sugar contents showed that a monosaccharide, 2-O-methyl-d-mannose, was present in all Frankia strains tested. This sugar may thus play a role in interactions and communications between Frankia and its hosts (Kucho et al., 2010). From the analysis of Frankia supernatants, it was also found that auxins including indole-3-acetic acid (IAA) and analogues are produced by Frankia strains (Hammad et al., 2003). Actinobacterial auxin possibly plays a role in plant cell expansion, cell-wall remodelling, induction of adventitious roots, and in increasing the level of auxin in nodule primordia (Péret et al., 2007; Perrine-Walker et al., 2010).

Infection process and nodule development in actinorhizal plants

Two modes of root infection by Frankia have been described that depend on the host plant (Fig. 1) (Wall, 2000; Franche and Bogusz, 2011; Pawlowski and Demchenko, 2012). In the order Fagales, infection proceeds intracellularly via root hairs, whereas in Rosales, actinobacteria enter the root intercellularly. In Datiscaceae and Coriariaceae, the infection process is poorly known due to the difficulty involved in obtaining pure cultures of the symbionts and in studying the early stages of the infection process.

Fig. 1.

Schematic representation of actinorhizal root infection by Frankia. Frankia penetrates via a root hair infection process in host plants from the families Betulaceae, Casuarinaceae and Myricaceae, and intercellularly in Eleagnaceae, Rosaceae and Rhamnaceae. Prenodule formation resulting from mitotic activity in the root cortical cells is observed only during the intracellular infection process. Nodule primordia arise from divisions in root pericycle cells, located opposite a protoxylem pole, and near the site of infection. Frankia hyphae progress either from cell to cell in the intracellular mode of infection, or apoplastically in a matrix secreted into the intercellular spaces. Frankia hyphae progress towards the nodule primordium where they will penetrate developing cortical cells intracellularly. Mature nodules consist of multiple lobes. Adapted from Franche and Bogusz (2011)).

Fig. 1.

Schematic representation of actinorhizal root infection by Frankia. Frankia penetrates via a root hair infection process in host plants from the families Betulaceae, Casuarinaceae and Myricaceae, and intercellularly in Eleagnaceae, Rosaceae and Rhamnaceae. Prenodule formation resulting from mitotic activity in the root cortical cells is observed only during the intracellular infection process. Nodule primordia arise from divisions in root pericycle cells, located opposite a protoxylem pole, and near the site of infection. Frankia hyphae progress either from cell to cell in the intracellular mode of infection, or apoplastically in a matrix secreted into the intercellular spaces. Frankia hyphae progress towards the nodule primordium where they will penetrate developing cortical cells intracellularly. Mature nodules consist of multiple lobes. Adapted from Franche and Bogusz (2011)).

Intracellular infection begins with the deformation of root hairs induced by currently unknown Frankia signals. Actinorhizal hyphae are entwined by curled root hairs and only a few of them penetrate at the site of folding. Pronounced deposition of wall material is associated with Frankia invasion and hyphae become embedded in a structure analogous to the infection thread found in legume/rhizobia symbioses. Within this structure, Frankia filaments are encapsulated within a plant-derived cell-wall matrix consisting of xylans, cellulose and pectins (Berg, 1999). Frankia penetration triggers cell divisions in the root cortex subadjacent to the infected root hair, forming a mitotically active zone called the prenodule. The infection threads grow toward the prenodule, penetrate the thin wall of the recently expanded cortical cells, and infect prenodule cells that enlarge and ultimately fix nitrogen. Unlike in legumes, nodule primordia do not arise from these actively dividing cells of the prenodule. The nodule primordium arises from cell divisions induced in pericycle cells located opposite a protoxylem pole and close to a prenodule. In C. glauca, the prenodule cells displayed the same differentiation pattern as the one observed in the corresponding Frankia-infected cells of the nodule, suggesting that it may be a remaining form of a common nodule ancestor for legumes and actinorhizal plants (Laplaze et al., 2000a). In the following stage of nodule ontogenesis, the nodule primordium grows and becomes infected by Frankia hyphae progressing from the infected cells of the prenodule. During the infection process, Frankia first grows as filamentous hyphae that proliferate in the newly infected host cell, and when the infected cell matures, the tips of the hyphae eventually differentiate vesicles that will fix nitrogen (Newcomb and Wood, 1987).

No root hair deformation is observed in the intercellular root invasion process. Frankia hyphae penetrate the middle lamella between adjacent cells of the root epidermis and progress apoplastically between cortical cells, within an electron-dense matrix secreted into the intercellular spaces (Wall and Berry, 2008), which might represent the equivalent of the encapsulation material described previously in infection threads. Cell divisions are induced in the root pericycle opposite a protoxylem pole, leading to the nodule primordium. Frankia hyphae infect primordium cells from the apoplast by intense branching of hyphae, concomitant with continuous invagination of the plant plasma membrane.

Mature actinorhizal nodules are multilobed structures, each lobe exhibiting a central vascular bundle surrounded by an endoderm, an expanded cortex and a periderm. Due to activity of the apical meristem, nodule lobes show indeterminate growth and developmental zonation with specific patterns of gene expression (Duhoux et al., 1996; Obertello et al., 2003; Franche and Bogusz, 2011; Pawlowski and Demchenko, 2012). Structural organization varies among actinorhizal root nodules, with some nodules such as those of C. glauca exhibiting a so-called nodular root at the apex of each lobe; this root is believed to facilitate the diffusion of gases in and out of the nodule lobe (Callaham and Torrey, 1977; Tjepkema, 1978; Schwintzer and Lancelle, 1983).

Molecular mechanisms underlying actinorhizal infection and nodulation

Whereas no tools are yet available in Frankia to perform functional analysis of candidate genes, genetic transformation procedures based on Agrobacterium tumefaciens and A. rhizogenes are available for some actinorhizal plants, providing a tool for promoter studies and down-regulation of plant genes by RNAi (Franche et al., 1997; Gherbi et al., 2008a, b; Svistoonoff et al., 2010).

Functional and transcriptome analyses revealed that the common SYM pathway shared by rhizobium–legume and arbuscular mycorrhizal (AM) symbioses also controls nodulation by Frankia (Gherbi et al., 2008a, b; Markmann et al., 2008; Hocher et al., 2011). This pathway includes a receptor-like kinase, nuclear pore proteins and potassium channels required for the induction of calcium oscillations. A putative calcium/calmodulin-dependent protein kinase (CCaMK) is also present and might thus recognize calcium ‘actinorhizal signatures’ (Singh and Parniske, 2012). Interestingly, transcriptome analysis also revealed the presence of genes linked to a ‘NOD’-specific pathway (not shared with AM symbiosis) used by legumes for the nodulation process with rhizobia. These interesting data suggest the possibility of a similar ‘NOD’ pathway between RNS. This overlapping of legume and actinorhizal RNS reinforces the hypothesis of a common genetic ancestor with a genetic predisposition for nodulation in the nitrogen-fixing clade Rosid I (Soltis et al., 1995).

In addition to sequences involved in the Frankia signalling pathway, genes Ag12 and Cg12, which encode subtilisin-like serine proteases in A. glutinosa and C. glauca, respectively, have also been studied (Ribeiro et al., 1995; Laplaze et al., 2000b). Subtilases are a superfamily of proteases thought to play a role in different aspects of plant development, including lateral root initiation and responses to pathogens. Using transgenic Casuarinaceae containing Cg12 promoter–reporter gene fusions, it has been shown that the expression of Cg12 starts very early during the symbiotic process and is specifically induced in cells infected by Frankia (Svistoonoff et al., 2003). When Cg12-reporter gene fusions were introduced in the legume M. truncatula, a similar pattern of expression was observed during the nodulation process with Mesorhizobium meliloti (Svistoonoff et al., 2004). The conservation of the expression profile in M. truncatula suggests that a signalling pathway independent of Nod factors, and conserved between the two systems, is activated specifically in cells infected by symbiotic bacteria. Intracellular infection mechanisms in legumes and actinorhizal plants were further investigated by introducing the Enod11 promoter from M. truncatula into C. glauca (Svistoonoff et al., 2010). In M. truncatula, MtEnod11 gene expression was shown to be correlated with both preinfection and infection events throughout nodulation (Journet et al., 2001). In C. glauca, activation of the ProMtEnod11::gus reporter was shown to be correlated with Frankia infection in root hairs, prenodules and nodules. These results suggest high conservation of regulatory pathways between legumes and actinorhizal plants in cells involved in bacterial infection and accommodation. However, ProMtEnod11 is not activated in transgenic Casuarina prior to infection, during the perception of Frankia signal(s), indicating that the pre-infection stage differs between actinorhizal and legume–rhizobium symbioses (Svistoonoff et al., 2010).

Since several plant hormones have been reported to regulate nodulation in legumes (Ding and Oldroyd, 2009), the C. glauca auxin influx carrier gene CgAux1 was characterized to investigate the role of auxin in actinorhizal symbiosis. Using a transcriptional fusion between the promoter region and the β-glucuronidase (gus/uidA) reporter gene, CgAux1 was shown to be expressed during nodule ontogenesis in all Frankia-infected cells, including root hairs (Péret et al., 2007). Moreover, accumulation of auxins was reported in Frankia-infected cells in actinorhizal nodules of C. glauca and this accumulation was shown to be driven by cell-specific expression of auxin transporters and by Frankia auxin biosynthesis in planta (Perrine-Walker et al., 2010). This localization raises the question of the role of auxin in infected cells. It was suggested that auxin might be involved in the cell wall remodelling processes that occur upon Frankia infection resulting from the growth of infection threads (Perrine-Walker et al., 2010). Jasmonic acid belongs to another class of signalling molecules that may be involved in root endosymbioses with AM fungi and rhizobia (reviewed in Hause and Schaarschmidt, 2009). However, recent results suggest that jasmonic acid does not play a role in nodulation of the model legume Medicago truncatula or in the two actinorhizal plant species, C. glauca and D. glomerata (Zdyb et al., 2011).

Among infection mechanisms leading to RNS, the intracellular infection pathway is probably the most ancestral but also one of the least characterized. To decipher the molecular mechanisms underlying intercellular infection with Frankia, and to enable a comparative analysis between intra- and intercellular processes of infection, some tools for functional analysis of candidate genes were recently developed in Discaria trinervis (Rhamnaceae), a shrub endemic to Patagonia. Intercellular infection by Frankia was first analysed in detail in a time-course experiment in D. trinervis (Valverde and Wall, 1999), and an efficient genetic transformation protocol for D. trinervis based on A. rhizogenes was then set up (Imanishi et al., 2011). First data indicate that ProMtEnod11 drives expression in the infection zone of D. trinervis nodules, thus suggesting conservation of the corresponding regulators among all plants able to enter RNS regardless of the infection mechanism involved. As mentioned previously, ProCg12 and ProCgAux1 are two other promoters that drive expression in infected root hairs and prenodules prior to nodule formation in C. glauca (Péret et al., 2007; Svistoonoff et al., 2003, 2004). Analyses of these two promoters are currently underway in the intercellularly infected actinorhizal plant D. trinervis. The expression patterns that will be observed during the different stages of the root infection and nodule ontogenesis should help to explore signalling mechanisms in symbioses with this ancestral mode of infection, as well as to identify both conserved and divergent regulatory mechanisms (Rhizogenesis laboratory, France and University of Quilmes, Argentina, unpubl. res.).

Unlike legume nodules, actinorhizal nodules are modified lateral roots. Since auxin is a key signal in lateral root initiation, development, emergence and meristem activation (Overvoorde et al., 2010), this raises the question of the role of auxin during actinorhizal nodule development. Interestingly, in studies on Alnus root nodules development, Angulo Carmona (1974) suggested that nodules do not form from pre-existing lateral root primordia. In addition, Wheeler et al. (1979) showed that the number of lateral root primordia initiated on Alnus roots following inoculation by Frankia was higher than on uninoculated control roots, indicating that the actinomycete can stimulate lateral root initiation. Further work is thus needed to characterize the role of auxin in symbiotic root development and to understand to what extent the lateral root developmental programme has been hijacked by Frankia to form an actinorhizal nodule. A first result was obtained by Sy et al. (2007) who showed that the cell-cycle promoter cdc2aAt from Arabidopsis thaliana has retained its ability to be induced by hormones in transgenic Allocasuarina verticillata roots and that, upon Frankia infection, this promoter was strongly induced in pericycle cells. These data suggests that, in response to Frankia, some pericycle cells in lateral roots can recover mitotic competence due to changes in their hormonal balance.

Recently, microarray expression analysis of transcripts in nodules versus uninfected roots showed that 1500 genes in A. glutinosa and 2000 in C. glauca are regulated or specifically induced in nodules (Hocher et al., 2011). The majority of regulated genes are involved in transport (e.g. DCAT, dicarboxylate transporter for delivery of photosynthates to the symbiont), metabolism (e.g. GS, glutamine synthetase for the assimilation of the ammonium fixed and its transfer to the plant) and protein synthesis machinery, as expected during a switch from root-specific to nodule-specific gene expression. The analysis also revealed regulated genes involved in cell wall structure, defence (defensins, chitinases) and response to stress (catalase, DnaJ, γ-expansin natriuretic peptide), which is consistent with the results of previous studies (Ribeiro et al., 2011).

O2 regulation and haemoglobin in actinorhizal nodules

Several reviews summarize many aspects of oxygen metabolism in actinorhizal symbioses (Silvester et al., 1990; Pawlowski, 2008). As mentioned above, in the free-living state, Frankia has its own oxygen protection mechanism: it forms specialized thick-walled cells (vesicles) that protect nitrogenase from oxygen (Torrey and Callaham, 1982). Actinorhizal plants have developed different strategies to reduce oxygen levels in nodules, thus reflecting the taxonomic diversity of the hosts and the different structural organization of the actinorhizal nodules. In Casuarina, where Frankia strains do not form vesicles within nodules, cell-wall lignification observed after Frankia infection is believed to provide a barrier against oxygen diffusion (Berg and McDowell, 1988) together with a large amount of symbiotic haemoglobin (class 2) (Fleming et al., 1987; Jacobsen-Lyon et al., 1995; Gherbi et al., 1997). In contrast, in Alnus and Myrica nodules where Frankia do not form vesicles, class 2 symbiotic haemoglobin was not found. However, nonsymbiotic class 1 haemoglobin was detected in A. glutinosa (Suharjo and Tjekema, 1995) and Myrica gale nodules (Pathirana and Tjepkema, 1995). The corresponding genes were cloned in A. firma (Sasakura et al., 2006) and M. gale (Heckmann et al., 2006). Alnus haemoglobin 1 was strongly induced in actinorhizal nodules by nitric oxide (NO) and cold stress, but not by hypoxia or osmotic stress (Sasakura et al., 2006). As reported by Hill (2012), class 1 nonsymbiotic haemoglobin is involved in reactive oxygen and NO metabolism. A truncated haemoglobin, classified in class 3 (Watts et al., 2001), was also been identified in the actinorhizal plant D. glomerata (Pawlowski et al., 2007). This truncated haemoglobin is induced upon plant infection by Frankia, leading the authors to suggest a role in NO detoxification. Although reactive oxygen (Tavares et al., 2007) and possibly NO production occur during interaction with Frankia, the role of class 1 and 3 haemoglobins in actinorhizal symbioses has not yet been fully elucidated.

NON-LEGUME ROOT ENDOSYMBIOSIS PARASPONIA-RHIZOBIUM

General features of the symbiosis

In the order Rosales, in addition to actinorhizal nitrogen-fixing plants, Parasponia species (family Cannabaceae,) also display an original nitrogen-fixing root symbiosis (Sytsma et al., 2002). Parasponia is the only non-legume host plant known to be nodulated by rhizobia (Trinick, 1973; Akkermans et al., 1978). The host plants are medium-sized tropical trees (up to 15 m in height), and pioneer species growing on nitrogen-poor and disturbed soils. These trees originated from the Malay Archipelago. Among the five nodulated species identified (Becking, 1992), P. andersonii/Rhizobium is the most widely studied symbiotic association.

Although different Rhizobium species, including some strains isolated from legume nodules, are capable of nodulating Parasponia species (Trinick and Galbraith, 1980; Trinick and Hadobas, 1988), they do not represent a specific lineage, thus suggesting the recent emergence of the ability of the host plants to be nodulated by rhizobia (Lafay et al., 2006). More recently, P. andersonii was tested for the symbiotic effectiveness of a wide range of Rhizobium species and was found to be nodulated by bacteria from four different genera harbouring highly diverse Nod factor biosynthesis genes (Op den Camp et al., 2012). Although Parasponia allows such symbiotic promiscuity of rhizobia endosymbionts, the efficiency of the symbiotic nitrogen fixation varies. Microscopy studies of nodules obtained with under-performing rhizobia indicate that the control of symbiotic association is less sophisticated than with legumes (Trinick and Hadobas, 1988; Op den Camp et al., 2012).

Rhizobium and plant host signalling

Like in most legume–rhizobium symbioses, the Parasponia–rhizobium symbiosis depends on Nod factors. Rhizobial Nod factors are lipochitooligosaccharides that consist of an acylated chitin oligomeric backbone with different functional group substitutions at the terminal or non-terminal residues. These Nod factors are key symbiotic signals and are indispensable in the specific-host rhizobium interaction and at later stages in the infection process and nodule organogenesis (Oldroyd and Downie, 2008). Recently, it was shown that a single gene closely related to lysin-motif (LysM) domain proteins involved in Nod factor perception in legumes, is required for both nodulation and mycorrhization in P. andersonii (Op den Camp et al., 2011). It was also shown that the common ‘SYM’ pathway described for AM, legume–rhizobium and actinorhizal symbioses is activated during P. andersonii nodule organogenesis. Together, these data reinforce the hypothesis of a common genetic ancestor of the nodulating clade with a genetic predisposition for nodulation (Soltis et al., 1995). While considerable information is available on the role of particular flavonoids in the rhizobium–legume symbiosis, their role during the different stages of Parasponia nodulation is not known. Because it is a Nod-dependent symbiotic interaction, it is likely that the role of flavonoids produced by Parasponia is similar to that played in legume nodulation.

Infection process and nodule structure

The infection process that leads to nodule development occurs via the so-called crack entry mode of infection (Sprent and Faria, 1988). Rhizobia enter the root between epidermal cells, this intercellular infection being concomitant with the formation of infection threads of plant origin and with the stimulation of cortical cell division leading to a subsurface swelling zone comparable with the prenodule in actinorhizal plants (Lancelle and Torrey, 1984a, b; Bender et al., 1987). As the infection threads continue to grow, a few pericycle cells in the immediate vicinity of the prenodule divide giving rise to a nodule-lobe primordium. Progressively, the threads intracellularly invade cells derived from the apical region of the nodule lobe primordium leading to a nodule containing an apical meristem and a central vascular cylinder, which is surrounded by a zone of infected tissue. As in actinorhizal plants, nodule primordium formation does not involve prenodule cells, and the function of prenodules is still not known. The rhizobia remain in threads throughout the symbiotic process and are not released from the threads unlike during bacteroid formation in rhizobium–legume symbioses (Trinick, 1979; Lancelle and Torrey, 1984b). In contrast to legumes, where nodule primordia are initiated in the cortex and have a stem-like anatomy with a peripheral vascular bundle, ontogenesis and the final structure of the Parasponia nodule lobe is similar to that observed in actinorhizal symbiosis and resembles lateral roots.

Control of oxygen levels in nodules

Symbiotic haemoglobin (Hb) proteins and genes have been isolated from Parasponia and characterized (Appleby et al., 1983; Landsmann et al., 1986). Parasponia andersonii was shown to possess a single Hb gene expressed in both nodules and in non-nodulated roots, suggesting symbiotic and non-symbiotic roles result from a single gene (Landsmann et al, 1986, 1988; Bogusz et al., 1988). Its oxygen-binding properties and cellular location in young rhizobia-infected cells are consistent with a role in oxygen transport to rhizobia within root nodules (Wittenberg et al., 1986; Gibson et al., 1989; Trinick et al., 1989). Furthermore, comparative analysis of promoter activity in transgenic Casuarina indicated that, in contrast to the lack of conservation of cell-specific expression of P. andersonii haemoglobin promoter in transgenic legume nodules (Bogusz et al., 1990), it retains its specific expression in bacteria-infected cells of actinorhizal nodules (Franche et al., 1998). As mentioned above, it is interesting that P. andersonii and C. glauca nodules have the same origin and structure as lateral roots. This suggests that in Rosales, beyond similar nodule structure and ontogeny, there is a non-legume key symbiotic gene that has identical regulatory mechanisms (Franche et al., 1998).

Along with legume–rhizobium and actinorhizal symbioses, Parasponia is a key species for studies of the accommodation of symbiotic bacteria in plant cells. A comparative analysis of these three symbiotic systems should help define strategies for transferring nitrogen-fixing ability to non-legume crops.

CYANOBACTERIAL–PLANT ENDOPHYTIC AND ENDOSYMBIOTIC ASSOCIATIONS

A wide range of cyanobacterial associations

Cyanobacteria are a diverse group of oxygenic photosynthetic prokaryotes that occur in marine, aquatic and terrestrial environments all over the world (Rippka et al., 1979). Some cyanobacteria have the ability to live in association with a wide range of plants from the divisions Bryophyta (liverworts and hornworts), Pteridophyta (the genus Azolla), gymnosperms (family Cycadaceae) and angiosperms (family Gunneraceae) (Table 1) (for reviews see Meeks, 1998; Adams, 2000; Rai et al., 2000, 2002; Adams et al., 2006; Bergman et al., 2007). A striking difference between cyanobacteria–plant associations, and the non-legumes actinorhiza–Frankia and Parasponia–Rhizobium symbioses where the bacteria are hosted in a root nodule, is that the plant structure colonized by the symbiotic cyanobacteria develops independently of cyanobacterial infection.

Table 1.

Main features of plant cyanobacterial symbiotic associations

Plant taxon Symbiotic host species Symbiotic Cyanobiont structure Proposed time for plant origin 
Angiosperm All known species of Gunnera Stem gland Intracellular Nostoc 80 Mya 
Gymnosperm All known cycads (150 species in 10 genera belonging to 3 families) Root zone Intercellular Nostoc or Calothrix 200–150 Mya 
Pteridophyte All species of the genus Azolla Cavities in each dorsal leaf Intercellular Nostocales obligatory symbiont 420 Mya for the ferns, 120 Mya for Azolla fossils 
Bryophyte Only two of the 330 genera of liverwort; four of the six genera of hornwort Cavities in the gametophyte Intercellular Nostoc 400–500 Mya 
Plant taxon Symbiotic host species Symbiotic Cyanobiont structure Proposed time for plant origin 
Angiosperm All known species of Gunnera Stem gland Intracellular Nostoc 80 Mya 
Gymnosperm All known cycads (150 species in 10 genera belonging to 3 families) Root zone Intercellular Nostoc or Calothrix 200–150 Mya 
Pteridophyte All species of the genus Azolla Cavities in each dorsal leaf Intercellular Nostocales obligatory symbiont 420 Mya for the ferns, 120 Mya for Azolla fossils 
Bryophyte Only two of the 330 genera of liverwort; four of the six genera of hornwort Cavities in the gametophyte Intercellular Nostoc 400–500 Mya 

Bryophytes are small, non-vascular land plants including liverworts (Hepaticae), horworts (Anthocerotae) and mosses (Musci), a relatively small number of which are able to form epiphytic or endophytic associations with cyanobacteria (Adams, 2002; Meeks, 2003; Adams and Duggan, 2008). Epiphytic associations with mosses are not discussed in this review. Two liverwort species, Blasia pusilla and Cavicularia densa (Blasiales, Marchantiophyta), and all hornworts (Anthocerophyta) are able to form an endosymbiotic association with cyanobacteria that generally belong to the genus Nostoc (Rodgers and Stewart, 1977; Adams and Duggan, 2008). Endosymbiont filaments are hosted in specialized auricles on the ventral surface in Blasiales and in slime cavities within the thallus in Anthicerotophyta such as Anthoceros and Phaeoceros. During the development of the thallus, new auricles are continuously formed and infected by cyanobacteria.

Cyanobacterial associations with pteridophytes are limited to the genus Azolla in the family Azollaceae. Azolla is a small floating aquatic fern with a worldwide distribution ranging from tropical to warm temperate regions. It has been exploited for many years as a source of nitrogen for agriculture and is extensively used as a green manure and biofertilizer for rice (Watanabe and Roger, 1984; Ladha et al., 2000; Van Hove and Lejeune, 2002). The nitrogen-fixing cyanobacteria are hosted in a highly specialized cavity located on the dorsal lobe of the leaves (Peters and Mayne, 1974; Zheng et al., 2009). An envelope lines the cavity where cyanobacterial filaments are localized in the periphery within a mucilaginous matrix surrounding a gaseous central region. Morphological analysis of the leaf cavity established that a pore remains open during leaf development, even when the leaf is mature, thus permitting gas exchanges (Veys et al., 1999). Throughout its life cycle, the symbiont remains associated with its host, and is automatically transmitted from generation to generation, including during sexual reproduction (Calvert et al., 1985; Peters and Meeks, 1989). So far, there are no confirmed reports of successful in vitro cultivation of the cyanobiont that belongs to the order Nostocales, making Azolla symbiosis the only known permanent symbiosis among cyanobacteria–plant associations (Lechno-Yossef and Nierwicki-Bauer, 2002; Pabby et al., 2003). Besides the cyanobiont, it has been shown that minor cyanobacterial and bacterial species coexist in the cavity (Gebhart and Nierzwicki-Bauer, 1991).

Cycads are the only known gymnosperms that have the ability to develop a nitrogen-fixing symbiosis through an intimate association with cyanobacteria. Cycads include approx. 156 species in nine genera that grow naturally in tropical and subtropical regions, and all of them possess a symbiotic cyanobacterium. Cyanobacteria are hosted in specialized coralloid roots that arise from the lateral roots and are formed by the plant before being invaded by the cyanobacteria (Costa and Lindblad, 2002). Filamentous cyanobacteria are located intercellularly in a zone filled with mucilage and comprise a large number of elongated cycad cells that interconnect two adjacent cortical layers in the coralloid roots. These cells may contribute to the transfer of metabolites between the symbionts and the host. In transverse root sections, cyanobacteria are visible as a green ring. Nostoc spp. are the most common cyanobionts in Cycadaceae, although Calothrix has occasionally been reported (Costa et al., 1999; Thajuddin et al., 2010).

Plants in the genus Gunnera are mainly distributed in the southern hemisphere with about 40 species, including small, stoloniferous species such as Gunnera magellanica and plants that can reach 3 m in height (e.g. Gunnera manicata). Natural populations are restricted to humid areas with heavy rainfall (Bergman et al., 1992a; Bergman, 2002). The cyanobiont identified as Nostoc enters Gunnera plants through specialized glands located on the stem and observed in conditions of nitrogen starvation, even in absence of cyanobacteria (Silvester and McNamara, 1976; Towata, 1985; Bonnett, 1990; Chiu et al., 2005). The glands secrete polysaccharide-rich mucilage that attracts specific symbiotic cyanobacteria (Nilsson et al., 2006), supports their growth on the gland surface, and contributes to their differentiation (Chiu et al., 2005). After entering the gland through existing channels, cyanobacteria induce divisions in the host cells lining the channel and are subsequently taken up into the host cells (Bergman et al., 1992a). Cyanobacteria become restricted to certain Gunnera cells intermixed with non-infected cells. Once inside the Gunnera cells, Nostoc filaments grow and divide and fill most of the host cell. As observed with Frankia in nodular structures, filaments are always surrounded by the host plasma membrane. After colonization, new glands continue to form on the stem at the base of each leaf and infection continues at the apex of the growing stem. In large Gunnera species, the symbiotic tissue is visible as blue-green patches along the rhizomatous stems or along stolons in smaller Gunnera species (Bergman, 2002).

The intimacy of the symbiosis Gunnera–Nostoc is therefore greater than in other cyanobacterial associations since the cyanobacteria are hosted intracellularly, whereas in bryophytes, Azolla and cycads, the cyanobionts occur as endophytes in mucus-filled cavities. With the exception of Azolla, the associations are facultative, the two partners can be isolated and cultivated independently, and the symbiosis can be easily reconstituted.

Diversity of cyanobacteria associated with plants

Cyanobionts are filamentous cyanobacteria and generally belong to the genus Nostoc, although a few other cyanobacteria such as Calothrix and Chlorogloeopsis have been reported. Cyanobacteria of the genus Nostoc belong to the order Nostocales and to Section IV of cyanobacteria (Rippka et al., 2001). They are all characterized by their ability to differentiate some nitrogen-fixing cells called heterocysts, some resting spores called akinetes, and some motile filaments called hormogonia, which constitute the infective units during the establishment of the symbiotic process and contribute to short distance dispersal in free-living conditions (Campbell and Meeks, 1989; Vagnoli et al., 1992; Adams and Duggan, 2008).

Molecular techniques have shown that, with the exception of the cyanobacteria living in the cavities of the water fern Azolla, there is high strain diversity both within and among the different plant hosts (Costa et al., 1999, 2001; Nilsson et al., 2000; Guevara et al., 2002; Rasmussen and Nilson, 2002; Zheng et al., 2002; Papaefthimiou et al., 2008; Rikkinen and Virtanen, 2008). For instance, by studying the 16S rRNA sequence of cyanobionts, a single coralloid root of Cycas revoluta was found to harbour up to three cyanobacterial strains and some diversity was also observed in multiple roots from a single plant (Gheringer et al., 2010; Yamada et al., 2012).

The limited genetic variation in cyanobacterial symbionts from Azolla was first revealed using RFLP analysis with nif gene probes and PCR fingerprinting (Franche and Cohen-Bazire, 1985, 1987; Plazinski et al., 1988; Zheng et al., 1999) and recently confirmed by a phylogenetic tree based on 16S rRNA gene sequences of 35 symbionts extracted directly from leaf cavities of Azolla species collected in different geographical regions. Nostoc azollae form a homogenous cluster separated from free-living cyanobacterial genera (Papaefthimiou et al., 2008). These data suggest that one cyanobacterial taxon originally infected an ancestor of the Azolla species and that the symbiosis ultimately became obligatory for the cyanobacterial partner. Such features suggest long-lasting co-evolution between the partners, potentially extending back as far as 140 million years, corresponding to the oldest fossil records of Azolla (Raven, 2002).

Except for N. azollae, plant–cyanobacteria associations can be reconstituted under laboratory conditions in bryophytes, Gunnera and, to a lesser extent, in cycads, offering a way to perform studies on the two partners alone and in symbiotic association. It is interesting to note that the cyanobacterial strains that form intracellular symbioses with Gunnera remain extracellular when they infect the hornwort Anthoceros, and extracellular symbionts that infect Anthoceros are able to become intracellular symbionts with Gunnera. The possibility for the same Nostoc strain such as N. punctiforme ATCC 29133 (also referred as PCC 73102) to infect a wide range of hosts, points to a lower degree of specificity in the cyanobacteria–plant associations than that observed in the highly specific Rhizobium–legume symbioses or even in actinorhizal symbioses. N. puctiforme was first isolated from the roots of the cycad Macrozamia sp. in Australia, and has the ability to form a symbiotic association with the angiosperm Gunnera (Johansson and Bergman, 1994) and the bryophyte A. punctatus (Enderlin and Meeks, 1983).

In addition to taxonomic studies on symbiotic cyanobacteria, data are accumulating on cyanobacterial genomes, with >40 genome sequences of cyanobacterial strains available and many genomes still to come (http://www.ncbi.nlm.nih.gov/genomes/lproks.cgi) (Hess, 2011). Genome sizes vary from 1·44 Mb to 9·05 Mb, with reported genes ranging from 1241 to 8462. Most genomes are circular and a small number of plasmids can be observed. The facultative symbiont N. punctiforme ATCC 29133 has one of the largest genomes: 9·05 Mb and 8462 reported genes (Meeks et al., 2001). For comparison, the free-living non-symbiotic reference strain Anabaena PCC 7120 genome is 7.13 Mb and contains 5610 ORFs. The sequence of N. punctiforme indicates a genome that is highly plastic and in a state of flux, with numerous insertion sequences and multilocus repeats, as well as encoding transposases and DNA modification enzymes (Meeks et al., 2001; Meeks, 2005a, 2009). In contrast to N. punctiforme, one obligatory symbiont of Azolla, N. azollae 0708, has a small genome of 5486 Mb with 5413 genes, among which 1689 are pseudogenes (Ran et al., 2010; Larsson et al., 2011). The number of intact coding sequences is among the lowest in filamentous cyanobacteria sequenced to date.

The role of hormogonia in the infection process

To establish a successful interaction, host plants must attract and internalize the cyanobacteria and then regulate their growth and differentiation. The cyanobacteria must avoid eliciting the plant defence response, and must adapt their metabolism to a new environment. Like in the previously described symbioses with actinorhizal plants and Parasponia, these events require sophisticated communication between the plant and the cyanobacteria (Fig. 2) (Gorelova, 2006; Adams and Duggan, 2011).

Fig. 2.

Schematic representation of the infection process in cyanobacteria–plant symbioses. In nitrogen-free medium, Nostoc sp. filaments consist of vegetative cells (V) and regularly spaced heterocysts (H) that fix nitrogen. A hormogonium inducing factor (HIF) produced by the host under nitrogen starvation conditions leads to differentiation of motile small-celled hormogonial structures. Following the exchange of appropriate recognition signals, hormogonia penetrate the host symbiotic cavities and revert to vegetative filaments with a large number of heterocysts. The repression of hormogonia is linked to a hormogonia repressing factor (HRF). In ageing symbiotic tissues, multiple contiguous heterocysts are observed that exhibit low nitrogen-fixation activity. Adapted from Rai et al. (2000) and Meeks (2005b).

Fig. 2.

Schematic representation of the infection process in cyanobacteria–plant symbioses. In nitrogen-free medium, Nostoc sp. filaments consist of vegetative cells (V) and regularly spaced heterocysts (H) that fix nitrogen. A hormogonium inducing factor (HIF) produced by the host under nitrogen starvation conditions leads to differentiation of motile small-celled hormogonial structures. Following the exchange of appropriate recognition signals, hormogonia penetrate the host symbiotic cavities and revert to vegetative filaments with a large number of heterocysts. The repression of hormogonia is linked to a hormogonia repressing factor (HRF). In ageing symbiotic tissues, multiple contiguous heterocysts are observed that exhibit low nitrogen-fixation activity. Adapted from Rai et al. (2000) and Meeks (2005b).

The conversion of vegetative filaments into motile and short-lived hormogonia is an essential step for the establishment of the symbiotic process and in laboratory conditions, and a cyanobacterial culture rich in hormogonia can increase the efficiency of plant infection (Uheda and Silvester, 2001). Hormogonia are short gliding filaments that lack heterocysts, with cells that are smaller than the cells in vegetative filaments (Duggan et al., 2007). The reduced cell size results from cell divisions that are not accompanied by an increase in cell biomass and a significant synthesis of DNA. Different environmental stimuli and/or plant factors released during nitrogen starvation can stimulate the induction of hormogonia (Meeks and Elhai, 2002; Adams et al., 2006). Hormogonia are in a transient, non-growth state, and they maintain their gliding activity for 48–72 h before reverting back to vegetative growth. During the interaction with a host plant, hormogonia revert to filaments with nitrogen-fixing heterocysts after entering the host.

Under low nitrogen conditions, certain plants exude one or several hormogonia-inducing factors (HIF), thereby dramatically increasing the frequency at which nearby Nostoc spp. filaments convert to hormogonia (Campbell and Meeks, 1989). As reviewed by Adams and Duggan (2008), HIF have been found in the hornwort A. punctatus (Meeks, 2003), in Blasia, cycads and the angiosperm Gunnera (Rasmussen et al., 1994; Bergman et al., 1996; Knight and Adams, 1996; Cohen and Meeks, 1997; Campbell et al., 1998; Ow et al., 1999). In Gunnera, the viscous mucilage secreted by the stem contains an HIF and extracts from other tissues or seeds have no effect on the cyanobacteria. In both Gunnera and A. punctatus, the putative signal has been identified as a heat labile compound, possibly a protein of <12 kDa (Campbell and Meeks, 1989; Rasmussen et al., 1994; Meeks, 2003). Cycad root extracts also promoted hormogonium formation in competent Nostoc. Surprisingly, HIF is not restricted to symbiotic plant extracts since hormogonium formation has also been detected in artificial associations between cyanobacteria and non-host plants such as wheat (Gantar et al., 1991, 1993; Gusev et al., 2002). The biochemical composition of the HIF(s) has not yet been determined.

Although our knowledge on the molecular events involved in the differentiation of hormogonia is still limited, several genes affecting this process have been identified. In contrast to Frankia where no genetic tools are available to create mutations, procedures for genetic analysis and transposon mutagenesis have been developed in the large host range Nostoc ATCC 29133, thus providing a valuable tool for investigating the function of putative symbiotic genes (Cohen et al., 1994). A mutation in NtcA, which encodes a transcription factor essential for nitrogen metabolism, resulted in a reduction in the frequency of hormogonia induced by the HIF and the resulting hormogonia did not infect Anthoceros (Wong and Meeks, 2002). Mutations in sigH and trpN which, respectively, encode an alternative group 2 sigma factor and a tetratricopeptide repeat protein, were also associated with increased symbiotic competence (Campbell et al., 1998). In 2008, Chapman et al. isolated two different mutants of N. punctiforme resulting from the insertion of a transposon in the cyaC gene encoding an adenylate cyclase which catalyses the formation of the intracellular messenger cyclic AMP. Following cocultivation experiments with the symbiotic partner B. pusilla, both mutant strains were found to form hormogonia about 12 h earlier than in the wild-type strain, and displayed reduced symbiotic competence. Recently knowledge on hormogonia differentiation has progressed thanks to transcriptome analyses by Campbell et al. (2007, 2008). Their data revealed that, although hormogonia do not grow, they are characterized by a high dynamic transcriptional state, with 1827 genes differentially transcribed in N. punctiforme 24 h after their induction. A plant extract of A. punctatus containing the still unknown HIF was found to down-regulate 345 genes and up-regulate 689 genes in 30 min (Campbell et al., 2008).

Hormogonium production is not sufficient for the establishment of the symbiosis, since some non-infective Nostoc strains are also capable of forming motile hormogonia in the presence of the angiosperm Gunnera spp. (Johansson and Bergman, 1994; Rasmussen et al., 1994). Rapid migration of hormogonia inside the symbiotic structure is a critical factor for the successful establishment of nitrogen-fixing associations (Nilsson et al., 2005). This step in the process involves a combination of motility and chemoattraction. The cell surface of hormogonia of symbiotically competent Nostoc is covered with pili that confer a form of surface motility called gliding. In contrast, immotile vegetative trichomes of mature Nostoc filaments are devoid of these specialized filaments (Duggan et al., 2007). Following the analysis of insertion mutants in N. punctiforme ATCC 29133, the inactivation of two ORF coding for the pilT and pilD coding for pilus-like structures was seen to alter the level of surface piliation and to reduce the symbiotic competency of N. punctiforme in Blasia (Duggan et al., 2007).

Chemoattraction of cyanobacteria was first shown with the host plant Blasia (Knight and Adams, 1996). Although hormogonium is a prerequisite for symbiosis, not all hormogonia-forming cyanobacteria are capable of infecting plants (Rasmussen et al., 1994) and chemoattraction is an important factor in the initiation of the association. Whereas factors stimulating the formation of hormogonia are not highly specific, the induction of chemotaxis could be a more specific event. In Blasia, a chemoattractant compound was characterized as a low molecular-weight (<1 kDa) compound, stable at temperatures up to 95 °C, and almost completely inactivated by acetic acid (Watts et al., 1999). Some sugars such as arabinose, glucose and galactose can also attract hormogonia (Nilsson et al., 2006). Although chemoattractants are responsible for some specificity in the interaction, they can also be secreted by non-host plants. Chemotactic attraction has been detected in wheat (Gantar et al., 1993; Gusev et al., 2002; Nilsson et al., 2002), in Trifolium rupens, and to a lesser extent in Arabidopsis thaliana and Oryza sativa (Nilsson et al., 2006).

Cyanobacteria in symbiotic status

Symbiosis causes modifications in both partners. For the host, symbiosis leads to an increase in size of the symbiotic structures and organs inhabited by the cyanobiont caused by local plant cell proliferation, the accumulation of mucus, and the formation of specialized plant cells penetrating cyanobacterial colonies that contribute to the exchange of metabolites. For the cyanobiont, after entering the host, hormogonia revert to non-motile vegetative filaments, the differentiation of nitrogen-fixing heterocysts is observed at an unusually high frequency, and vegetative cells show altered morphology, i.e. are enlarged and irregular in shape comparedwith their free-living counterparts (Meeks and Elhai, 2002).

The repression of hormogonia is linked to the release by the host of an unidentified hormogonia-repressing factor. This factor induces genes such as hrmA belonging to the hormogonium-regulating locus hrmRIUA, which plays a central role in the repression of hormogonia formation and exhibits high similarity with sugar uronate metabolism operons of bacteria (Meeks, 2003). hrmA is induced by an aqueous extract of Anthoceros tissue, leading to the suggestion that a factor in the extract prevents hormogonium formation (Cohen and Meeks, 1997). In addition to the flavonoids, naringenine and, to a lesser degree, neohesperidine and prunine, can induce the expression of hrmA (Cohen and Yamasaki, 2000). hrmA was also induced by aqueous extracts of fronds of Azolla pinnata and A. filiculoides, and this induction was correlated with the amount of deoxyanthocyanin contained in the extract, even though pure deoxyanthocyanin had only a weak effect on the induction process (Cohen et al., 2002). Recently, quantitative analyses of soluble sugars in the mucilage of Gunnera revealed that low levels of soluble sugars help attract the cyanobiont N. punctiforme, but are not involved in the formation of hormogonia. Conversely, high levels of soluble sugars in plant cells help prevent further development of hormogonia once Nostoc is inside the plant cells (Khamar et al., 2010).

The development of heterocysts after plant infection is essential for a functional nitrogen-fixing symbiosis. Whereas in a free-living Nostoc strain, the average heterocyst frequency in a nitrogen-free medium is 5–10 %, heterocysts appear at an average frequency of 35 % in plant associations, and a gradient in the number of heterocysts is observed (Meeks and Elhai, 2002). In the glands of Gunnera, 5–10 % of heterocysts were observed in the young glands and up to 75 % in older glands (Bergman et al., 1992a; Zhang et al., 2006). At the molecular level, differentiation from a vegetative cell to a heterocyst is a complex process that is regulated by several important genes (Buikema and Haselkorn, 1991; Adams and Duggan, 1999; Haselkorn, 2007). The expression of four cyanobacterial genes connected to signalling (ntcA and glnB (PII)), heterocyst differentiation (hetR) and dinitrogen fixation (nifH) was monitored in cyanobacteria from two Gunnera species at eight different stages of development from newly infected tissues at the apex to more mature tissues (Wang et al., 2004). The hetR gene was highly expressed and correlated positively with an increase in heterocyst frequency and with ntcA expression; nifH expression was already high in the apical part of Gunnera and glnB expression decreased from the apex along the stem. Analysis of Nostoc mutants revealed that the mutant in ntcA that encodes a global nitrogen regulator (Marcozzi et al., 2009) has lost the ability to infect the hornwort Anthoceros (Wong and Meeks, 2002).

When the proteome of Nostoc sp. freshly isolated from stem glands of G. manicata was compared with the proteome obtained in the same strain grown in a free-living state, several proteins were shown to be affected in the symbiotic process, with 23 proteins being up-regulated and ten down-regulated (Ekman et al., 2006, 2008). When the cyanobacteria enter the host, the normally photoautotrophic genus Nostoc is exposed to specific conditions and has to adapt to intracellular conditions, darkness and a microoxic environment. Despite all these physiological changes, data obtained by Ekman et al. (2006) revealed that most of the proteins present in symbiosis are also present in the free-living state, indicating that most cellular functions remain unmodified in the host. However, some adaptations to the symbiotic state were observed. These included up-regulation of the genes involved in nitrogen fixation such as nifH coding for dinitrogenase reductase, whose level was about four times higher in the symbiotic state than in the free-living culture. Because nitrogen fixation is a highly energy-demanding process, enzymes such as ATP synthase were up-regulated in the cyanobiont. Conversely, enzymes involved in the Calvin cycle such as phosphoribulokinase were down-regulated, together with Vipp1, a protein needed for thylakoid biosynthesis. Several proteins differentially expressed in the cyanobiont were involved in exopolysaccharide synthesis, along with surface, and membrane-associated proteins, suggesting adaptation of the surface for the establishment of the symbiosis and the exchange of nutrients. A third class of modifications was observed in the composition of phycobiliproteins, with allophycocyanin and phycocyanin being up-regulated in symbiosis, while phycoerythrin was down-regulated. These differential levels of expression are likely to be linked to the dark interior of the gland tissues.

ASSOCIATIVE AND ENDOPHYTIC NITROGEN FIXATION IN RICE, MAIZE AND WHEAT

Associative and endophytic nitrogen-fixing bacteria

Members of the Poaceae family do not naturally form symbiotic nitrogen-fixing associations. However, it has been shown that they can derive a substantial part of their nitrogen from BNF. Although the amount of fixed nitrogen is not as large as that measured in legumes nodulated by rhizobia, in actinorhizal plants or in cyanobacterial associations, increases in yields have been reported in the field (Dobbelaere et al., 2003; Vessey, 2003; Verma et al., 2010; Bhattacharyya and Jha, 2012).

Bacteria that colonize the rhizosphere are called rhizobacteria, and rhizobacteria with beneficial effects on plant development are referred to as plant-growth-promoting rhizobacteria (PGPR) (Kloepper and Beauchamp, 1992). Some of these PGPR are diazotrophic bacteria and have the ability to develop root associations with different plants including grasses. When they are found in close association with roots, they are usually designated ‘associative’ nitrogen-fixing bacteria (Elmerich, 2007). ‘Endophytic’ nitrogen-fixing bacteria have been defined as bacteria detected inside surface-sterilized plants or extracted from inside plants, having no visible harmful effects on the plants, fixing nitrogen, and proved by microscopic evidence to be located inside the plant (Hallmann et al., 1997; Reinhold-Hurek and Hurek, 1998). However, the frontier between associative and endophytic plant colonization is not always clear, since associative bacteria can also be observed in plant tissues, although they are less abundant than strains originally classified as endophytic (Elmerich, 2007). In contrast to endosymbioses, no differentiated structures in the roots are induced by these bacteria and, although endophytic bacteria invade plant tissues, they cannot be regarded as endosymbionts that reside intracellularly in living plant cells. Endophytic diazotrophs may have an advantage over root-surface associative diazotrophs, as they colonize the interior of plant roots and can establish themselves in niches that provide more appropriate conditions for effective nitrogen fixation and subsequent transfer of the fixed nitrogen to the host plant (Reinhold-Hurek and Hurek, 1998, 2011).

Diazotrophic rhizobacteria have been identified in several genera of alpha- and beta-proteobacteria including Acetobacter, Azoarcus, Azospirillum, Azotobacter, Burkholderia, Enterobacter, Herbasprillum, Glucenobacter and Pseudomonas (reviewed in Baldani et al., 1986; Döbereiner et al., 1993; Vessey, 2003; Schmid and Hartmann, 2007; Cocking, 2009; Richardson et al., 2009). Among these, Azoarcus spp., Herbaspirillum seropedicae and Glucenobacter are recognized as endophytes. They differ from other rhizobacteria such as Azospirillum and Azotobacter, in that they are tightly associated with plants and do not survive well in soil (Reinhold-Hurek and Hurek, 1998). Some of the main bacteria that can live in association with maize, rice and wheat and contribute to improved plant growth are presented in Table 2.

Table 2.

Association of cereals and nitrogen-fixing PGPR

Cereals Diazotroph inoculant Benefits % increase References 
Rice Azoarcus 16 (total dry weight) Reinhold-Hurek and Hurek, 1997,Engelhard et al., 2000 
Burkholderia 68 (shoot biomass) 19 (seed biomass) Baldani et al., 2000 
B. vietnamiensis 13–22 (yield)* Trân Van et al., 2000 
Gluconacetobacter diazotrophicus 30 (total dry weight) Muthukumarasamy et al., 2005 
Herbaspirillum seropedicae 37.6 (plant dry weight)James et al., 2002 
Serratia marcescens 23 (total dry weight) Gyaneshwar et al., 2001 
Maize Burkholderia sp. 5.9–6.3 (yield)* Estrada et al., 2005 
Azospirillum brasilense 13–25 (yield) Riggs et al., 2001 
 33 (grain yield)* Dobbelaere et al., 2001 
Azotobacter  Pandey et al., 1998 
H. seropedicae 19.5 (yield)* Riggs et al., 2001 
Pseudomonas sp. 11.7 (total biomass) Shaharoona et al., 2006 
Wheat H. seropedicae 49–82 (total biomass) Riggs et al., 2001 
Azospirillum sp.  Boddey et al., 1986 
Azotobacter sp.  Mrkovacki and Milic, 2001 
Cereals Diazotroph inoculant Benefits % increase References 
Rice Azoarcus 16 (total dry weight) Reinhold-Hurek and Hurek, 1997,Engelhard et al., 2000 
Burkholderia 68 (shoot biomass) 19 (seed biomass) Baldani et al., 2000 
B. vietnamiensis 13–22 (yield)* Trân Van et al., 2000 
Gluconacetobacter diazotrophicus 30 (total dry weight) Muthukumarasamy et al., 2005 
Herbaspirillum seropedicae 37.6 (plant dry weight)James et al., 2002 
Serratia marcescens 23 (total dry weight) Gyaneshwar et al., 2001 
Maize Burkholderia sp. 5.9–6.3 (yield)* Estrada et al., 2005 
Azospirillum brasilense 13–25 (yield) Riggs et al., 2001 
 33 (grain yield)* Dobbelaere et al., 2001 
Azotobacter  Pandey et al., 1998 
H. seropedicae 19.5 (yield)* Riggs et al., 2001 
Pseudomonas sp. 11.7 (total biomass) Shaharoona et al., 2006 
Wheat H. seropedicae 49–82 (total biomass) Riggs et al., 2001 
Azospirillum sp.  Boddey et al., 1986 
Azotobacter sp.  Mrkovacki and Milic, 2001 

*, , Experiments in fields (*) or in controlled conditions ().

Plant colonization process

The plant–bacterial interaction takes place in the rhizosphere where PGPR are stimulated by plant root exudates and attracted by root mucilage (reviewed in Vanbleu and Vanderleyden, 2007; Raaijmakers et al., 2009; Compant et al., 2010). The composition of root exudates depends on the type of soil, the availability of nutrients, the plant genotype and growth stage, and environmental biotic and abiotic stresses. In addition, there are differences in root exudation patterns along the root system that result in differences in the composition of the associated communities of bacteria. Some studies have shown that, as in the endosymbiotic process between legumes and rhizobia (Zhang et al., 2009) and actinorhizal plants and Frankia (Abdel-Lateif et al., 2012), flavonoids seem to be important plant signals for interaction with the bacteria. Some flavonoids were found to stimulate the colonization of wheat by Azospirillum brasilense and to be responsible for an almost 100 % increase in the number of lateral root cracks colonized in Arabidospsis by Herbaspirillum seropedicae (Webster et al., 1998). In addition, the flavanone naringenin was found to regulate genes of H. seropedicae predicted to be involved in the colonization process (Tadra-Sfeir et al., 2011).

Root colonization involves migration towards the plant roots, adsorption and anchoring onto the root system, as well as microbial proliferation and the formation of microcolony/biofilm structures at the surface of roots (Reinholdt et al., 1986; Zhu et al., 2002; Alexandre and Zhulin, 2007; Vanbleu and Vanderleyden, 2007; Compant et al., 2010; Reinholdt-Hurek and Hurek, 2011). Diazotrophic PGPR probably employ an array of distinct mechanisms, either alone or in combination, to colonize successfully the plant roots and compete with other soil microorganisms. Among the mechanisms concerned, chemotaxis resulting from the presence of flagella will allow the bacteria to get into contact with roots, together with type IV pili and twitching motility. Twitching motility is based on a mechanism which includes pilus extrusion, surface attachment of the pilus tip, and pilus retraction to convey the bacterial cell to the point of attachment (Böhm et al., 2007). In Azoarcus, type IV pili were shown to be involved in adherence to plant surfaces and the pilA, pilB and pilT genes were essential for root-surface colonization and for infection of plant tissues in rice (Dörr et al., 1998; Krause et al., 2006; Böhm et al., 2007).

As observed in different plant–bacteria associations (Downie, 2010; Gough and Cullimore, 2011), surface polysaccharides such as exopolysaccharides and lipopolysaccharides (LPS) are involved in the colonization of roots. In a Tn5 mutant of A. brasilense affected in the biosynthesis of dTDP-rhamnose, LPS composition was modified and resulted in impaired attachment of the mutant to maize roots and reduced root colonization (Jofré et al., 2004). Additional studies recently undertaken on H. serepedicae by Balsanelli et al. (2010) confirmed these data. Two knock-out mutants were obtained in the genes rfbB (dTDP-d-glucose 3,5-epimerase) and rfbC (dTDP-4keto-l-rhamnose reductase) involved in the biosynthetic pathway of rhamnose. The ability of the knock-out mutants to attach to the surface of the maize root was 100-fold lower than that of the wild type, and the number of bacteria colonizing the internal plant tissues was also 100-fold lower. In addition to LPS, a major outer membrane protein from A. brasilense strain Cd was purified and shown, by in vitro adhesion assays, to bind to roots of wheat, corn and sorghum seedlings (Burdman et al., 2001). In addition to its involvement in root adsorption, this protein acted on cell aggregation of Azospirillum. Bacterial cell aggregation was also modified in mutants of A. brasilense Sp7 affected in the chemotaxis Che1 pathway (Edwards et al., 2011). Moreover, an outer membrane lectin that specifically recognizes and binds to the extracellular exopolysaccharides from Azospirillum is thought to play a role in cell-to-cell adhesion (Mora et al., 2008).

In endophytes, undifferentiated tissues above the root tips and the points of emergence of lateral roots are the sites for primary colonization and entry into the plant (Reinhold-Hurek and Hurek, 1998). It has been suggested that cellulolytic and pectinolytic enzymes contribute to the infection process by degrading plant cell walls, thus providing a means to pass through the endoderm and to continue colonization inside the plant (Reinholdt-Hurek et al., 1993; Kovtunovych et al., 1999; Adriano-Anaya et al., 2005). This hypothesis was confirmed by some studies on the role of an endoglucanase gene referred as eglA in Azoarcus BH72 (Reinhold-Hurek et al., 2006). Whereas in rice plants inoculated with the control Azoarcus, bacteria were observed inside root epidermis cells 3 weeks after inoculation, the number of colonized cells was considerably decreased with an eglA mutant and the level of nifH mRNA was reduced in rice plants. The mutation also had an impact on the spreading into the shoot, thus leading to the conclusion that the cellulolytic degradation of plant cell walls plays an important role in plant colonization by Azoarcus BH72 and may contribute to systemic infection. However, since genes encoding cell wall-degrading enzymes have not been found in all endophytic PGPR, some of them may passively enter the root system, using disrupted endodermal cell layers resulting from the emergence of developing lateral roots.

After penetration, some endophytes may then colonize nutrient-rich intercellular spaces of the root cortex, move towards the xylem, and spread into stems and leaves (Olivares et al., 1996). The interaction can develop rapidly with some endophytes such as H. seropedicae, which was observed in cortical cell layers of maize 12 h after inoculation and in xylem after 24 h (Monteiro et al., 2008). Endophytic bacteria are found in roots, stems, leaves and seeds; nevertheless, in most plants, roots have the higher number of endophytes than above-ground tissues. Up to 108 colony-forming units per gram of root fresh weight have been reported (Barraquio et al., 1997), whereas bacterial densities in leaves may reach 103–104 colony-forming units of fresh weight with active colonizers (reviewed in Compant et al., 2010). However, it should be noted that recent metagenomic studies revealed a wide range of endophytes that are adapted to proliferate and spread in field-grown plants, although some have not yet been cultured or are difficult to isolate as pure cultures (Ikeda et al., 2010; Sessitsch et al., 2012).

Factors involved in plant growth promotion

Most of the PGPR isolates significantly increase plant height, root length and dry-matter production in agricultural crops like maize, wheat and rice. This plant growth promotion is the result of many different factors that can act directly or indirectly (reviewed in Dobbelaere et al., 2003; Rosenblueth and Martinez-Romero, 2006; Saharan and Nehra, 2011).

Since mutants of A. brasilense and Azoarcus sp. deficient in nitrogenase activity were shown to retain their ability to promote plant growth, these data initially raised the question of the relative contribution of nitrogen fixation to increasing plant growth in grasses (Hurek et al., 1994; Pedrosa and Elmerich, 2007). The contribution of associative and endophytic nitrogen fixation has now been clearly established, even though it appears to be highly variable, depending on the bacterial strain, the plant genotype and growth stage, and environmental conditions (Table 2). Evidence for nitrogen fixation was demonstrated by a number of techniques including acetylene reduction assays, 15N dilution studies, immunogold labelling with antibodies against the iron protein of nitrogenase, expression of transcriptional fusion between nifH and reporter genes, and qRT-PCR on transcripts encoding the nitrogenase complex. These techniques revealed nitrogenase activity in several endophytic bacteria of grasses including G. diazotrophicus (Sevilla et al., 2001), Azoarcus sp. strain BH72 in rice (Hurek et al., 2002), Herbasprillum sp. in rice (Elbeltagy et al., 2001; James et al., 2002; Roncato-Maccari et al., 2003) and Klebsiella sp. in maize and wheat (Chelius and Triplett, 2000; Iniguez et al., 2004). Nevertheless, in some associations such as Klebsiella sp. and rice, or Azospirillum and maize, the addition of a supplementary carbon source such as sodium malate was necessary to observe significant nitrogenase activity, suggesting a shortage of suitable carbon sources during the nitrogen-fixation process (Egener et al., 1999; Wouters et al., 2000; Saikia and Jain, 2007).

In PGPR and, in particular, the well-known Azospirillum, the production of phytohormones rather than nitrogen fixation is considered to be a major factor for plant growth promotion (Fulchieri et al., 1993; Dobbelaere et al., 2001; Baca and Elmerich, 2007; Spaepen et al., 2007, 2008). The production of the auxin IAA together with cytokinin has been reported in numerous rhizobacterial strains. These two hormones play a central role in regulating plant development, including processes that determine root architecture, such as root pole establishment during early embryogenesis, root meristem maintenance, root gravitropism and lateral root organogenesis (Kramer and Bennett, 2006). Phytohormones produced by bacteria thus enhance root branching and root elongation, which in turn favour the uptake of soil water and minerals and has a positive effect on plant growth (Steenhoudt and Vanderleyden, 2000). However, the effect of exogenous IAA on the root system can vary from growth stimulation to inhibition and is usually a function of the amount of IAA that is available to the plant and the sensitivity of the plant tissues to changes in IAA concentration (Keyeo et al., 2011). Recent transcriptome analyses in A. brasilense revealed that interfering with IAA biosynthesis led to broad transcriptional changes in the bacteria, suggesting that IAA is an important signalling molecule involved in the plant–PGPR communication process (Van Puyvelde et al., 2011). Besides auxin and cytokinin, the synthesis of gibberellin and, to a lesser extent ethylene, can also be observed. Gibberellin produced by Azospirillum was found to play an important role in the early stages of plant growth in Graminae by enhancing shoot and root growth and increasing root-hair density.

Although ethylene is essential for normal growth and development in plants, and is required for the induction of systemic resistance and in defence pathways in plants, at high concentrations it can be harmful and can reduce plant performance. PGPR produce the enzyme 1-aminocyclopropane-1-carboxylate deaminase, whose activity can divert 1-aminocyclopropane-1-carboxylate from the ethylene biosynthesis pathway (Blaha et al., 2006; Desbrosses et al., 2009). Rhizobacteria may thus reduce the accumulation of ethylene and re-establish a healthy root system. Production of NO due to the activity of nitrate reductase has also been observed in PGPR such as A. brasilense Sp245 (Pothier et al., 2007, 2008). This NO may have some positive effects on root development since it is a key signal molecule that controls root growth, stimulates seed germination and is involved in plant defence responses against pathogens (Fernández-Marcos et al., 2012).

Other direct effects of PGPR include the production of siderophores, vitamins and the solubilization of phosphorous. Iron is essential for plant growth and, in microorganisms, it acts as a global regulator of many cellular, metabolic and biosynthetic processes and, in bacteria, is a key element for nitrogen-fixation activity. Since in nature, iron is not readily available, microorganisms produce a wide range of small high-affinity chelating molecules called siderophores for its acquisition (Saha et al., 2012). Microbial siderophores may stimulate plant growth directly by increasing the availability of iron in the soil surrounding the roots or by inhibiting pathogen growth. Concerning vitamins, the production of thiamine, biotin, riboflavin and niacin has been documented in some strains of Azosprillum and Azotobacter (reviewed in Richardson et al., 2009). This exogenous supply may also stimulate root development. Phosphorous is one of the most essential nutrients for plants, but the majority of soil P is present in insoluble forms, while the plants can only absorb it in the soluble forms HPO42– and H2PO4. Phosphate-solubilizing bacteria such as Azospirillum and Burkholderia convert insoluble phosphorous into soluble form through acidification, secretion of organic acids or protons and chelation, thereby helping to improve phosphate nutrition in the associated plants (Sturz and Nowak, 2000; Richardson et al., 2009).

For some PGPR, positive effects on plant growth are indirect and result from mechanisms involving antagonism toward phytopathogens and the induction of systemic resistance pathways in the plant (Verhagen et al., 2004; Bally and Elmerich, 2007; Raaijmakers et al., 2009). These beneficial bacteria can help suppress a broad spectrum of viral, bacterial and fungal pathogens (reviewed in Sahan and Nehra, 2011). For instance, the inoculation of rice plants with the Azospirillum strain sp. B510 enhanced disease resistance to virulent rice blast fungus and to the bacterial pathogen Xanthomonas oryzae (Yasuda et al., 2009). The biocontrol of phytopathogens in the root zone can also be achieved through the production of antifungal or antibacterial agents, siderophore production or through competition for colonization sites and nutrients (Rienhold-Hurek and Hurek, 2011). Interestingly PGPR can also confer tolerance to a number of abiotic stresses and can stimulate plant growth even in areas affected by drought (Alvarez et al., 1996; Creus et al., 1996), salt (Creus et al., 1997; Jofré et al., 1998; Bacilio et al., 2004), and in soils polluted by heavy metals (Belimov and Dietz, 2000).

As the above-mentioned factors act in combination, a prerequisite for using these bacteria as nitrogen biofertilizers is to find the most appropriate combination of diazotrophic PGPR strain–plant cultivars to achieve a significant increase in plant biomass in the field (Table 1) (Roesh et al., 2006; Bhattacharjee et al., 2008). Some Azospirillum, Glucenobacter and Acetobacter inoculants are available for a variety of crops in Europe and Africa and from 5 % to 30 % increases in yields have already been reported (Vessey, 2003; Karthikeyan et al., 2007; Bhattacharyya and Jha, 2012). In rice and maize, associative nitrogen fixation can supply 20–25 % of total nitrogen requirements and, in wheat, inoculation with A. brasilense significantly increased the yields of foliage, grain and branching of root hairs (reviewed in Okon and Labandera-Gonzalez, 1994; Saikia and Jain, 2007; Mano and Morisaki, 2008; do Vale Barreto Figueiredo et al., 2010; Montanez et al., 2012). However, the impact of associative nitrogen fixation and plant-growth promotion is usually more marked in soils with poor fertility.

Molecular mechanisms underlying the association with cereals

So far, our knowledge of bacterial endophytes has come from the study of model PGPR, via genomic and functional analysis of candidate genes. As reviewed by Reinhold-Hurek and Hurek (2011), several genomes of endophytes are now available, including Azoarcus sp. BH72 (Krause et al., 2006), Klesiella pneumoniae 342 (Fouts et al., 2008), Pseudomonas stutzeri A1501 (Yan et al., 2008), Gluconacetobacter diazotrophicus Pal5 (Bertalan et al., 2009), Azospirillum sp. B510 (Kaneko et al., 2010) and Herbaspirillum seropedicae SmR1 (Pedrosa et al., 2011). The exploration of these genomes revealed a number of characteristics that are important for rhizosphere competence. Gene-encoding products with relevant functions linked to plant–microbe interactions were identified, such as nitrogen fixation, production of hormones and degradation of ethylene intermediate, iron transport, flagella, pili and quorum sensing that modulate functions related to rhizosphere competence and adaptation.

The growing number of available genomes of rhizobacteria makes it possible to use comparative analyses to improve our understanding of the common or specific features of plant diazotrophic endophytes. For example, comparison of the genome of G. diazotrophicus Pal5 and that of Azoarcus sp. BH72 showed that these two bacteria have evolved different strategies to colonize plants (Bertalan et al., 2009). Gluconacetobacter diazotrophicus is capable of growing on a wide variety of carbon sources and has a larger number of transport systems, whereas Azoarcus has complex signalling mechanisms to communicate with its host plant. Comparisons can also be extended to closely related beneficial and pathogenic endophytes. In 1997, Olivares et al. reported a genomic comparison of the endophyte H. seropedicae SmR1 and the phytopathogen Herbaspirillum rubrisubalbicans M1, which causes mottled stripe disease in sugarcane (Olivares et al., 1997). Suppressive subtractive hybridization, together with direct comparison of genome sequences, revealed some structural differences in LPS and adhesins between the two strains, suggesting that molecular determinants of bacterial cell surface structure may be responsible for their contrasted phenotypic behaviour as an endophyte or phytopathogen (Monteiro et al., 2012). These comparisons are expected to improve our understanding of contrasted phenotypes such as endophytic associations and pathogenic life styles.

On the plant side, global approaches based on PGPR have targeted plant transcriptomics (Cartieaux et al., 2003; Verhagen et al., 2004), proteomics (Miché et al., 2006; Cheng et al., 2010) and metabolomic approaches (Walker et al., 2011). First results of ‘omics’ approaches were reported with the model plant A. thaliana, followed by studies in rice (Oryza sativa), the model plant for monocot species, and more recently in maize. Transcriptomic studies initially helped reveal that, when A. thaliana was colonized by a PGPR, an accumulation of transcripts involved in the responses to pathogens and abiotic stresses was observed (Cartieaux et al., 2003), thus conferring resistance to subsequent infection by pathogenic bacteria. More recently, proteome analysis of O. sativa was used to examine the extent to which a defence response might be involved in the interaction with an Azoarcus sp. with different rice cultivars (Miché et al., 2006). Root proteomes of two rice cultivars with differing degrees of compatibility with the endophytic bacteria, were compared in response to jasmonate and to inoculation with the Azoarcus strain BHNG3.1. Jasmonate is a key signalling phytohormone in numerous plant responses to stresses such as pathogen attacks, and plays an important role in rice defence mechanisms (Rakwal et al., 1999). Data suggested that plant defence involving jasmonate plays a role in restricting colonization when the host–bacterium interaction is less compatible (Miché et al., 2006). Consequently, to obtain optimal plant–bacteria interactions, in the future it will be important to determine which factors suppress the defence response in a compatible variety.

EXTENDING NITROGEN-FIXING ABILITY TO NON-LEGUME CROPS: WHAT CAN BE LEARNED FOR NON-LEGUME SYMBIOSES?

In recent years, new insights into rhizobium–legume, rhizobium–Parasponia, actinorhizal and AM symbioses led to renewed interest in the possibility of transfering nitrogen-fixing ability to non-legume crops (Charpentier and Oldroyd, 2010; Beatty and Good, 2011; Geurts et al., 2012). It has been known for several years that several components (SYMs) of the legume symbiotic signalling pathway acting downstream from Nod factor receptors play a role in both nodulation and in the more ancient AM symbiosis. The demonstration that the common SYM gene SymRK is also required for actinorhizal nodulation (Gherbi et al., 2008a, b; Markmann et al., 2008) raised the question as to what extent the nodulation signalling pathway is conserved in legumes and non-legume actinorhizal plants. In addition, Hocher et al. (2011) highlighted the fact that, beyond SymRK, the whole array of compounds of the Nod factor signal transduction pathway is shared between RNS in legumes and actinorhizal plants. The fact that a series of well-characterized symbiotic genes in legumes exhibit similar expression patterns in actinorhizals lends credibility to a common ‘SYM’ pathway for endosymbioses and, for the first time, points to the possibility of a similar ‘NOD’ pathway between RNS. This overlapping of legume and actinorhizal RNS reinforces the hypothesis of a common genetic ancestor of the nodulating clade with a genetic predisposition for nodulation (Soltis et al., 1995).

AM symbioses appeared some 400 Mya (Remy et al., 1994), while nitrogen-fixing RNS evolved approx. 60–70 Mya (Doyle, 1998), supporting the hypothesis that the symbiotic signalling mechanisms involved in rhizobium–legume associations derived in the course of evolution from pre-existing mycorrhizal signalling components. The majority of land plants, including cereals, can form an AM association with fungi belonging to the phylum Glomeromycota. Most of the genes closely related to those involved in the signalling pathways leading to nodulation or AM symbiosis (i.e. SymRK/DMI2, CCaMK/DMI3, Cytokinin receptor, NIN) have been identified in the rice genome. Moreover, it has been shown that rice CCaMK is able to restore AM symbiosis in a Medicago dmi3 mutant and a rice ccamk mutant fails to develop AM (Chen et al., 2007). The rice CCaMK is also able to restore nodulation in the dmi3 Medicago mutant, although nodules mostly did not contain rhizobia or bacteria were not released from infection threads (Godfroy et al., 2006). These data show that elements of the rice AM genetic programme can trigger the appropriate downstream signalling pathway, thus paving the way for strategies to engineer nitrogen-fixing symbiosis in cereals by redirecting the evolutionary conserved common symbiotic pathway.

In addition to the use of knowledge accumulated on model legumes, we highlight the fact that non-legume actinorhizal and Parasponia symbioses could be more suitable models to obtain nitrogen-fixing cereals. Several steps including recognition, infection, intracellular accommodation of nitrogen-fixing endosymbionts, and nodule organogenesis are necessary to establish highly efficient nitrogen-fixing cereals. Because of the absence of the common nodABC genes in the genomes from clusters I and III strains (Normand et al., 2007), the fact that Frankia strains can interact with non-legumes belonging to eight angiosperm families raises the question of whether Frankia is not a better choice than Rhizobia to infect cereals. In 75 % of legume genera, rhizobia enter the root hairs via an intracellular infection, as observed in the two model legumes M. truncatula and L. japonicus. In contrast, in 75 % of actinorhizal genera and in Parasponia, endosymbionts form nitrogen-fixing nodules via intercellular invasion (Sprent, 2007; Wall, 2000). In evolutionary terms, intracellular infection via root hairs would be the most recent and sophisticated mechanism and intercellular infection probably the most primitive mode of root colonization (Madsen et al., 2010). Since much less is known about intercellular infection than about root hair infection, Imanishi et al. (2011) recently developed an efficient genetic transformation protocol for Discaria trinervis, an actinorhizal plant belonging to the Rosales order, in order to understand this ancestral and simpler infection mode. The study of Parasponia–Rhizobium and actinorhizal symbioses are thus of great interest in the context of engineering non-legume crops to allow infection by rhizobia or Frankia.

As previously mentioned, actinorhizal and Parasponia–rhizobia symbioses exhibit the same origin and structure as lateral roots. Interestingly, in Parasponia and in actinorhizal plants in the order Fagales, infection by endosymbionts led to cell division in the cortex, resulting in a small external protuberance called the prenodule (Angulo Carmona, 1974; Callaham and Torrey, 1977). Although nodule primordium formation does not involve prenodule cells, Laplaze et al. (2000a) showed that, in the actinorhizal plant C. glauca, the prenodule represents a very simple symbiotic organ where both Frankia and plant cells differentiate into their symbiotic condition. Before considering the engineering of a fully developed symbiotic nodule in non-legume crop, an intermediate step similar to the prenodule could be considered. Since Parasponia and actinorhizal nodules are modified lateral roots, we can expect a common subset of phytohormones and genes in the control of their development. Recently, Péret et al. (2007) and Perrine-Walker et al. (2010) showed that auxin accumulated in Frankia-infected cells, suggesting a role for auxin in the establishment of actinorhizal symbiosis. Further work is needed to explain how Frankia and rhizobia trigger the lateral root development programme of the host root system. This could be a clue on how to initiate the formation of a nodule primordium in a non-legume crop.

For some unknown reasons, angiosperms appear better adapted for developing intracellular symbiosis, since the angiosperm Gunnera is the only host plant in which intracellular symbiosis with cyanobacteria is observed. Whereas the need for a pre-existing gland restricts the possible transfer of this colonization process outside Gunneraceae, understanding the physiological conditions and molecular mechanisms underlying the intracellular penetration of Nostoc sp. in the angiosperm could provide valuable information, for example in the context of a comparative analysis of intracellular colonization in root and nodule cortical cells by Frankia and rhizobia. The advantage of using cyanobacteria to create new symbioses with agricultural plants is the broad host range of some strains, such as those belonging to the genus Nostoc, which can infect different plant organs. Furthermore, nitrogen fixation is accomplished in heterocyst cells that naturally protect the nitrogenase complex from inactivation by oxygen. The associative competence of symbiotic Nostoc strains isolated from Gunnera and Anthoceros was studied in rice roots by Nilsson et al. (2002, 2005). The association obtained was tight and the cyanobacteria could not be removed by washing or by sonication. When associated with rice roots, the Nostoc strains increased their nitrogen fixation and their presence appeared to improve root and shoot growth, and increased the weight of the rice grains (Nilsson et al., 2002).

In contrast to extensive molecular knowledge of rhizobium–legumes interactions, there are still only limited data available on the molecular aspects and signalling in the interactions leading to associative and/or endophytic interactions. Although the interactions with PGPR appear to be less complex than endosymbioses, they require the exchange of appropriate signals between the two partners to achieve successful colonization and phytostimulation, and for the bacteria to escape plant defence mechanisms. An intensive search for plant and bacterial signals, the receptors involved, cellular mediators and target genes in both partners is a primary goal to improve our understanding of these non-legume symbioses. This will provide additional knowledge leading to a broad view of the plant and microbial genes that could be manipulated to engineer new nitrogen-fixing plants.

CONCLUSIONS

A number of non-legume plants have evolved multiple strategies in association with diazotrophs to deal with N deficiency. The most sophisticated associations are root nodule endosymbioses between Frankia and actinorhizal plants, rhizobium and Parasponia sp., and cyanobacteria that associate with Gunnera sp. in cells of the specialized stem gland. In recent years, a major breakthrough has been the demonstration of a common genetic basis for plant root endosymbioses with AM fungi, rhizobia and Frankia bacteria in both legumes and non-legumes. This finding strengthens the hypothesis of a single origin for all nitrogen-fixing root nodule endosymbioses, and that RNS could have been partially recruited from the more ancient AM.

Compared with legumes, important questions remain to be answered including whether Frankia signals are structurally similar to Nod and Myc-factors, whether signals from actinorhizal plant and plant-hosting cyanobacteria are flavonoids – as in legumes and probably in Parasponia – and whether the conservation of the signalling pathway in Parasponia and actinorhizal plants goes beyond the common legume/rhizobium AM pathway. Progress in the knowledge of the basic mechanisms underlying symbiotic and endophytic associations in non-legumes has been generally slow, mainly due to the difficulties encountered in designing tools for the identification of candidate genes and their functional analysis. The value of comparative genomic approaches to help identifying, in addition to nif genes, common conserved gene functions specific to endosymbiotic and/or endophytic bacteria has been demonstrated. On the plant side, tools for ‘omics’ approaches and high-throughput sequencing technologies to finely explore transcriptomes are expected to provide new opportunities to decipher the molecular mechanisms underlying successful associations with diazotrophs.

The creation of artificial symbioses or associations between nitrogen-fixing microorganisms and plants of great agricultural importance is a primary goal in agriculture to reduce the demand for chemical nitrogen fertilizers. Since much of the basic work and major breakthroughs have been done on model legumes, strategies to expand the genetic capacity to fix nitrogen in symbiosis are currently based on that knowledge (Charpentier and Oldroyd, 2010; Beatty and Good, 2011). Recent advances in the understanding of endosymbiotic and endophytic nitrogen fixation with non-legume plants may represent original and alternative new avenues for engineering non-legume nitrogen-fixing crops.

ACKNOWLEDGEMENTS

Research conducted in the Rhizogenesis laboratory on actinorhizal plants was supported by the Institut de Recherche pour le Développement (IRD), Montpellier University 2, the Agence Nationale de la Recherche (ANR) Blanc project NewNod (ANR-06-BLAN-0095) and SESAM (BLAN-1708-01), the ECOS-Sud and the PHC-IMHOTEP. We thank Jocelyne Bonneau (IRD) for help with the manuscript.

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